Abstract
Background: No previous in vitro studies have tested radio frequency radiation for at least one full cell cycle in culture. The aim was to test if exposure used in mobile phones and wireless network technologies would induce DNA damage in cultured human lymphocytes with and without a known clastogen. Materials and Methods: Lymphocytes from six donors were exposed to 2.3 GHz, 10 W/m2 continuous waves, or 2.3 GHz, 10 W/m2 pulsed waves (200 Hz pulse frequency, 50% duty cycle). Mitomycin C was added to half of the cultures. DNA synthesis and repair were inhibited in one experiment. Results: No statistically significant differences were observed between control and exposed cultures. A weak trend for more chromosomal damage with the interaction of pulsed fields with mitomycin C compared to a constant field was observed. Conclusion: Exposure during the whole cell cycle in inhibited cultures did not resulted in significant differences in chromosomal aberrations as compared to controls.
In recent years there have been considerable advances in the development of devices emitting microwaves and radiofrequency electromagnetic radiation (RFR) such as mobile phones. Sienkiewicz and Kowalczuk (1) in a review of 26 previous reviews found that most of the reports reached a similar conclusion, namely that exposure to low level RFR may cause a variety of subtle biological effects. However, the possibility of exposure causing adverse health effects remained unproven. Gøtrik et al. (2), in a common approach for the Nordic competent authorities, agreed that there is no scientific evidence for any adverse health effects from mobile telecommunication systems, but concluded that a knowledge gap existed that justified more research in this field. Moulder et al. (3) in a recent review concluded that the current evidence for a causal association between cancer and exposure to RFR is weak and unconvincing. When epidemiological evidence is weak, partly due to short follow-up periods, laboratory studies are essential and in vitro studies are a good supplement if the results are replicable and consistent across different methods of testing.
Several recent in vitro studies on genotoxic effects of RFR have been published. A few of them reported that exposure of cultured cells to mobile phone RFR caused DNA damage in some assays (4, 5), while other in vitro studies reported no effect on DNA (6-10). Based on a meta-analysis of 63 publications from 1990 to 2005, Vijayalaxmi and Prihoda (11) concluded that the overall genotoxicity indices were similar in the RFR-exposed and control groups when in vitro investigations were conducted under the recommended safety guidelines. A direct genotoxic action is generally thought to be unlikely because of the low energy of microwave photons (12), but several studies have suggested that RFR exposure may enhance the genotoxic effects of other cancer-inducing substances (3, 13, 14).
What kind of exposure is of interest? The third generation of mobile phones, UMTS (Universal Mobile Telecommunication) was introduced in Europe in 2002, using frequencies around 2.0-2.2 GHz. Blue-tooth equipment (2.4 GHz) is used in hands-free equipment for mobile phones and many workplaces have installed equipment for wireless local area network (LAN) communication in rooms and corridors operating in the frequency range 2-2.4 GHz, employing modulated fields. Thus, an exposure for 2.0-2.4 GHz electromagnetic fields might be of particular interest. Since these frequencies have only been used extensively since 2002, the short latency period reduces the chances of an effect being seen in cancer studies. Chromosomal aberrations in lymphocytes have proven to be associated with cancer risk in humans (15-17) and are considered informative indicators of exposure; moreover they can be used in a more immediate screening for adverse effects.
The aim of the present pilot study was therefore to investigate the clastogenetic effects of 2.3 GHz radiation, both pulsed and continuous exposures, alone or in combination with the known clastogen mitomycin C (MMC) on cultured human lymphocytes. In order to enhance the likelihood of discovering a possible effect, we chose to use lymphocytes from several donors, to expose the cells during the whole in vitro period and in one experimental setup to inhibit DNA synthesis and repair in the G2-phase of the cell cycle.
The level of exposure we have chosen corresponds roughly to the safety limit recommended by the International Commission on Non-Ionizing Radiation Protection (ICNIRP) (13). Safety limits are based on induced currents in the human body, and the difference in geometry and material properties makes it difficult to compare exposures in an in vitro experiment to the human body. This problem can be investigated by detailed finite difference method calculations. Since our aim is not to discuss safety guidelines, but rather to perform a pilot study to identify potentially interesting effects that could be followed up more closely in further study, we have not carried out such detailed theoretical calculations. However, we have given enough information about the experimental design so that others can perform finite difference calculations.
