- Email: [email protected]
Genotoxic, cytotoxic, developmental and survival effects of tritiated water in the early life stages of the marine mollusc, Mytilus edulis
Aquatic Toxicology 74 (2005) 205–217
Genotoxic, cytotoxic, developmental and survival effects of tritiated water in the early life stages of the marine mollusc, Mytilus edulis Josephine A. Hagger, Franck A. Atienzar 1 , Awadhesh N. Jha ∗ School of Biological Sciences, University of Plymouth, Drake Circus, Devon, Plymouth PL4 8AA, UK Received 12 January 2005; received in revised form 27 May 2005; accepted 28 May 2005
Abstract Using an integrated approach linking different levels of biological organisation, the genotoxic, cytotoxic, developmental and survival impact of tritiated water (HTO) were investigated in the embryo-larvae of marine mollusc Mytilus edulis. One-hour-old embryos were exposed to a range of concentrations (0.37–370 kBq ml−1 ) of HTO, which delivered a dose between 0.02 and 21.41 mGy over the exposure period for different end points. Detrimental effects, if any, were monitored at different levels of biological organisation (i.e. DNA, chromosomal, cellular and individual). Genotoxic effects were assessed using molecular and cytogenetic approaches which included analysis of random amplified polymorphic DNA (RAPD), induction of sister chromatid exchanges (SCEs) and chromosomal aberrations (Cabs). Cytotoxic effects were evaluated by determining the proliferative rate index (PRI) of the embryo-larval cells. Developmental and survival effects were also monitored every 24 h up to 72 h. Results in general indicated that HTO significantly increased cytogenetic damage, cytotoxicity, developmental abnormalities and mortality of the embryo-larvae as a function of concentration or radiation dose. The analysis of RAPD profiles also revealed qualitative effects in the HTO exposed population compared to controls. However, while the embryo-larvae showed dose or concentration dependent effects for mortality, developmental abnormalities and induction of SCEs, the dose-dependent effects were not apparent for Cabs and PRI at higher doses. The study contributes to our limited understanding of the impact of environmentally relevant radionuclides on non-human biota and emphasises the need for further investigations to elucidate potentially long term damage induced by persistent, low levels of other radionuclides on commercially and ecologically important species, in order to protect human and ecosystem health. © 2005 Elsevier B.V. All rights reserved. Keywords: Tritium; Genotoxicity; RAPD assay; Developmental effects; Marine mussels
1. Introduction ∗
Corresponding author. Tel.: +44 1752 232978; fax: +44 1752 232970. E-mail address: [email protected] (A.N. Jha). 1 Present address: UCB S.A. Pharma Sector, Chemin du Foriest, B-1420 Braine-l’Alleud, Belgium. 0166-445X/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.aquatox.2005.05.013
In 1977 the International Commission on Radiological Protection stated that “If humans are protected from the effects of ionising radiation, then flora and fauna
206
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
are also adequately protected” (ICRP, 1984). However, this statement is often questioned by the general public and regulatory authorities as: (a) different species vary in their sensitivity to toxicants including radiation; (b) different life stages of the same species can show varying levels of response to contaminants; (c) there are areas where wildlife may be exposed to higher levels of radiation than are permissible to humans and (d) endangered species with longer generation times or isolated populations might need special consideration, in order to maintain ecosystem quality. At present, an internationally accepted method for assessing the environmental impact of ionising radiation does not exist (Pentreath, 1998). Attempts are therefore being made to link together the release scenario of radioactive substances, potential exposure pathways, dose to ecologically relevant, commercially important and critical target organisms and subsequently effects at different levels of biological organisation (ERICA, 2005). As a result of nuclear establishments around coastal areas, the marine environment is often the ultimate recipient of radioactive contaminants. In this context, tritium (3 H), a naturally occurring isotope of the element hydrogen, is manufactured commercially and is also released as a by-product of nuclear reactions. An overview of historic trends in liquid and aerial discharges of radioactive substances in the UK, carried out between 1979 and 1998, showed that there has been a sustained increase in the discharge of 3 H from different nuclear facilities (Department of the Environment, Transport and the Regions, 2000). As HTO can move freely in the biosphere it could be a threat to human health as it may be ingested not only through drinking water but also by food, especially from fish and shellfish that are known to bioaccumulate some toxicants (Kirchmann et al., 1973). In this context, elevated activity concentrations of 3 H have been reported in sediment and at different trophic levels of biota from the Severn estuary, UK (McCubbin et al., 2001). It was suggested that the high activity concentrations in these materials, relative to that in seawater, was due to the presence of bioavailable organic 3 H-labelled compounds in radiochemical wastes discharged in the vicinity. Despite recent moves to protect non-human biota from exposures to ionising radiations and the growing scientific and public concern over the presence of 3 H in the environment (Environment Agency, 2001), very little work has been carried out to evaluate the radiobio-
logical or toxicological impact of 3 H on commercially and/or ecologically important species. In particular, there has been a lack of research on potential detrimental effects of ionising radiation on invertebrates, which constitute over 90% of the existing species and play a major role in the maintenance and function of every ecosystem. Consequently, the aim of the present study was to assess the genotoxic, cytotoxic and developmental effects of -radiation emitted by HTO on embryolarvae of an ecologically and commercially important invertebrate, the marine bivalve mollusc Mytilus edulis.
2. Materials and methods 2.1. Exposure scenario HTO was obtained from ICN Pharmaceuticals, Inc., UK at a concentration of 3.7 GBq ml−1 . The range of concentrations of HTO used in the present study (0.37–370 kBq ml−1 ) was adopted from previous work carried out on fish and barnacle larvae (Strand et al., 1977; Abbott and Mix, 1979) and gave a dose between 0.02 and 21.42 mGy over the exposure period as described in Section 3.2. Fig. 1 shows the schematic representation of the general protocol used for assessing the genotoxic [random amplified polymorphic DNA (RAPD)], sister chromatid exchanges (SCEs) and chromosomal aberration (Cabs) assays, cytotoxic proliferative rate index (PRI), developmental and survival effects of HTO on M. edulis embryo-larvae. In brief, following the induction of spawning, as described in detail elsewhere (Jha et al., 2000a), eggs from two female mussels and sperm from one male were collected. The number of eggs was adjusted to 50 eggs ml−1 prior to adding sperm solution at approximately 107 sperm ml−1 . Fertilisation (rate 92%) was carried out at 15 ± 1 ◦ C, whereafter excess sperm was removed by filtering through a sieve (30 m) and the embryo density adjusted to approximately 30 embryos ml−1 as per standard guidelines for acute toxicity testing of environmental contaminants (ASTM, 1989). Following fertilisation, the embryos were allowed to grow for 1 h, after which they were divided into different exposure vessels (glass beakers) where the final volume was adjusted to 500 ml with seawater. Temperature, salinity and pH were controlled throughout the exposure and air was supplied
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
207
(Jha et al., 2000a,b). HTO concentrations were made using a serial dilution with seawater from the original stock. One hundred microlitres of each dilution was added to 500 ml of seawater containing the embryos. MMS was also made using a serial dilution with seawater and 100 l of sample was added to achieve the desired concentration for the volume of seawater. In order to obtain sister chromatid differential staining (SCD) a minimal concentration (10−5 M) of 5-bromo2-deoxyuridine (BrdU) was added to all the vessels at 12 h post-fertilisation. 2.2. Random amplified polymorphic DNA (RAPD) assay As described in Fig. 1, along with Cabs and SCEs assays, a subset of embryo-larvae were processed for analysis using the random amplified polymorphic DNA assay. Genomic DNA was extracted and purified using a conventional phenol/chloroform method as described elsewhere (Atienzar et al., 1999). DNA concentrations were determined by electrophoresis by comparison with known concentrations of Lambda phage DNA (Sigma, Poole, UK). RAPD was performed with 5 ng of genomic DNA as described elsewhere in details (Atienzar et al., 2002). The 10-mer primers used were OPA9, OPB1, OPB5, OPB6, OPB7, OPB8, OPB10, OPB11, OPB12, OPB14, or OPB17 (Operon Technologies, Southampton, UK; for the sequences see Atienzar and Jha, 2004). Thermal cycling parameters and analysis of PCR products by electrophoresis have been described elsewhere in details (Atienzar et al., 2002; Atienzar and Jha, 2004). The GeneRulerTM 100 bp DNA ladder plus (Immunogen International, Sunderland, UK) was used. Fig. 1. Flow chart illustrating the experimental protocol adopted to evaluate genotoxic, cytotoxic, developmental and survival effects of tritiated water (HTO) on the embryo-larvae of M. edulis.
