Mutagenesis, Vol. 14, No. 5, 457-462,
September 1999
© 1999 UK Environmental Mutagen Society/Oxford University Press
Genotoxic activity of chlorohydroxyfuranones in the microscale micronucleus test on mouse lymphoma cells and the unscheduled DNA synthesis assay in rat hepatocytes
Department of Organic Chemistry, Åbo Akademi University, Biskopsgatan 8, FIN-20500 Turku/Åbo, Finland and 1 Laboratory of Toxicology, Institut Pasteur Lille, 1 rue Professeur Calmette, BP 245, F-59019 Lille Cedex, France
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Abstract |
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Chlorohydroxyfuranones (CHFs) are mutagenic disinfection by-products found in chlorine-treated drinking water. In the current study, the genotoxicity of four CHFs, 3,4-dichloro-5-hydroxy-2(5H)-furanone (MCA), 3-chloro-4-methyl-5-hydroxy-2(5H)-furanone (MCF), 3-chloro-4-(chloromethyl)-5-hydroxy-2(5H)-furanone (CMCF) and 3-chloro-4-(dichloromethyl)-5-hydroxy-2(5H)-furanone (MX), was determined. Two in vitro assays, the microscale micronucleus assay on L5178Y mouse lymphoma cells and the unscheduled DNA synthesis assay on a hepatocyte primary culture from Fisher F344 rats, were carried out. All four CHFs demonstrated genotoxic effects in both assays. In the two systems used, CMCF was the most genotoxic compound, followed by MCA, MX and MCF, respectively. This work was the first study of the DNA damaging properties of all four CHFs in two in vitro genotoxicity tests. These new data expand the range of genetic damages induced by this group of compounds.
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Introduction |
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Chlorohydroxyfuranones (CHFs) have drawn growing attention over the last decade. The most studied CHF, 3-chloro-4-(dichloromethyl)-5-hydroxy-2(5H)-furanone (MX), was identified in drinking water (Hemming et al., 1986
) and was shown to be one of the strongest direct acting mutagens known in the Ames test (Meier et al., 1987a; Tikkanen and Kronberg, 1990; Lalonde et al., 1991). MX is found in chlorine-disinfected drinking water at concentrations up to 67 ng/l (Kronberg and Vartiainen, 1988) and is produced in the reaction of chlorine with natural organic material present in the water (Meier et al., 1987b; Horth, 1990). Other CHFs, such as 3-chloro-4-(chloromethyl)-5-hydroxy-2(5H)-furanone (CMCF), 3-chloro-4-methyl-5-hydroxy-2(5H)-furanone (MCF) and 3,4-dichloro-5-hydroxy-2(5H)-furanone (MCA) are also formed and observed in drinking water at concentrations up to 60 ng/l (Kronberg and Franzén, 1993; Franzén and Kronberg, 1994; Smeds et al., 1997). CMCF, MCF and MCA are also bacterial mutagens, although their mutagenic potency is lower than that of MX (Ishiguro et al., 1988; Kronberg and Franzén, 1993; Smeds et al., 1997).
CHFs, especially MX, induce genetic alterations in numerous biological systems in vitro. In cultured rodent cells, they cause chromosomal aberrations (Meier et al., 1987a; Jansson et al., 1993; Mäki-Paakkanen et al., 1994; Harrington-Brock et al., 1995), sister chromatid exchanges (Brunborg et al., 1991; Jansson et al., 1993; Mäki-Paakkanen et al., 1994) and DNA damage (Brunborg et al., 1991). MX also induces DNA strand breaks (Chang et al., 1991; Nunn and Chipman, 1994; Marsteinstredet et al., 1997a) in cultured human cells.
