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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Mech Ageing Dev. Author manuscript; available in PMC 2009 July 1.
Published in final edited form as:
PMCID: PMC2704552

Increased ROS generation in subsets of OGG1 knockout fibroblast cells


Oxoguanine DNA glycosylase (OGG1) is a major base excision repair protein responsible for excision of the mutagenic 8-oxoguanosine (8-oxoG) lesions from the genome. Despite OGG1’s importance, the moderate phenotype of Ogg1-null (Ogg1−/−) mice is not well understood. This study addresses a mechanism by which Ogg1−/− cells limit accumulation of 8-oxoG in their genome. Our data reveal that a subset of Ogg1−/− cells shows higher ROS levels (HROS cells), while ~85% of Ogg1−/− cells exhibit physiological levels of ROS (LROS cells). Ogg1−/− cells were sorted based on their DCF fluorescence intensity to obtain LROS and HROS cell cultures. LROS cultures proliferated at a rate comparable to Ogg1+/+ and gradually accumulated cells exhibiting increased ROS and 8-oxoG levels. LROS cells show a 2.8-fold increase in 8-oxoG level vs. HROS cells (7- to 27-fold). Mitochondria of HROS cells released more H2O2 than LROS and Ogg1+/+ cells and were eliminated by apoptotic-like processes. These findings suggest that in the absence of OGG1, a surveillance system is activated that removes cells with extreme 8-oxoG levels from Ogg1−/− cultures. Whether similar mechanism exists in tissues of Ogg1−/− mice is the focus of future investigations.

Keywords: Ogg1 null fibroblast, ROS, 8-oxoguanine, mitochondria


Steady-state increases in oxidative DNA base modifications in the nuclear and mitochondrial DNA of aged cells and tissues have been extensively documented both in animal models and human subjects, as reviewed previously (Hamilton et al., 2001b; Van Remmen et al., 2003). Among DNA bases, guanine has the lowest redox potential, and so it is easily oxidized to 8-oxo-7,8-dihydroguanine (8-oxoG) via two-electron oxidation (Shukla et al., 2004). The key repair enzyme that excises 8-oxoG from cellular DNA (8-oxoG: C base pairs) is the 8-oxoguanine DNA glycosylase (OGG1) (Hazra et al., 1998). If the damaged base is not removed, 8-oxoG may pair with adenine, causing G:C → T:A transversions after DNA replication (Cheng et al., 1992; Mitra et al., 1997). These mutations are often found in coding regions of tumor-promoting and tumor suppressor genes (Bos, 1989; Sigal and Rotter, 2000). Mutations may occur in the human Ogg1 gene leading to polymorphisms and altered 8-oxoG and FapyG (ring-opened guanine) excision capacity of OGG1, thereby OGG1 appears to be etiologically involved in accumulation of mutations in the coding regions of DNA in head, neck, lung and kidney cancers (Blons et al., 1999; Shinmura and Yokota, 2001). The lack of efficient removal of 8-oxoG from DNA in tissues of patients with Cockayne syndrome implicates 8-oxoG in accelerated aging (Stevnsner et al., 2002).

Recently, Ogg1−/− mice were generated (Klungland et al., 1999; Minowa et al., 2000). It has been shown that the levels of 8-oxoG in the nuclear and mitochondrial DNA of OGG1-deficient mice increases continuously over the whole life-span, especially under oxidative stress conditions (Arai et al., 2002; Klungland et al., 1999; Minowa et al., 2000; Nishimura, 2002; Osterod et al., 2001). Despite the increase in this potentially mutagenic, miscoding DNA lesion, Ogg1−/− mice are only moderately proned to cancer, and show no marked age-associated pathological changes (Klungland et al., 1999; Sakumi et al., 2003). There are no significant differences in life-span between wild-type and OGG1-difficient mice, which appear phenotypically normal (Klungland et al., 1999).

Fibroblast cultures developed from Ogg1−/− mice showed no major morphological changes and their cell cycle dynamics were similar to those of wild-type counterparts (Osterod et al., 2001). Up to date there are no studies addressing how Ogg1−/− cells maintain steady state ~3-fold increase in 8-oxoG level in their nuclear DNA in the absence of OGG1, although they are continuously being damaged by exogenous/endogenous ROS. Here we show that in cultures of Ogg1−/− there is a balance between low- and high-ROS-containing cells. High-ROS cells containing 7- to 27-fold increase in 8-oxoG levels exhibit mitochondrial dysfunction, and are eliminated by process(es) that share characteristics with apoptosis. These results suggest that a surveillance mechanism(s) is activated in response to supraphysiological levels of ROS and 8-oxoG, which limit the lifespan of Ogg1−/− cells. These data raise the possibility that similar mechanism(s) could exist in tissues of OGG1 knockout mice.

