A multi-biomarker approach to study the effects of smoking on oxidative DNA damage and repair and antioxidative defense mechanisms
A.Besarati Nia,
F.J. Van Schooten,
P.A.E.L. Schilderman,
T.M.C.M. De Kok1,
G.R. Haenen1,
M.H.M. Van Herwijnen,
E. Van Agen,
D. Pachen and
J.C.S. Kleinjans2
Department of Health Risk Analysis and Toxicology, Maastricht University, PO Box 616, 6200 MD, Maastricht, The Netherlands and
1 Department of Pharmacology, Maastricht University, PO Box 616, 6200 MD, Maastricht, The Netherlands
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Abstract
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We investigated the effects of smoking-induced oxidative stress in healthy volunteers (21 smokers versus 24 non-smokers) by quantifying various markers of oxidative DNA damage and repair, and antioxidative defense mechanisms. Lymphocytic 7-hydroxy-8-oxo-2'-deoxyguanosine (8-oxo-dG) levels measured by high performance liquid chromatography with electrochemical detection, were significantly lower in smokers as compared with non-smokers (38.6 ± 5.2 versus 50.9 ± 4.6/106 dG, P = 0.05). The levels of oxidized pyrimidine bases in lymphocytes of smokers quantified by the endonuclease III-modified comet assay were non-significantly lower than those of non-smokers (% DNA in tail: 13 ± 3 versus 14 ± 2; tail length: 69 ± 13 versus 96 ± 10; tail moment: 6416 ± 1220 versus 7545 ± 1234). Urinary excretion levels of 8-hydroxy-2'-deoxyguanosine (8-OH-dG) assessed by enzyme-linked immunosorbent assay did not differ significantly between smokers and non-smokers (197 ± 31 versus 240 ± 33 ng/body mass index, P = 0.3). Overall DNA repair activity expressed as unscheduled DNA synthesis in blood leukocytes, was not significantly different between smokers and non-smokers (2.9 ± 0.3 versus 3.3 ± 0.3, P = 0.4). Plasma antioxidative capacity measured by the Trolox equivalent antioxidant capacity assay was slightly higher in smokers as compared with non-smokers (440 ± 16 versus 400 ± 15 µM Trolox equivalent, P = 0.09), and it was significantly related to lymphocytic 8-oxo-dG levels (r = 0.4, P = 0.001). Genotyping of human 8-OH-dG glycosylase/apurinic lyase and glutathione S-transferase M1 showed that a polymorphism in either or both of the two genes does not affect any of the quantified biomarkers. We conclude that oxidative stress imposed by cigarette smoking has a low impact upon certain pathways involved in DNA damage and the antioxidative defense system.
Abbreviations: ABAP, 2,2'-azobis-2-amidinopropane; BER, base excision repair; dG, 2'-deoxyguanosine; ELISA, enzyme-linked immunosorbent assay; GSTs, glutathione S-transferases; OGG1, 8-hydroxy-2'-deoxyguanosine-glycosylase/apurinic lyase; HPLC-ECD, high performance liquid chromatography-electrochemical detection; NER, nucleotide excision repair; 8-OH-dG, 8-hydroxy-2'-deoxyguanosine; 8-oxo-dG, 7-hydroxy-8-oxo-2'-deoxyguanosine; 8-oxo-G, 8-oxo-7,8-dihydroguanine; PBL, peripheral blood lymphocytes; PBS, phosphate buffered saline; PCR, polymerase chain reaction; ROS, reactive oxygen species; TEAC, Trolox equivalent antioxidant capacity; UDS, unscheduled DNA synthesis.