The results obtained in vivo were, however, less clear. Mäki-Paakkanen and Jansson (1995) reported the micronuclei inducing potency of MX in rat lymphocytes after oral administration, while Meier et al. (1987a) and Tikkanen and Kronberg (1990) failed to detect this effect in mouse bone marrow after oral and i.p. administration, respectively. In a very recent communication, Jansson (1998) reported a lack of induction of micronuclei in bone marrow of rats exposed to MX for 2 years in a carcinogenicity bioassay. Furihata et al. (1992) observed DNA damage (measured by alkaline elution) in rat stomach after oral exposure to MX, whereas Brunborg et al. (1991) failed to detect any effect in several organs of mouse after oral and i.p. administration. MX caused sister chromatid exchanges in rat peripheral lymphocytes and kidney after oral administration (Jansson et al., 1993; Mäki-Paakkanen and Jansson, 1995). In mice exposed orally to MX, DNA damage (measured with the Comet assay) was observed in the liver and other organs (Sasaki et al., 1997), but unscheduled DNA synthesis (UDS) was not detected in liver cells (Nunn et al., 1997).
After oral exposure, MX was shown to induce nuclear anomalies in the gastrointestinal tract of the B6C3F1 mouse, while MCA gave suggestive evidence of genotoxic activity (Daniel et al., 1991). Moreover, MX was shown to be a direct acting in vitro teratogen in the rat (Teramoto et al., 1998) and to induce apoptosis in promyelocytic human leukaemic cells in vitro (Marsteinstredet et al., 1997b). Several epidemiological studies associated some human cancers with the consumption of chlorine-disinfected drinking water (Cantor et al., 1987; Morris et al., 1992; Morris, 1995) and with the mutagenicity of chlorinated drinking water (Koivusalo et al., 1995). MX was recently demonstrated to be a multi-site carcinogen in rat (Komulainen et al., 1997) and is considered to be a drinking water contaminant that may impair public health (Melnick et al., 1997).
L5178Y mouse lymphoma cells were used in a study by Harrington-Brock et al. (1995) which showed that MX induces mutations at the thymidine kinase locus and causes clastogenicity. This strain of cells was used in our laboratory to develop a new micromethod for the in vitro detection of micronuclei (Nesslany and Marzin, 1999). In a recent work, Nunn et al. (1997) demonstrated that MX induces UDS in cultured rodent hepatocytes. In the current study, we have tested the genotoxicity of all four chlorohydroxyfuranones (MCA, MCF, CMCF and MX) by using two in vitro assays, the microscale micronucleus assay on L5178Y mouse lymphoma cells and the UDS assay on rat hepatocytes.
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Materials and methods |
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Chemicals
MCA was purchased from Aldrich Chemical (Steinheim, Germany) and was 99% pure. MCF, CMCF and MX were synthesized and purified according to the method of Franzén and Kronberg (1995). The structures of these compounds are presented in Figure 1
. The purities of MCF, CMCF and MX were at least 98%, as estimated by 1H NMR and GC. 3-[4,5-Dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) was obtained from Sigma (St Louis, MO). Fisher medium, William medium and horse serum from Gibco BRL were supplied by Life Technologies (Cergy Pontoise, France). Coverslips were from Nunc (A/S, Roskilde, Denmark), [3H]thymidine was from Amersham Life Science (Amersham, UK) and 2-AAF, purity at least 95%, was from Sigma (Aldrich-Sigma Chemical Co, l'Isle d'Abeau Chesnes, France).
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Microscale micronucleus assay on mouse lymphoma cells
The procedure for this new micromethod for in vitro micronucleus assay has been described in detail by Nesslany and Marzin (1999). A brief account of the main features of the assay is given below.
Cells. L5178Y mouse lymphoma cells (strain TK+/– 3.7.2c), originally obtained from the European Collection of Animal Cell Cultures (Porton Down, Salisbury, UK), were stored frozen and an aliquot was thawed for each experiment. The cells were maintained in culture in FM10 medium, i.e. Fisher medium supplemented with 200 U/ml penicillin, 50 µg/ml streptomycin, 2.5 µg/ml amphotericin B, 200 µg/ml L-glutamine, 200 µg/ml sodium pyruvate, 500 µg/ml pluronic acid and 10% (v/v) heat-inactivated horse serum. Cell cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2. Each new batch of cells was tested to confirm the absence of mycoplasma contamination. As in the mutation assay, each batch of cells was also treated with methotrexate to avoid the presence of spontaneous TK–/– mutants.
