Mitochondrial DNA mutation frequencies in experimentally irradiated compost worms, Eisenia fetida

Mutation Research 603 (2006) 56–63

Mitochondrial DNA mutation frequencies in experimentally irradiated compost worms, Eisenia fetida Craig S. Wilding a,∗ , Michael Z. Trikic a , Joanne L. Hingston b,c , David Copplestone b,c , E. Janet Tawn a b

a Genetics Department, Westlakes Research Institute, Moor Row, Cumbria CA24 3JY, UK Jones Building, School of Biological Sciences, University of Liverpool, Liverpool L69 3GS, UK c Environment Agency, Richard Fairclough House, Knutsford Road, Warrington WA4 1HG, UK

Received 18 July 2005; received in revised form 21 October 2005; accepted 27 October 2005 Available online 27 December 2005

Abstract The compost worm Eisenia fetida is routinely used in ecotoxicological studies. A standard assay to assess genetic damage in this species would be extremely valuable. Since mitochondrial DNA (mtDNA) is known to exhibit an increased mutation rate following exposure to ionising radiation we assessed the validity of a mtDNA-based assay for measuring increases in mutation rate in laboratoryirradiated compost worms. To this end the mutation frequency in the mtDNA of the compost worm E. fetida was quantified following in vivo ␥-irradiation of adult worms in three dose groups. Five adult worms exposed to 1.4 mGy/h for 55 days (total dose 1.85 Gy), five adult worms exposed to 8.5 mGy/h for 55 days (total dose 11.22 Gy) and five adult control worms were used to assess the effect of irradiation on mtDNA mutation induction. DNA samples extracted from irradiated adult worms were used in high-fidelity PCR of a 486 bp region of mtDNA spanning the ATPase 8 gene, chosen for its high spontaneous mutation rate. PCR products were cloned and sequenced to identify mutations, with 89–102 clones successfully sequenced per individual. A significant elevation in mtDNA mutation frequency (p = 0.032) was seen in worms exposed at the higher dose rate (8.5 mGy/h, total dose 11.22 Gy; mutation frequency 27.98 ± 4.85 × 10−5 mutations/bp) in comparison to controls (mutation frequency 12.68 ± 3.06 × 10−5 mutations/bp), but no elevation in mutation frequency (p = 0.764) was seen for the lower dose rate (1.4 mGy/h, total dose 1.85 Gy; mutation frequency 13.74 ± 1.29 × 10−5 mutations/bp) compared with controls. This indicates that although the technique has the potential to detect an elevation in mutation frequency, it does not have sufficient sensitivity at the doses likely to be encountered in environmental monitoring scenarios. © 2005 Elsevier B.V. All rights reserved. Keywords: Annelid; Biomarker; Compost worm; Ionising radiation; Mitochondrial DNA; Mutation

1. Introduction Biomarkers provide a direct measurement of biological response to an environmental pollutant and are

∗

Corresponding author. Tel.: +44 1946 514158; fax: +44 1946 514057. E-mail address: [email protected] (C.S. Wilding). 1383-5718/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.mrgentox.2005.10.011

capable of measuring both exposure and effect [1]. The application of genetic biomarkers of exposure to genotoxicants such as ionising radiation can provide valuable information on the health of the organism [1,2] and be utilisable as a biological dosimeter where exposures are uncertain or not measured. Quantification of DNA damage at the sequence level as a biomarker of ionising radiation exposure in biota has proven difficult, and there are no data on increased muta-