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Introduction
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Reactive oxygen species (ROS) is a collective term used for compounds containing partially reduced oxygen and possessing high reactivity with biomolecules, e.g. DNA, proteins and lipids (15). The ROS are generated during cellular respiration, intra-cellular signal transduction, phagocytosis and metabolism of xenobiotics (3,612). Oxidative stress occurs when the cell is subjected to an increased amount of endogenous and/or exogenous ROS (3,9). To counteract the oxidative stress, aerobic organisms have evolved various defense mechanisms (13). Plasma antioxidants, which scavenge the ROS prior to their interactions with cellular components, are the first line of defense against exogenous oxidative stress (1315). Plasma antioxidants are comprised of water-soluble antioxidants: ascorbic acid, uric acid, protein thiols and bilirubin; and lipid-soluble antioxidants:
-,
-tocopherols, ubiquinol, lycopene, carotenoids and oxycarotenoids (16). A triple enzyme, superoxide dismutase, catalase and glutathione peroxidase, along with a phase II family of enzymes, glutathione S-transferases (GSTs), constitute the secondary defensive system against endogenous and/or exogenous oxidative stress. The system reduces ROS to less reactive metabolites and then, to excretable endproducts (15,17). Theoretically, polymorphisms of genes encoding for the above-mentioned enzymes may account for inter-individual variability in handling oxidative stress. In humans, a notable polymorphism is GSTM1 homozygous allelic loss, GSTM1 null genotype, which is prevalent in ~50% of Caucasians and associated with several types of cancer (1822). Lastly, a complementary repair system exists in case ROS escape the first two defense lines and assault the cellular targets (23,24). An example of this system is the 8-hydroxy-2'-deoxyguanosine-glycosylase/apurinic lyase gene (OGG1) which repairs 8-oxo-7,8-dihydroguanine (8-oxo-G), the most abundant and highly mutagenic ROS-induced DNA lesion (2533). In Escherchia coli, polymorphic variants of OGG1 vary in their enzymatic effectivity to suppress spontaneous mutagenesis (34). It is assumed that in humans, polymorphism of the homologue gene (hOGG1) might also have a functional impact on the repair of 8-oxo-G (34,35). Of most interest is the SerCys polymorphism of hOGG1 at codon 326, which results in encoded enzymes with varying activities (36).
In the present study, we investigated the effects of oxidative stress imposed by cigarette smoking, a well-documented source of ROS (3739), on various pathways involved in DNA damage and repair as well as antioxidative defense mechanisms. For this purpose, we quantified the following in a group of smokers as compared with non-smokers. (i) Levels of 7-hydroxy-8-oxo-2'-deoxyguanosine (8-oxo-dG) in peripheral blood lymphocytes (PBL) by means of reversed phase high performance liquid chromatography with electrochemical detection (HPLC-ECD). (ii) Levels of oxidized pyrimidine bases by means of endonuclease III-modified comet assay. (iii) Levels of the repair product of 8-hydroxy-2'-deoxyguanosine (8-OH-dG) excreted in urine by means of a competitive enzyme-linked immunosorbent assay (ELISA). (v) Overall DNA repair activity in blood leukocytes by means of unscheduled DNA synthesis (UDS) assay. (iv) The scavenging capacity of plasma antioxidants by means of Trolox equivalent antioxidant capacity (TEAC) assay. (vi) Genetic polymorphisms of GSTM1 and hOGG1 by means of the polymerase chain reaction (PCR) technique.
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Materials and methods
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Study population
Volunteers were recruited by advertising in the local newspapers. Upon enrollment, every individual signed an informed consent and filled out a comprehensive questionnaire on his/her age, gender, occupation, lifestyleincluding smoking and drinking behavioursfresh fruit and vegetable consumption as well as vitamin supplementation, medical history of disease and family history of cancer. The study population consisted of two groups: healthy smokers (n = 21) and lifelong non-smokers (n = 24) with an average age of 39 ± 2.4 years (range: 2454) and 34 ± 1.7 years (range: 2343), respectively. The participants had no occupational exposure to known or suspected sources of ROS, and received no dietary or medicinal vitamin supplementation. Detailed characteristics of the study population are presented in Table I
. The study was approved by the Medical Ethical Commission of Maastricht University.
Peripheral blood
Thirty millilitres of venous blood were drawn into heparinized Venoject®II tubes (Terumo Europe N.V., Leuven, Belgium). Plasma and lymphocyte fractions were isolated according to the standardized gradient centrifugation procedure (40). Part of the isolated lymphocytes was used for immediate analyses, and the remainder was pelleted and preserved at 80°C until DNA isolation.