Cell treatment. The assay was carried out without metabolic activation because this class of chemicals (for instance MX) are known to be direct mutagens. The cells were treated for 24 h, then incubated in normal medium for 20 h and finally harvested. Each treatment was performed in duplicate and was coupled to a cytotoxicity assessment. Exponentially growing cells were added to FM10 medium containing the compound to be tested at the appropriate concentration. The final concentration was 4x105 cells/ml. The mixture was distributed (0.1 ml/well) in a 96-well V-bottom microplate. The assay was conducted without cytochalasin B because preliminary experiments showed that treatment with 2 µg/ml cytochalasin B for 8 h induced DNA fragmentation and pycnotic nuclei.
At each step, microplates were centrifuged for 5 min at 900 r.p.m. and the supernatant discarded by gentle pouring off. The cells were first washed (0.2 ml Fisher medium, 0.1% pluronic acid) and gently resuspended before hypotonic treatment (4 min with 0.2 ml Fisher medium diluted 1:1 in distilled water + 0.1% pluronic acid). The cells were then fixed by addition of 0.1 ml of ethanol:acetic acid (3:1 v/v) for at least 10 min. The cells were finally resuspended by drawing and expelling using a Pasteur pipette, dropped onto clean glass slides and allowed to dry at room temperature. After 24 h, the air-dried slides were stained for 10 min in 2% Giemsa water solution, rinsed and coded before analysis. The slides were analysed under a microscope (500x magnification) by two scorers, one for each series of slides. Micronuclei were analysed in at least 1000 mononucleated cells per culture in two parallel cultures, i.e. at least 2000 mononucleated cells were analysed for each dose. Micronuclei were identified according to criteria described by Miller et al. (1995). The statistical significance of differences between groups was determined using the 
2 test. The criteria for determining a positive result were a concentration-related increase in the number of micronucleated cells and a statistically significant increase over the spontaneous level in at least one treatment. A positive control (mitomycin C at 25 ng/ml) was included in the assay.
Cytotoxicity assay.
Cytotoxic effects were evaluated using the MTT colourimetric method (Borenfreund et al., 1988). Five concentrations (usually with a 2-fold step) were retained for the genotoxicity assay; the highest concentration should induce a significant reduction in MTT incorporation.
UDS assay on rat hepatocyte primary culture
Hepatocyte culture.
The hepatocyte cell suspension was obtained by perfusing (HEPES buffer, then HEPES/collagenase buffer) the liver of anaesthetized male Fisher F344 rats (IFFA CREDO, Saint-Germain sur l'Abresles, France). The suspension was washed by centrifugation (1 min at 40 g) and resuspension in complete William E medium (WE-C), i.e. William E medium supplemented with 200 U/ml penicillin, 50 µg/ml streptomycin, 2.5 g/ml amphotericin B, 200 µg/ml L-glutamine and 10% (v/v) heat-inactivated fetal calf serum. The percentage of viable cells was determined using the trypan blue technique and a Malassez haemocytometer. The percentage viability of the final cell suspension must be >50% (Fautz et al., 1993). The culture was diluted in order to obtain 1.5x105 viable cells/ml and distributed in 6-well microplates containing round plastic coverslips. Beside the wells intended to assess the UDS, a few (without coverslips) were used to determine survival. Some wells were seeded for the solvent control and others for the positive control. In order to enable cell attachment, the microplates were incubated for ~90 min at 37°C in an atmosphere containing 5% CO2.
Treatment and radiolabelling of hepatocyte cultures. After cell attachment, the incubation medium was sucked away from the wells and the monolayers were washed with incomplete William E medium (WE-I), with the same composition as WE-C but without fetal calf serum. The WE-I was then replaced by WE-I containing 10 µCi/ml [3H]thymidine and the test compound. Cultures for determination of survival were treated similarly except that [3H]thymidine was omitted. The cultures were incubated for 17–20 h at 37°C in an atmosphere of 5% CO2. A positive control (2-acetamidofluorene at 6.5 µM) was included in each assay.