Materials and Methods
Lymphocyte donors. After informed consent, four non-smoking healthy individuals, two females (25 and 44 years old) and two males (40 and 44 years old), and two smoking females (41 and 55 years old) were chosen as blood donors. The hospital authorities approved the study, otherwise the research was exempted from review since the blood samples were only used for in vitro studies and the results were registered in an anonymous database. All donors were genotyped for polymorphisms in the glutathione-S-transferase (GST) genes as polymorphisms in these detoxifying enzymes are common in the Caucasian population (18) and are therefore important to control for (19). Individuals with GSTM1 null and GSTP1 heterozygote or wild-type alleles were chosen as donors. Polymorphisms of the GSTM1 (null), GSTT1 (null) and GSTP1 (Ile105Val) genes were analyzed simultaneously by multiplex PCR as described by Nedelcheva Kristensen et al. (20) with minor modifications.
Cytogenetic analysis. Phytohemagglutinin-stimulated whole blood in RPMI medium (1640; BioWhittaker Cambrex, Verviers, Belgium) was cultured using methods as described elsewhere (21). A cell density of 1×106 cells/ml of medium was ensured. Before the experiments were initiated, cultures with different concentrations of mitomycin C (MMC) (Sigma-Aldrich, Oslo, Norway) were performed and harvested at different time intervals. After 48 hours of incubation, very few cells were in mitosis. For the experiments Colcemid (Sigma-Aldrich, Oslo, Norway) (0.3 μg /ml) was added three hours before harvesting at 53 hours. Previous experiences in handling blood from adults have suggested a similar delay in first cell division in our assay system. The cells were harvested and the slides stained with Giemsa according to conventional methods. In addition, chromosomal aberrations were scored in cells where DNA synthesis and repair were inhibited in vitro with hydroxyurea (Sigma-Aldrich) and caffeine (Sigma-Aldrich) both added at a concentration of 7.5×10−2 M three hours before harvesting together with Colcemid (0.3 μg /ml) (22, 23). For the uninhibited cultures, 200 cells per test were scored blind on coded slides by the same three microscopists. For the inhibited cultures, 50 cells per culture were scored due to the increased aberration frequencies in this assay. The same microscopist scored both the exposed and control cultures from the same blood donor. The scoring criteria used were as described by Brøgger et al. (24) and Savage (25) and harmonized among scorers. The effect parameters in the present study were number of cells with aberrations (CA), cells with aberrations and gaps (CAG), chromosome type aberrations (CSA), chromosomal breaks (CAB), chromatid type aberrations (CTA), chromatid breaks (CTB), and chromosome and chromatid gaps (CSG and CTG).
Study design. Positive and negative controls were included in each experiment. Eight cultures of 10 ml of medium were set up for each donor for the uninhibited cultures. Half of the cultures were exposed for 53 hours to RFR either as continuous or pulsed-field exposure. As a positive control and to test if the combination of a known clastogen and RFR exposure would be more detrimental to the cells, MMC (100 ng/ml) was added to half of the cultures with and without RFR exposure for the last 23 hours in vitro. Each experimental setup included cultures from two blood donors (either two females or two males). For each donor, two cultures with and two without MMC were exposed, and two cultures with and two without MMC were controls using 10 ml of medium per culture flask (EasYFlasks™ Nunclon™ flask; Nunc A/S, Roskilde, Denmark). For the setups where inhibition of DNA synthesis and repair were applied, the two cultures for each treatment regime were replaced with four cultures of 3 ml of medium in culture tubes (TubeNunclon™; Nunc A/S) otherwise the setups were identical. For the uninhibited cultures blood, samples from two males and two non-smoking females were exposed to continuous RFR. For the exposure to pulsed RFR, blood from two smoking females were included in the study. The inhibited cultures from all six donors were exposed to pulsed RFR only.