Urine
Twenty-four-hour urine was collected in polyethylene bottles. The volume of the sample was measured, and after agitation, aliquots (2x1 ml) of the homogenized urine were kept at 80°C until further analysis.
DNA isolation
DNA isolation was performed as described previously (41). Briefly, the cell pellet was lysed with Proteinase K (0.2 µg/µl) in 1% SDS/1 mM EDTA, pH 8.0, for 30 min at 37°C. To prevent artifactual formation of 8-OH-dG during extraction procedure, 0.1% 8-hydroxy-quinoline was added to phenol. Nucleic acids were repeatedly extracted with phenol, phenol:chloroform:isoamyl alcohol (25:24:1) and chloroform:isoamyl alcohol (24:1), precipitated with 0.1 vol 1 M sodium acetate, pH 6.0, in 2 vol 100% ice-cold ethanol, washed with 1 vol 70% ethanol and then, dissolved in 5 mM Tris/1 mM EDTA, pH 7.4. The resultant was treated with an RNase mixture (RNase A, 100 µg/ml and RNase T1, 50 U/ml) for 30 min at 37°C and subsequently, the DNA content of the solution was recovered using the extraction, precipitation and dissolving procedures as described above. Quality and quantity of the DNA were determined spectrophotometrically (A230/260: ~0.4, A260/280: ~1.8) and, ultimately, its concentration was adjusted to 1 mg/ml.
HPLC-ECD for determining 8-oxo-dG in PBL
HPLC-ECD of lymphocytic 8-oxo-dG was performed as described previously (42). Briefly, DNA was digested into deoxyribonucleosides after treatment with nuclease P1 (0.02 U/µl) and alkaline phosphatase (0.01 U/µl). The digest was then injected into a Spectroflow 480 isocratic pump (Gynkotek, München, Germany) coupled with a Spectroflow 783 injector (Kratos, Ramsey, NJ) and connected to a SupelcosilTM LC-18S column (250x4.6 mm) and an electrochemical detector CU-04-AZ (Antec, Leiden, the Netherlands). The mobile phase consisted of 10% aqueous methanol containing 25 mM sodium acetate, 12.5 mM citric acid, 30 mM sodium hydroxide and 10 mM acetic acid. Elution was performed at a flow rate of 1.0 ml/min with a lower detection limit of 40 fmol absolute for 8-oxo-dG, or 1.5 residues/106 2'-deoxyguanosine (dG). dG was simultaneously monitored at 260 nm. Results were expressed as the ratio of the determined 8-OH-dG to dG.
Endonuclease III comet analysis
The endonuclease III-modified comet assay was performed according to the method of Collins et al. (43). Briefly, PBL (1x105) were embedded in a layer of low melting point agarose on a microscope slide and subsequently, lysed with 2.5 M NaCl, 0.1 M Na2EDTA, 10 mM TrisHCl and 1% Triton X-100, pH 10.0 for 1 h at 4°C. The slides were rinsed with endonuclease buffer [40 mM HEPES, 0.1 M KCl, 0.5 mM EDTA and bovine serum albumin (0.2 mg/ml, pH 8.0)] and then treated with either endonuclease III (1:500), or the endonuclease buffer for 30 min at 37°C (a non-treated control slide was also included). After alkaline electrophoresis, neutralization and ethidium bromide staining, the slides were evaluated under a fluorescence microscope. Quantification was done by means of an Image Processing and Analysis System (Quantimet 500, Leica, Cambridge, UK) which measured the percentage of DNA in tail, tail length and tail moment (% DNA in tail multiplied by tail length) in 50 cells per slide. To correct for background levels, all measured parameters in endonuclease buffer-treated slides were subtracted from respective values in endonuclease III-treated slides. Results were expressed in arbitrary units established by the Image Analyzer.
