Mutagenesis, Vol. 14, No. 5, 449-456,
September 1999
© 1999 UK Environmental Mutagen Society/Oxford University Press
Detection of mitomycin C-induced genetic damage in fish cells by use of RAPD
Toxicology Department CNA, I.S. Carlos III, E-28220 Majadahonda, Madrid and 1 Division of Environmental Toxicology, CISA-INIA, E-28130, Madrid, Spain
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Concern about genetic alterations in fish populations arising from anthropogenic activities has led to the adaptation and/or development of new tests and techniques that shed light on these alterations. The high number and the reduced size of chromosomes and the long cell cycle associated with most fish species preclude the use of most accepted genotoxicity assays. The purpose of this work was to study the capability of the randomly amplified polymorphic DNA technique to show genotoxic effects induced by chemicals in fish cells. To do that we studied the effect of 0.5 µg/ml mitomycin C (MMC) on an established rainbow trout cell line (RTG-2). To increase the sensitivity of detecting altered copies of DNA and to avoid the presence of false positives and a lack of reproducibility, the amounts of DNA template and primer present in amplification reactions were studied and optimized after comparison between the control and exposed fingerprints for 4, 6 and 8 h. Results show that 5 ng of DNA template and 4 pM chosen primer were optimum to show differences between control and exposed cells and to obtain reproducible results. The results obtained, after optimum conditions were established, show that this system could be useful for the assessment of DNA alterations in in vitro genotoxicity studies.
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Introduction |
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TopAbstractIntroductionMaterials and methodsResults and conclusionsDiscussionReferences |
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Sublethal exposure to environmental genotoxic agents arising from human activity causes different forms of alterations to exposed aquatic populations, which may lead to the structure and function of ecosystems being altered (Anderson et al., 1994
).
These derived effects of genotoxins directly or indirectly affect genome integrity due to mutations in germinal and/or somatic cells, leading to an increase in the incidence of different types of tumours (Leblanc and Bain, 1997) and, in the long term, to alterations in the genetic variability of the exposed populations (Theodorakis et al., 1998).
These facts, and for risk assessment purposes, make it imperative that prior laboratory tests are carried out to establish if a particular pollutant or a complex mixture can cause adverse affects on the genetic material of the exposed populations before its liberation into the aquatic environment.
However, reproducing the effects on a fish population of long-term exposure to genotoxic agents, or even short-term exposure but with long-term expression of the effects, requires the availability of large laboratory facilities, personnel specialized in maintaining the species and a large volume of samples, all of which make it very costly.
In vitro tests using fish cell lines are an alternative to the use of fish and have big advantages in ecotoxicology studies (Babich and Borenfreund, 1991; Castaño et al., 1994). The RTG-2 cell line derived from rainbow trout (Oncorhynchus mikiss) has been widely used in cytotoxicity studies and shows a good correlation with in vivo acute fish bioassays (Bols et al., 1985; Castaño et al., 1996). Owing to its capacity to metabolize xenobiotics without requiring the addition of metabolic activation systems (Smolarek et al., 1988; Araujo et al., 1998), it has also been shown to be appropriate for genotoxicity studies (Kocan et al., 1985).
The OECD Guidelines (1998) list a series of widely used techniques, both in vivo and in vitro, for use in risk assessment procedures associated with genotoxins for humans. However, the application of these traditional techniques to the detection in DNA of both macro- and micro-damage (sister chromatid exchange, chromosomal aberrations, etc.) are particularly tedious when applied to fish, due to the high number of chromosomes and the long cell cycle found in fish cells. The appearance of new assays in the last few years, such as the Comet assay (McKelvey et al., 1993; Pandangi et al., 1996), automatic scoring techniques for micronuclei assays (OCDE, 1998) and 32P-post-labelling for the detection of adducts (Phillips et al., 1997), together with recent advances in molecular biology, such as DNA fingerprinting and gene amplification by PCR, offer new possibilities for detecting DNA damage, circumventing these limitations.