Exposure. The experiments were performed in a large anechoic chamber (5×4×4 m) with temperature stabilized at 37°C (details below). A diagram with a detailed description of the instrumental setup is shown in Figure 1 and the system is illustrated in Figure 2. Both exposed and control samples were kept in the same room, but well separated. The exposure of the control samples was approximately 40 dB lower than that of the exposed samples. The RFR exposure was 2.3 GHz, 10 W/m2 incident continuous wave, or 2.3 GHz, 10 W/m2 incident pulsed-wave (200 Hz pulse frequency, 50% duty cycle). Time averaged values are given. The cells were exposed to the RFR through a rectangular horn antenna (Narda 615, Odd Tvedt & Co, Bergen, Norway). The opening of the antenna was approximately 6×8 cm and was directed upwards, placed approximately 1.2 m beneath the cultures. The exposed area (ca. 30 cm diameter) was constructed to cover only the number of exposed cultures. The culture flasks were placed horizontally on 5-6 mm thick cardboard. The samples were oriented so that the magnetic field was parallel to the long side of the flask or tube, with the electric field horizontally and perpendicular to this direction. The exposure level was determined with a Wandel & Goltermann EMR 300 instrument with a Type 9C probe (Odd Tvedt, Bergen, Norway). For the continuous wave, the electric field was adjusted to 61 V/m rms just outside the cell growth medium containing the cells. This corresponds to 10 W/m2 for a far field situation, which we expect is valid for our geometry. The intensity varied by 1.5 dB across the area where the samples were placed, with the highest intensity in the middle. To compensate for the fact that exposure is somewhat unevenly distributed within the flask and to some degree depends on the position of the flasks within the exposed area, the cultures were stirred and replaced randomly in the exposed area three times during the culture period. The cells were placed in EasY Flasks Nunclone™ culture flasks with 25 cm2 culture area (4 cm wide), or flat-bottomed Nunc Tubes Nunclon™ culture tubes with 5.5 cm2 culture area (1.0 cm wide). The level of the medium was 4 mm in the flat-bottomed flasks and ranged from 2 to 9 mm within the tubes. The flasks or tubes were positioned relatively close to each other as indicated in Figure 2.
Block diagram for the experimental setup for the continuous-wave experiment. For the pulsed-wave, the setup was similar. The temperature regulation setup is not included in the drawing. The incident power desity was measured with a Wandel & Goltermann EMR300 instrument with a 9C probe. The probe was used to map the exposure level across the full area where the samples were placed. The probe was removed during the exposure of the cells. The control samples are not included in this drawing. LF, Low frequency; HF, high frequency.
The anechoic chamber was temperature stabilized to 37.0±0.3°C and the temperature of the samples was determined by the all-room regulation only. The temperature was measured and recorded continuously with two thermocouples interfaced to an INTAB data logger AAC-2 (Intab Interface-teknik AB, Sternkulle, Sweden). The absolute precision of the temperature registration was better than 0.5°C, and the resolution in the recording was 0.1°C. Each probe was placed in an additional culture flask with comparable amount of water as the samples, and placed near the exposed and control area. The temperature recordings showed 37.0±0.3°C throughout the culture period. The standard deviation given represents the fluctuation of the temperature only. Separate control measurements revealed that the temperature in the exposed samples was 0.5°C higher than in the control samples. This difference was not compensated for in this study. The heating elements and fans circulating the room air generated a background magnetic field (50 Hz) of 0.03 μT at the position of the cells. This magnetic field was determined with a Wandel & Goltermann EMR 300 instrument with an EFA-3 probe. The calculated depth of penetration (skin depth) in the vertical direction for the frequency used in this study was approximately 1 cm in the culture medium.
Statistics. For statistical comparisons of the mean values, the Mann-Whitney non-parametric test was performed. The Kruskal-Wallis test was performed to test for possible differences between the control cultures for each individual. A probability level of 0.05, two-tailed, was used to indicate statistical significance. Anova was used to analyze differences between exposed and control cultures for all conventional cultures with gender, MMC, pulsed and continuous field exposure in the model. SPSS 12.0 for windows (SPSS, Chicago, USA) was used for the statistical analyses.