ELISA for determining 8-OH-dG in urine
Urinary 8-OH-dG was quantified by means of an ELISA kit (Japan Institute for the Control of Aging, Shizuoka Pref., Japan) the validity and comparability of which to HPLC-ECD had already been verified (4447). Briefly, samples and standards of 8-OH-dG (0.5, 2, 8, 20, 80 and 200 ng/ml) were placed in microtiter plates (8x12 wells, split type) pre-coated with 8-OH-dG. A monoclonal primary antibody recognizing 8-OH-dG was then added and the plates were incubated for 1 h at 37°C. They were then rinsed with phosphate buffered saline (PBS, pH 7.4) and subsequently incubated with an enzyme-labelled secondary antibody for 1 h at 37°C. After rinsing with PBS, the chromatic substrate, 3,3',5,5'-tetramethylbenzidine, was added and incubation was carried out for 15 min at 37°C in the dark. The reaction was terminated by adding 1 N phosphoric acid and after a lag time of ~3 min, the plates were read by a Microplate Reader measuring the absorbance at 450 nm. A standard curve was established by plotting the measured absorbance versus log concentration of the 8-OH-dG standards. Results were expressed in ng/body mass index (BMI).
UDS for determining overall DNA repair
UDS analysis of the samples was performed as described previously (48). Briefly, unstimulated blood leukocytes were cultured in the presence of [methyl-3H]thymidine (10 µCi/ml) (Amersham, Buckinghamshire, UK) for 24 h in a humidified incubator at 37°C under 5% CO2. The cells were hypotonized with 75 mM KCl for 10 min, fixed with methanol:acetic acid (3:1) and subsequently, dropped onto microscope slides. The slides were dipped into Illford K2 Scientific Emulsion (Illford, Cheshire, UK), drained on an absorbent paper and then, kept for a period of 1014 days at 4°C in the dark. Afterwards, the slides were developed in photo emulsion at 21°C, fixed [fixative: sodium thiosulfate (200 g) + potassium metabisulfite (20 g) + potassium chromosulfate (10 g)/l] and finally, counterstained with hematoxin. The slides were evaluated under a Zeiss Axioskop microscope (Zeiss, Oberkochen, Germany) equipped with a CHOU high-performance CCD camera (Leica) and coupled to an Artek CounterTM 880 (New Brunswick Scientific, New York, NY). Quantification was based on the number of grains scored in 250 nuclei per slide (corrected for extra nucleus grains as background).
TEAC for determining scavenging capacity of plasma antioxidants
The TEAC assay was performed as described previously (49). Briefly, after generation of the long-lived radical anion of ABTS in the presence of a thermolabile azo compound, 2,2'-azobis-2-amidinopropane (ABAP), plasma was added and subsequently, absorbance was monitored at 734 nm over a period of 6 min. The decrease in absorbance after addition of the plasma was plotted on a calibration curve established by application of known concentrations of Trolox, an analogue of vitamin E, as standard. Results were expressed as µM Trolox equivalents.
PCR for genotyping GSTM1 and hOGG1
Genotype determination for GSTM1 and hOGG1 was done according to the published PCR-based procedures (50,51). In the case of hOGG1, the (Ser/Ser) variant has been shown to encode for the enzyme with the highest effectivity (36) and so we used this genotype as the reference.
Statistical analysis
Results were expressed as mean ± standard error throughout. All variables were compared between the smoking group and the non-smoking group using the MannWhitney U-test. The relationship between different variables was evaluated by Spearman rank correlation analysis. To assess the impact of various independent variables on a dependant variable, a multiple linear regression analysis was performed using the logarithmically-transformed dependant variable and up to three independent variables. Statistical significance was considered at P
0.05.
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Results
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Table II
summarizes the quantified biomarkers used in this study. Lymphocytic 8-oxo-dG levels in smokers were significantly lower than in non-smokers (38.6 ± 5.2 versus 50.9 ± 4.6/106 dG, P = 0.05) (Figure 1
). In smokers and non-smokers, respectively, the levels of lymphocytic 8-oxo-dG varied between the range of 1775/106 dG and 2883/106 dG. In smokers, there was no significant relationship between lymphocytic 8-oxo-dG levels and questionnaire-derived smoking indices (cigarettes/day and pack years). In both smokers and non-smokers, the levels of lymphocytic 8-oxo-dG were negatively associated with age (r = 0.5, P = 0.07 and r = 0.5, P = 0.02, respectively). Also, lymphocytic 8-oxo-dG levels in female smokers and non-smokers were higher than those in male respectives (48.4 ± 6.9 versus 24.6 ± 4.2/106 dG, P = 0.02 and 52.4 ± 5.1 versus 45.4 ± 10.5/106 dG, P = 0.5, respectively). Multiple regression analysis with age, gender and smoking status as independent variables showed that the levels of lymphocytic 8-oxo-dG can be best predicted by age (r = 0.6, P = 0.003). After adjustment for age, smokers still had significantly lower levels of lymphocytic 8-oxo-dG than non-smokers (P = 0.03) (Figure 1
).