Determination of survival. The trypan blue technique was used to determine the viability of the cells. The proportion of viable cells was determined for each concentration tested and expressed as a percentage of the solvent control viability. The highest concentration usually retained for the UDS test showed 50–75% survival in comparison with the solvent control.
Autoradiography. The slides (coverslips glued on normal microscopic slides) were coated in Kodak D19 liquid emulsion. After air drying, the slides were incubated in a light-tight box and left refrigerated for 10–14 days. After this period, the emulsion was developed and fixed. The cell nuclei and cytoplasm were stained with Meyers hemalun. Slides were dehydrated in ethanol, cleaned in xylene and mounted with coverslips for microscopic examination.
Autoradiographic analysis and grain counting. Grain counting was performed using an image analysis system (Visilog, Noesis, France). Fifty cells per slide and three slides per concentration were examined. Only cells with normal morphology were scored. Isolated nuclei with no surrounding cytoplasm, cells with unusual staining or heavily labelled cells in S phase were not scored. One cell could not be counted more than once. Nuclear grain counts (NC) and cytoplasmic (CC) grain counts were recorded and the net nuclear grain (NNG) per cell was determined (NNG = NC – CC).
Expression of the results and criteria for genotoxic activity. For each slide and concentration, the following end-points were calculated: average NNG, percentage of cells in repair, average CC and NC and number of cells in S phase. Data were analysed using the non-parametric U rank Mann–Whitney test.
The compound under study was considered as genotoxic in this system if: (i) at any concentration tested, group mean value is >0 NNG and 20% or more of cells are in repair (NNG values >5); (ii) compared with the control, an increase is observed in both NNG and the percentage of cells in repair; (iii) a dose-related increase is seen both in NNG and in percentage of cells in repair.
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Results |
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The results of the tests are reported in Tables I and II
and summarized in Table III.
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In the micronucleus assay on mouse lymphoma cells, all four CHFs showed significant genotoxic effects. MCA induced the formation of micronuclei only at the highest concentration tested (25 µM). CMCF, MX and MCF increased the frequency of micronuclei in the concentration ranges 6.25–25, 50–100 and 100–200 µM, respectively. CMCF caused the formation of micronuclei at a much lower concentration than the other CHFs.
All four compounds were genotoxic in the rat hepatocyte UDS assay. MCA, CMCF and MX were clearly positive: the NNG was >5 at the highest dose and an obvious concentration-dependent increase was observed for the NNG count as well as for the percentage of cells in repair. CMCF induced UDS at a much lower concentration than MCA and MX. MCF gave only a weak positive response in this test system: the NNG count and the percentage of cells in repair increased with concentration, but the NNG count, although positive, did not exceed 5 at the highest dose. In summary, CMCF, MCA, MX and MCF induced UDS at 1.5–6.25, 12.8–20, 16.7–56 and 100 µM, respectively.
If we consider the lowest concentration inducing a genotoxic effect, CMCF clearly appears as the most potent genotoxicant in both mouse lymphoma cells and rat hepatocytes. For both tests, the following order of genotoxic potency can be established: CMCF > MCA > MX > MCF.
Taking into account the highest concentration retained in the genotoxicity assay, the cytotoxicity of the compounds appears to follow the same pattern as the genotoxicity: CMCF was the most toxic compound, followed by MCA, MX and MCF, respectively.
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Discussion |
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To our knowledge, an in vitro study on mammalian cells has not previously been carried out to assess the micronucleus induction potency of CHFs. MX and MCA were once evaluated for their ability to cause micronucleus formation, but the assay was performed on plants, i.e. pollen mother cells of Tradescantia (Helma et al., 1995
). Both chemicals caused a dose-dependent increase in the frequency of micronuclei when applied directly to the inflorescences, but not when applied on the stems. Our choice of the L5178Y cell, a line of cancer cells, in a test with micronuclei as the end-point might be questioned, particularly considering the possibility of instability of chromosome number. The low spontaneous micronucleus formation rate and the clear micronucleus response to treatment with the CHFs rule out any concern on that issue.