Randomly amplified polymorphic DNA (RAPD), developed by Williams et al. (1990), is a technique that involves the amplification of random segments of genomic DNA using the PCR methodology. This method does not require prior sequence information and arbitrarily chosen short primers are used at low stringency to amplify multiple segments from genomic DNA to any species (Williams et al., 1990). The majority of such fragments are identical between individuals or strains, which represents a tremendous potential for application in different research fields: the study of genetic variation in natural populations (Welsh and McClelland, 1991; Theodorakis et al., 1998), genetic variability studies (Paffeti et al., 1996; Endtz et al., 1997) and mutation detection in in vivo studies (Kubota et al., 1992, 1995; Theodorakis and Shugart, 1997; Atienzar et al., 1998).
In the same way, cell populations exposed in vitro to genotoxins suffer DNA alterations in a certain number of cells, which are reflected as variations in the fingerprint obtained for the control population. These are defined as band losses and/or gains as well as alterations in the intensity of amplification of some of them. Such alterations in vivo are considered mutations that are produced by changes to, deletions of or insertions into the pair bases (Muralidharan and Wakeland, 1993).
In a previous work, the characteristic fingerprint of the RTG-2 cell line was established in order to use this methodology for the detection of DNA damage after exposure of cell populations to genotoxic agents (Ferrero et al., 1998).
The RAPD technique has been associated with some limitations concerning lack of reproducibility and the appearance of `ghost' bands which are difficult to interpret. In this work we have studied some variables in order to avoid these limitations. For that, a well-known clastogenic agent was used, mitomycin C (MMC), an inducer of DNA interstrand crosslinks (Sehlmeyer et al., 1996), DNA adducts and mutations (Sanderson and Shield, 1996), both in vivo (Fahring, 1977; Ehling, 1978; Cao et al., 1993; Grawé et al., 1993) and in vitro (Kato and Shimada, 1975; Singh and Gupta, 1983; Davies et al., 1993; Salvadori et al., 1994).
To increase the sensitivity of detection of altered copies of DNA and to avoid the presence of false positives and lack of reproducibility, the amounts of DNA template and primer present in amplification reactions were studied and optimized after comparison between the control and exposed fingerprints for different exposure periods. The final purpose of this work was thus to minimize the limitations of this technique in order to be able to apply it to the detection of genomic DNA alterations caused by environmental chemicals to exposed cells.
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Materials and methods |
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Cell cultures
RTG-2 cells (CCL 55; American Type Culture Collection, Rockville, MD), an established fibroblastic cell line derived from rainbow trout (Oncorhynchus mykiss) were grown in Eagle's minimum essential medium with Earle's salts (EMEM), supplemented with 10% fetal serum, 100 U/ml penicillin, 100 µg/ml streptomycin, 1.25 µg/ml fungizone and 2 mM L-glutamine, in PVC tissue culture flasks (Costar, Corning, NY) and incubated at 20 ± 1°C in a 5% CO2/air atmosphere.
Exponentially growing cells were exposed to 0.5 µg/ml MMC (Sigma, St Louis, MO) for 4, 6 or 8 h. After each exposure period, the medium was removed and the cells washed with phosphate-buffered saline (PBS). Fresh medium was then added and the cells were left to grow for a further 72 h (~1.5 cell cycles).
Two 75 cm2 flasks were used for the control and for each one of the exposure periods. The experiment was repeated twice on different days.
DNA extraction and RAPD reaction
Cells were dissociated with trypsin–EDTA (Flow Laboratories, Rickmansworth, UK), collected in PBS, pH 7.3, and their DNA isolated by phenol extraction (Ferrero et al., 1998). The integrity of the extracted genomic DNA was checked by 0.8% agarose electrophoresis using the 
phage as molecular weight marker (Eurobio, Paris, France). Amplification was performed in a 25 µl volume containing buffer solution, 4 mM MgCl2, 2 U Stoffel fragment (Perkin-Elmer, Branchburg, NY) and 0.2 mM each dNTP (Pharmacia, Barcelona, Spain). DNA template was added at two different concentrations, 20 and 5 ng.