2. Materials and Methods

2.1. Cell cultures

Ogg1−/− and Ogg1+/+ primary mouse embryo fibroblast (MEF) cultures were established by standard procedures from individual C57BL/6J 129 embryos on embryonic day 13.5 (Klungland et al., 1999). Cells were cultured in DMEM/Ham’s F-12 (3:1) with 10% FBS, genotyped by Southern hybridization, and permanent cell lines were established from individual transformed clones arising spontaneously after repeated passage in culture (Klungland et al., 1999). Immortalized Ogg1−/− and Ogg1+/+ cell lines were kindly provided by Dr. Barnes (Imperial Cancer Research Fund, Clare Hall Laboratories, South Mimms, Hertfordshire EN6 3LD, United Kingdom). NTH1 (thymine glycol-DNA glycosylase)-knockout (Nth1−/−) MEF cells were derived from individual C57BL/6J 129 Nth1−/− embryos and continuously passaged to obtain spontaneously immortalized cells (Takao et al., 2002). Nth1−/− and Nth1+/+ cell lines were kindly provided by Dr. Elder (Cancer Research UK, Carcinogenesis Group, Paterson Institute for Cancer Research, Christie Hospital NHS Trust, Manchester M20 4BX, United Kingdom). In our laboratory the cells were cultured in DMEM/Ham’s F-12 (3:1) (Invitrogen, Carlsbad, CA) medium supplemented with 10% fetal calf serum, 100 U/ml penicillin, 100 μg/ml streptomycin and 0.2 mM glutamine (Elder and Dianov, 2002; Takao et al., 2002). Ogg1−/− and Ogg1+/+ cells were cloned to establish single cell colonies that were further propagated to develop permanent cell lines. Individual clonal cell lines were further characterized. There were no differences between parent unselected Ogg1−/− fibroblast cultures and clonal cell lines in morphology, replication dynamics, cellular ROS and 8-oxoG levels. Ogg1−/− LROS and HROS cell cultures were isolated by sorting (FACSaria, Becton Dickinson, Mountain View, CA) as we described below (2.5. third paragraph). These cells were maintained in DMEM/Ham’s F-12 (3:1) (Invitrogen, Carlsbad, CA) medium supplemented with 10% fetal calf serum, 100 U/ml penicillin, 100 μg/ml streptomycin and 0.2 mM glutamine. At passage second the doubling time of LROS and HROS cells were 18.2±1.2 and 19.3±0.7 h, respectively. Since we did not know the exact number of population doubling levels (PDLs), the initial PD was arbitrarily set at zero. Cultures were initiated by seeding 2 x104 cells per cm2 and at 90% confluence, the cells were removed by trypsinization, counted, and PDs calculated (PD = log(number of cells obtained at subculture per 2 ×104)/log 2) (Szczesny et al., 2003).

2.2. Western blot analysis

Equal amounts of protein-containing cell lysates were electrophoresed on SDS-PAGE; transferred to a nitrocelullose membrane PROTRAN (Schleicher and Schuell BioScience, Keene, NH), and blocked in TBST (20 mM Tris–HCl {pH 7.6}, 137 mM NaCl, and 0.5 % Tween 20) containing 5% dry milk. The membranes were incubated with primary antibody against mouse OGG1 (Szczesny et al., 2003), at 1:500 dilutions in TBST+5% milk. After washing in TBS-T horseradish peroxidase (HRP)-conjugated secondary antibody (1:2000 anti-rabbit IgG-HRP; Amersham Biosciences, Piscataway, NJ) was added. Subsequently, membranes were washed and incubated in ECL Western blotting detection reagent (Amersham Biosciences). Chemiluminograms were analyzed using Image Quant software (Molecular Dynamics, Sunnyvale, CA).

2.3. Immunohistochemistry

Cells were grown to 50% confluences on glass cover slips (25 mm), washed with PBS, dried, and fixed in acetone-methanol 1:1 (v/v). After exposure to pre-immune serum (0.1 μg in PBS containing 0.05% Tween 20 and 0.5% BSA; PBS-T), cells were incubated with primary antibody (anti-OGG1 or anti-FLAG) for 60 min. A fluorescein-conjugated secondary antibody (Santa Cruz Biotechnology, Inc) was then applied to cells for 60 min at 37°C and washed in PBS-T. Nuclei of cells were stained for 15 min with DAPI (4′6-diamidino-2-phenylindole dihydrochloride; 10 ng/ml, Molecular Probes, Eugene, OR). Extensively PBS-T-washed cells were dried and then mounted in anti-fade medium (Dako Inc. Carpinteria, CA) on microscope slides (Boldogh et al., 2001). Fluorescent images were captured at magnification of ×60 using a Photometrix CoolSNAP Fx digital camera mounted on a NIKON Eclipse TE 200 UV microscope.

2.4. 8-oxoG assays

The 8-oxoG in nuclear DNA was quantified as we previously described (Bhakat et al., 2006). Briefly, cells on coverslip cultures were washed with PBS, air-dried, and fixed in acetone-methanol (1:1), rehydrated in PBS for 15 min, then sequentially treated with 100 μg/ml pepsin in 0.1 N HCl for 15 to 30 min at 37°C, 1.5 N HCl for 15 min, and sodium borate for 5 min. The cells were incubated with nonimmune IgG (100 μg/ml) for 30 min and washed in PBS containing 0.5% bovine serum albumin, 0.1% Tween 20 (PBS-T). After incubation with anti-8-oxoG antibody (Trevigen, Gaithersburg, MD; 1:200 dilutions) (Bespalov et al., 1999) for 30 min, the cells were washed three times with PBS-T for 15 min then exposed to fluorescein-conjugated secondary antibody (Santa Cruz Biotechnology, Santa Cruz, CA) for 60 min. Cells were washed with PBS-T for 15 min (3-times) and their DNA stained with DAPI (10 ng/ml) for 15 min. The cells were air dried and mounted in anti-fade medium (Dako North America, Inc., Carpinteria, CA) on a microscope slide. The fluorescence intensities of a minimum of 40 cells per plate were determined using a Zeiss LSM510 META system, operated via MetaMorph software version 6.06r (Universal Imaging Corporation, Downingtown, PA).

The comet assay was performed according to the manufacturer’s instructions with some modifications as we described previously (Bacsi et al., 2005; Boldogh et al., 2003). Briefly, cells were suspended in 0.75% low-melting point agarose (Trevigen, Inc., Gaithersburg) in PBS and spread on microscope slides (Trevigen, Inc., Gaithersburg). The cells in solidified agarose were lysed for 1 h at 4°C in lyses buffer (10 mM Tris-HCl, pH 10, 2.5 M NaCl, 100 mM EDTA, 1% Triton X-100). DNA was released in the presence of NaI (Hamilton et al., 2001a) and treated with bacterial DNA formamidopyrimidine DNA glycosylase (Fpg, 1 μg/ml; New England Biolabs, Beverly, MA) in digestion buffer (40 mM HEPES, pH 8.0, 100 mM KCl, 0.5 mM EDTA, 0.2 mg/ml BSA) for 1 h at 37°C (Trzeciak et al., 2004). Electrophoresis was performed in alkaline electrophoresis buffer at 4°C for 30 min at 1 V/cm. The slides were then neutralized (0.4 M Tris-HCl, pH 7.5), washed, air-dried, and stained with 10 ng/ml of SYBR Green I (Trevigen, Inc., Gaithersburg). The comet moment images were recorded with a Nikon TE200 epifluorescence UV microscope photomicrographic system (Photometric CoolSNAP Fx camera). Fifty or more images were randomly selected for each data points and the tail moments were analyzed.

