Ecotoxicology DOI 10.1007/s10646-015-1432-x

Zebrafish genome instability after exposure to model genotoxicants Maja Sˇrut • Anamaria Sˇtambuk • Jean-Paul Bourdineaud Go¨ran I. V. Klobucˇar

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Accepted: 13 February 2015 Ó Springer Science+Business Media New York 2015

Abstract Sublethal exposure to environmental genotoxicants may impact genome integrity in affected organisms. It is therefore necessary to develop tools to measure the extent and longevity of genotoxicant-induced DNA damage, and choose appropriate model organisms for biomonitoring. To this end, markers of DNA damage were measured in zebrafish larvae and adults following exposure to model genotoxicants (benzo[a]pyrene and ethyl methanesulfonate). Specifically, we assessed primary DNA damage and the existence of potentially persistent genomic alterations through application of the comet assay, quantitative random amplified polymorphic DNA (qRAPD) and amplified fragment length polymorphism (AFLP) assays. Furthermore, expression of genes involved in DNA repair, oxidative stress response and xenobiotic metabolism was evaluated as well. Additionally, the AFLP method was applied to adult specimens 1 year after larval exposure to the genotoxicants to evaluate the longevity of the observed DNA alterations. Large numbers of DNA alterations were detected in larval DNA using the comet assay, qRAPD and AFLP, demonstrating that zebrafish larvae are a sensitive model for revealing genotoxic effects. Furthermore, some of these genomic alterations persisted into adulthood, Electronic supplementary material The online version of this article (doi:10.1007/s10646-015-1432-x) contains supplementary material, which is available to authorized users. M. Sˇrut (&)  A. Sˇtambuk  G. I. V. Klobucˇar Department of Zoology, Faculty of Science, University of Zagreb, Rooseveltov trg 6, 10000 Zagreb, Croatia e-mail: [email protected] J.-P. Bourdineaud Arcachon Marine Station, CNRS, UMR EPOC 5805, University of Bordeaux, Place du Docteur Peyneau, 33120 Arcachon, France

indicating the formation of stable genomic modifications. qRAPD and AFLP methods proved to be highly sensitive to genotoxic effects, even in cases when the comet assay indicated a lack of significant damage. These results thus support the use of zebrafish larvae as a sensitive model for monitoring the impact of genotoxic insult and give evidence of the longevity of genomic modifications induced by genotoxic agents. Keywords Danio rerio  Comet assay  Quantitative RAPD  AFLP  DNA repair genes

Introduction Environmental pollution results in continuous exposure to a range of toxicants that may directly or indirectly induce DNA damage, with long-term consequences for the health and survival of organisms. To evaluate DNA damage many studies employ the comet assay (Collins 2004). However, this assay typically measures DNA lesions that can be easily repaired and is frequently used for monitoring of DNA repair processes (Azqueta et al. 2014). Therefore, this assay may have limited value in the evaluation of longterm consequences of exposure to environmental genotoxicants. As a result, a more robust approach would be to use a range of methodologies to characterize pollution-induced DNA damage. In particular, these methods could include RAPD (random amplified polymorphic DNA) and AFLP (amplified fragment length polymorphism) markers, which both screen for changes in DNA polymorphisms and which have been previously used to evaluate genomic instability. RAPD analysis has been used to determine the genotoxicity of various chemical and physical agents in vitro