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Fig. 1. Distribution of the levels of 8-oxo-dG in peripheral blood lymphocytes of smokers and non-smokers. The lower and upper edges of the boxes are the 25th and the 75th percentiles, respectively. The black ellipses and the lines within the boxes are the means and the medians, respectively. The lower and upper bars are the 10th and the 90th percentiles, respectively. Individual values below the 10th or above the 90th percentiles are shown as (o). 1Adjusted for age and expressed in arbitrary units. 2Statistically significant as compared with non-smokers, P = 0.05. 3Statistically significant as compared with non-smokers, P = 0.03.
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In the simple comet assay, there was no difference in any of the measured parameters between smokers and non-smokers (% DNA in tail: 16 ± 0.4 versus 16 ± 0.9; tail length: 173 ± 19 versus 174 ± 17; tail moment: 2710 ± 294 versus 2938 ± 406). In the endonuclease III-modified comet assay, the differences were more pronounced, yet non-significant (% DNA in tail: 13 ± 3 versus 14 ± 2; tail length: 69 ± 13 versus 96 ± 10 and tail moment: 6416 ± 1220 versus 7545 ± 1234). Neither simple nor endonuclease III-modified comet data correlated with the data on smoking status or gender or age. Overall, endonuclease III-modified comet parameters tended to be associated with lymphocytic 8-oxo-dG levels (adjusted for age) (r = 0.3, P = 0.2).
Urinary excretion levels of 8-OH-dG did not differ between smokers and non-smokers (197 ± 31 versus 240 ± 33 ng/BMI, P = 0.3). The levels of urinary 8-OH-dG ranged from 78 to 449 ng/BMI in smokers and from 60 to 472 ng/BMI in non-smokers. Urinary excretion levels of 8-OH-dG in smokers were non-significantly related to smoking index (cigarettes/day: r = 0.4, P = 0.07). Also, there was an inverse correlation between urinary 8-OH-dG of smokers and their lymphocytic 8-oxo-dG levels (r = 0.4, P = 0.09, adjusted for age) (Figure 2
). No association was found between urinary excretion of 8-OH-dG and comet parameters, or gender or age.

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Fig. 2. Relationships between lymphocytic 8-oxo-dG and urinary 8-OH-dG (r = 0.4, P = 0.09) and scavenging capacity of plasma antioxidatives (r = 0.7, P = 0.006) in smokers. 1Adjusted for age and expressed in arbitrary units. 2Trolox equivalent antioxidant capacity.
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There was no significant difference in UDS between smokers and non-smokers (2.9 ± 0.3 versus 3.3 ± 0.3, P = 0.4). In smokers and non-smokers, UDS levels ranged from 1.6 to 5.0 and from 1.3 to 5.5, respectively. The UDS of smokers did not relate to smoking indices. Nor did UDS correlate with urinary 8-OH-dG or lymphocytic 8-oxo-dG (adjusted for age), or comet parameters, or gender or age.