via gentle aeration. Embryos were exposed to four different activity concentrations of HTO (0.37, 3.7, 37 and 370 kBq ml−1 ), 1 negative control (i.e. 10 m filtered seawater) and 1 positive control containing methyl methane sulphonate (1.0 × 10−3 M MMS) as a reference genotoxic agent. This single concentration of MMS was selected based on our earlier studies
2.3. Slide preparation for cytogenetic (i.e. PRI, SCEs and Cabs) analyses Following addition of BrdU, after 1 21 (Cabs) and 2 cell cycles (SCEs and PRI) two subsets of embryos were removed (1 cell cycle = 3.8 h; Jha et al., 2000a,b). One subset was placed into colchicine dissolved in seawater (0.025%, w/v) for 30 min and then exposed to a series of hypotonic (seawater/0.075 M KCl) treatments. After the hypotonic treatments, the embryos were transferred into centrifuge tubes and were centrifuged for 5 min at 200 × g. After centrifugation
208
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
the supernatant was discarded and a few drops of cold (4 ◦ C) Carnoys fixative (1:3 (v/v) glacial acetic acid/methanol) was added. The cells were then processed to produce metaphase spreads, stained with Giemsa (for the analysis of Cabs), or processed for sister chromatid differential staining (for the analysis of SCEs and PRI) as previously described (Harrison and Jones, 1982; Jha et al., 1995a,b, 1996, 2000a,b) for microscopic analysis. Prior to scoring, slides were randomised, coded by an independent disinterested individual and then scored blind. Complete metaphase spreads (2n = 28) were examined using a bright field microscope (final magnification ×1000). Details of classification of PRI stages, scoring of Cabs and SCEs have been described in earlier studies (Jha et al., 1995a, 1996, 2000a, 2000b). In brief, metaphases were classified, as first (M1), second (M2) and third or subsequent (M3+) division cells. For PRI at least 200 cells per treatment were scored, 100 each from two replicates. PRI was calculated using the formula PRI = ((1 × M1) + (2 × M2) + (3 × M3))/ number of cells analysed (Lamberti et al., 1983). Only second (M2) division metaphases were analysed for the occurrence of SCEs. Fifty metaphases per treatment were analysed for SCEs, 25 from each of two replicates, and the results were expressed as the SCEs frequency per cell. For Cabs 200 cells from each treatment were scored (100 from each of two replicates) as per the recommendation of the United Kingdom Environmental Mutagen Society (UKEMS) (Kirkland, 1990). Cabs were expressed as the percent aberrant cells (percentage of cells that contained one or more aberrations including heavily damaged cells) and total aberrations (the number of chromosome plus chromatid type aberrations per 100 cells excluding gaps). 2.4. Analyses of developmental and mortality/ survival effects Developmental morphology, behaviour and mortality of the growing embryo-larvae were assessed at 24, 48 and 72 h post-fertilisation. Fig. 2 shows various stages of normal and abnormal development. The development of M. edulis consists of an embryonic phase followed by a larval phase (Bayne, 1976). At the experimental temperature (15 ± 1 ◦ C), the first larvae stage known as trochophores appears at approximately 24 h. This stage is defined as free swimming larvae,
with a circular band of short cilia (prototroch) and a flagellum (Fig. 2a). Abnormal larvae are usually misshapen (not spherical in shape or symmetrical) and may lack cilia and/or the flagella, resulting in abnormal swimming behaviour (Fig. 2b). The second larvae stage is the veliger, which in M. edulis is usually formed after 48 h (15 ◦ C). A dorsal thickening of the ectodermis, secreted by the shell gland, forms an initial organic cuticle that spreads over the entire body. A straight dorsal hinge gives the larvae the characteristic capital D shape, hence the name “D-larva” (Fig. 2c). At this stage the larvae are approximately 60–70 m in size (His et al., 1999). Dead trochophores appear as a mass of degenerating cells with no cilia or flagellum movement. Dead D-larvae may also lack ciliary movement, lose clarity of internal organs and become transparent (His et al., 1999). Pollutants present in the marine environment may lead to morphologically abnormal trochophores and D-larvae (Fig. 2b and d, respectively). 2.5. Water quality parameters and measurement of radioactivity Water quality parameters (pH, temperature and salinity) were monitored at the start and end of exposure and were relatively stable throughout (mean and S.E.; pH 8.07 ± 0.05; salinity 35 ± 1, 15.5 ± 0.5 ◦ C). Pre- and post-exposure levels of radioactivity were determined using 100 l of test solution in 5 ml of liquid scintillation cocktail, Packard Ultima Gold (Sigma–Aldrich Ltd., USA). Five replicates were counted from each activity concentration. Samples were read for 10 min (detection limits range from 0.06 to 14.5 keV) in a scintillation counter (Beckman LS6500, USA). Readings were displayed as counts per minute (CPM), then automatically converted to disintegrations per minute (DPM) using percentage efficiency for 3 H and then converted to kBq ml−1 (37 kBq equivalent to 2,220,000 DPM). The scintillation counter has a built in quench correction and 14 C and 3 H standards were used to calibrate the counter regularly. The lower detection limit for 3 H was 0.00962 kBq ml−1 . 2.6. Statistical analyses Statistical analysis were carried out using the statistical software package STATGRAPHICS Plus for Windows 3.1©from Statistical Graphics Corp. 1994–1997.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
209
Fig. 2. The developmental stages of M. edulis embryo-larvae. Photos a and c represent normal embryo-larvae at 24 h and >48 h, respectively. Photos (b) and (d) represent morphologically abnormal embryo-larvae at 24 h and >48 h, respectively.
One-way analysis of variance (ANOVA) was carried out on normalised data. If required, transformation of data for developmental effects was carried out using Arc SIN and square root as recommended by Burgeot et al. (1995) whereas SCEs, Cabs and PRI values were transformed either via log or square root conversions. Non-normally distributed data were analysed using the non-parametric Kruskal–Wallis test (Sparks, 2000).
was neither significant variation in the percentage of radioactivity lost during the exposure period (difference between pre- and post-exposure), nor any trend in the percentage loss for different concentrations of HTO used. Each vessel containing HTO lost similar percentages of radioactivity although the samples containing larger amounts showed a greater loss in terms of the actual concentration over the exposure period.
3. Results
3.2. Calculation of dose received by embryo-larvae
3.1. Determination of radioactivity Table 1 presents the levels of radioactivity in different test vessels for the exposure scenario. There
The estimated dose to growing embryo-larvae exposed to HTO was calculated by a method used by Strand et al. (1977) on fish larvae. The dose equation is as follows: Dß = 2.13ε C, where, Dß = dose
210
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
Table 1 Nominal and measured radioactivity (kBq ml−1 ) of tritiated water (HTO) in seawater before and after exposure of M. edulis embryo-larvae Nominal Concentration (kBq ml−1 )
Measured pre-exposure activity (kBq ml−1 )
Seawater (SW) SW + MMS 0.37 3.7 37 370
0.003 0.003 0.496 2.460 30.200 284.180
± ± ± ± ± ±
Measured post-exposure activity (kBq ml−1 )
0.001 0.001 0.010 0.025 0.300 2.67
0.002 0.002 0.361 2.320 22.610 220.200
± ± ± ± ± ±
0.001 0.001 0.008 0.023 0.220 1.960
Percentage loss 33.33 33.33 27.20 5.70 25.13 22.51
Mean ± S.E. from five replicate counts from different beakers at each concentration.
in rads per hour, ε = average beta-ray energy (3 H = 0.00569 MeV), C = concentration of 3 H (in Ci ml−1 : where 1 Ci is equal to 37 kBq) and 2.13 is the conversion factor. These calculations were based on the assumptions that: (a) 3 H was uniformly distributed within the embryo-larvae over the exposure period and (b) that no concentration of 3 H above a water equilibrium level occurred in the embryos. The predicted dose received during the exposure to 3 H ranged between 0.02 and 27.83 mGy as presented in Table 2. 3.3. Analysis of RAPD results Rather than showing individual changes in RAPD patterns related to HTO treatments, we decided to show some variations in profiles that occurred at different doses. Fig. 3a and b are an illustration of such changes. For instance with primer OPB6, the band 6-1 was more intense after 1.5 and 2 cell cycles than the control at all doses of 3 H and even in the positive control (MMS) (Fig. 3a). RAPD profiles generated by primer OPB12 show that band 12-1 present in the control profile nearly disappeared after 1.5 and 2 cell cycles in embryos exposed to 3.7, 37 and 370 kBq ml−1 (Fig. 3b). The use
of two DNA concentrations (5 and 20 ng) confirmed the reproducibility of the RAPD profiles with primer OPB6 and OPB12 (data not presented). Finally, not all primers showed differences in RAPD profiles generated by the HTO treated population in comparison to control patterns (data not shown). 3.4. Analysis of sister chromatid exchanges The induction of SCEs in the embryo-larvae of M. edulis is shown in Fig. 4a. The frequency of SCEs increased with increasing activity concentration of HTO. Statistical analysis using Kruskal–Wallis test showed that there was a significant difference between the controls and the samples treated with HTO (P < 0.005). The controls were not significantly different to the lowest activity concentration (0.37 kBq ml−1 ) but there was significant difference among the controls and the other three concentrations of HTO. It was also suggested that there was a significant difference between all activity concentrations of HTO, with 0.37 being different to 3.7, 37 and 370 and 3.7 being different to 37 and 370 kBq ml−1 . In addition, there was a significant difference for the induction of SCEs between
Table 2 Dose (mGy) received by M. edulis embryo-larvae for different exposure period and endpoints following exposure to tritiated water (HTO) Concentration (kBq ml−1 )
Cabs (exposure time = 16.7 h) (mGy)
SCEs andPRI (exposure time = 18.6 h) (mGy)
Developmental and survival effects (exposure time = 23 h) (mGy)
Seawater (0.00) MMS (0.00) 0.49 (0.37) 2.46 (3.70) 30.20 (37.00) 284.18 (370.00)
0.00 (0.00) 0.00 (0.00) 0.03 (0.02) 0.13 (0.21) 1.65 (2.14) 15.54 (21.42)
0.00 (0.00) 0.00 (0.00) 0.03 (0.02) 0.15 (0.24) 1.84 (2.37) 17.31 (23.72)
0.00 (0.00) 0.00 (0.00) 0.04 (0.03) 0.19 (0.28) 2.27 (2.78) 21.41 (27.83)
Actual based on measured mean HTO concentrations presented in Table 1. Values in parentheses represent the predicated values.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
211
Fig. 3. RAPD profiles of embryo-larvae M. edulis exposed to tritiated water (HTO) and MMS. RAPD profiles were generated using 10-mer primers OPB6: (a) and OPB12 (b). M: DNA molecular size marker (GeneRulerTM 100 bp DNA ladder plus, Immunogen International). The molecular sizes (in kilobases) of selected bands are indicated on the left of each panel. Bands visualised were from top to bottom 3000, 2000, 1500, 1200, 1031, 900, 800, 700, 600, 500, 400, 300, 200 and 100 bp. (−): No DNA control (negative control). The number of cell cycles are indicated in both panels. Embryos were exposed to 0 (line 1), 0.37 (line 2), 3.7 (line 3), 37 (line 4) and 370 kBq ml−1 (line 5) as well as 1.0 × 10−3 M of MMS. Selected changes are indicated by arrows.
embryo-larvae exposed to 37 and 370 kBq ml−1 HTO. 3.5. Analysis of chromosomal aberrations Fig. 5 represents the induction of Cabs in embryolarvae of M. edulis as a function of dose delivered by
each concentration of HTO over the exposure period. ANOVA indicated that there was a statistically significant increase for the induction of both aberrant cells (P < 0.001) and the number of total aberrations (P < 0.005) in HTO exposed samples compared to seawater controls. However, there was no significant difference among the three highest concentrations of HTO
212
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
Fig. 4. (a) Induction of sister chromatid exchanges (SCEs) and (b) Proliferative rate index (PRI) in M. edulis embryo-larvae after exposure to tritiated water (HTO). Error bars represent standard error. (*) Indicates means significantly different from control (P < 0.05).
for the induction of aberrations. The baseline level of aberrations in the controls was 7 ± 2 (mean and S.E.) aberrant cells containing 11 ± 6 (mean and S.E.) aberrations in total. MMS exposed cells contained 31 ± 2 aberrant cells and 32.5 ± 15 total aberrations (data not included in Fig. 5). The frequency of aberrant cells following MMS exposure was greater compared to HTO, however, the overall frequency of total aberrations was greater in the HTO exposed embryo-larvae. Thus, HTO exposed embryo-larvae had less cells containing aberrations than MMS exposed embryo-larvae but HTO exposed cells that did have aberrations contained more
aberrations than MMS exposed cells. As the majority of the aberrations in this study were acentrics (chromosome/chromatid breaks) the distribution pattern was over dispersed for acentrics as have been observed for most studies carried out for external low LET radiation (Lloyd et al., 1986). In the (untreated) controls, 31.8% of aberrations observed were chromatid type aberrations while in the HTO exposed cells the frequency of chromatid type aberrations ranged from 45.2 to 67.9%. There was no trend for the ratio of chromosome and chromatid type aberrations for different doses.