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and in vivo in various systems (Atienzar et al. 2000; Castan˜o and Becerril 2004; Jin et al. 2009; C¸ulcu et al. 2010), including zebrafish (Zhiyi and Haowen 2004; Rocco et al. 2010; 2012). In response to criticism of this method relating to lack of reproducibility and the presence of spurious amplification products in negative control reactions (Atienzar and Jha 2006), a novel version of this methodology called quantitative RAPD (qRAPD), has recently been proposed. qRAPD employs quantitative PCR to analyse the hybridization efficiency of selected RAPD primers (Cambier et al. 2010). The method has been applied to zebrafish exposed to different stressors, including single chemicals and environmental samples (Cambier et al. 2010; Orieux et al. 2011; Geffroy et al. 2012; Lerebours et al. 2013), and was able to detect genotoxicity caused by these agents, even in cases when the comet assay did not reveal any genotoxic effects (Orieux et al. 2011). AFLP has high power in revealing genomic polymorphisms and it has been used to evaluate the impact of toxins in both plant and animal models. For instance, genotoxic effects of potassium dichromate, chromium and volatile organic contaminants have been recorded in algae Pseudokirchneriella subcapitata (Labra et al. 2007; 2010). Brassica napus and Trifolium repens proved to be sensitive models for evaluating genotoxic effects of air pollution, chemical exposure and salt stress by AFLP markers (Labra et al. 2004; Aina et al. 2006; Piraino et al. 2006; Lu et al. 2007; Zeng et al. 2010). Furthermore, AFLP was applied on mussel (Mytilus galloprovincialis) in order to assess the influence of anionic surfactants and on boar (Sus scrofa) and black sea bream (Acanthopagrus schlegeli) for evaluation of sperm genetic integrity following cryopreservation (Fraser et al. 2008; Hsu et al. 2008; Liu et al. 2010; Thurston et al. 2002). Additionally, AFLP was successfully applied to the RTG-2 and PAC2 cell lines and detected genomic alterations caused by benzo[a]pyrene (B[a]P) and ethyl methanesulfonate (EMS), even in cases where the comet assay indicated complete repair (Sˇrut et al. 2013, 2015). However, its sensitivity has yet to be compared with the comet assay in vivo. The consequences of DNA damage are diverse and usually adverse. Acute effects include changes in DNA metabolism that may trigger cell cycle arrest or cell death, whereas long-term effects include irreversible mutations that can contribute to cancer development (Hoeijmakers 2001). DNA repair reduces the frequency of genotoxininduced mutations, although incorrect repair can also generate new mutational events. Evaluation of the expression levels of DNA repair genes can aid in understanding the occurrence of DNA alterations and their fixation in the genome, and therefore complements the tools used for the assessment of genotoxic events. In this study expression of several genes belonging to different

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pathways of DNA repair was monitored. Nucleotide excision repair (NER) genes xpa, xpc and hr23b are involved in recognition and verification of DNA damage and initiation of open complex formation prior to the excision of damaged DNA (Notch et al. 2007). Base excision repair (BER) gene apex encodes for AP endonuclease which generates a single strand break by cleaving the phosphodiester backbone 50 to the AP site (Powell et al. 2005). Homologous recombination (HR) gene rad51 is involved in formation of a nucleoprotein filament on single stranded DNA regions. Furthermore, it catalyses the search for homologous sequences, strand pairing and strand exchange (Helleday 2003). Mismatch repair (MMR) genes msh2 and msh6 encode for proteins which recognise single base–base mispairs and insertion/deletion loops of 1–2 nucleotides (Hsu et al. 2010). In addition to evaluating various forms of acute DNA damage following genotoxic insult, predicting the persistence and longevity of the detected damage is of paramount importance. This issue has not been well studied thus far, as most ecotoxicological studies have employed crosssectional as opposed to longitudinal designs. Nevertheless, a number of studies have assessed DNA damage several months after the end of the exposure period. For instance, EMS-induced alkylated damage in mussels was monitored with the comet assay more than 2 months after the end of exposure (Sˇtambuk et al. 2008), while irradiation-induced micronuclei were studied in cultured human cells and mice up to 3 months after exposure (Hande et al. 1996; Ramı´rez et al. 1999). If persistent alterations occur in gametic cells they could lead to transgenerational effects, which have been described in both invertebrates and vertebrates (Atienzar and Jha 2004; Brevik et al. 2012). Zebrafish (Danio rerio) is commonly used in toxicology research, including genetic toxicology investigations using the comet assay and RAPD analysis (Jarvis and Knowles 2003; Boettcher et al. 2011; Ha¨feli et al. 2011; Osterauer et al. 2011). Zebrafish embryos and larvae are particularly valuable in toxicology due to their ease of use, small size, and the small volumes of chemicals and media required for exposure. Larvae have also been found to be more sensitive to toxicants than adults, although further studies are necessary to adequately explore the larval window of vulnerability as well as to define the long-term effects of toxicant exposure (Oliveira et al. 2009; Domingues et al. 2010). To test the hypothesis that zebrafish larvae are more sensitive to genotoxins than adults, and to explore the longterm persistence of DNA damage, we exposed zebrafish embryos and adults to the genotoxicants B[a]P and EMS. Primary DNA damage was measured with the comet assay, while the existence of potentially more permanent DNA alterations was assessed via qRAPD and AFLP analyses. The long-term persistence of the DNA damage was