On the other hand, numerous in vitro studies have demonstrated that MX causes a range of chromosomal and DNA damages in mammalian cells; in each case MX was positive without metabolic activation. Chromosomal aberrations were induced in Chinese hamster ovary cells (Meier et al., 1987a; Mäki-Paakkanen et al., 1994), rat peripheral lymphocytes (Jansson et al., 1993) and mouse L5178Y lymphoma cells (Harrington-Brock et al., 1995). MX increased the frequency of sister chromatid exchanges in rat peripheral lymphocytes (Jansson et al., 1993), in V79 cells (Brunborg et al., 1991) and Chinese hamster ovary cells (Mäki-Paakkanen et al., 1994). This compound caused DNA damage (measured by alkaline elution) in Chinese hamster V79 cells and in rat liver and testicular cells (Brunborg et al., 1991). MX also induced DNA strand breaks in human lymphoblastoid cells (Chang et al., 1991) and human white blood cells (Nunn and Chipman, 1994), measured by DNA alkaline unwinding, and in human promyelocytic leukaemic cells (Marsteinstredet et al., 1997a), measured by alkaline elution or single cell gel electrophoresis. It was recently proposed that at least some of the DNA strand breaks observed in MX-treated cells may be explained by alkali-labile apurinic/apyrimidinic sites (Hyttinen and Jansson, 1995).
Only one other CHF has been studied for its DNA damaging potency: MCA induced DNA strand breaks in human lymphoblastoid cells (Chang et al., 1991) and caused cleavage of plasmid DNA (Lalonde and Ramdayal, 1997). Until today, the DNA damaging properties of CMCF and MCF had not yet been tested, which is certainly related to the fact that these compounds are not commercially available and their synthesis is difficult.
The lowest MX concentrations inducing genotoxicity in previous studies on mammalian cells were 3.47 µM for chromosomal aberrations (Harrington-Brock et al., 1995), 0.87 µM for sister chromatid exchanges (Mäki-Paakkanen et al., 1994), 30 µM for DNA damage (Brunborg et al., 1991) and 1 µM for DNA strand breaks (Nunn and Chipman, 1994). These values are lower than the MX lowest concentration inducing micronuclei in the current study (50 µM). This fact can be explained by differences in the test procedures and in biological system sensitivity. Moreover, micronucleus formation probably involves more complex and drastic events than chromosomal aberration, sister chromatid exchange, DNA damage or DNA strand breakage. Another explanation is that the L5178Y cell line is less sensitive to MX than other cell lines.
In the assays of mutagenicity in bacteria, MX was several orders of magnitude more potent than the other CHFs (Smeds et al., 1997). The results of the current study indicate that, when tested on mammalian cells (mouse lymphoma cells and rat hepatocytes), the genotoxic potency of MX was relatively decreased, as it was not the most genotoxic CHF in these biological systems.
Reports in the literature concerning the effects of CHFs on DNA synthesis or repair are quite rare. MCA, CMCF and MX caused differential DNA repair in the bacterium Escherichia coli (Fekadu et al., 1994). In a recent study by Nunn et al. (1997), MX was shown to cause UDS in rat hepatocytes in vitro at 1–10 µM. These values are lower than the genotoxic concentrations observed in our study (16.7–56 µM). The strain of rat utilized as a source of hepatocytes in the work of Nunn and co-workers was Wistar rats, whereas it was Fisher F344 rats in our study. The use of two different rat strains as the source of hepatocytes can lead to significant differences in the DNA repair test. McQueen and Way (1991) demonstrated that the lowest concentration of aflatoxin B1 inducing DNA repair was modified by a factor of 104. Hepatocytes from Wistar rats might thus be more sensitive to the UDS enhancing capacity of MX than hepatocytes from Fisher rats. Concerning the sex of the animals, which was also recognized as an important factor (McQueen and Way, 1991), male rats were utilized in our study and that of Nunn et al. Furthermore, reproducibility of the results in the UDS assay can be affected by significant inter-animal variation (Fautz et al., 1993; Madle et al., 1994). In the work of Nunn et al., (1997), MX did not induce UDS in mouse hepatocytes in vivo.