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tion frequency at the DNA sequence level in exposed invertebrates. To a large extent this is due to the dearth of sequence information available for such animals. A standard assay for measuring DNA damage in invertebrates following genotoxic exposure could prove extremely valuable. A promising candidate for the development of an assay system for measurement of genotoxic damage is the study of mutations in mitochondrial DNA (mtDNA). In addition to the nuclear genome, cells contain a small circular DNA molecule within the mitochondria. This molecule encodes 37 genes: 13 protein-coding genes, 22 transfer-RNA genes and 2 ribosomal RNA genes. The mtDNA of a wide range of species has now been sequenced and these genes are universally found in all species, although the relative orders of the constituent genes do differ between species [3]. The spontaneous mutation rate of the mtDNA is higher than that of the majority of the nuclear genome [4]. This is attributed to the absence of histones protecting the coiled DNA, the presence of reactive oxygen species within the mitochondrion and an underdeveloped mitochondrial DNA-repair repertoire in comparison with that of the nuclear genome [5]. Each mitochondrion contains multiple copies of mtDNA and each cell thousands of mitochondria. It is therefore feasible for somatic mutations of mtDNA to give rise to mutant copies that coexist with the wild-type form. Heteroplasmy, or the presence of different mtDNA copies within the same individual is indeed tolerated, with phenotypic consequences (in humans) only resulting when the frequency of the mutant copy rises above approximately 70% [6]. Since somatic mutations can arise and be tolerated by the organism without detriment, mtDNA may be able to provide an integrated measure of exposure to genotoxicants. Both ␥-irradiation [7] and X-irradiation [8] have been shown to damage mtDNA in vitro. Following in vivo exposure, an increase in mtDNA mutation rate has also been seen in humans exposed to therapeutic doses of radiation [9] and mutations have been detected in family pedigrees of individuals inhabiting naturally radioactive areas [10]. Studies have also been made of mtDNA mutations in rodents inhabiting, or enclosed in, areas contaminated by radionuclides following the Chernobyl disaster. However, although an initial study [11] on a native population of voles suggested a 100-fold increase in mtDNA mutation rate for voles inhabiting a radionuclide-contaminated area in comparison with those from a non-contaminated site, this was subsequently retracted [12] and later studies have failed to demonstrate an effect [13–15]. Similarly, no effect on

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mtDNA mutation rate was found in waterfowl inhabiting a contaminated nuclear fuel reactor-cooling pond [16]. The compost worm Eisenia fetida is routinely used in ecotoxicological assays [17]. Genetic biomarkers used in studies of Eisenia include the comet assay [18–20] and DNA-adduct analysis [21]. To develop assays of mtDNA mutation rate for this species it is necessary to obtain sufficient sequence data from which to design PCR primers for the region of interest. Limited partial gene sequences of Eisenia have been utilised for phylogenetic analyses [22–24]. However, since the complete mitochondrial sequence is available for another Clittelid annelid, the earthworm Lumbricus terrestris [25] it is possible to develop PCR assays to target any region of interest in the mtDNA utilising sequence information from this species. Here, we wished to test whether ionising radiation causes elevations in mtDNA mutations in compost worms irradiated in a laboratory setting with known dosimetry. We have analysed large numbers of mitochondrial copies from each individual animal using DNA sequencing of a section of mtDNA containing the ATPase 8 gene, two tRNAs and a partial CoIII sequence. Animals irradiated in this study were also examined for effects on reproduction, morbidity and mortality (results published elsewhere [26]) thus, as has been advocated [27,28], permitting the observed DNA damage to be related to individual health and reproduction, and hence to effects at the population level. 2. Materials and methods 2.1. Irradiation of worms Experimental animals, E. fetida were obtained from a commercial supplier (Blades Biological, Cowden, Edenbridge, Kent TN8 7DX, United Kingdom). Worms were housed in 3.5 L tanks containing rehydrated topsoil overlain with rehydrated horse manure and irradiated with a 137 Cs source [26]. Worms were irradiated at either 1.4 or 8.5 mGy/h for 55 days to give total doses of 1.85 and 11.22 Gy, respectively, with a further control tank exposed only to background irradiation. Dosimetry was confirmed using thermoluminescence dosimeters. Following exposure, worms were preserved in ethanol. 2.2. DNA extraction Genomic DNA was extracted from small (10 mm) sections of adult worms. Since PCR was used to amplify defined sections of mtDNA it was not necessary to isolate mtDNA away from the nuclear DNA. Body sections were first rehydrated in distilled water, then genomic DNA was isolated from tissue sections. Initial attempts at DNA purification using stan-