Fig. 5. Induction of chromosomal aberrations (Cabs) in M. edulis embryo-larvae as a function of predicted radiation dose. Table 2 shows details of actual and predicted doses. Line of best fit = logarithmic.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
213
Fig. 6. Percentage of: (a) normal and (b) dead M. edulis embryo-larvae following exposure to tritiated water (HTO). Error bars represent standard deviation. (*) Indicates means significantly different from control (P < 0.05).
3.6. Analysis of cytotoxic effects (i.e. proliferative rate index or PRI) Fig. 4b shows the values for PRI after exposure to HTO in comparison with controls. Statistical analysis of the results using ANOVA showed that all four concentrations of HTO have statistically significantly lower values for PRI compared to the controls. However, there was no significant difference among different concentrations of HTO except between 0.37 and 370 kBq ml−1 . 3.7. Analysis of developmental and survival/mortality effects Fig. 6a and b represent the percentage of normal and dead M. edulis embryo-larvae after 72 h postfertilisation following exposure to HTO and MMS. A dose dependent decrease for the percentage of normal embryo-larvae occurred at 24, 48 and 72 h (P < 0.001 for all times) with all the concentrations of HTO and MMS being significantly different only from the control as indicated using ANOVA. By 72 h only the lowest concentration of HTO (0.37 kBq ml−1 ) contained a small frequency of normal embryo-larvae and all the other concentrations were devoid of normality. The percentage of dead embryo-larvae in the seawater controls did not vary greatly over the three days with approximately 10% of the embryos being dead. At 72 h there was a large increase in mortality in embryo-larvae
exposed to HTO and MMS. With all the concentrations of HTO and the MMS being significantly different from the control (P < 0.001). Furthermore, the lowest concentration (i.e. 0.37 kBq ml−1 ) was also significantly different from all the other concentrations and the MMS. There was no difference in mortality between 3.7 and 37 kBq ml−1 but they were statistically different from both the 370 kBq ml−1 and MMS exposed larvae. The regression coefficient indicated a moderate dose–response relationship (R2 = 42.69%). One hundred percent mortality was exhibited at 72 h in the highest HTO concentration as well as the MMS concentration. Consequently, the 72 h LC50 for M. edulis could not be predicted. The predicted 48 h LC50 was, however, 0.27 Bq ml−1 with a received dose of 0.9 mGy.
4. Discussion The analysis of RAPD profiles in the present study showed changes in patterns due to 3 H induced DNA alterations even at the lowest HTO concentration (i.e. 0.37 kBq ml−1 ). Although it is difficult to decipher the exact mechanisms responsible for the changes in RAPD profiles following HTO exposure, it has been demonstrated that DNA damage and mutations can induce changes in RAPD patterns after exposure to genotoxic agents (Atienzar et al., 2002). For instance, damaged DNA can block or reduce the processivity of the Taq DNA polymerase leading to disappearance
214
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
of bands in the RAPD profiles. Alternatively, conformational changes in the DNA could also improve the access of the primer(s) to the binding site(s) leading to appearance of new bands in the corresponding RAPD patterns. In fact, such changes in the RAPD profiles have been correlated with other ecotoxicological parameters (i.e. growth, development and reproductive success) in aquatic invertebrates (Atienzar et al., 1999, 2001; Atienzar and Jha, 2004). Combined with robust protocols, such molecular approaches could play an increasingly important role in determining the impact of genotoxins in the aquatic environment (Jha, 2004). Despite the fact that low-LET ionising radiations delivered by external sources are suggested to be poor inducer of SCEs (Natarajan et al., 1993), in the present study a clear dose-dependent increase for SCEs was found between 3.7 and 370 kBq ml−1 of HTO compared to controls. Straume and Carsten (1993) reported a statistically significant increase for the incidence of SCEs on bone marrow cells of mice maintained on 111 kBq ml−1 of ingested HTO for 81–216 days. Since induction of SCEs is S-phase dependent phenomenon, it could be suggested that 3 H interferes either with the replication process or inhibits the actions of associated enzymes, in common with other chemical genotoxins (Jha et al., 1992, 1996, 2000a,b). Suyama et al. (1981) reported increased frequency of Cabs (i.e. chromosome bridges) when eggs of the freshwater teleost, Oryzias latipes, were treated with HTO delivering doses higher than 0.19 Gy. This is slightly higher than the dose that induced significantly elevated levels of Cabs in the embryo-larvae of M. edulis in the present study. While a dose–response relationship for SCEs, developmental abnormalities and mortality, were observed in the present study, there was a lack of clear dose-response relationships for Cabs and PRI. The reasons for this apparent lack of dose–response relationships are not clear. It could be hypothesized that only critical sites or sensitive cells are targets for radiation action, when these small-sized organisms, undergoing cellular differentiation come into contact or uptake the radionuclides in common with higher organisms (Gossner, 2001). It could also be speculated that in the very sensitive embryo-larval life stages, highly damaged cells leading to death are eliminated from analysis, affecting both the yield of aberrations and analysis of cells in different stages of division. In other words, the absence of a clear dose-
dependent increase for Cabs and PRI in the very early life stages of aquatic organisms could be related to their known capacity to undergo apoptosis, thereby efficiently eliminating cells with high frequencies of Cabs, as observed in mammalian cells (Jha et al., 1995b). In addition, it is also possible that if highly damaged cells are eliminated from the analysis, only the ‘bystander effects’ are analysed (Morgan, 2003). Furthermore, apart from unstable Cabs as observed in the present study, intake of HTO can also induce stable or balanced Cabs that could be detected several years after intake by humans (Lloyd et al., 1998). In parallel with human health risk assessment, it is therefore important to take into account such stable genetic damage and potential delayed effects (Mothersill and Seymour, 1998) to protect the environment. As well as alterations in RAPD profile, induction of SCEs and Cabs, HTO exposed embryo-larvae also experienced lower values of PRI due to a reduction in cell cycle progression probably caused by an increase in toxicity. However, as mentioned above, no marked difference for the PRI values could be found in the embryo-larvae exposed to different concentrations of HTO. With respect to this apparent lack of dose–response relationship for PRI, in addition to possible explanations provided above, it should be pointed out that although originally proposed by Lamberti et al. (1983) for application to actively proliferating and highly differentiated mammalian cells in vitro, the PRI has been found to provide an useful means to quantify the cytotoxicity in embryo-larval stages of aquatic organisms following exposure to different toxicants (Jha et al., 1996, 2000a,b; Hagger et al., 2002). However, the PRI for the embryo-larvae probably represents an average figure, based on a mixed population of highly differentiating cells with differing mitotic rates over the exposure period described (Jha et al., 1996). Therefore, even a slight perturbation in the rate of cell cycle progression, cellular differentiation and programmed cell death, for which as mentioned above these cells have a well known capacity, could lead to significant changes in the PRI values. This is, however, in contrast to our earlier studies using chemical genotoxins, where PRI values showed significant differences from controls only at the higher concentrations of test compounds (e.g. methyl methane sulphonate, benzo (a) pyrene, tributyltin, contaminated sediment samples) in the embryo-larvae of M. edulis and P.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
dumerilii (Jha et al., 2000a,b; Hagger et al., 2002). A lower plateau-value for PRI at higher concentrations of HTO in the present study therefore appears to be related to the mechanisms of action of ionising radiation, which, in contrast to chemicals, could impart their effect continuously the very moment cells come in contact with the radionuclides. In contrast to induction of Cabs and PRI values, compared to unexposed controls, the study showed a significant dose-dependent increase for mortality in HTO exposed embryo-larvae by the end of the experiment. Abbott and Mix (1979) also observed similar effects on survival in the larval stage of goose barnacles following exposure to seawater containing only 0.2 Bq ml−1 of HTO. In contrast to the sexually mature adults, the embryo-larval stages of aquatic organisms are considered to be several orders of magnitude more sensitive in terms of toxicological injury. The highest detrimental effects of radiation in both mammals and aquatic organisms have been observed in experiments when irradiation occurred immediately after fertilisation. It has also been suggested that the irradiation of cells immediately after fertilisation may also encompass the sensitive period of second mitosis (Anderson and Harrison, 1990). It is therefore not surprising that 1h old embryo showed extensive developmental abnormalities and mortality following exposure to 3 H, an internal emitter. In this context, Knowles and Greenwood (1997) also found that the development and survival of eggs to larvae of the polychaete worm, Ophryotrocha diadema, was significantly reduced when exposed to 2.24 MBq ml−1 of HTO but was not affected by similar doses of ␥radiation delivered by 137 Cs. Indeed, -rays from HTO have been shown to be more effective than ␥-rays emitted by 90 Sr, 90 Y (in solution), 60 Co and X-rays (external sources) in inducing Cabs in the blastula stage of Oryzias latipes (freshwater teleost) eggs, potentially leading to an increase in abnormality and mortality (Suyama et al., 1981), as observed in the present study. While having a global view of all the concentration or dose-dependent responses in the present study, it is interesting to observe that while significantly decreased values of normal embryo-larvae (correspondingly increased frequencies of deads) and PRI (which had a ‘plateau level’ effect at all the higher concentrations) were found at the lowest concentra-
215
tion (0.37 kBq ml−1 ) of HTO, no significant increase for SCEs and aberrant cells were found at this concentration. These discrepancies in ‘concentration–effect’ relationships for the lowest concentration of HTO appear to be odd. In fact, studies in the literature simultaneously evaluating concentration dependent effects at different levels of biological organisation (i.e. chromosomal, cellular and individual) are very scarce. From the observations made in the present study, it appears that there is not a direct, linear relationship between observed effects at lower (i.e. chromosomal and cellular) and higher (e.g. developmental abnormalities and mortality) levels of biological organisation. It is also to be remembered here that while cytogenetic effects (i.e. SCEs, Cabs and PRI) were evaluated soon after HTO exposure (one and half to two cell cycles), the developmental abnormality and survival effects were analysed at later stages (data presented for 72 h post-fertilisation period). It appears therefore that induced damage at lower levels of biological organisation, which could have knock-on effects as a function of time, could precipitate cumulatively and thus be manifested later during the developmental process at the individual level. In conclusion, our integrated study linking genetic and cellular biomarkers with embryo-larval bioassay suggests that extremely low doses of -radiation emitted by HTO can induce genetic damage in the embryolarval stages of marine mussels and that this damage potentially accumulates at cellular and individual levels. Such an approach could be adopted to early life stages of other invertebrates to establish dose–response relationships since estimation of dose received by the biota and resultant effects are important in radiological protection criteria for the natural environment (Pentreath, 1998; ERICA, 2005). Our study goes some way towards achieving this goal.