The RAPD protocol was previously described by Ferrero et al. (1998). Briefly, an initial denaturing step at 92°C was followed by 45 cycles of annealing at 36°C for 75 s and extension at 72°C for 6 min. The thermal cycler used was a Perkin-Elmer model 2400.
The primer used was AA-82 (5'-GATCCATTGC-3'; Biopolymers Department, CNBCR, I.S. Carlos III, Madrid, Spain). This primer was selected because with this cell line it generates different amplification products within a wide range of molecular weights.
Amplification products (22 µl) were resolved electrophoretically on 2.1% agarose gels and stained with ethidium bromide. The image was recorded by the Grab-it program (UVP, Uplad, CA).
Optimization of primer concentration
In order to avoid the presence of artefacts, we carried out 20 amplifications on different days, without DNA template and using three different primer concentrations (2.5, 4 and 5 pM) following the above described amplification conditions.
Analysis of the band pattern of control and exposed cells
The genomic DNA extract in the two different flasks was obtained and amplified at least twice on different days. The control and exposed cells for each experiment were individually amplified but developed together in the same gels, using the molecular weight markers 
X174-HaeIII and /EcoRI-HindIII (Eurobio). The agarose gels were analysed by densitometry (Gelworks 1D; UVP).
Qualitative analysis was performed by comparing the percentage appearance of each of the peaks of the control and exposed cells. After eliminating the background, quantitative differences were studied using three types of parameters: height, volume and percentage of the amplified band. Additionally, the individual data obtained on the height, volume and percentage of amplified bands were grouped together according to the following criteria: bands of high molecular weight (>800 bp), bands of intermediate molecular weight (between 600 and 800 bp) and bands of low molecular weight (<600 bp).
For statistical analysis all individual data on control and treated cells from the different experiments were considered together. The data were analysed using Student's t-test and the Mann–Whitney U-test.
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Results and conclusions |
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Primer concentration
In five of the 20 amplifications carried out with 5 pM of primer and without template, the appearance of `ghost' bands of different molecular weights was noted as a result of polymerization of the primer with itself. When assays were performed with 2.5 and 4 pM, no artefacts were observed, so 4 pM primer was selected as the optimum for later assays.
Differences in the band pattern of control and treated cells
Qualitative analysis. The assays carried out with 5 and 20 ng of control genomic DNA resulted in six common amplification products, three considered as having high molecular weights (i.e. >800 bp), one with an intermediate molecular weight (700 bp) and two with low molecular weights (i.e. <600 bp). When the assay was carried out with 20 ng, apart from those described, three bands with a low appearance rate (erratic bands) were observed and therefore were not included in the pattern (Figure 1). Assays with 2.5 ng of genomic DNA generated the characteristic bands already defined, but they varied greatly in their intensity and frequency of appearance, so this concentration was discarded in later studies. In summary, in the assay conditions of 4 pM primer and 5 or 20 ng of DNA template, a constant pattern was established by those bands whose percentage of appearance was always >80% (Table I).
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In genomic DNA extracts of the exposed cells, qualitative differences were observed in both concentrations when compared with the control cells, although these were greater when using 5 ng of genomic DNA. A drop in the appearance rate of bands was observed in treated cells at 5 ng for all the exposure periods. Band D was particularly affected, going from being present in 84% of the control assays to 50% at 4 h and 40% at 6 and 8 h of exposure, respectively. Five extra bands were also detected: band D1 appeared at all exposure periods, band F1 at 6 and 8 h, band A0 at 4 h and band C1 at 8 h. With 20 ng of template, a drop in the appearance rate of band D was also observed, decreasing by up to 74% after 6 and 8 h of treatment. No new bands were observed with this concentration (Table II
).