2.5. Measurement of Reactive Oxygen Species

2′,7′-dichlorodihydro-fluorescein diacetate (H2DCF-DA; Invitrogen Co., Carlsbad, CA), a redox-sensitive probe was used to determine cellular ROS levels (Bacsi et al., 2005; Boldogh et al., 2003). Cells on microscope cover-slips were placed in a thermo-controlled microscopic chamber and loaded with 10 μM (final concentration) H2DCF-DA for 15 min. Cells were washed with medium pre-warmed to 37°C and fluorescent images captured using a Photometrix CoolSNAP Fx digital camera mounted on a NIKON Eclipse TE 200 UV microscope. Fluorescence intensities of >200 cells were determined by Metamorph Version 6.06r software (Universal Imaging Corporation).

To define the intracellular site(s) of ROS generation the cells were loaded with 2 μM dihydroethidium (H2Et; Invitrogen Co., Carlsbad, CA) for 10 min and placed in a thermo-controlled microscopic chamber (Bacsi et al., 2006). MitoTracker Red (Invitrogen Co., Carlsbad, CA), a cell-permeable fluorescent probe that accumulates in active mitochondria was used to stain mitochondria at a final concentration of 25 nM. Images were captured with Photometrix CoolSNAP Fx digital camera mounted on a NIKON Eclipse TE 200 UV microscope. The microscope was operated and superimpositions were performed with Metamorph Version 6.06r software.

In selected experiments cells in suspension were loaded with H2DCF-DA (2.5 μM, final concentration) for 15 min and changes in fluorescence intensities were then determined by flow cytometry (FACScanto, Becton Dickinson, Mountain View, CA). 15,000 cells were collected and analyzed for each sample. To generate nearly homogenous HROS and LROS cultures, Ogg1−/− cells were loaded with 2.5 μM H2DCF-DA and sorted using Becton-Dickinson FACSAria cell sorter (Becton Dickinson, Mountain View, CA). Cells were cultured and purities of populations were assessed after 2 population doublings by flow cytometry.

Amplex Red assays were used to measure H2O2 production of isolated mitochondria (Bacsi et al., 2006). Amplex® Red (10-acetyl-3,7-dihydroxyphenoxazine; Molecular Probes) reacts with H2O2 in the presence of horseradish peroxidase to generate a stable fluorescent product, resorufin (Zhou et al., 1997). Briefly, isolated mitochondria (100 μg/ml) were suspended in 50 μl (per well) reaction buffer containing pyruvate (5 mM) plus malate (5 mM) as well as succinate (10 mM) and incubated at room temperature (25°C) for 15 min with 0.25 U/ml of Amplex® Red (determined in preliminary studies) and 1 U/ml of horseradish peroxidase. The change in fluorescence (excitation 563 nm; emission 587 nm) was measured using a microplate reader (SpectraMass M2, Molecular Devices Corporation, Downingtown, PA). In selected experiments, 3-nitropropionic acid (3-NPA; 1 mM, pH balanced) – inhibitor succinate dehydrogenase in complex II, rotenone (5 μM) – inhibitor of NADH-decylubiquinone reductase activity of complex I and antimycin A (0.4 μM) – inhibitor of cytochrome b reoxidation in complex III, were added alone or in combination to the reaction mixtures. In control, the H2O2 production was linear with mitochondrial protein concentration. Reactions were carried out ±exogenously added superoxide dismutase (SOD) that converts superoxide anion (O2•−) into H2O2. The addition of catalase (400 U/ml) decreased H2O2 levels by ~90 percent. As a positive control, increasing dilutions (0 to 4000 pM) of H2O2 were used.

2.6. Simultaneous analysis of cell cycle and cellular ROS levels

To sort out whether the increased ROS levels were associated with a particular cell cycle stage we loaded cells with Vybrant DyeCycle Violet stain (Invitrogen Co., Carlsbad, CA) (Telford et al., 2007) and H2DCF-DA simultaneously as previously described (Boldogh, 2003). Vybrant DyeCycle Violet stain (excitation/emission maxima ~396/437 nm) is cell membrane permeant dye and nonfluorescent until bound to double-stranded DNA. Briefly, actively replicating Ogg1+/+; Ogg1−/−, HROS and LROS cells (2 × 106 cells per ml) were suspended in Hanks’ balanced saline solution containing 2% FBS and 2 mM HEPES (pH: 7.4) and loaded with Vybrant DyeCycle Violet (10 μg per ml) and H2DCF-DA (5 μM, final concentration) for 15 min. Extracellular probes were removed by washing cells in Hank’s solution and cells were analyzed by Flow cytometry (FACScanto, Becton Dickinson, Mountain View, CA). 30,000 cells were collected and analyzed for each sample.

2.7. Mitochondria Isolation

Cells were propagated in large volumes, collected by centrifugation (800 × g), and mitochondria were isolated as we described previously (Bacsi et al., 2006). Briefly, cell pellets were incubated in 10× volume of hypotonic buffer (10 mM KCl, 20 mM MOPS, and 1 mM EGTA; ethylene glycol-bis (β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid) for 20 min. Sucrose (200 mM) and mannitol (50 mM) were added to the swollen cells, which were then Dounce-homogenized. The homogenate was centrifuged at 800 × g and the supernatants re-centrifuged at 10,000 × g to collect mitochondria. Mitochondrial pellets were washed, and resuspended in 10 mM KCl, 20 mM MOPS and 1 mM EGTA containing 200 mM sucrose, 50 mM mannitol. In selected experiments fresh mitochondrial suspensions were purified on a continuous sucrose gradient (0.25M to 1.5M) and used immediately for determining the levels of released H2O2.