Acknowledgements This work was supported financially, mainly by the Natural Environmental Research Council (NERC), Environment Agency (EA) (GT22-1998-EA-0003) in association with Devonport Royal Dockyard (DML), Devon, UK. The RAPD work was supported by the Marie Curie Grant of the European Commission (ERB4001-GT-97-0136).
216
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
References Abbott, D.T., Mix, M.C., 1979. Radiation effects of tritiated seawater on development of the Goose Barnacle, Pollicipes polymerus. Health Phys. 36, 283–287. Anderson, S.L., Harrison, F.L., 1990. Guidance document for prepermit bioassay testing of low level radioactive waste. United States Environmental Protection Agency, pp. 50. ASTM, 1989. Standard guide for conduction static acute toxicity tests starting with embryos of four species of saltwater bivalve molluscs. E724-89, America Society for Testing of Materials, Philadelphia. Atienzar, F.A., Jha, A.N., 2004. The random amplified polymorphic DNA (RAPD) assay to determine DNA alterations, repair and transgenerational effects in B(a)P exposed Daphnia magna. Mutat. Res. 552, 125–140. Atienzar, F.A., Conradi, M., Evenden, A.J., Jha, A.N., Depledge, M.H., 1999. Qualitative assessment of genotoxicity using random amplified polymorphic DNA: comparison of genomic template stability with key fitness parameters in Daphnia magna exposed to benzo[a]pyrene. Environ. Toxicol. Chem. 18, 2275– 2282. Atienzar, F.A., Cheung, V.V., Jha, A.N., Depledge, M.H., 2001. Fitness parameters and DNA effects are sensitive indicators of copper induced toxicity in Daphnia magna. Toxicol. Sci. 59, 241–250. Atienzar, F.A., Venier, P., Jha, A.N., Depledge, M.H., 2002. Evaluation of the random amplified polymorphic DNA (RAPD) assay for the detection of DNA damage and mutations. Mutat. Res. 521, 151–163. Bayne, B.L., 1976. The biology of mussel larvae. In: Bayne, B.L. (Ed.), Marine Mussels, their Ecology and Physiology. Cambridge University Press, Cambridge, pp. 81–119. Burgeot, T., His, E., Galgani, F., 1995. The micronucleus assay in Crassostrea gigas for the detection of seawater genotoxicity. Mutat. Res. 342, 125–140. Department of the Environment, Transport and the Regions, 2000. UK strategy for radioactive discharges, 2001–2020. Consultation document. DETR, London, pp. 1–74. Environment Agency, 2001. Proposed Decision Document on the Application made by Devonport Royal Dockyard Limited to Dispose of Radioactive Wastes from: Devonport Royal Dockyard Plymouth. Environment Agency, UK, p. 288. ERICA, 2005. Environmental risk from ionising contaminants; www.erica-project.org (Accessed January 2005). Gossner, W., 2001. Target cells for internal dosimetry. Radiat. Prot. Dosim. 105, 39–42. Hagger, J.A., Fischer, A.S., Hill, S.J., Depledge, M.H., Jha, A.N., 2002. Genotoxic, cytotoxic and ontogenetic effects of tri-nbutyltin on the marine worm, Platynereis dumerilii (Polychaeta: Nereidae). Aquat. Toxicol. 57, 243–255. Harrison, F.L., Jones, I.M., 1982. An in vivo sister chromatid exchange assay in the larvae of the mussel Mytilus edulis: response to three mutagens. Mutat. Res. 105, 235–242. His, E., Beiras, R., Seaman, M.N.L., 1999. The assessment of marine pollution—bioassays with bivalve embryos and larvae. Adv. Mar. Biol. 37, 1–178.
ICRP. Statement from the 1983 meeting of the International Commission on Radiological Protection. 1984. ICRP Publications 39, Ann. ICRP 14:1, 1–8. Jha, A.N., 2004. Genotoxicological studies in aquatic organisms: an overview. Mutat. Res. 552, 1–17. Jha, A.N., Noditi, M., Nilsson, R., Natarajan, A.T., 1992. Genotoxic effects of sodium arsenite on human cells. Mutat. Res. 284, 215–221. Jha, A.N., Hutchinson, T.H., Mackay, J.M., Elliott, B.M., Pascoe, P.L., Dixon, D.R., 1995a. The chromosomes of Platynereis dumerilii (Polychaeta: Nereidae). J. Mar. Biol. Assoc. U.K. 75, 551–562. Jha, A.N., Hande, P.M., Mullenders, L.H.F., Natarajan, A.T., 1995b. Mimosine is a potent clastogen in primary and transformed hamster fibroblasts but not in primary or transformed human lymphocytes. Mutagenesis 10, 385–391. Jha, A.N., Hutchinson, T.H., Mackay, J.M., Elliott, B.M., Dixon, D.R., 1996. Development of an in vivo genotoxicity assay using the marine worm Platynereis dumerilii (Polychaeta: Nereidae). Mutat. Res. 359, 141–150. Jha, A.N., Cheung, V.V., Foulkes, M.E., Hill, S.J., Depledge, M.H., 2000a. Detection of genotoxins in the marine environment: adoption and evaluation of an integrated approach using the embryolarval stages of the marine mussel, Mytilus edulis. Mutat. Res. 464, 213–228. Jha, A.N., Hagger, J.A., Hill, S.J., 2000b. Tributyltin induces cytogenetic damage in the early life stages of the marine mussel, Mytilus edulis. Environ. Mol. Mutag. 35, 343–350. Kirchmann, R., Koch, G., Adam, V., van Den Hoek, J., 1973. Studies on the food chain contamination by tritium. In: Moghissi, A.A., Carter, M.W. (Eds.), Tritium. Messenger graphics, Nevada. Kirkland, D.J., 1990. Basic mutagenicity tests: UKEMS recommended procedures. Cambridge University Press, pp. 1–143. Knowles, J.F., Greenwood, L.N., 1997. A comparison of the effect of long-term - and ␥-irradiation on the reproductive performance of a marine invertebrate Ophryotrocha diadema (Polychaete Dorvilleidae). J. Environ. Radioact. 34, 1–7. Lamberti, L., Bigatti, P., Ardito, G., 1983. Cell kinetics and sister chromatid exchange frequency in human lymphocytes. Mutat. Res. 120, 193–199. Lloyd, D.C., Edwards, A.A., Prosser, J.S., 1986. Chromosome aberrations induced in human lymphocytes by in vitro acute X and gamma radiation. Radiat. Prot. Dosim. 15, 83–88. Lloyd, D.C., Moquet, J.E., Oram, S., Edwards, A.A., Lucas, J.N., 1998. Accidental intake of tritiated water: a cytogenetic followup case on translocation stability and dose reconstruction. Int. J. Radiat. Biol. 73, 543–547. McCubbin, D., Leonard, K.S., Bailey, T.A., Williams, J., Tossell, P., 2001. Incorporation of organic tritium (3 H) by marine organisms and sediment in the Severn Estuary/Bristol Channel (UK). Mar. Pollut. Bull. 42, 852–863. Morgan, W.F., 2003. Is there a common mechanism underlying genome instability, bystander effects and other-targeted effects of exposure to ionising radiation? Oncogene 22, 7094–7099. Mothersill, C., Seymour, C.B., 1998. Mechanisms and implications of genomic instability and other delayed effects of ionising radiation exposure. Mutagenesis 13, 421–426.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217 Natarajan, A.T., Darroudi, F., Jha, A.N., Meijers, M., Zdzienicka, M.Z., 1993. Ionising radiation induced DNA lesions which lead to chromosomal aberrations. Mutat. Res. 299, 297–303. Pentreath, R.J., 1998. Radiological protection criteria for the natural environment. Radiat. Prot. Dosim. 75, 175–179. Sparks, T., 2000. Statistics in Ecotoxicology. John Wiley and Sons Ltd., Wiltshire, UK, 229 p. Strand, J.A., Fujihara, M.P., Burdett, R.D., Poston, T.M., 1977. Suppression of the primary immune response in Rainbow
217
Trout, Salmo gairdneri, sublethally exposed to tritiated water during embryogenesis. J. Fish. Res. Board Can. 34, 1293– 1304. Straume, T., Carsten, A.L., 1993. Tritium radiobiology and relative biological effectiveness. Health Phys. 65, 657–672. Suyama, I., Etoh, H., Maruyama, T., Kato, Y., Ichikawa, R., 1981. Effects of ionising radiation on the early development of Oryzias eggs. J. Radiat. Res. 22, 125–133.