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Quantitative analysis. The control amplification products obtained for both DNA concentrations showed a constant fluorimetric profile and for the three quantitative parameters studied, band B presented the highest average values and band D the lowest. The values of these parameters behaved in the same way: Bands A, B, D and E coincide and differences observed in bands C and F are directly and inversely proportional to the genomic DNA concentration, respectively (Figure 2
). In exposed cells, significant statistical differences (P < 0.05) versus the control were found using 5 ng of template for most fingerprint bands at all exposure periods as a function of the parameter analysed. The most significant differences (P < 0.0001) were observed at 8 h of exposure, when considering the percentage of the amplified band (Table III). These results contrast with those obtained when the concentration of genomic DNA used was 20 ng. No band showed significant overall differences in the parameters considered and, furthermore, the values obtained for the percentage of amplified bands coincide, for all exposure periods, with those of the control pattern (Table IV).
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When each band is analysed individually, it is noted that the results for the control and exposed data are greatly dispersed. However, if data are grouped according to their molecular weights, this variability decreases and the differences between the control and treated cells are seen more clearly. When using this method with 5 ng of template, differences are noticeable at 6 and 8 h of treatment and the 5:2 ratio between the high and low molecular weight bands of the control was modified to a value of 1:1 (Table V
). The differences were seen as a decrease in the amplification of the high molecular weight peaks in favour of the lower molecular weights. Conversely, at 4 h the differences were seen as a decrease in amplification of the low molecular weight peaks in favour of the high molecular weight peaks (bands C and F).
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Statistically significant differences (P < 0.01) were obtained for all exposure periods in the low molecular weight band group (<600 bp) for all parameters, as well as in the high molecular weight band group (>800 bp), but only when the band percentage is considered. Although the parameters used for the quantitative valuation are all related, the height varies due to the electrophoretic process itself and the volume has a great dispersion of data. As the band percentage is a value relative to the total amplification of each reaction, we consider this latter as the most appropriate parameter for quantifying our results as it allows for a better comparison between gels. No differences were found with 20 ng of template even when the bands were grouped according to their molecular weights.
In conclusion, qualitative and quantitative differences were found in RTG-2 cells after exposure to 0.5 µg/ml MMC, but only when the template concentration was lowered to 5 ng of genomic DNA (Figure 3).
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Discussion |
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Template concentration affects the electrophoretic band pattern and is the major source of non-reproducibility in RAPD fingerprint analysis, as previously described by other authors using different species (Muralidharan and Wakeland, 1993
; Sakallah et al., 1995). In our case, 5 and 20 ng of genomic DNA from RTG-2 cells show variations in the intensity of amplification of two of the bands (C and F) and in the appearance of erratic bands at the higher concentration (Table I
and Figure 2). The template concentration is also a key factor in showing differences in the band pattern between control and exposed cells. While some qualitative differences occur at both concentrations, such as a decrease in the frequency of the appearance of some bands, the appearance of new bands as well as quantitative alterations can be observed only when the template concentration is reduced (Figure 3).
Amongst the new bands observed in the exposed cells, band D1 (645 bp) was detected for all the exposure periods and the frequency of its appearance increased with the length of exposure. This leads one to think that these bands are the result of a MMC-directed action on specific points on the genomic DNA. This is in accordance with the specificity of MMC in CG-rich areas (Kumar et al., 1997; Palom et al., 1998; Warren et al., 1998).
Quantitative differences in control cells can be observed in the intensity of amplification by decreasing the amount of genomic DNA from 20 to 5 ng. This same effect, but much more noticeable, is found when comparing the control and exposed genomic DNA using 5 ng.