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dard proteinase K extractions followed by phenol–chloroform extraction resulted in co-purification of a PCR-inhibiting compound. Following trials, consistently clean preparations with no PCR inhibition were obtained using the Wizard® SV genomic DNA purification system (Promega). All genomic DNA extracts were standardised to 100 ng/␮l prior to use. 2.3. Amplification of mtDNA The ATPase 8 gene was chosen as the target gene due to its high evolutionary rate indicative of a higher inherent mutation rate [29]. Primers based on conserved regions of mtDNA biased to the Lumbricus terrestris sequence were used in initial amplifications. An 1186 bp section of mtDNA was then cloned and sequenced, and from this primers were designed to amplify a 486 bp section of mtDNA containing the ATPase 8 gene, tRNAtyr , tRNAgly and a section of the CoIII gene. These primers were named CoIIend (5 -TCT GGT GTT CTT CTA TGC CTC AC-3 ) (bases 2336–2358 of the Lumbricus sequence) and CoIII (5 -CTC GGA TTA CAT CTC GTC ATC A-3 ) (bases 2797–2818 of the Lumbricus sequence). The mitochondrial fragment was amplified in 50 ␮l volumes containing 1 × PCR buffer, 200 mM dNTPs, 25 pmol each primer, 2.5U PfuUltra high-fidelity DNA polymerase (Stratagene) and 100 ng genomic DNA. The use of the thermostable proofreading polymerase PfuUltra which has extremely high fidelity [30,31] minimised PCR-generated mutations. PCR products were purified using the StrataPrep® PCR purification kit (Stratagene) according to the manufacturer’s instructions. Purified PCR products were cloned into the standard PCR product cloning vector PCR-scriptTM AmpSK(+) (Stratagene) and transformed into high competency XL-10-Gold® cells. Recombinant plasmids were identified by blue–white colour selection. Presence/absence of inserts was confirmed through PCR screening of individual colonies. Colonies containing inserts were plated onto 96-well plates containing LB-amp and sent for commercial plasmid isolation and sequencing (MWG Biotech, Ebersberg, Germany). 96–103 plasmids were sequenced per individual animal. 2.4. Sequence analysis Sequences were aligned against a standard Eisenia reference sequence using ClustalX [32]. Discrepant positions were verified through manual comparison with the electropherogram. Homoplasmic mutations were identified in some animals as differences from the reference sequence in every clone. Additionally, singleton mutations were observed and these were counted and classified. Mutation frequency was calculated as number of singleton mutations/(number of clones sequenced × 441) where the length of the insert DNA without primers is 441 bp. Observed mutations were classified according to type (e.g. C–T, A–G, indel, etc.). The numbers of different mutational types were compared between treatment groups (control, 1.4 and 8 mGy/h) using Fisher’s exact test with an alpha level of p < 0.05.

2.5. Procedural error rate In a separate study [33] we have determined the procedural error rate for this assay, indicative of the number of mutations induced by the DNA polymerase used. DNA extracted from a single clone containing a 443 bp fragment of the human mtDNA HVR1 region [9] was PCR-amplified and subsequently cloned. Plasmid DNA was extracted and sequenced from 93 clones.

3. Results 3.1. Procedural error rate One mutation was detected in 93 clones generated from PCR amplification of DNA from a single clone (insert size minus primers = 405 bp) [33]. Therefore, the baseline mutation frequency of this system is extremely low (one mutation in 37,665 bp or 2.65 × 10−5 ). 3.2. Mitochondrial DNA sequence variability The sequence of the 486 bp mtDNA fragment amplified is shown in Fig. 1. This was the standard sequence in 7/15 animals. Six of the remaining eight animals differed from this sequence by the presence of a G > A transition at position 66. Two animals (both in the control group) exhibited fragment sequences in every clone with numerous differences from that shown in Fig. 1 (65/441 and 45/441 bp). These are differences more typical of inter-specific comparisons. However, E. fetida is easily recognisable due to its red/yellow-striped pattern and the worms were provided by a reputable supplier of these animals. We therefore believe that we have not analysed DNA from a species other than E. fetida and that this indicates substantial intra-specific variability. An alternative possibility is that these sequences represent nuclear pseudogenes [34]. We do not believe this to be the case since all copies within these two animals contained these sequences and the pattern of variability between these sequences and the alternative E. fetida sequences was typical of functional copies (data not shown). The measurement of mutation frequency is not hampered or devalued by this since we are still able to count singleton mutations within these groups of clones. 3.3. Mutation distribution by treatment A total of 1416 plasmids (≡mitochondrial copies), representing 624,456 bp of mitochondrial sequence were successfully sequenced and 113 mutations detected (following rejection of sequences with poor-quality

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Fig. 1. Alignment of the 486 bp amplicon sequence from Eisenia with sequence from Lumbricus terrestris. There are four indel events between the sequences. Where the indel event shows missing sequence in the Eisenia sequence this base is omitted from the numbering to allow for consecutive numbering of the Eisenia sequence. Indel events with missing sequence in the Lumbricus sequence are not omitted. Thus, Lumbricus has a CC deletion at bases 131–132 and an AC deletion at bases 284–285 of the Eisenia sequence and an inserted C between bases 228–229 and an inserted T between bases 255–256 of the Eisenia sequence.