Genotoxic, cytotoxic, developmental and survival effects of tritiated water in the early life stages of the marine mollusc, Mytilus edulis Josephine A. Hagger, Franck A. Atienzar 1 , Awadhesh N. Jha ∗ School of Biological Sciences, University of Plymouth, Drake Circus, Devon, Plymouth PL4 8AA, UK Received 12 January 2005; received in revised form 27 May 2005; accepted 28 May 2005
Abstract Using an integrated approach linking different levels of biological organisation, the genotoxic, cytotoxic, developmental and survival impact of tritiated water (HTO) were investigated in the embryo-larvae of marine mollusc Mytilus edulis. One-hour-old embryos were exposed to a range of concentrations (0.37–370 kBq ml−1 ) of HTO, which delivered a dose between 0.02 and 21.41 mGy over the exposure period for different end points. Detrimental effects, if any, were monitored at different levels of biological organisation (i.e. DNA, chromosomal, cellular and individual). Genotoxic effects were assessed using molecular and cytogenetic approaches which included analysis of random amplified polymorphic DNA (RAPD), induction of sister chromatid exchanges (SCEs) and chromosomal aberrations (Cabs). Cytotoxic effects were evaluated by determining the proliferative rate index (PRI) of the embryo-larval cells. Developmental and survival effects were also monitored every 24 h up to 72 h. Results in general indicated that HTO significantly increased cytogenetic damage, cytotoxicity, developmental abnormalities and mortality of the embryo-larvae as a function of concentration or radiation dose. The analysis of RAPD profiles also revealed qualitative effects in the HTO exposed population compared to controls. However, while the embryo-larvae showed dose or concentration dependent effects for mortality, developmental abnormalities and induction of SCEs, the dose-dependent effects were not apparent for Cabs and PRI at higher doses. The study contributes to our limited understanding of the impact of environmentally relevant radionuclides on non-human biota and emphasises the need for further investigations to elucidate potentially long term damage induced by persistent, low levels of other radionuclides on commercially and ecologically important species, in order to protect human and ecosystem health. © 2005 Elsevier B.V. All rights reserved. Keywords: Tritium; Genotoxicity; RAPD assay; Developmental effects; Marine mussels
1. Introduction ∗
Corresponding author. Tel.: +44 1752 232978; fax: +44 1752 232970. E-mail address: [email protected] (A.N. Jha). 1 Present address: UCB S.A. Pharma Sector, Chemin du Foriest, B-1420 Braine-l’Alleud, Belgium. 0166-445X/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.aquatox.2005.05.013
In 1977 the International Commission on Radiological Protection stated that “If humans are protected from the effects of ionising radiation, then flora and fauna
206
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
are also adequately protected” (ICRP, 1984). However, this statement is often questioned by the general public and regulatory authorities as: (a) different species vary in their sensitivity to toxicants including radiation; (b) different life stages of the same species can show varying levels of response to contaminants; (c) there are areas where wildlife may be exposed to higher levels of radiation than are permissible to humans and (d) endangered species with longer generation times or isolated populations might need special consideration, in order to maintain ecosystem quality. At present, an internationally accepted method for assessing the environmental impact of ionising radiation does not exist (Pentreath, 1998). Attempts are therefore being made to link together the release scenario of radioactive substances, potential exposure pathways, dose to ecologically relevant, commercially important and critical target organisms and subsequently effects at different levels of biological organisation (ERICA, 2005). As a result of nuclear establishments around coastal areas, the marine environment is often the ultimate recipient of radioactive contaminants. In this context, tritium (3 H), a naturally occurring isotope of the element hydrogen, is manufactured commercially and is also released as a by-product of nuclear reactions. An overview of historic trends in liquid and aerial discharges of radioactive substances in the UK, carried out between 1979 and 1998, showed that there has been a sustained increase in the discharge of 3 H from different nuclear facilities (Department of the Environment, Transport and the Regions, 2000). As HTO can move freely in the biosphere it could be a threat to human health as it may be ingested not only through drinking water but also by food, especially from fish and shellfish that are known to bioaccumulate some toxicants (Kirchmann et al., 1973). In this context, elevated activity concentrations of 3 H have been reported in sediment and at different trophic levels of biota from the Severn estuary, UK (McCubbin et al., 2001). It was suggested that the high activity concentrations in these materials, relative to that in seawater, was due to the presence of bioavailable organic 3 H-labelled compounds in radiochemical wastes discharged in the vicinity. Despite recent moves to protect non-human biota from exposures to ionising radiations and the growing scientific and public concern over the presence of 3 H in the environment (Environment Agency, 2001), very little work has been carried out to evaluate the radiobio-
logical or toxicological impact of 3 H on commercially and/or ecologically important species. In particular, there has been a lack of research on potential detrimental effects of ionising radiation on invertebrates, which constitute over 90% of the existing species and play a major role in the maintenance and function of every ecosystem. Consequently, the aim of the present study was to assess the genotoxic, cytotoxic and developmental effects of -radiation emitted by HTO on embryolarvae of an ecologically and commercially important invertebrate, the marine bivalve mollusc Mytilus edulis.
2. Materials and methods 2.1. Exposure scenario HTO was obtained from ICN Pharmaceuticals, Inc., UK at a concentration of 3.7 GBq ml−1 . The range of concentrations of HTO used in the present study (0.37–370 kBq ml−1 ) was adopted from previous work carried out on fish and barnacle larvae (Strand et al., 1977; Abbott and Mix, 1979) and gave a dose between 0.02 and 21.42 mGy over the exposure period as described in Section 3.2. Fig. 1 shows the schematic representation of the general protocol used for assessing the genotoxic [random amplified polymorphic DNA (RAPD)], sister chromatid exchanges (SCEs) and chromosomal aberration (Cabs) assays, cytotoxic proliferative rate index (PRI), developmental and survival effects of HTO on M. edulis embryo-larvae. In brief, following the induction of spawning, as described in detail elsewhere (Jha et al., 2000a), eggs from two female mussels and sperm from one male were collected. The number of eggs was adjusted to 50 eggs ml−1 prior to adding sperm solution at approximately 107 sperm ml−1 . Fertilisation (rate 92%) was carried out at 15 ± 1 ◦ C, whereafter excess sperm was removed by filtering through a sieve (30 m) and the embryo density adjusted to approximately 30 embryos ml−1 as per standard guidelines for acute toxicity testing of environmental contaminants (ASTM, 1989). Following fertilisation, the embryos were allowed to grow for 1 h, after which they were divided into different exposure vessels (glass beakers) where the final volume was adjusted to 500 ml with seawater. Temperature, salinity and pH were controlled throughout the exposure and air was supplied
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
207
(Jha et al., 2000a,b). HTO concentrations were made using a serial dilution with seawater from the original stock. One hundred microlitres of each dilution was added to 500 ml of seawater containing the embryos. MMS was also made using a serial dilution with seawater and 100 l of sample was added to achieve the desired concentration for the volume of seawater. In order to obtain sister chromatid differential staining (SCD) a minimal concentration (10−5 M) of 5-bromo2-deoxyuridine (BrdU) was added to all the vessels at 12 h post-fertilisation. 2.2. Random amplified polymorphic DNA (RAPD) assay As described in Fig. 1, along with Cabs and SCEs assays, a subset of embryo-larvae were processed for analysis using the random amplified polymorphic DNA assay. Genomic DNA was extracted and purified using a conventional phenol/chloroform method as described elsewhere (Atienzar et al., 1999). DNA concentrations were determined by electrophoresis by comparison with known concentrations of Lambda phage DNA (Sigma, Poole, UK). RAPD was performed with 5 ng of genomic DNA as described elsewhere in details (Atienzar et al., 2002). The 10-mer primers used were OPA9, OPB1, OPB5, OPB6, OPB7, OPB8, OPB10, OPB11, OPB12, OPB14, or OPB17 (Operon Technologies, Southampton, UK; for the sequences see Atienzar and Jha, 2004). Thermal cycling parameters and analysis of PCR products by electrophoresis have been described elsewhere in details (Atienzar et al., 2002; Atienzar and Jha, 2004). The GeneRulerTM 100 bp DNA ladder plus (Immunogen International, Sunderland, UK) was used. Fig. 1. Flow chart illustrating the experimental protocol adopted to evaluate genotoxic, cytotoxic, developmental and survival effects of tritiated water (HTO) on the embryo-larvae of M. edulis.