2.8. Oxygen Consumption

The oxygen consumption rates of mitochondria were determined at 30°C with a Clark-type oxygen electrode (Strathkelvin Oxygen System Model 782, Strathkelvin Instruments, United Kingdom) as we previously described (Bacsi et al., 2006). Purified mitochondria (0.2 mg) were suspended in 1 ml respiration medium (125 mM KCl, 20 mM HEPES; pH 7.4), 5 mM potassium phosphate, ±substrates (5 mM pyruvate plus 5 mM malate as well as 10 mM succinate). The signal from the oxygen sensor was recorded on a computer at sampling intervals of 0.5 seconds with the aid of software from Strathkelvin Instruments (782 System V3.0). Respiration was measured without ADP (state IV) and with 0.5 mM ADP (state III) (Bacsi et al., 2006; Chen et al., 2003). Mitochondria suspension showing a higher than 3 respiratory control ratio were used.

2.9. Transfection

cDNA encoding human OGG1 was originally cloned into E. coli expression plasmid pRSETB (Dr. Sankar Mitra, University of Texas Medical School, Galveston, TX). OGG1 encoding sequences were re-cloned into the pcDNA3.0-neo expression plasmid (He et al., 2002) and kindly provided by Dr. MR Kelley (Herman B. Wells Center for Pediatric Research, and Biochemistry and Molecular Biology, Indiana University Medical School, Indianapolis, Indiana). Sequence identity of OGG1 was confirmed by direct sequencing (Molecular Genomic Core Facility University of Texas Medical School, Galveston, TX). At 75% confluence, Ogg1−/− and Ogg1+/+ cells were transfected with 0.25 μg of OGG1-encoding expression vector. As control, parallel cultures of cells were transfected with 0.25 μg of empty plasmid DNA (pCDNA.3.0-neo). The cells were subcultured 24 h after transfection and fresh medium containing 350 μg/ml of G418 (Invitrogen Co., Carlsbad, CA) was added. Ogg1−/− cell colonies resistant to G418 were grown and screened for OGG1 expression by immunostaining.

2.10. Annexin V Assay

Flow cytometric analyses with annexin V-PE and 7AAD-stained cells were performed as we described previously (Boldogh et al., 2003). Briefly, cells were trypsinized, washed in PBS and resuspended in cold annexin V binding buffer (10 mM HEPES pH 7.4, 0.14 M NaCl, 0.25 mM CaCl2·2H2O, 0.1% BSA (w/v); anti-annexin V-PE-conjugated antibody (Becton Dickinson) was then added to the cells for 30 min. Cells were sedimented and 7AAD was added (5 μg/ml per sample). The cells were analyzed on a FACScanto flow cytometer (Becton Dickinson, Mountain View, CA). Each data point represents the mean fluorescence for 12,000 cells from three or more independent experiments; expressed ±SEM.

2.11. Assessment of ATP levels

Cellular ATP levels were measured by a bioluminescence assay employing the luciferase enzyme using the ATP Determination Kit (Invitrogen Co., Carlsbad, CA). Prior to measurement of ATP levels annexin V positive cells were removed from the cell population by flow cytometric sorting. The cells (3 × 106/ml) were collected by centrifugation and ATP was then extracted by adding 100 μl of boiling distilled water to the pellet and heated at 100°C (Isotemp 125 D, Fisher Scientific, Pittsburg, PA) for 3 min. The cell lysates were then centrifuged at 12,000 × g for 5 min at 4°C (Silent SPIN, Continental Lab Products, Edison, NJ). Luminescences corresponding to ATP concentrations in the supernatant fluid were determined in a Veritas Microplate Luminometer (Turner Biosystems, Sunnyvale, CA).

2.12. Caspase activity measurement

Cells were grown to 80% confluence washed and lysed in lysis buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% {v/v} Triton X-100 and 10 mM sodium pyrophosphate). The lysates were incubated with 20 μM Ac-DEVD-AMC (Becton-Dickinson) in reaction buffer (20 mM HEPES, 10% (v/v) glycerol, 2 mM dithiothreitol) at 37°C for 1 h. Changes in fluorescence intensity were determined at 380 nm (excitation) and 435 nm (emission) wavelengths in an FLx800 microplate reader (Bio-Tek Instruments, Winooski, VT). Caspase-3-like activities were expressed as pmoles of AMC (7-amino-4-methyl coumarin) released from milligram of protein per 60 min (Boldogh, 2003).

2.13. Reagents

Anti-OGG1 antibody (OGG1-2672) was generated against the purified OGG1 polypetide (Alpha Diagnostic International, San Antonio, TX). This affinity-purified antibody reacts with both human and mouse OGG1 in Western blot and immunohystochemical analysis (Bhakat et al., 2006; Szczesny et al., 2003). Antimycin A, ATP, catalase, H2O2, malate, pyruvate, rotenone, succinate and SOD were purchased from Sigma-Aldrich (St. Louis, MO). 3-NPA (Aldrich Chemical Co., Milwaukee, WI) was dissolved in PBS and the pH was adjusted to 7.4 with 1 M NaOH (Bacsi et al., 2006). The osmolarity of pH-balanced 3-NPA solution was determined using an Advanced Osmometer (SIM International Co).