Zebrafish genome instability

monitored in 1-year-old adult zebrafish that had been exposed during embryonic and early larval stages. Furthermore, to understand the mechanism underlying the development of mutations and their fixation in the genome in more detail, the expression of several genes involved in various DNA repair pathways was evaluated as well, along with the expression of oxidative stress response and xenobiotic metabolism genes.

Materials and methods Exposure and sampling of adult zebrafish The zebrafish used in this study were obtained through full sib-pair mating of the WIK strain through ten generations to obtain genetically homogenous stock. Prior to exposure, 33–43 adult zebrafish (body weight: 0.47 ± 0.09 g; length: 3.76 ± 0.21 cm; 1:2 ratio between females and males) were placed in each of the 5 glass aquaria and acclimatized for 2 weeks in 24 L of activated carbon-filtered, aerated tap water, conditioned by letting it set (temperature: 26 ± 2 °C, pH 7.0–7.5, alkalinity: 95–140 mg/L CaCO3, carbonate hardness: 5.3–7.8 °dH). During the acclimatization period, fish health was evaluated by observing their physical appearance and behaviour. Individuals demonstrating unusual behaviour such as erratic swimming, prolonged resting on the tank bottom or floating at the surface were immediately discarded from the aquaria. During acclimatization, four fish were removed from the tanks. After acclimatization, the zebrafish were exposed to nominal concentrations of the model genotoxicants: B[a]P (SigmaAldrich, USA) (33 and 41 zebrafish were exposed to 0.1 and 1 lM B[a]P, respectively) and EMS (Sigma-Aldrich, USA) (33 and 41 zebrafish were exposed to 0.5 and 1 mM EMS, respectively). The B[a]P solution was prepared in dimethyl sulfoxide (DMSO, Kemika, Croatia). The maximal DMSO concentration in the assay was 0.005 %. Zebrafish exposed to filtered tap water were used as a control (42 zebrafish). Control exposure with DMSO was not performed because previous studies have demonstrated that DMSO concentrations up to 1 % have no effects on DNA integrity in in vitro and in vivo models, as assessed by the comet assay (Kosmehl et al. 2007, 2008; Sˇrut et al. 2011, 2013). During the experiment, the fish were fed once a day with dry feed (SAK 55, Exot Hobby s.r.o., Czech Republic). Exposure was carried out for 6 days, and during that period, 1/3 (8 L) of the water containing the toxicant was renewed daily. After the exposure period, eight zebrafish from each tank were sampled for the comet assay and gene expression analysis. Fish were euthanized by incubation in ice water (Cambier et al. 2010; Orieux et al. 2011; Geffroy et al. 2012). Blood samples from eight individuals were