via gentle aeration. Embryos were exposed to four different activity concentrations of HTO (0.37, 3.7, 37 and 370 kBq ml−1 ), 1 negative control (i.e. 10 m filtered seawater) and 1 positive control containing methyl methane sulphonate (1.0 × 10−3 M MMS) as a reference genotoxic agent. This single concentration of MMS was selected based on our earlier studies
2.3. Slide preparation for cytogenetic (i.e. PRI, SCEs and Cabs) analyses Following addition of BrdU, after 1 21 (Cabs) and 2 cell cycles (SCEs and PRI) two subsets of embryos were removed (1 cell cycle = 3.8 h; Jha et al., 2000a,b). One subset was placed into colchicine dissolved in seawater (0.025%, w/v) for 30 min and then exposed to a series of hypotonic (seawater/0.075 M KCl) treatments. After the hypotonic treatments, the embryos were transferred into centrifuge tubes and were centrifuged for 5 min at 200 × g. After centrifugation
208
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
the supernatant was discarded and a few drops of cold (4 ◦ C) Carnoys fixative (1:3 (v/v) glacial acetic acid/methanol) was added. The cells were then processed to produce metaphase spreads, stained with Giemsa (for the analysis of Cabs), or processed for sister chromatid differential staining (for the analysis of SCEs and PRI) as previously described (Harrison and Jones, 1982; Jha et al., 1995a,b, 1996, 2000a,b) for microscopic analysis. Prior to scoring, slides were randomised, coded by an independent disinterested individual and then scored blind. Complete metaphase spreads (2n = 28) were examined using a bright field microscope (final magnification ×1000). Details of classification of PRI stages, scoring of Cabs and SCEs have been described in earlier studies (Jha et al., 1995a, 1996, 2000a, 2000b). In brief, metaphases were classified, as first (M1), second (M2) and third or subsequent (M3+) division cells. For PRI at least 200 cells per treatment were scored, 100 each from two replicates. PRI was calculated using the formula PRI = ((1 × M1) + (2 × M2) + (3 × M3))/ number of cells analysed (Lamberti et al., 1983). Only second (M2) division metaphases were analysed for the occurrence of SCEs. Fifty metaphases per treatment were analysed for SCEs, 25 from each of two replicates, and the results were expressed as the SCEs frequency per cell. For Cabs 200 cells from each treatment were scored (100 from each of two replicates) as per the recommendation of the United Kingdom Environmental Mutagen Society (UKEMS) (Kirkland, 1990). Cabs were expressed as the percent aberrant cells (percentage of cells that contained one or more aberrations including heavily damaged cells) and total aberrations (the number of chromosome plus chromatid type aberrations per 100 cells excluding gaps). 2.4. Analyses of developmental and mortality/ survival effects Developmental morphology, behaviour and mortality of the growing embryo-larvae were assessed at 24, 48 and 72 h post-fertilisation. Fig. 2 shows various stages of normal and abnormal development. The development of M. edulis consists of an embryonic phase followed by a larval phase (Bayne, 1976). At the experimental temperature (15 ± 1 ◦ C), the first larvae stage known as trochophores appears at approximately 24 h. This stage is defined as free swimming larvae,
with a circular band of short cilia (prototroch) and a flagellum (Fig. 2a). Abnormal larvae are usually misshapen (not spherical in shape or symmetrical) and may lack cilia and/or the flagella, resulting in abnormal swimming behaviour (Fig. 2b). The second larvae stage is the veliger, which in M. edulis is usually formed after 48 h (15 ◦ C). A dorsal thickening of the ectodermis, secreted by the shell gland, forms an initial organic cuticle that spreads over the entire body. A straight dorsal hinge gives the larvae the characteristic capital D shape, hence the name “D-larva” (Fig. 2c). At this stage the larvae are approximately 60–70 m in size (His et al., 1999). Dead trochophores appear as a mass of degenerating cells with no cilia or flagellum movement. Dead D-larvae may also lack ciliary movement, lose clarity of internal organs and become transparent (His et al., 1999). Pollutants present in the marine environment may lead to morphologically abnormal trochophores and D-larvae (Fig. 2b and d, respectively). 2.5. Water quality parameters and measurement of radioactivity Water quality parameters (pH, temperature and salinity) were monitored at the start and end of exposure and were relatively stable throughout (mean and S.E.; pH 8.07 ± 0.05; salinity 35 ± 1, 15.5 ± 0.5 ◦ C). Pre- and post-exposure levels of radioactivity were determined using 100 l of test solution in 5 ml of liquid scintillation cocktail, Packard Ultima Gold (Sigma–Aldrich Ltd., USA). Five replicates were counted from each activity concentration. Samples were read for 10 min (detection limits range from 0.06 to 14.5 keV) in a scintillation counter (Beckman LS6500, USA). Readings were displayed as counts per minute (CPM), then automatically converted to disintegrations per minute (DPM) using percentage efficiency for 3 H and then converted to kBq ml−1 (37 kBq equivalent to 2,220,000 DPM). The scintillation counter has a built in quench correction and 14 C and 3 H standards were used to calibrate the counter regularly. The lower detection limit for 3 H was 0.00962 kBq ml−1 . 2.6. Statistical analyses Statistical analysis were carried out using the statistical software package STATGRAPHICS Plus for Windows 3.1©from Statistical Graphics Corp. 1994–1997.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
209
Fig. 2. The developmental stages of M. edulis embryo-larvae. Photos a and c represent normal embryo-larvae at 24 h and >48 h, respectively. Photos (b) and (d) represent morphologically abnormal embryo-larvae at 24 h and >48 h, respectively.
One-way analysis of variance (ANOVA) was carried out on normalised data. If required, transformation of data for developmental effects was carried out using Arc SIN and square root as recommended by Burgeot et al. (1995) whereas SCEs, Cabs and PRI values were transformed either via log or square root conversions. Non-normally distributed data were analysed using the non-parametric Kruskal–Wallis test (Sparks, 2000).
was neither significant variation in the percentage of radioactivity lost during the exposure period (difference between pre- and post-exposure), nor any trend in the percentage loss for different concentrations of HTO used. Each vessel containing HTO lost similar percentages of radioactivity although the samples containing larger amounts showed a greater loss in terms of the actual concentration over the exposure period.
3. Results
3.2. Calculation of dose received by embryo-larvae
3.1. Determination of radioactivity Table 1 presents the levels of radioactivity in different test vessels for the exposure scenario. There
The estimated dose to growing embryo-larvae exposed to HTO was calculated by a method used by Strand et al. (1977) on fish larvae. The dose equation is as follows: Dß = 2.13ε C, where, Dß = dose
210
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
Table 1 Nominal and measured radioactivity (kBq ml−1 ) of tritiated water (HTO) in seawater before and after exposure of M. edulis embryo-larvae Nominal Concentration (kBq ml−1 )
Measured pre-exposure activity (kBq ml−1 )
Seawater (SW) SW + MMS 0.37 3.7 37 370
0.003 0.003 0.496 2.460 30.200 284.180
± ± ± ± ± ±
Measured post-exposure activity (kBq ml−1 )
0.001 0.001 0.010 0.025 0.300 2.67
0.002 0.002 0.361 2.320 22.610 220.200
± ± ± ± ± ±
0.001 0.001 0.008 0.023 0.220 1.960
Percentage loss 33.33 33.33 27.20 5.70 25.13 22.51
Mean ± S.E. from five replicate counts from different beakers at each concentration.
in rads per hour, ε = average beta-ray energy (3 H = 0.00569 MeV), C = concentration of 3 H (in Ci ml−1 : where 1 Ci is equal to 37 kBq) and 2.13 is the conversion factor. These calculations were based on the assumptions that: (a) 3 H was uniformly distributed within the embryo-larvae over the exposure period and (b) that no concentration of 3 H above a water equilibrium level occurred in the embryos. The predicted dose received during the exposure to 3 H ranged between 0.02 and 27.83 mGy as presented in Table 2. 3.3. Analysis of RAPD results Rather than showing individual changes in RAPD patterns related to HTO treatments, we decided to show some variations in profiles that occurred at different doses. Fig. 3a and b are an illustration of such changes. For instance with primer OPB6, the band 6-1 was more intense after 1.5 and 2 cell cycles than the control at all doses of 3 H and even in the positive control (MMS) (Fig. 3a). RAPD profiles generated by primer OPB12 show that band 12-1 present in the control profile nearly disappeared after 1.5 and 2 cell cycles in embryos exposed to 3.7, 37 and 370 kBq ml−1 (Fig. 3b). The use
of two DNA concentrations (5 and 20 ng) confirmed the reproducibility of the RAPD profiles with primer OPB6 and OPB12 (data not presented). Finally, not all primers showed differences in RAPD profiles generated by the HTO treated population in comparison to control patterns (data not shown). 3.4. Analysis of sister chromatid exchanges The induction of SCEs in the embryo-larvae of M. edulis is shown in Fig. 4a. The frequency of SCEs increased with increasing activity concentration of HTO. Statistical analysis using Kruskal–Wallis test showed that there was a significant difference between the controls and the samples treated with HTO (P < 0.005). The controls were not significantly different to the lowest activity concentration (0.37 kBq ml−1 ) but there was significant difference among the controls and the other three concentrations of HTO. It was also suggested that there was a significant difference between all activity concentrations of HTO, with 0.37 being different to 3.7, 37 and 370 and 3.7 being different to 37 and 370 kBq ml−1 . In addition, there was a significant difference for the induction of SCEs between
Table 2 Dose (mGy) received by M. edulis embryo-larvae for different exposure period and endpoints following exposure to tritiated water (HTO) Concentration (kBq ml−1 )
Cabs (exposure time = 16.7 h) (mGy)
SCEs andPRI (exposure time = 18.6 h) (mGy)
Developmental and survival effects (exposure time = 23 h) (mGy)
Seawater (0.00) MMS (0.00) 0.49 (0.37) 2.46 (3.70) 30.20 (37.00) 284.18 (370.00)
0.00 (0.00) 0.00 (0.00) 0.03 (0.02) 0.13 (0.21) 1.65 (2.14) 15.54 (21.42)
0.00 (0.00) 0.00 (0.00) 0.03 (0.02) 0.15 (0.24) 1.84 (2.37) 17.31 (23.72)
0.00 (0.00) 0.00 (0.00) 0.04 (0.03) 0.19 (0.28) 2.27 (2.78) 21.41 (27.83)
Actual based on measured mean HTO concentrations presented in Table 1. Values in parentheses represent the predicated values.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
211
Fig. 3. RAPD profiles of embryo-larvae M. edulis exposed to tritiated water (HTO) and MMS. RAPD profiles were generated using 10-mer primers OPB6: (a) and OPB12 (b). M: DNA molecular size marker (GeneRulerTM 100 bp DNA ladder plus, Immunogen International). The molecular sizes (in kilobases) of selected bands are indicated on the left of each panel. Bands visualised were from top to bottom 3000, 2000, 1500, 1200, 1031, 900, 800, 700, 600, 500, 400, 300, 200 and 100 bp. (−): No DNA control (negative control). The number of cell cycles are indicated in both panels. Embryos were exposed to 0 (line 1), 0.37 (line 2), 3.7 (line 3), 37 (line 4) and 370 kBq ml−1 (line 5) as well as 1.0 × 10−3 M of MMS. Selected changes are indicated by arrows.
embryo-larvae exposed to 37 and 370 kBq ml−1 HTO. 3.5. Analysis of chromosomal aberrations Fig. 5 represents the induction of Cabs in embryolarvae of M. edulis as a function of dose delivered by
each concentration of HTO over the exposure period. ANOVA indicated that there was a statistically significant increase for the induction of both aberrant cells (P < 0.001) and the number of total aberrations (P < 0.005) in HTO exposed samples compared to seawater controls. However, there was no significant difference among the three highest concentrations of HTO
212
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
Fig. 4. (a) Induction of sister chromatid exchanges (SCEs) and (b) Proliferative rate index (PRI) in M. edulis embryo-larvae after exposure to tritiated water (HTO). Error bars represent standard error. (*) Indicates means significantly different from control (P < 0.05).
for the induction of aberrations. The baseline level of aberrations in the controls was 7 ± 2 (mean and S.E.) aberrant cells containing 11 ± 6 (mean and S.E.) aberrations in total. MMS exposed cells contained 31 ± 2 aberrant cells and 32.5 ± 15 total aberrations (data not included in Fig. 5). The frequency of aberrant cells following MMS exposure was greater compared to HTO, however, the overall frequency of total aberrations was greater in the HTO exposed embryo-larvae. Thus, HTO exposed embryo-larvae had less cells containing aberrations than MMS exposed embryo-larvae but HTO exposed cells that did have aberrations contained more
aberrations than MMS exposed cells. As the majority of the aberrations in this study were acentrics (chromosome/chromatid breaks) the distribution pattern was over dispersed for acentrics as have been observed for most studies carried out for external low LET radiation (Lloyd et al., 1986). In the (untreated) controls, 31.8% of aberrations observed were chromatid type aberrations while in the HTO exposed cells the frequency of chromatid type aberrations ranged from 45.2 to 67.9%. There was no trend for the ratio of chromosome and chromatid type aberrations for different doses.