Results
The results are presented as mean values per 100 cells, with standard deviation for the two or four parallel cultures for CA, CSA, CTA and CTG for each experimental setup. The results for the uninhibited cultures exposed to continuous-wave radiation are presented in Table I, with males and non-smoking females separate and summarized. In Table II, the results for cultures exposed to pulsed-wave radiation are presented. In this experiment, two smoking female blood donors were included. There was a significant difference for all types of aberrations between cultures with and without MMC as expected (p<0.05 for CSA, p=0.001 for CA, CTA, CTG). No statistically significant differences were found between control and RFR-exposed cultures regardless of MMC and RFR treatment for either gender nor for combined results using the Mann-Whitney test. No difference between control and RFR-exposed cultures were observed for the remaining effect parameters either (data not shown).
The culture flasks or tubes were placed relatively closely to each other. This figure, in combination with the text, provides sufficient information if one wants to investigate the field pattern inside the samples further, using finite difference methods or similar numerical techniques. The diameter of the outer circle is slightly more than 30 cm. LF, Low frequency; HF, high frequency.
The effect of inhibition of DNA synthesis and repair in the G2-phase of the cell cycle was studied for cultures exposed to pulsed RFR with and without MMC for all six blood donors. The results are presented in Table III. Compared to the uninhibited cultures (Table II), a significant increase was observed for all types of aberrations, except for CSA in cultures without MMC, despite the fact that only 50 cells per culture were scored in the inhibited cultures. No statistical significant differences were observed for any of the chromosomal parameters for controls compared to RFR-exposed cultures, in spite of the increase in number of aberrations.
A comparison of the effect of continuous contra-pulsed RFR exposure was possible for the uninhibited cultures for the non-smoking blood donors. The mean frequencies combined for all donors are presented in Table IV for the exposed cultures, for all the chromosomal parameters scored. A consistent trend for a slightly non-significant increase in mean frequencies for all chromosomal parameters for pulsed-RFR exposure combined with MMC treatment was found. To include the control cultures in the statistical evaluation of continuous contra-pulsed-RFR exposure, the frequency difference between the exposed and control culture was calculated for each individual for each experimental setup separately because non-significant variations were observed for control cultures from the same individual in the different experimental settings using the Kruskal-Wallis test (data not shown). The mean frequency difference for each experimental setting was then used as the effect parameter in an Anova analysis including gender, MMC, pulsed- and continuous-field exposure in the model. None of the included parameters had a significant impact on the effect parameters. A weak trend towards more damage from pulsed-rather than constant-field exposure was observed, as well as a trend towards more damage from the interaction of pulsed fields and MMC. This was observed for all the chromosomal parameters (Table IV).
Discussion
The primary focus of this in vitro pilot study was to evaluate if exposure to continuous or pulsed 2.3 GHz electromagnetic radiation of the type used in different mobile phone and wireless network technology would induce chromosomal damage in human lymphocytes.
Inhibition of DNA synthesis and repair in the G2-phase of the cell cycle was carried out in one experimental setup in order to test if an increase in aberrations would be a more sensitive method for detecting possible differences between control and exposed cultures. The focus was on whether RFR could cause chromosomal damage either alone or in combination with the known clastogenic agent MMC when RFR was applied during a full cell cycle in culture. We used lymphocytes from both smoking and non-smoking individuals in order to see if smoking habits would influence the sensitivity of our test system.
Test cultures were performed before the experiments were initiated. Since few mitoses were observed in the early cultures, most of the cells collected in mitosis from 50 to 53 hours were assumed to be in the first cell division. Even if a few cells should be in the second cell division at 53 hours, the cells were continuously exposed and aberrations could also be expected to appear during the second cell division. Exact cell proliferation measurements were not performed.
Aberrations recorded for uninhibited cultures exposed to 2.3 GHz continuous-wave (CW) radiation. Mean frequencies per 100 cells (standard deviation in brackets) are given for two parallel cultures for two male (M) and two female (F) blood donors.
Aberrations recorded for uninhibited cultures exposed to 2.3 GHz pulse-wave (PW) radiation. Mean frequencies per 100 cells (standard deviation in brackets) are given for two parallel cultures for two male (M), two non-smoking female (F) and two smoking female (FS) blood donors.