The scavenging capacity of plasma antioxidants in smokers was non-significantly higher than that in non-smokers (440 ± 16 versus 400 ± 15 µM Trolox equivalent, P = 0.09). The scavenging capacities of plasma antioxidants were in the range of 358545 µM Trolox equivalent in smokers, and 307488 µM Trolox equivalent in non-smokers. In smokers, the scavenging capacity of plasma antioxidants was not related to smoking indices. The scavenging capacities of plasma antioxidants in male smokers and non-smokers were higher than those in female smokers and non-smokers, respectively (474 ± 27 versus 416 ± 15 µM Trolox equivalent, P = 0.1 and 479 ± 18 versus 373 ± 14 µM Trolox equivalent, P = 0.002, respectively). Overall, there was an inverse relationship between plasma antioxidative capacity and age (r = 0.5, P = 0.003, adjusted for gender). Multiple regression analysis with gender and age and smoking status as independent variables showed that the scavenging capacity of plasma antioxidants can only be significantly predicted by gender (r = 0.6, P = 0.0007). Adjusting the data for gender, there was no difference in plasma antioxidative capacity between smokers and non-smokers (P = 0.9). Overall, there was a significant relationship between plasma antioxidative capacity and lymphocytic 8-oxo-dG levels (r = 0.4, P = 0.01, adjusted for gender and age); multiple regression analysis revealed that this relationship had a major impact on the correlation between urinary 8-OH-dG and lymphocytic 8-oxo-dG in smokers (Figure 2
). No association was found between the scavenging capacity of plasma antioxidants and UDS, or urinary 8-OH-dG or comet parameters. In smokers, multiple regression analysis with plasma antioxidative capacity, urinary 8-OH-dG and UDS as independent variables showed that the levels of lymphocytic 8-oxo-dG can be best predicted by the scavenging capacity of plasma antioxidants (r = 0.9, P = 0.007).
Prevalence of GSTM1 null genotype, GSTM1(/), in smokers, in non-smokers and smokers and non-smokers overall was 65, 62 and 54%, respectively. Overall, GSTM1(/) individuals did not differ from GSTM1(+) genotypes, regarding age, gender and smoking status. GSTM1 genotyping data were not associated with the data on the scavenging capacity of plasma antioxidants (adjusted for gender), UDS, comet analysis, urinary 8-OH-dG or lymphocytic 8-oxo-dG (adjusted for age) (Figure 3
).

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Fig. 3. Association between GSTM1 and hOGG1 genotypes and lymphocytic 8-oxo-dG and urinary 8-OH-dG and UDS. The horizontal bars within the scattered values represent the mean of the variables in each group. 1Adjusted for age and expressed in arbitrary units.
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Prevalence of hOGG1 (Ser/Ser), (Ser/Cys) and (Cys/Cys) genotypes was: 57.1, 42.9 and 0.0% in smokers; 65.2, 30.4 and 4.4% in non-smokers; and 61.4, 36.4 and 2.2%, smokers and non-smokers overall. There was no significant difference between individuals with the hOGG1 (Ser/Ser) genotype and those with the (Ser/Cys) or (Cys/Cys) genotypes in terms of age, gender and smoking status. Overall, there was no association between hOGG1 genotyping data and the data on the scavenging capacity of plasma antioxidants (adjusted for gender and age), UDS, comet analysis, urinary 8-OH-dG or lymphocytic 8-oxo-dG (adjusted for cigarette/day and age, respectively) (Figure 3
). Possession of hOGG1 (Ser/Ser) and GSTM1(+) genotypes did not affect additively or multiplicatively any of the above-mentioned parameters.
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Discussion
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In this study, we observed lower levels of the well-established marker of oxidative DNA damage, 8-oxo-dG, in lymphocytes of smokers as compared with non-smokers. This is in agreement with our previous observation (41) as well as with one recent report by van Zeeland et al. (52). However, two other studies in leukocytes and lung tissues have shown the opposite result (53,54). As we demonstrated, lymphocytic 8-oxo-dG levels are mainly dependent upon age and smoking status, whereas the contradicting lung study provided no indication of age adjustment in its analysis (54). In addition, the exposure variable may have also contributed to this discrepancy because the lung in that study, as opposed to the PBL in the present study, has relatively higher and more direct exposure to smoking materials. In the other contradictory report (53), the investigated population consisted of heavy smokers with intense smoking behaviours (high lifetime consumption of cigarettes with an average of 34 cigarettes/day). Our study population was, however, made up of moderate smokers. Interestingly, the characteristics of the study populations in both confirming reports are comparable to those in our report (41,52). Furthermore, we found a tendency towards lower levels of oxidized pyrimidine bases in comet analysis of lymphocytes of smokers versus non-smokers, which is in contrast to two other reports (43,55). This latter discrepancy may also be ascribed to the apparent differences in the characteristics of the examined populations. At the same time, the less pronounced difference in comet-quantified parameters versus lymphocytic 8-oxo-dG between smokers and non-smokers questions the sensitivity of the comet assay for detecting smoke-related oxidative lesions. Taken together, it seems that a modulation occurs in smoking-induced oxidative DNA damages as a result of up-regulation of the primary defense antioxidative system and/or complementary DNA repair system.