Fig. 5. Induction of chromosomal aberrations (Cabs) in M. edulis embryo-larvae as a function of predicted radiation dose. Table 2 shows details of actual and predicted doses. Line of best fit = logarithmic.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
213
Fig. 6. Percentage of: (a) normal and (b) dead M. edulis embryo-larvae following exposure to tritiated water (HTO). Error bars represent standard deviation. (*) Indicates means significantly different from control (P < 0.05).
3.6. Analysis of cytotoxic effects (i.e. proliferative rate index or PRI) Fig. 4b shows the values for PRI after exposure to HTO in comparison with controls. Statistical analysis of the results using ANOVA showed that all four concentrations of HTO have statistically significantly lower values for PRI compared to the controls. However, there was no significant difference among different concentrations of HTO except between 0.37 and 370 kBq ml−1 . 3.7. Analysis of developmental and survival/mortality effects Fig. 6a and b represent the percentage of normal and dead M. edulis embryo-larvae after 72 h postfertilisation following exposure to HTO and MMS. A dose dependent decrease for the percentage of normal embryo-larvae occurred at 24, 48 and 72 h (P < 0.001 for all times) with all the concentrations of HTO and MMS being significantly different only from the control as indicated using ANOVA. By 72 h only the lowest concentration of HTO (0.37 kBq ml−1 ) contained a small frequency of normal embryo-larvae and all the other concentrations were devoid of normality. The percentage of dead embryo-larvae in the seawater controls did not vary greatly over the three days with approximately 10% of the embryos being dead. At 72 h there was a large increase in mortality in embryo-larvae
exposed to HTO and MMS. With all the concentrations of HTO and the MMS being significantly different from the control (P < 0.001). Furthermore, the lowest concentration (i.e. 0.37 kBq ml−1 ) was also significantly different from all the other concentrations and the MMS. There was no difference in mortality between 3.7 and 37 kBq ml−1 but they were statistically different from both the 370 kBq ml−1 and MMS exposed larvae. The regression coefficient indicated a moderate dose–response relationship (R2 = 42.69%). One hundred percent mortality was exhibited at 72 h in the highest HTO concentration as well as the MMS concentration. Consequently, the 72 h LC50 for M. edulis could not be predicted. The predicted 48 h LC50 was, however, 0.27 Bq ml−1 with a received dose of 0.9 mGy.
4. Discussion The analysis of RAPD profiles in the present study showed changes in patterns due to 3 H induced DNA alterations even at the lowest HTO concentration (i.e. 0.37 kBq ml−1 ). Although it is difficult to decipher the exact mechanisms responsible for the changes in RAPD profiles following HTO exposure, it has been demonstrated that DNA damage and mutations can induce changes in RAPD patterns after exposure to genotoxic agents (Atienzar et al., 2002). For instance, damaged DNA can block or reduce the processivity of the Taq DNA polymerase leading to disappearance
214
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
of bands in the RAPD profiles. Alternatively, conformational changes in the DNA could also improve the access of the primer(s) to the binding site(s) leading to appearance of new bands in the corresponding RAPD patterns. In fact, such changes in the RAPD profiles have been correlated with other ecotoxicological parameters (i.e. growth, development and reproductive success) in aquatic invertebrates (Atienzar et al., 1999, 2001; Atienzar and Jha, 2004). Combined with robust protocols, such molecular approaches could play an increasingly important role in determining the impact of genotoxins in the aquatic environment (Jha, 2004). Despite the fact that low-LET ionising radiations delivered by external sources are suggested to be poor inducer of SCEs (Natarajan et al., 1993), in the present study a clear dose-dependent increase for SCEs was found between 3.7 and 370 kBq ml−1 of HTO compared to controls. Straume and Carsten (1993) reported a statistically significant increase for the incidence of SCEs on bone marrow cells of mice maintained on 111 kBq ml−1 of ingested HTO for 81–216 days. Since induction of SCEs is S-phase dependent phenomenon, it could be suggested that 3 H interferes either with the replication process or inhibits the actions of associated enzymes, in common with other chemical genotoxins (Jha et al., 1992, 1996, 2000a,b). Suyama et al. (1981) reported increased frequency of Cabs (i.e. chromosome bridges) when eggs of the freshwater teleost, Oryzias latipes, were treated with HTO delivering doses higher than 0.19 Gy. This is slightly higher than the dose that induced significantly elevated levels of Cabs in the embryo-larvae of M. edulis in the present study. While a dose–response relationship for SCEs, developmental abnormalities and mortality, were observed in the present study, there was a lack of clear dose-response relationships for Cabs and PRI. The reasons for this apparent lack of dose–response relationships are not clear. It could be hypothesized that only critical sites or sensitive cells are targets for radiation action, when these small-sized organisms, undergoing cellular differentiation come into contact or uptake the radionuclides in common with higher organisms (Gossner, 2001). It could also be speculated that in the very sensitive embryo-larval life stages, highly damaged cells leading to death are eliminated from analysis, affecting both the yield of aberrations and analysis of cells in different stages of division. In other words, the absence of a clear dose-
dependent increase for Cabs and PRI in the very early life stages of aquatic organisms could be related to their known capacity to undergo apoptosis, thereby efficiently eliminating cells with high frequencies of Cabs, as observed in mammalian cells (Jha et al., 1995b). In addition, it is also possible that if highly damaged cells are eliminated from the analysis, only the ‘bystander effects’ are analysed (Morgan, 2003). Furthermore, apart from unstable Cabs as observed in the present study, intake of HTO can also induce stable or balanced Cabs that could be detected several years after intake by humans (Lloyd et al., 1998). In parallel with human health risk assessment, it is therefore important to take into account such stable genetic damage and potential delayed effects (Mothersill and Seymour, 1998) to protect the environment. As well as alterations in RAPD profile, induction of SCEs and Cabs, HTO exposed embryo-larvae also experienced lower values of PRI due to a reduction in cell cycle progression probably caused by an increase in toxicity. However, as mentioned above, no marked difference for the PRI values could be found in the embryo-larvae exposed to different concentrations of HTO. With respect to this apparent lack of dose–response relationship for PRI, in addition to possible explanations provided above, it should be pointed out that although originally proposed by Lamberti et al. (1983) for application to actively proliferating and highly differentiated mammalian cells in vitro, the PRI has been found to provide an useful means to quantify the cytotoxicity in embryo-larval stages of aquatic organisms following exposure to different toxicants (Jha et al., 1996, 2000a,b; Hagger et al., 2002). However, the PRI for the embryo-larvae probably represents an average figure, based on a mixed population of highly differentiating cells with differing mitotic rates over the exposure period described (Jha et al., 1996). Therefore, even a slight perturbation in the rate of cell cycle progression, cellular differentiation and programmed cell death, for which as mentioned above these cells have a well known capacity, could lead to significant changes in the PRI values. This is, however, in contrast to our earlier studies using chemical genotoxins, where PRI values showed significant differences from controls only at the higher concentrations of test compounds (e.g. methyl methane sulphonate, benzo (a) pyrene, tributyltin, contaminated sediment samples) in the embryo-larvae of M. edulis and P.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
dumerilii (Jha et al., 2000a,b; Hagger et al., 2002). A lower plateau-value for PRI at higher concentrations of HTO in the present study therefore appears to be related to the mechanisms of action of ionising radiation, which, in contrast to chemicals, could impart their effect continuously the very moment cells come in contact with the radionuclides. In contrast to induction of Cabs and PRI values, compared to unexposed controls, the study showed a significant dose-dependent increase for mortality in HTO exposed embryo-larvae by the end of the experiment. Abbott and Mix (1979) also observed similar effects on survival in the larval stage of goose barnacles following exposure to seawater containing only 0.2 Bq ml−1 of HTO. In contrast to the sexually mature adults, the embryo-larval stages of aquatic organisms are considered to be several orders of magnitude more sensitive in terms of toxicological injury. The highest detrimental effects of radiation in both mammals and aquatic organisms have been observed in experiments when irradiation occurred immediately after fertilisation. It has also been suggested that the irradiation of cells immediately after fertilisation may also encompass the sensitive period of second mitosis (Anderson and Harrison, 1990). It is therefore not surprising that 1h old embryo showed extensive developmental abnormalities and mortality following exposure to 3 H, an internal emitter. In this context, Knowles and Greenwood (1997) also found that the development and survival of eggs to larvae of the polychaete worm, Ophryotrocha diadema, was significantly reduced when exposed to 2.24 MBq ml−1 of HTO but was not affected by similar doses of ␥radiation delivered by 137 Cs. Indeed, -rays from HTO have been shown to be more effective than ␥-rays emitted by 90 Sr, 90 Y (in solution), 60 Co and X-rays (external sources) in inducing Cabs in the blastula stage of Oryzias latipes (freshwater teleost) eggs, potentially leading to an increase in abnormality and mortality (Suyama et al., 1981), as observed in the present study. While having a global view of all the concentration or dose-dependent responses in the present study, it is interesting to observe that while significantly decreased values of normal embryo-larvae (correspondingly increased frequencies of deads) and PRI (which had a ‘plateau level’ effect at all the higher concentrations) were found at the lowest concentra-
215
tion (0.37 kBq ml−1 ) of HTO, no significant increase for SCEs and aberrant cells were found at this concentration. These discrepancies in ‘concentration–effect’ relationships for the lowest concentration of HTO appear to be odd. In fact, studies in the literature simultaneously evaluating concentration dependent effects at different levels of biological organisation (i.e. chromosomal, cellular and individual) are very scarce. From the observations made in the present study, it appears that there is not a direct, linear relationship between observed effects at lower (i.e. chromosomal and cellular) and higher (e.g. developmental abnormalities and mortality) levels of biological organisation. It is also to be remembered here that while cytogenetic effects (i.e. SCEs, Cabs and PRI) were evaluated soon after HTO exposure (one and half to two cell cycles), the developmental abnormality and survival effects were analysed at later stages (data presented for 72 h post-fertilisation period). It appears therefore that induced damage at lower levels of biological organisation, which could have knock-on effects as a function of time, could precipitate cumulatively and thus be manifested later during the developmental process at the individual level. In conclusion, our integrated study linking genetic and cellular biomarkers with embryo-larval bioassay suggests that extremely low doses of -radiation emitted by HTO can induce genetic damage in the embryolarval stages of marine mussels and that this damage potentially accumulates at cellular and individual levels. Such an approach could be adopted to early life stages of other invertebrates to establish dose–response relationships since estimation of dose received by the biota and resultant effects are important in radiological protection criteria for the natural environment (Pentreath, 1998; ERICA, 2005). Our study goes some way towards achieving this goal.