Table 1 Mutation frequency in compost worms following sequencing of the 486 bp fragment of mtDNA from five control compost worms (subjected to background irradiation), five compost worms exposed to 1.4 mGy/h for 55 days (total dose 1.85 Gy) and five compost worms exposed to 8.5 mGy/h for 55 days (total dose 11.22 Gy) Treatment group

Sample

Control

1 2 3 4 5

Average Total 1.4 mGy/h

1 2 3 4 5

Average Total 8.5 mGy/h

Average Total

1 2 3 4 5

Number of sequences

Total bp examined

Number of mutations

Mutation frequency (×10−5 )

98 95 89 91 91

43218 41895 39249 40131 40131

4 7 2 9 4

92.8 464

40924.8 204624

5.2 26

44982 41895 41454 41895 41895

4 6 6 6 7

96.2 481

42424.2 212121

5.8 29

13.74 ± 1.29 13.67

98 98 89 91 95

43218 43218 39249 40131 41895

5 16 (14)* 13 (11)* 9 15

11.56 37.02 (32.39) 33.12 (28.02) 22.42 35.8

94.2 471

41542.2 207711

11.6 (10.8) 58 (54)*

27.98 ± 4.85 (26.04 ± 4.26) 27.92 (25.99)

102 95 94 95 95

9.25 16.7 5.09 22.42 9.96 12.68 ± 3.06 12.7 8.89 14.32 14.47 14.32 16.7

Six animals (2, 4, and 5 from the 1.4 mGy/h group and 1–3 from the 8.5 mGy/h group) exhibited homoplasmic base changes in comparison with Fig. 1 (G66A). Two animals (4 and 5 from the control group) exhibited numerous homoplasmic base changes in comparison with Fig. 1 (see text). * Individuals with two identical mutations in two different sequences. Numbers in parentheses discount these heteroplasmic mutations from calculations.

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Table 2 Mutation frequency in compost worms following sequencing of the 486 bp fragment of mtDNA from five control compost worms (subjected to background irradiation: BG1–BG5), five compost worms exposed to 1.85 Gy at 1.4 mGy/h (1.85-1–1.85-5) and five compost worms exposed to 11.22 Gy at 8.5 mGy/h (11.22-1–11.22-5) Sample

Mutn−

Gene

Posn−

A acid

Sample

Mutn−

Gene

Posn−

A acid

Sample

Mutn−

Gene

Posn−

A acid

BG1

C62A C111A C311A C452A

ATPase 8 ATPase 8 CoIII CoIII

3 1 3 3

F-L H-N R-R I-M

1.85-1

C176A C259A C311A G411A

ATPase 8 tRNAgly CoIII CoIII

3

S-S

11.22-1

P-T S-Y

R-R G-S

ATPase 8 ATPase 8 tRNAtyr tRNAgly CoIII

1 2

3 1

C30A C175A G237T G254T G337A

2

S-N

G64T C121A C152A T255C G310T C340A A409G

ATPase 8 ATPase 8 ATPase 8 tRNAgly CoIII CoIII CoIII

2 2 3

W-L S-Y T-T

1.85-2

E-D W-R

R-L P-Q H-R

ATPase 8 ATPase 8 tRNAtyr tRNAgly tRNAgly CoIII

3 1

2 2 2

G158T T165C G187T G241T C298A† G407A†

3

G-G

ATPase 8 ATPase 8 tRNAgly CoIII CoIII CoIII

1 3

L-M S-S

G64T C76A C176A C180A† C312A† C323A C346T C353A G397T

C114A G125T G248T C340A C356A G397T

S-STOP T-N F-L S-STOP S-STOP N-N T-N

tRNAgly CoIII

2 3 2

P-Q S-S W-L

C140A C231A delA292 T335A C371A C452A

ATPase 8 tRNAtyr tRNAgly CoIII CoIII CoIII

3

N-K

ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 Atpase 8 tRNAtyr tRNAtyr tRNAgly CoIII CoIII CoIII CoIII CoIII CoIII

2 2 3 2 2 3 2

C285A G305T

C49A C85A C119A§ C124A* C124A* ‡ C140T C151A† G224T C230A§ G241T C308A† G330T C340A C345A§ G364T C380A‡

3 1 2 1 1 3

I-M E-STOP P-Q P-T A-S A-A

3 3 3

Y-STOP F-L I-M

C88A C121A C308T G366T

C88A C142A G156T† G223T† C259A G286T G421T†

ATPase 8 ATPase 8 ATPase 8 tRNAtyr tRNAgly tRNAgly CoIII

2 2 1

S-Y A-K E-STOP

W-L N-K F-L F-L G-G Q-L P-P

C-F

ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 tRNAtyr tRNAtyr tRNAgly tRNAgly CoIII CoIII