2.14. Statistical Analysis

Results were analyzed for significant differences using ANOVA procedures and Student’s t-tests (Sigma Plot 6.0). Data are expressed as the mean ±SE. Results were considered significant at p<0.05. (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001)

3. Results

3.1. A subset of Ogg1−/− cultures show extreme levels of 8-oxoG

Ogg1−/− cells showed a 3.2±0.2-fold increase in 8-oxoG levels in the nuclear DNA compared to the corresponding wild-type Ogg1+/+ cells. In Nth1−/− cells, which are devoid of NTH1 glycosylase involved in the repair of oxidized pirimidines (Ikeda et al., 1998) the 8-oxoG level was similar to Ogg1+/+ cells (Fig. 1A). These results are consistent with the permanent absence of OGG1 (Fig. 1A, inset) in Ogg1−/− cells. Interestingly, 8-oxoG levels were increased by 7- to 27-fold in subsets (12.4±1.5%) of Ogg1−/− cells compared to the Ogg1+/+ cells. Representative images of 8-oxoG-mediated fluorescence are shown in Fig. 1B. The low- and high 8-oxoG-containing cells appear to be in equilibrium, because their percentages did not change with increasing passage number. These data raise the possibility that Ogg1−/− fibroblast cultures are heterogeneous in regard to maintenance of 8-oxoG in their genome. Therefore, we isolated cell lines of single cell origin. In all eight clonal Ogg1−/− cell lines, 11 to 12.5 percent of the cells showed increased 8-oxoG levels (7- to 27-fold, compared to Ogg1+/+) similarly to the parent Ogg1−/− cells as determined by fluorescent imaging (data not shown). Data shown are from experiments with the parent Ogg1−/− fibroblast cells.

Figure 1Figure 1
Steady-state levels of 8-oxoG in the DNA of Ogg1−/− fibroblasts

To confirm increased levels of 8-oxoG in DNA of Ogg1−/− cells, we used single cell DNA electrophoresis (Boldogh et al., 2003; Olive et al., 1991) combined with Fpg digestion (Banath et al., 1999; Trzeciak et al., 2004). In control (without Fpg), only small differences were observed between comet tail moments of Ogg1−/− and Ogg1+/+ cells (Fig. 1C). Treatment of DNA with Fpg resulted in 4.2-fold difference between Comet tail moments of Ogg1−/− and Ogg1+/+ cells (Fig. 1C). These data suggest that the increased tail moment was due to 8-oxoG and FapyG (2,6-diamino-4-hydroxy-5-foramidopyriminidine; a ring opened 8-oxoG product) lesions in the DNA and not to increased levels of DNA single-strand breaks in the Ogg1−/− cells. In repeated experiments, 11.4±2% of Comet tail moments were larger (Fig. 1D, double arrow head) compared to the majority of Comet moments of DNA from Ogg1−/− cells (Fig. 1D, single arrow head). When compared to DNA from Ogg1+/+ cells the Comet tails moments showed 7 to 27-fold increase (Fig. 1E). Data from Comet assays are consistent with the increased reactivity of anti-8-oxoG antibody with the oxidized guanine base lesions in the DNA.

3.2. Increased ROS levels in subsets of Ogg1−/− cells

To investigate whether the excessive levels of 8-oxoG in subsets of Ogg1−/− cultures were due to increased levels of ROS, cells were loaded with H2DCF-DA and fluorescence intensities were assessed microscopically. We observed that 16±5% of the cells showed increased DCF fluorescence (high-ROS: HROS cells), while ~84% of the cells had lower oxidative stress levels (low-ROS: LROS cells), which were similar to Ogg1+/+ cells (Fig. 2A,B). The percentage of HROS and LROS cells did not significantly change during subculturing. In Nth1−/− and Ogg1+/+ cultures only <1% of cells showed increased DCF fluorescence (Fig. 2B). When ROS levels were assessed by flow cytometry there were no significant differences in overall ROS levels between Ogg1−/− and Ogg1+/+ cells (Fig. 2C and D upper panel). However; there was a distinct Ogg1−/− cell population showing increased DCF fluorescence. Frequencies of HROS cells were nearly identical in the parent Ogg1−/− and in sub-cell lines of single cell origin. The increased intracellular oxidative stress levels showed no correlation with cell cycle stages as determined by simultaneous measurements of DNA content and DCF fluorescence (data not shown).

Figure 2Figure 2Figure 2Figure 2
Increased ROS levels in subsets of Ogg1−/− cells

LROS and HROS cells were separated by cell-sorting based on DCF fluorescence intensity. ROS levels of sorted cells were assessed by flow cytometry at PDL 2. There was an approximately 2-fold increase in oxidative stress levels in HROS vs. LROS cells (Fig. 2C and Fig. 2D lover panel). When HROS cells were propagated further they showed 2.2- and 2.7-fold increase in ROS levels, at PDL-6 and PDL-11, respectively compared to Ogg1+/+ cells (Fig. 2E). At PDL-17 the ROS levels in HROS exceeded 3.5-fold of Ogg1+/+ cells. HROS cells proliferated slowly and cultures were lost between PDL-19 and -23. In cells maintained in N-acetyl-L-cysteine (5 mM)-containing media cellular ROS levels were significantly lower (Fig. 2E). Addition of BSO (100 μM), a known inhibitor of GSH synthesis (Anderson, 1998) to the culture medium increased intracellular levels of ROS and decreased cell viability (data not shown). At PDL-2 HROS cells were exhibiting more then 7-fold increase in 8-oxoG levels, while at PDL-17, 8-oxoG levels were more than 22-times higher than in Ogg1+/+ cells (data not shown). The increase in 8-oxoG levels in HROS cultures correlated well with increased Comet tail moments (Fig. 2F).

In LROS cultures, the percentage of cells showing increased DCF fluorescence increased as a function of PDL. For example, at PDL-12, 6.3%; PDL-25, 11.7%; and PDL-32, ~14% of cells showed increased oxidative stress levels (Fig. 2G). Additional culturing of these cells resulted in no further increase in the percentage of HROS cells, and now they were similar to that of unsorted Ogg1−/− cell cultures. LROS subsets at PDL-2 exhibited 2.8-fold elevation in 8-oxoG levels compared to Ogg1+/+ cells. The number of cells showing >7-fold elevation in 8-oxoG levels was proportional to the percentage of HROS cells. These data together suggest that HROS-high 8-oxoG containing cells are continuously generated (and eliminated), leading to an equilibrium between LROS and HROS cells.