obtained via tail clipping and withdrawing approximately 1 lL of blood using heparinized tips. Blood samples were subsequently mixed with 100 lL of PBS (phosphate-buffered saline, 150 mM NaCl, 6 mM Na2HPO4, 4 mM KH2PO4, Kemika, Croatia). Among these eight individuals, the gills of four male individuals were dissected, placed in 500 lL of RNA later stabilization reagent (Qiagen, Germany) to prevent RNA degradation, frozen at -80 °C and subsequently used for the gene expression analysis. The remainder of the exposed zebrafish were transferred to clean water and left for an additional 6-day recovery period. After the recovery period, eight zebrafish per group were sampled for the comet assay and for gene expression analysis, as indicated above, and an additional 10 zebrafish per group were frozen at -80 °C and used for DNA isolation and genetic variation analysis. Exposure and sampling of zebrafish embryos/larvae Approximately 8-month-old adult zebrafish of both sexes were placed in three 10 L aquaria, each containing four males and two females for spawning. The fish were maintained under a 12 h day/12 h night light regime and were fed daily with dry feed. The bottom of each aquarium was covered with small glass balls to prevent egg cannibalism. Fertilized eggs were collected each morning 2–3 h after fertilization, separated from the unfertilized and damaged eggs and placed in 24-well plates. Two eggs per well were placed in 2 mL of toxicant solution prepared with previously aerated solution according to ISO standards (ISO 1996). Originally, embryos were exposed to the same toxicant concentrations as the ones used for the adult zebrafish, but due to the high mortality observed after the B[a]P treatment, the concentrations were decreased to 1 and 10 nM. The B[a]P solution was prepared in DMSO. The maximal DMSO concentration in the assay was 0.0002 %. The EMS concentrations were the same as those used for adult zebrafish exposure (nominal concentrations of 0.5 and 1 mM). Eggs exposed to aerated ISO solution (ISO 1996) were used as a control. Exposure was carried out for 6 days. For the first 3 days, the embryos were exposed in 24-well plastic plates (Nunc A/S, Denmark), and 1/3 of the toxicant solution was exchanged daily. After 3 days, when the larvae hatched, they were transferred to glass aquaria containing 1.5 L of toxicant solution for 3 additional days, and during that period, 1/3 of the toxicant solution was exchanged daily. A total of 20–40 larvae were exposed in aquaria for 6 days in the exposure experiment and 35–60 for the exposure and recovery experiment. For the AFLP and qRAPD analyses, all of the larvae used were offspring from the same parents (two females and four males). After the 6-day exposure period, eight larvae per group were collected for the comet assay, and four larvae

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per group were placed in microcentrifuge tubes containing 100 lL of RNA Later stabilization reagent (Qiagen, Germany) to prevent RNA degradation, then frozen at -80 °C and subsequently used for the gene expression analysis. The remainder of the exposed larvae were transferred to 1.5 L of clean aerated tap water and left for an additional 6-day recovery period. After the recovery period, larvae for the comet assay and gene expression analysis were sampled as described above, and an additional 10 larvae per group were frozen at -80 °C and subjected to genetic variation analysis. To monitor the persistence of DNA alterations, several larvae exposed to 0.01 lM B[a]P were allowed to grow for 1 year, and after that period, 10 individuals were sacrificed and used for AFLP analysis. To obtain the cell suspension for the comet assay, larvae were placed in 1.5 mL Eppendorf tubes and dilacerated with a Potter–Elvehjem tissue homogenizer (Braun Biotech, Sartorius, Goettingen, Germany) in 500 lL of PBS. The homogenates obtained in this manner were filtered through a 70-lm sieve, centrifuged at 2009g for 10 min at 4 °C, resuspended in 400 lL of PBS, centrifuged again at 1809g for 10 min at 4 °C and finally resuspended in 60 lL of PBS. The comet assay The alkaline comet assay (single cell gel electrophoresis assay) was performed according to the basic procedure of Singh et al. (1988), with slight modifications according to Sˇrut et al. (2011). For adult zebrafish, 5 lL of a blood cell suspension was mixed with 95 lL of 0.5 % low melting point agarose (LMP, Sigma-Aldrich, USA) and transferred to microscope slides precoated with 1 % normal melting point agarose (NMP, Sigma-Aldrich, USA), whereas for the zebrafish larvae, 50 lL of a cell suspension was mixed with 50 lL of 0.5 % LMP agarose and transferred to microscope slides precoated with 1 % NMP. After solidification for 2.5 min in a freezer, a third layer consisting of 80 lL of 0.5 % LMP agarose was added (approximately 40 °C) and left to solidify as described above. The cells were lysed in freshly made lysis solution (2.5 M NaCl, 100 mM EDTA, 10 mM Tris–HCl, 10 % DMSO, 1 % Triton X-100 (all chemicals are from Kemika, Croatia, except Triton X-100 which is from Sigma-Aldrich, USA), pH 10) for 1 h at 4 °C. After rinsing with redistilled water, the slides were placed in a horizontal gel box, covered with the cold alkaline buffer (0.3 M NaOH, 1 mM EDTA, Kemika, Croatia) pH [13) and left for 20 min. Electrophoresis was performed in the same buffer at 25 V (0.83 V/cm) at 300 mA for 20 min at 4 °C. After electrophoresis, the slides were neutralized in cold neutralization buffer (0.4 M Tris–HCl, pH 7.5) for 2 9 5 min, then fixed in methanol:acetic acid (Kemika, Croatia) (3:1) for