Acknowledgements This work was supported financially, mainly by the Natural Environmental Research Council (NERC), Environment Agency (EA) (GT22-1998-EA-0003) in association with Devonport Royal Dockyard (DML), Devon, UK. The RAPD work was supported by the Marie Curie Grant of the European Commission (ERB4001-GT-97-0136).
216
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217
References Abbott, D.T., Mix, M.C., 1979. Radiation effects of tritiated seawater on development of the Goose Barnacle, Pollicipes polymerus. Health Phys. 36, 283–287. Anderson, S.L., Harrison, F.L., 1990. Guidance document for prepermit bioassay testing of low level radioactive waste. United States Environmental Protection Agency, pp. 50. ASTM, 1989. Standard guide for conduction static acute toxicity tests starting with embryos of four species of saltwater bivalve molluscs. E724-89, America Society for Testing of Materials, Philadelphia. Atienzar, F.A., Jha, A.N., 2004. The random amplified polymorphic DNA (RAPD) assay to determine DNA alterations, repair and transgenerational effects in B(a)P exposed Daphnia magna. Mutat. Res. 552, 125–140. Atienzar, F.A., Conradi, M., Evenden, A.J., Jha, A.N., Depledge, M.H., 1999. Qualitative assessment of genotoxicity using random amplified polymorphic DNA: comparison of genomic template stability with key fitness parameters in Daphnia magna exposed to benzo[a]pyrene. Environ. Toxicol. Chem. 18, 2275– 2282. Atienzar, F.A., Cheung, V.V., Jha, A.N., Depledge, M.H., 2001. Fitness parameters and DNA effects are sensitive indicators of copper induced toxicity in Daphnia magna. Toxicol. Sci. 59, 241–250. Atienzar, F.A., Venier, P., Jha, A.N., Depledge, M.H., 2002. Evaluation of the random amplified polymorphic DNA (RAPD) assay for the detection of DNA damage and mutations. Mutat. Res. 521, 151–163. Bayne, B.L., 1976. The biology of mussel larvae. In: Bayne, B.L. (Ed.), Marine Mussels, their Ecology and Physiology. Cambridge University Press, Cambridge, pp. 81–119. Burgeot, T., His, E., Galgani, F., 1995. The micronucleus assay in Crassostrea gigas for the detection of seawater genotoxicity. Mutat. Res. 342, 125–140. Department of the Environment, Transport and the Regions, 2000. UK strategy for radioactive discharges, 2001–2020. Consultation document. DETR, London, pp. 1–74. Environment Agency, 2001. Proposed Decision Document on the Application made by Devonport Royal Dockyard Limited to Dispose of Radioactive Wastes from: Devonport Royal Dockyard Plymouth. Environment Agency, UK, p. 288. ERICA, 2005. Environmental risk from ionising contaminants; www.erica-project.org (Accessed January 2005). Gossner, W., 2001. Target cells for internal dosimetry. Radiat. Prot. Dosim. 105, 39–42. Hagger, J.A., Fischer, A.S., Hill, S.J., Depledge, M.H., Jha, A.N., 2002. Genotoxic, cytotoxic and ontogenetic effects of tri-nbutyltin on the marine worm, Platynereis dumerilii (Polychaeta: Nereidae). Aquat. Toxicol. 57, 243–255. Harrison, F.L., Jones, I.M., 1982. An in vivo sister chromatid exchange assay in the larvae of the mussel Mytilus edulis: response to three mutagens. Mutat. Res. 105, 235–242. His, E., Beiras, R., Seaman, M.N.L., 1999. The assessment of marine pollution—bioassays with bivalve embryos and larvae. Adv. Mar. Biol. 37, 1–178.
ICRP. Statement from the 1983 meeting of the International Commission on Radiological Protection. 1984. ICRP Publications 39, Ann. ICRP 14:1, 1–8. Jha, A.N., 2004. Genotoxicological studies in aquatic organisms: an overview. Mutat. Res. 552, 1–17. Jha, A.N., Noditi, M., Nilsson, R., Natarajan, A.T., 1992. Genotoxic effects of sodium arsenite on human cells. Mutat. Res. 284, 215–221. Jha, A.N., Hutchinson, T.H., Mackay, J.M., Elliott, B.M., Pascoe, P.L., Dixon, D.R., 1995a. The chromosomes of Platynereis dumerilii (Polychaeta: Nereidae). J. Mar. Biol. Assoc. U.K. 75, 551–562. Jha, A.N., Hande, P.M., Mullenders, L.H.F., Natarajan, A.T., 1995b. Mimosine is a potent clastogen in primary and transformed hamster fibroblasts but not in primary or transformed human lymphocytes. Mutagenesis 10, 385–391. Jha, A.N., Hutchinson, T.H., Mackay, J.M., Elliott, B.M., Dixon, D.R., 1996. Development of an in vivo genotoxicity assay using the marine worm Platynereis dumerilii (Polychaeta: Nereidae). Mutat. Res. 359, 141–150. Jha, A.N., Cheung, V.V., Foulkes, M.E., Hill, S.J., Depledge, M.H., 2000a. Detection of genotoxins in the marine environment: adoption and evaluation of an integrated approach using the embryolarval stages of the marine mussel, Mytilus edulis. Mutat. Res. 464, 213–228. Jha, A.N., Hagger, J.A., Hill, S.J., 2000b. Tributyltin induces cytogenetic damage in the early life stages of the marine mussel, Mytilus edulis. Environ. Mol. Mutag. 35, 343–350. Kirchmann, R., Koch, G., Adam, V., van Den Hoek, J., 1973. Studies on the food chain contamination by tritium. In: Moghissi, A.A., Carter, M.W. (Eds.), Tritium. Messenger graphics, Nevada. Kirkland, D.J., 1990. Basic mutagenicity tests: UKEMS recommended procedures. Cambridge University Press, pp. 1–143. Knowles, J.F., Greenwood, L.N., 1997. A comparison of the effect of long-term - and ␥-irradiation on the reproductive performance of a marine invertebrate Ophryotrocha diadema (Polychaete Dorvilleidae). J. Environ. Radioact. 34, 1–7. Lamberti, L., Bigatti, P., Ardito, G., 1983. Cell kinetics and sister chromatid exchange frequency in human lymphocytes. Mutat. Res. 120, 193–199. Lloyd, D.C., Edwards, A.A., Prosser, J.S., 1986. Chromosome aberrations induced in human lymphocytes by in vitro acute X and gamma radiation. Radiat. Prot. Dosim. 15, 83–88. Lloyd, D.C., Moquet, J.E., Oram, S., Edwards, A.A., Lucas, J.N., 1998. Accidental intake of tritiated water: a cytogenetic followup case on translocation stability and dose reconstruction. Int. J. Radiat. Biol. 73, 543–547. McCubbin, D., Leonard, K.S., Bailey, T.A., Williams, J., Tossell, P., 2001. Incorporation of organic tritium (3 H) by marine organisms and sediment in the Severn Estuary/Bristol Channel (UK). Mar. Pollut. Bull. 42, 852–863. Morgan, W.F., 2003. Is there a common mechanism underlying genome instability, bystander effects and other-targeted effects of exposure to ionising radiation? Oncogene 22, 7094–7099. Mothersill, C., Seymour, C.B., 1998. Mechanisms and implications of genomic instability and other delayed effects of ionising radiation exposure. Mutagenesis 13, 421–426.
J.A. Hagger et al. / Aquatic Toxicology 74 (2005) 205–217 Natarajan, A.T., Darroudi, F., Jha, A.N., Meijers, M., Zdzienicka, M.Z., 1993. Ionising radiation induced DNA lesions which lead to chromosomal aberrations. Mutat. Res. 299, 297–303. Pentreath, R.J., 1998. Radiological protection criteria for the natural environment. Radiat. Prot. Dosim. 75, 175–179. Sparks, T., 2000. Statistics in Ecotoxicology. John Wiley and Sons Ltd., Wiltshire, UK, 229 p. Strand, J.A., Fujihara, M.P., Burdett, R.D., Poston, T.M., 1977. Suppression of the primary immune response in Rainbow
217
Trout, Salmo gairdneri, sublethally exposed to tritiated water during embryogenesis. J. Fish. Res. Board Can. 34, 1293– 1304. Straume, T., Carsten, A.L., 1993. Tritium radiobiology and relative biological effectiveness. Health Phys. 65, 657–672. Suyama, I., Etoh, H., Maruyama, T., Kato, Y., Ichikawa, R., 1981. Effects of ionising radiation on the early development of Oryzias eggs. J. Radiat. Res. 22, 125–133.