3.3. Increased ROS production from mitochondria

To identify the site of ROS production, cells were placed in a microscopic culture chamber and loaded with 2 μM dihydroethidium (H2Et) (Bacsi et al., 2006). The green fluorescence mediated by H2Et/superoxide reaction products (Zhao et al., 2003) co-localized with MitoTracker Red suggesting that the mitochondria are the sites of ROS generation (Fig. 3A). Suspensions of Ogg1−/− cultures were sorted for LROS and HROS subsets and the individual cultures were further propagated. Mitochondria were isolated at PDL-10 to determine the amount of H2O2 originating from O2•− dismutation by SOD. Intact mitochondria from unsorted Ogg1−/− and Ogg1+/+ cells oxidizing from pyruvate/malate (Complex I substrates) plus succinate (Complex II substrate) released nearly equal amounts of H2O2 (Fig. 3B). On the other hand, mitochondria from LROS generated quantitatively less H2O2 than mitochondria from HROS subsets (Fig. 3B). Addition of rotenone, an inhibitor of NADH-decylubiquinone reductase activity of complex I, or 3-NPA, an inhibitor of succinate dehydrogenase to mitochondrial suspension only decreased (data not shown), while addition of rotenone and 3-NPA together inhibited H2O2 production from mitochondria of LROS and HROS cells (Fig. 3B). On the other hand, antimycin A, an inhibitor of cytochrome b reoxidation in complex III generated significantly higher amounts of H2O2 from the mitochondria of HROS cells compared to LROS cells (Fig. 3B). Addition of catalase decreased H2O2 to an undetectable level under our assay conditions.

Figure 3
Increase in mitochondrial ROS generation in a subset of Ogg1−/− cells

3.4. HROS cells are eliminated from cell cultures

Microscopic observation of cultures showed significantly higher cell death in Ogg1−/− cultures than in Ogg1+/+ or Nth1−/− ones. Results show that 9.2±2 % of unsorted Ogg1−/− cells bound annexin V (Fig. 4Aa) and of these 24.5±1.1% also incorporated 7-amino actinomycin D (7AAD; data not shown), an indicator for loss of cell membrane integrity. Less than 1% of Nth1−/− and Ogg1+/+ cells bound annexin V and ~0.1% incorporated 7AAD (Fig. 4Aa). Next, Ogg1−/− cells were sorted for LROS and HROS subsets. Percentage of annexin V positive cells was ~1 % in LROS cell populations at the time of sorting. Further subculturing of LROS cells, resulted in a gradual increase in annexin V positive cells (Fig. 4Ab). For example, at PDL-16 there were 7.5%, while at PDL-24 and higher PDLs it increased to the levels of unsorted Ogg1−/− (Fig. 4Ab). When HROS cells were further cultured the frequency of annexin V positive cells increased reaching more then 60% by PDL-17 (data not shown). At this or higher PDLs cells did not replicate further and the cultures were gradually lost.

Figure 4Figure 4Figure 4
HROS cells are eliminated from Ogg1−/− cultures by apoptosis-like processes

Next, we determined changes in caspase-3 activity of HROS and LROS cells at increasing PDLs. Results summarized in Fig. 4B illustrates that unsorted Ogg1−/− cell cultures showed no difference compared to Ogg1+/+ cells in caspase-3 activity. On the other hand, caspase-3 activity was significantly higher in HROS cultures at PDL-12 than at PDL-2 (Fig. 4B). Near the end of the HROS cells’ life-span (PDL-17), their caspase-3 activity was nearly 12.5-fold higher than in unsorted Ogg1−/− cells and 14-fold higher than in Ogg1+/+ cultures (Fig. 4B). From PDL-12 on, an increasing percentage of HROS cells displayed enlargement, irregular, bizarre morphology that did not resemble typical apoptosis (Fig. 4C, left and middle panel). For example, no cellular shrinkage or nuclear fragmentation could be observed (the latter shown by nuclear staining with DAPI, a DNA-binding fluorophore - data not shown). Fig. 4C (right panel) shows the morphology of Ogg1+/+ cells. Since typical apoptotic processes require ATP (Nicotera and Leist, 1997; Skulachev, 2006), we determined whether a decrease in ATP levels is related to this unusual cell elimination. Figure 4D shows that Ogg1−/−, Ogg1+/+, LROS (PDL-6, PDL-10) and HROS (PDL-6, PDL-10) cells contain physiological levels of ATP (62±6 nmol per mg protein). However, HROS cells close to the end of their life-span (PDL-17) contained significantly less ATP (Fig. 4D). Prior to measurement of ATP levels annexin V positive cells were discarded after flow cytometric sorting of cell suspensions.

3.5. Transgenic expression of OGG1 decreases 8-oxoG and ROS levels

Parallel cultures of LROS Ogg1−/− and Ogg1+/+ cells were transfected with OGG1 expressing or empty vectors and G418-resistant clones were isolated and individually maintained. OGG1 polypeptide primarily localized to nuclei of cells (Fig. 5A, lower panels). Unexpectedly, a large number of OGG1-transfected clones were lost expressing OGG1 protein. A few clones of OGG1-expressing cells were viable up to PDL-8 and then underwent crises, and were lost or reverted back to OGG1 non-expressing phenotype (Ogg1−/−). Overexpression of OGG1 in Ogg1+/+ cells or vector alone in Ogg1−/− did not affect cell viability. The cause of this unexpected behavior of OGG1-transfected Ogg1−/− clones is currently being investigated. Cells displaying normal morphology were examined for 8-oxoG and ROS levels. The ROS levels in OGG1-expressing cells (PDL-6, -7 and -8) were comparable to Ogg1+/+ cell cultures as observed by microscopic imaging of H2DCF-DA-loaded cells (data not shown). In the DNA of individual OGG1-expressing cells, 8-oxoG levels were nearly identical to the level of 8-oxoG found in Ogg1+/+ cells (Fig. 5B) in line with the physiological levels of ROS.