Regarding possible relationships between the structure (i.e. the nature of the substituent on C-4) and the genotoxicity of the CHFs, the results showed that the absence of a chlorine atom in the methyl substituent (in the case of MCF) caused a lower DNA damaging activity. Concerning the influence of the number of chlorine substituents, the compound with a chloromethyl group on C-4 (CMCF) was more genotoxic than that with a dichloromethyl group on C-4 (MX).
When the effects of CHFs in the micronucleus and UDS assays were compared, MCA, CMCF and MX yield similar results in both tests. At the given concentration of 25 µM, MCA clearly induced both micronuclei in mouse lymphoma cells and UDS in rat hepatocytes. Likewise, CMCF and MX were clearly genotoxic at 6.25 and 50 µM, respectively. On the other hand, at 100 µM, MCF significantly induced the formation of micronuclei but only caused weak DNA repair. This attempt to compare activity in the two experimental systems may, however, be criticized, as the micronucleus microwell assay was performed on cells devoid of an efficient metabolic activation system and the UDS assay on metabolizing cells.
It was proposed that DNA repair induced in the UDS assay might be related to the excision repair of DNA adducts and this potential pattern was recently considered by Nunn et al. (1997) in their study on MX. As a matter of fact, recent results from our laboratory indicated that adducts were formed in calf thymus DNA after exposure to CHFs. At present, all CHFs adducts identified in DNA have been 2'-deoxyadenosine (dA) adducts: cyclic dA derivatives with an ethenoformyl or a propenoformyl bridge between N-1 and N6 or dA derivatives including a propenal or a formylbutenoic acid moiety bound to N6 (Le Curieux et al., 1997; Munter et al., 1998). These findings demonstrate that, under conditions similar to physiological, MX reacts covalently with adenine bases in DNA and forms stable adducts. These genetic alterations might be important (pre)mutagenic events in some specific biological systems. The UDS may also be explained by the formation of apurinic/apyrimidinic sites, as reported by Hyttinen and Jansson (1995), in MX-exposed supercoiled PM2 DNA. The high DNA damaging potency of CMCF observed in the current work is consistent with very recent findings of our laboratory (T.Munter et al., in preparation) showing that CMCF reacts with nucleosides and with DNA to produce several adducts at relatively high yield, compared with other CHFs.
Considering our results, as well as the literature data available at present, our belief is that CHFs are at least clastogens. They indeed induce sister chromatid exchange, chromosome aberrations, DNA single-strand breaks, apurinic sites and DNA repair. All these genetic events are typically induced by clastogens. Until now, no study has indicated that CHFs could also be aneugens, but the available literature does not rule out that possibility either.
The present work is the first study of the DNA damaging properties of all four chlorohydroxyfuranones (MX, CMCF, MCF and MCA) in two in vitro genotoxicity tests. The individual results obtained for each compound are in agreement with the findings of previous studies. All four CHFs were positive in the mouse lymphoma cell micronucleus assay without metabolic activation and the rat hepatocyte UDS assay and these data expand the range of genetic damage induced by this class of compounds. The relatively high genotoxicity observed for CMCF is a new finding which needs to be further documented, particularly in in vivo tests, for instance in the UDS assay, as MX failed to demonstrate genotoxic properties in vivo (Nunn et al., 1997).
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Acknowledgments |
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This work was supported by a research grant from Åbo Akademi University (T.M.) and by the European Commission (Marie Curie Fellowship, contract no. ERBFMBICT961394; F.L.C.).
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2 To whom correspondence should be addressed. Tel: +33 3 20 87 79 14; Fax: +33 3 20 87 73 10; Email: daniel.marzin{at}pasteur-lille.fr
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References |
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Received on December 8, 1998; accepted on April 16, 1999.
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