2 3 3 3 3 2 3

2

G97T T107G† C119A* C119A* G128T A133T C164A C206A C230T A252G C285A† C312T G411T

1 1

Q-STOP G-W

G172T A182G† G187T G310A A362T C373A C408A† G432A C437A

ATPase 8 tRNAtry tRNAtyr CoIII CoIII CoIII CoIII CoIII CoIII

2

W-L

2 3 2 1 1 3

R-H M-I T-N H-N A-T A-A

G64A C91A G97T ClllA† C132A C142A G208T A264G C308T T347A T357C C358A† C379A C391A C408A

ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 ATPase 8 tRNAtyr tRNAgly CoIII CoIII CoIII CoIII CoIII CoIII CoIII

2 2 2 1 3 2

W-STOP T-N W-L H-N V-V T-K

3 3 1 2 2 2 1

I-I P-P S-P S-Y A-D A-D H-N

BG2

1.85-3 BG3

BG4

BG6

3

M-I

ATPase 8 ATPase 8 ATPase 8 tRNAtyr CoIII CoIII CoIII CoIII CoIII

2 2 3

W-L S-Y S-S

1 3 2 3 2

Q-K H-Q P-L T-T W-L

ATPase 8 ATPase 8 CoIII CoIII

2 2 3 1

S-Y S-Y I-I A-S

1.85-4

1.85-5

11.22-2

11.22-3

11.22-4

11.22-5

Mutation positions are relative to Fig. 1. Posn− : codon position and A. acid: amino acid change (where relevant). †, ‡ or §: substitutions found in same clone. * Identical mutations encountered in separate clones. (See text for details).

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sequencing traces). The number of mutations detected in each treatment group is shown in Table 1. Between two and nine mutations were detected in the control group, four–seven mutations in the 1.4 mGy/h dose-rate group and 5–16 in the 8.5 mGy/h dose-rate group. The mutation frequency in the three categories is then 12.68 ± 3.06 × 10−5 , 13.74 ± 1.29 × 10−5 and 27.98 ± 4.85 × 10−5 mutations/bp for the background, 1.4 and 8.5 mGy/h dose-rate categories, respectively. The mutation frequency in the 8.5 mGy/h dose-rate group differs significantly from both the control and the 1.4 mGy/h groups (p = 0.032 and 0.036, respectively), but the mutation frequency in the 1.4 mGy/h group is not significantly different from that in the controls (p = 0.764). Two samples from the 8.5 mGy/h dose group (samples 2 and 3) revealed two independent clones with an identical mutation (C124A and C119A, respectively; Table 2). These may represent independent somatic mutations classifiable as two separate singleton mutations, or they may be due to heteroplasmy within those individuals and therefore not induced by exposure to ionising radiation. If the two instances where identical mutations are encountered in two independent colonies are classed as heteroplasmy and not singleton mutations and removed from the analysis, then the 8.5 mGy/h mutation frequency is reduced to 26.04 ± 4.26 × 10−5 mutations/bp. This remains significantly different from both the control and 1.4 mGy/h groups (p = 0.038 and 0.040). 3.4. Mutation distribution by gene Mutations were distributed evenly across the genes within the amplicon, with 37% being located within ATPase 8 (representing 35% of the sequence), 23% within the two tRNA genes (28% of the sequence) and 40% within CoIII (37% of the sequence). The posi-

Fig. 2. Mutational spectra in experimental animals: all animals (black), control animals (white), animals exposed at 1.4 mGy/h (grey) and animals exposed at 8.5 mGy/h (diagonal hatching). Mutations refer to changes from the wild-type state to the mutated type, e.g. C–A is a C to A transversion. DEL: deletion.