Regarding the DNA repair pathway, we observed no significant difference in urinary excretion levels of 8-OH-dG between smokers and non-smokers. Apparently, previous investigations on urinary excretion of this repair product in smokers have not been conclusive either (5659). Also, we did not find any significant difference in DNA repair activity, expressed as UDS, between smokers and non-smokers. It should be emphasized that our UDS analysis was performed in blood leukocytes comprising several cell types with varying DNA repair activity rates (60,61). However, we measured the extent of oxidative DNA damage only in lymphocytes, which comprise ~35% of the total white blood cells (62). Besides, the quantified UDS is a collective index for both base excision repair (BER) and nucleotide excision repair (NER) activities (48), whereas most oxidative DNA lesions are repaired via the BER pathway alone (24). In our study, polymorphism of the specified BER gene, hOGG1, could not impact upon the corresponding markers. It should be noted that although in in vitro systems multi-variant products of hOGG1 have been shown to differently repair 8-oxo-G (34), there has not been any association between the polymorphic variants of this gene and the level of 8-oxo-dG in in vivo studies. Nor have hOGG1-encoded enzymes been associated with susceptibility to lung, kidney or gastric cancer (34,35,51,63,64). It seems that although hOGG1 constitutes a major pathway of repair for 8-oxo-G, other genes such as hMYH and hMTH may have considerable impacts on the repair of this lesion (32,65).
Thus far, there have been controversial reports on the effects of oxidative stress on the scavenging capacity of plasma antioxidants (16,6670). For example, the trapping activity of plasma antioxidants has been shown to be inversely/positively related to strenuous exercise and physical activities, smoking, lung cancer and neonatal prematurity, all imposing a massive influx of ROS. In the present study, we observed an up-regulation of this primary defense system in males, which may relate to the fact that males are under greater oxidative burden due to their higher metabolic rate (71), as well as in smokers as compared with females and non-smokers, respectively. We then found that such a trend was mostly gender-related because adjustment of data for gender bridged the gap between smokers and non-smokers. This together with the positive linearity between lymphocytic 8-oxo-dG and plasma antioxidants scavenging capacity imply that the antioxidative defense system is only marginally affected by smoking-induced oxidative stress. In other words the down-regulation of oxidative DNA damage in smokers is likely to be due to modulatory mechanisms other than plasma antioxidants scavenging capacity.
With regard to the ROS detoxifying system, we did not find an influence of GSTM1 on any of the quantified biomarkers. It is conceivable that GSTM1 is only one of the genes whose product is involved in detoxification of ROS. Therefore, it is possible that other genes encoding for the influential detoxifying enzymes, e.g. superoxide dismutase, catalase and glutathione peroxidase, in combination with or separate to GSTM1, play a more crucial role in response to oxidative stress (15,17).
In summary, we have demonstrated that smoking-induced oxidative stress may impact upon certain pathways involved in DNA damage and antioxidative defense mechanisms. However, given the marginal effects of smoking and contributory effects of host factors, e.g. gender and age, on these pathways, it is unlikely to observe an explicit adaptive response towards smoking, especially in light and moderate smokers. To better understand the effects of smoking-induced oxidative stress in humans, future large-scale research is needed to elucidate the role of relevant polymorphic genotypes specifically, encoding for DNA repair enzymatic networks.
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Notes
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2 To whom correspondence should be addressed Email: j.kleinjans{at}grat.unimaas.nl 
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Acknowledgments
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We would like to thank Prof. Ch.P.Wild, Dr L.J.Hardie, and Ms J.A.Briggs of the Molecular Epidemiology Unit, University of Leeds, for performing the hOGG1 analysis.
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Received September 7, 2000;
revised November 6, 2000;
accepted November 7, 2000.