Figure 5
Expression of OGG1 lowers 8-oxoG levels in Ogg1−/− cells

4. Discussion

8-oxoG and FapyG are the most abundant forms of ROS-induced guanine base damage in the DNA. The relatively high abundance of these DNA base lesions is due to guanine’s low redox potential (Candeias and Steenken, 2000). Accumulation of oxidatively damaged guanines in the DNA has been proposed to play a major role in aging-associated diseases and the aging process itself (Hamilton et al., 2001b; Shigenaga et al., 1994; Van Remmen et al., 2003). Extracts from tissues of homozygous Ogg1−/− mice do not excise 8-oxoG so it is accumulated in a tissue- and cell-type specific manner in both nuclear and mitochondrial DNA (Klungland et al., 1999; Minowa et al., 2000). Fibroblast cultures developed from Ogg1−/− animals contain a steady-state levels of 8-oxoG; however, there are no studies addressing how the steady-levels of 8-oxoG in nuclear DNA are maintained in the absence of OGG1. Our data show that the Ogg1−/− cell line consists of and can be separated into the predominant LROS and HROS subsets, the latter showing up to 27-fold increase in 8-oxoG levels (compared to the Ogg1+/+ cells). HROS cells showed mitochondrial dysfunction, and a limited life-span. LROS Ogg1−/− cells proliferate with a rate comparable to Ogg1+/+ and gradually regenerate cells exhibiting HROS phenotype with high 8-oxoG levels.

Due to environmental and endogenous oxidative stress we would expect continuously increasing 8-oxoG levels in the DNA of the Ogg1−/− cell populations. Our in depth investigations showed that a subset of the Ogg1−/− cells exhibit much higher (up to 27-fold) levels of 8-oxoG in their DNA than the majority (85%) of cells. 8-oxoG levels were determined by two independent but complementary methods. In individual cells, 8-oxoG levels were assessed by a fluorescent immunocytochemical procedure, which employs a well-characterized, 8-oxoG DNA lesion-specific monoclonal antibody (Bespalov et al., 1999). These results were confirmed by comet assay carried out on DNA treated with Fpg, which predominantly excises 8-oxoG and FapyG (Banath et al., 1999).

In the majority of Ogg1−/− cells 8-oxoG levels are maintained at a relatively low (~3-fold over Ogg1+/+) level. This raises the possibility that NEIL1 and/or NEIL2, recently discovered DNA glycosylases (Dou et al., 2003; Hazra et al., 2002) have a role in maintaining steady-state 8-oxoG/FapyG levels in the DNA. NEIL1 activity has been shown to be present in various organs of Ogg1−/− mice (Hu et al., 2005). However, NEIL1’s (replication coupled repair protein) activity in cultured Ogg1−/− fibroblast may be dispensable because there was no significant accumulation of the 8-oxoG/FapyG observed in Ogg1−/− cells when they were kept confluent and proliferation was arrested (Osterod et al., 2001). Moreover, removal of oxidative base modifications induced by photosensitization were 4-fold slower in Ogg1−/− than in Ogg1+/+ cells and no differences were found between confluent and proliferating fibroblasts (Osterod et al., 2001). These results suggest that the NEIL1- or NEIL2-initiated replication or transcription-coupled repair of 8-oxoG and/or FapyG residues is not significant in these cells. Thus, steady-state levels of oxidized guanines cannot be fully explained by the existence of these alternative repair pathway(s) in the absence of OGG1 (Osterod et al., 2001; Russo et al., 2006).

Increased DCF fluorescence allowed us to sort Ogg1−/− cells to establish cultures containing low or high ROS levels. The fluorescence intensity in LROS cells were 2-times lower than in HROS ones. When the nearly homogenous LROS cells were serially passaged, unexpectedly, the HROS cells appeared again with significantly higher 8-oxoG levels (up to 27-fold) and their percentage reached a level similar to unsorted Ogg1−/− cells by PDL-32. Microscopic imaging showed that fluorescence mediated by H2Et/superoxide reaction products (Zhao et al., 2003) were localized primarily to mitochondria. Consistent with these results, we found a markedly increased H2O2 release from mitochondria of HROS cells vs. LROS and unsorted Ogg1−/− or Ogg1+/+ cells. The amounts of H2O2 released from mitochondria of unsorted Ogg1−/− and Ogg1+/+ cells were nearly identical. These latter observations are consistent with the functionally normal mitochondria isolated from livers and hearts of Ogg1−/− mice, despite their 20-fold higher levels of 8-oxoG (de Souza-Pinto et al., 2001). Thus it appears that without isolating subsets of cells from organs or tissues, mitochondrial respiratory dysfunction cannot be detected.

An increased ROS level alone is sufficient to trigger intrinsic pathways that culminate in mitochondrial dysfunction, release of cytochrome c and other factors leading to cell death (Liu et al., 1996). NAC in the culture medium decreased, while BSO increased ROS levels and the frequency of lost cells, which strongly suggest implication of ROS in cell death processes. HROS subset of Ogg1−/− cultures exhibited mitochondrial dysfunction indicated by increased formation of H2O2. These cells showed increased caspase-3 activity; further implicating ROS in the elimination of these cells from the population. However, these HROS cells were not eliminated by typical apoptotic processes, in which caspase-mediated proteolityc and DNase activities result in shrinkage of cells and nuclear fragmentation as shown previously (Zakeri and Ahuja, 1997). Therefore, we termed this type of cell death, apoptosis-like. Apoptosis-like cell death in response to oxidative stress and DNA damage have been described in yeast and mammalian cells as well (Burhans et al., 2003; Malecki, 2001).