MMC is a well-known mutagenic and clastogenic agent; base substitutions, sister chromatid exchanges, chromosomal aberrations and micronuclei, both in vivo and in vitro, are genetic effects that can be observed as a function of dose and exposure period (Pelt et al., 1991; Rudd et al., 1991; Salvadori et al., 1994). The dosage selected for this work was in accordance with previous studies of micronucleus induction in our laboratory on the same cell line (Llorente et al., 1998) and is also in the range for the positive control in in vitro genotoxicity assays (Ribas et al., 1998). Although MMC provokes alterations in the DNA of the cell population, with the concentration and exposure periods used in this work a certain proportion of DNA copies remain intact. In the case of 20 ng, the number of unaffected copies is sufficient that with the concentration of primer used, it finds sections where it can hybridize and therefore the characteristic band pattern is not altered. This is supported by the fact that only the scarcest bands (bands D and E, which represent 2.5 and 8% of the total amplified, respectively) are affected at the longer exposure periods (Table V). Nevertheless, when the amount of template is lowered to 5 ng, despite the fact that in relative terms the proportion of intact genomic DNA is the same, the number of original copies to be amplified is now much lower and insufficient, which increases the probability of the altered genomic DNA showing itself. The result is that a band pattern different to that of the control is obtained.
Smaller but significant quantitative differences between control and exposed cells for the shorter studied period can be observed as an opposite tendency when compared with longer exposure periods. In the RAPD reaction a high number of different sequences are co-amplified in the first cycles and the products from the most stable primer–template unions are the `winners' of this process (Davin-Reglin et al., 1995). The observed tendency for 4 h could suggest that short exposures (4 h represents 1/12 of the population doubling time for this cell line) may lead to the production of mutations with changes in base pairs (Srikanth et al., 1994; Maccubbin et al., 1997) which can give the primer the opportunity to find new hybridization sites and obtain products of similar molecular weight but with different sequences (Reisenberg, 1996). Attention should also be paid to the qualitative analyses. After 4 h, a new high molecular weight band appears and the frequency of appearance of band C decreases by up to 90%, so that new and possibly unstable hybridizations may be suspected. Longer exposure periods lead to a decrease in the amplification of high molecular weight bands as a result of the clastogenic action of MMC (Pelt et al., 1991; Channarayappa and Ong, 1992).
Although RAPD assays are usually associated with inherent problems of the technique which prevent its routine application (Hadrys et al., 1993), if the conditions of the assay were optimized these inconveniences can be solved, i.e. appearance of `ghost' bands, reproducibility, etc. (Theodorakis et al., 1998). By optimizing the RAPD conditions, and bearing in mind that RAPD are PCR products, they can be cloned and sequenced to obtain a wider knowledge of genotoxicant action mechanisms (Theodorakis et al., 1998).
The results of this work shows that the RAPD technique could be a promising tool in the in vitro detection of alterations in DNA produced by genotoxic agents, allowing us to see the mechanisms of action of the agent in greater detail and also permitting areas or `hotspots' within the affected genomic DNA to be seen. Moreover, the practically unlimited number of informative primers provides a good overall coverage of genomic DNA (Ramser et al., 1996) so that the choice of a set of primers with different sequences will allow increased sensitivity of the assay in the detection of low frequency mutation events.
The purpose of this work was to study the capability of this technique to show genotoxic effects induced by chemicals in species where a long cell cycle, low ability of colony formation and small or high numbers of chromosomes (Babich and Borenfreund, 1991; Al-Sabty and Metcalfe, 1995) makes application of most mutagenicity assays impractical. Obviously, a large number and types of genotoxic agents must be tested to establish sensitivity limits before routine application of this assay.
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Acknowledgments |
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This work was supported by CICYT projects AMB94-0655-CO2 and AMB97-0431-CO2
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2 To whom correspondence should be addressed. Tel: +34 91 5097900; Fax: +34 91 5097926; Email: cbeceril{at}isciii.es
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References |
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Received on November 10, 1998; accepted on April 29, 1999.
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