Blood samples were collected twice with a month's interval for the uninhibited cultures and with a time lap of more than a year before the inhibited cultures were initiated. Repeated analysis of peripheral blood lymphocytes from the same individual demonstrated good reproducibility of the assay for the control cultures, although a non-significant variation was observed, stressing the importance of including both positive and negative controls in each experiment. Higher frequencies in the control culture corresponded to higher frequencies in the RFR culture. Glutathione-S-transferases (GSTs) encode a family of detoxifying phase II enzymes catalyzing the conjugation of glutathione to electrophilic compounds. The GSTM1, GSTT1 and GSTP1 genes are known to be polymorphic, and some in vitro experiments with human lymphocytes include the influence of the genotype of the donors on the background level of DNA damage (19, 26, 27). Therefore, to secure equal ability to detoxify possible chemical exposures, all blood donors were of GSTM1 null genotype, GSTP1 heterozygote or wild type, and only one donor was GSTT1 null genotype.
Aberrations recorded for inhibited cultures exposed to 2.3 GHz pulsed-wave radiation. Mean frequencies per 100 cells (standard deviation in brackets) are given for four parallel cultures for two male (M), two non-smoking female (F) and two smoking female (FS) blood donors.
The exposed samples experienced a temperature approximately 0.5°C higher than that for the control samples. We believe that this should not have a significant effect on the results, since in a similar experimental set up with 16.5 GHz exposure we tested cultures at both 37°C and 38°C. The temperature difference had very little effect on the chromosomal aberration frequencies Hansteen et al. (28). In the present experiment where RFR was applied during all phases of the cell cycle no significant differences were observed between control and RFR-exposed cultures for any of the chromosomal parameters for each blood donor, regardless of gender (results not shown), or for all results combined. This is consistent with previous reports for different RFR wavelength exposure for different cytogenetic assays (6-8, 11, 14, 28-31) even for intermittent exposures at different stages of the cell cycle (32). Other studies reported that exposure to RFR signals under extended conditions were capable of inducing DNA damage in human lymphocytes in vitro (5, 33), and Tice et al. (5) and Zotti-Martelli et al. (4) reported individual differences between blood donors.
In the present study, no cytogenetic effect of 2.3 GHz radiation combined with MMC treatment was documented. RFR exposure with or without treatment with MMC gave no increase in chromosomal damage, neither in lymphocytes from non-smoking nor from smoking individuals. This is consistent with some earlier studies where no synergistic effects were observed for a range of genotoxicity endpoints of a combined exposure of cells to RFR and cytotoxic drugs (34-37). Maes et al. (38) reported a weak synergistic effect of RFR with co-exposure to MMC on cytogenetic endpoints, but this was not reproduced in a later study (39). Zhang et al. (40) also reported a synergistic increase in both micronuclei and DNA damage in human lymphocytes upon co-exposure to MMC.
Aberrations compared for 2.3 GHz continuous and pulsed field exposure to lymphocytes from non-smoking males and females combined. Mean frequencies per 100 cells (standard deviation in brackets) are shown*.
Lai and Sing (41) and d'Ambrosio et al. (42) suggested a different response to continuous- and pulsed-wave RFR exposure. We did not find statistically significant results in our experiment, but the Anova analysis showed a weak trend towards more damage with the pulsed- than the continuous-field exposure and a higher damage rate from the interaction between pulsed fields and MMC. As seen by the mean frequencies listed in Table IV, the increase is not dramatic and the implication on health effects is uncertain (3).
In conclusion, no statistically significant differences were observed between control and the 2.3 GHz radiation-exposed cultures, with or without addition of MMC, for any of the chromosomal parameters for any of the blood donors in the uninhibited assay, or in the cultures where DNA synthesis and repair were inhibited. The increase in chromosomal damage due to unrepaired aberrations was not a more sensitive method for detecting possible differences between control and exposed cultures in this setting. An Anova analysis was performed with gender, MMC, pulsed- and continuous-field exposure in the model, and the difference between the control and exposed aberration frequencies as effect parameter. None of the included parameters had a significant impact on the effect parameter. A weak trend was observed towards more chromosomal damage from pulsed-than constant-field exposure, especially from the interaction of pulsed fields and MMC.
Footnotes
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↵+ Present address: StatoilHydro ASA, EPN ONS HSE, Stavanger, Norway.
- Received June 5, 2009.
- Revision received September 25, 2009.
- Accepted September 28, 2009.
- Copyright© 2009 International Institute of Anticancer Research (Dr. John G. Delinassios), All rights reserved
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