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tion and nature of the mutations encountered is given in Table 2 and Fig. 2. No significant differences were found between the three treatments with respect to the spectrum of mutations. 4. Discussion This study demonstrates that tissues of compost worms E. fetida exhibit an increase in the mutation frequency of mitochondrial DNA (mtDNA) following delivery of a high dose of radiation (11.22 Gy delivered at 8 mGy/h), thus validating the use of this technique for examining mutation induction. This dose and dose rate are substantially higher than would be encountered in most environmental settings. No increase in mtDNA mutation frequency is seen when the animals were irradiated at 1.4 mGy/h with a total dose of 1.85 Gy. In a parallel study, no measurable effects on morbidity, mortality, reproduction (number of offspring or number of cocoons) or histopathology were found for either dose regime [26]. Therefore, although the study of mtDNA is not sensitive enough to detect an elevation in mutation frequency after an exposure of 1.85 Gy at 1.4 mGy/h, the fact that an increase in mtDNA mutation frequency is seen following delivery of a dose of 11.22 Gy at a higher dose rate – which causes no measurable detrimental consequences to either the individual or the population – indicates that the technique can provide a biomarker of population health among compost worms; if mtDNA mutation frequency is measured in environmental samples of worms and no increase in mutation frequency is seen there are unlikely to be measurable effects on any parameters of ecological or biological significance. In studies of laboratory strains of radiosensitive mice experimentally enclosed at Chernobyl, no increase in mtDNA mutation rate of the cytochrome b gene was seen following a chronic exposure totalling 1.2–1.6 Gy [15]. Taken together, these studies contribute to mounting evidence that the technique is not as sensitive as initially anticipated [13–15] and therefore is of little use as an environmental biomarker of low-dose radiation exposure. Studies of human populations exposed to occupational and environmental low-dose radiation indicate that stable chromosome translocation frequencies can provide an integrated measure of exposure in the case of lifetime exposures of 0.5 Gy and above [35] and it has recently been suggested that the technique be extended to additional species inhabiting radionuclide-contaminated environments [28]. Since E. fetida does have a karyotype that is amenable to cytogenetic analysis [36] the use of

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this technique may be more appropriate for monitoring low-dose exposure. As E. fetida is an animal widely utilised in genotoxicology [17], development of such an approach may be justified. In this study, the majority of substitutions in all three groups are transversions. The reason for this is unclear but may be related to codon usage. Interestingly, these mutations were distributed evenly across the genes within the sequenced amplicon (ATPase 8, tRNAtyr , tRNAgly and a section of the CoIII gene). The ATPase 8 gene was targeted for its inherently high mutation rate, but this seems to have been unnecessary since all genes within the amplified region have accumulated mutations at the same rate. If future studies of other species are to be undertaken this is of relevance since it means that genes amplifiable with universal primers [37] situated in conserved regions of the mtDNA (and therefore potentially with lower mutation rates) can be utilised, such that primers for rapidly evolving regions do not need to be designed for individual species. Acknowledgements Irradiation was undertaken at the CEFAS laboratory, Lowestoft and was funded by the Environment Agency. This work was funded by British Nuclear Fuels Ltd. CW would like to thank Dr. Ken Halanych of the Life Sciences Department, Auburn University, USA for helpful discussion on annelid mitochondrial DNA sequence. References [1] D. Copplestone, S. Bielby, S.R. Jones, D. Patton, P. Daniel, I. Gize, Impact Assessment of Ionising Radiation on Wildlife, Environment Agency R&D Publication 128, 2001. [2] E.J. Tawn, Monitoring for environmental mutagenesis in wild animals-lessons from human studies, J. Radiol. Prot. 19 (1999) 333–338. [3] J.L. Boore, Animal mitochondrial genomes, Nucleic Acids Res. 27 (1999) 1767–1780. [4] C. Richter, J.W. Park, B.N. Ames, Normal oxidative damage to mitochondrial and nuclear DNA is extensive, Proc. Natl. Acad. Sci. U.S.A. 85 (1988) 6465–6467. [5] L.A. Marcelino, W.G. Thilly, Mitochondrial mutagenesis in human cells and tissues, Mutat. Res. 434 (1999) 177–203. [6] P.F. Chinnery, Modulating heteroplasmy, Trends Genet. 18 (2002) 173–176. [7] A. May, V.A. Bohr, Gene-specific repair of gamma-ray-induced DNA strand breaks in colon cancer cells: no coupling to transcription and no removal from the mitochondrial genome, Biochem. Biophys. Res. Commun. 269 (2000) 433–437. [8] G. Singh, W.W. Hauswirth, W.E. Ross, A.H. Neims, A method for assessing damage to mitochondrial DNA caused by radiation and epichlorohydrin, Mol. Pharmacol. 27 (1985) 167–170. [9] T.M. Wardell, E. Ferguson, P.F. Chinnery, G.M. Borthwick, R.W. Taylor, G. Jackson, A. Craft, R.N. Lightowlers, N. Howell, D.M.

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