It is well established that apoptosis requires ATP for the execution of steps in its ordered program (Nicotera and Leist, 1997). In the presence of dATP apoptosomes are formed, which in turn activate caspase-9 and caspase-3 (Liu et al., 1996). Our data showed significantly lower ATP levels in HROS cells near the end of their life-span (e.g., PDL-16, -18). Therefore the absence of typical morphological apoptotic changes (e.g., nuclear fragmentation) is not surprising. It is possible that in the presence of decreased ATP levels in HROS cells; the death-pathway share initial common events with apoptosis, such as increase in caspase-3 activity (at higher PDLs of HROS cells) until they reach energy exhaustion as shown in other systems (Liu et al., 1996; Nicotera and Leist, 1997).

8-oxoG is one of the most mutagenic lesions in DNA resulting in genomic instability, which may lead to reversal of immortalization in subsets of Ogg1−/− cells or accumulation of tumorigenic mutation(s) both inducing senescence-like response (Reed, 1999; Sager, 1991). Indeed, near the end of their lifespan HROS cells were enlarged and underwent cell cycle arrest, much like senescent cells. It has been shown that senescent cells become resistant to apoptotic signals (Campisi, 2003), so it is possible that HROS subset of Ogg1−/− cultures containing extreme levels of 8-oxoG and increased ROS levels is eliminated by senescence-related apoptotic processes.

It has been shown that transgenic expression of OGG1 in Ogg1−/− cells restored the wild-type phenotype in response to all pro-oxidants tested (Smart et al., 2006). In our hand, overexpression of OGG1 decreased 8-oxoG levels in the DNA and prevented development of HROS cells in the surviving clones of OGG1 expressing cells. These data suggest that supraphysiological levels of 8-oxoG in the DNA could be responsible for increased levels of ROS and cell death. Our most intriguing finding was that we were not able to develop stably-transfected OGG1 expressing cell lines from Ogg1−/− cells. These OGG1-expressing cells undergo typical apoptotic processes (including e.g., shrinkage of cells, nuclear fragmentation). On the other hand, the viability of Ogg1+/+ cells was not affected when stably transfected with the pcDNA3.0-neo expressing OGG1. Similarly, G418 resistant Ogg1−/− cells were viable after transfection with empty pcDNA3.0-neo vector. We speculate that OGG1 protein resulted in supraphysisological levels of nicks and single-strand breaks in the DNA of Ogg1−/− cells, which cause apoptosis and so led to loss of cell clones. This hypothesis is supported by data showing DNA damage-induced apoptosis (Zabkiewicz and Clarke, 2004). Another possibility is that 8-oxoG excised from DNA of OGG1-expressing cells may induce apoptosis. In support of the latter possibility, it has been reported that KG-1 human acute leukemia cells that have mutational loss of OGG1 activity and accumulate supraphysisological level of 8-oxoG into DNA like Ogg1−/− cells undergo G1-phase cell cycle arrest and apoptotic death after treatment with 8-oxoG (Hyun et al., 2006). However, it appears that increased 8-oxoG levels in the DNA of KG1 and Ogg1−/− cells do not interfere with transcription and DNA replication (Larsen et al., 2004). This assumption was supported by Arai et al., 2002, who demonstrated that KBrO3-treated Ogg1−/− liver cells showing more than a 70-fold increase in oxidative guanine lesions regenerated normally (Arai et al., 2002). In line with these observations, 8-oxoG failed to block RNA polymerase II at the lesion in experiments with purified transcription factors in vitro (Larsen et al., 2004). Similarly, 8-oxoG only slightly inhibited T7 RNAPol II-mediated transcription in vitro and complete lesion bypass was observed when the 8-oxoG lesion was positioned in the non-transcribed strand (Tornaletti et al., 2004).

In conclusion, we show here that Ogg1−/− cultures consist of two subpopulations LROS and HROS cells. HROS subsets of Ogg1−/− cells show supraphysiological levels of 8-oxoG in their DNA, mitochondrial dysfunction, decreased ATP synthesis and elimination by apoptosis-like processes, which could be responsible for preventing unlimited accumulation of 8-oxoG in DNA of Ogg1−/− cell cultures. The most intriguing question is how the HROS subset of Ogg1−/− cells arises when the cells are clonal in nature. We propose that in the absence of OGG1, Ogg1−/− cells are genetically more unstable than Nth1−/− or Ogg1+/+ cells due to accumulation of oxidized guanines that are considered one of the most pro-mutagenic lesions. These mutations are randomly distributed and their particular combinations may cumulate in altered phenotypes such as HROS cells. Altered availability and/or changes in structure and functions of mutated proteins in the respiratory chain result in increased ROS production and thereby more oxidative DNA damage to guanines (and other DNA bases) and additional mutations. Increased ROS production and excessive DNA damage is sufficient to induce processes required for elimination of cells via apoptosis or apoptosis-like processes as described in other cell culture models (Dobson et al., 2000; Ruchko et al., 2005). In Ogg1 knock out mice, oxidative damage to guanines are accumulating in a tissue dependent manner especially under oxidative stress conditions (Arai et al., 2002; Klungland et al., 1999; Minowa et al., 2000; Nishimura, 2002; Osterod et al., 2001), thus further studies are required to explore whether similar phenomena exist in vivo in tissues and organs of these animals.


This work was supported by P01 AG 021830 (I.B., D.K., T.H) from the NIH/NIA, P01 AI062885-01 (I.B) from the NIAID, NIH/NCI RO1 CA102271-01A (T.H), and NIEHS Center Grant, EOS 006677.


3-nitropropionic acid
antimycin A
base excision repair
formamidopyrimidine DNA glycosylase
2′,7′-dichlorodihydro-fluorescein diacetate
thymine glycol-DNA glycosylase
8-oxoguanine DNA glycosylase
population doubling level
superoxide dismutase
reactive oxygen species
cells showing physiological ROS levels
cells showing supraphysiological levels of ROS


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