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5 min and stored in the dark at room temperature. Prior to examination, the slides were rehydrated, stained with 10 lg/mL ethidium bromide (Sigma-Aldrich, USA) and examined using a Zeiss Axioplan epifluorescence microscope. The extent of DNA migration was determined as the percentage of DNA in the tail (% tDNA) using the Komet 5 image analysis system (Kinetic Imaging Ltd., UK). DNA extraction DNA from adult zebrafish was extracted from the whole organism according to the protocol described by Miller et al. (1988), with slight modifications. The fish were weighed, and their whole tissue was sliced with a scalpel blade and digested overnight at 50 °C in DNA extraction buffer (10 mL per g of tissue) and 200 lg/mL of proteinase K (Sigma-Aldrich, USA). The DNA extraction buffer consisted of 10 mM Tris (pH 8), 100 mM EDTA (pH 8), 0.1 M NaCl and 0.5 % SDS (Sigma-Aldrich, USA). Following overnight incubation, 100 lg/mL RNase A solution (Qiagen, Germany) was added to the mixture, followed by incubation for 2 h at 37 °C. After digestion was complete, 4 mL of the homogenate was mixed with 1 mL of saturated NaCl (approximately 6 M), and the mixture was shaken vigorously for 15 s and centrifuged at 40,0009g for 15 min. The precipitated protein pellet was left at the bottom of the tube, and the supernatant containing the DNA was transferred to a 15 mL polypropylene tube. Exactly 2 volumes of room temperature absolute ethanol were added, and the tubes were inverted several times until the DNA precipitated. The precipitated DNA strands were removed with a plastic spatula and transferred to a 1.5 mL microcentrifuge tube containing 1 mL of 70 % ethanol. Tubes were then shaken for several seconds and left for 30 min at room temperature. Ethanol was subsequently removed, and the tubes were left open for several minutes to allow for complete ethanol evaporation. DNA was dissolved in 500 lL of TE buffer (10 mM Tris–HCl, 0.2 mM EDTA), facilitated by shaking and incubation for 1 h at 40 °C. DNA from zebrafish larvae was extracted using a QIAmp DNA Mini Kit (Qiagen, Germany) according to the manufacturer’s recommendations. The DNA was quantified by measuring the absorbance at 260 nm using a Lumat spectrophotometer (Berthold, Germany). The ratio of the DNA absorbance at 260 nm to that at 280 nm (A260/A280 ratio) fell within the acceptable range of values that indicate a pure preparation. qRAPD qRAPD was performed according to Cambier et al. (2010) using a Lightcycler automate (Roche, Switzerland) and

Zebrafish genome instability

LightCycler FastStart DNA Master SYBR Green I PCR mix (Roche, Switzerland). Each reaction had a total volume of 20 lL, containing 16 ng of genomic DNA, 2 lL of primers (6 lM) and 16 lL of PCR mix. After 10 min at 95 °C, DNA fragments were amplified through 50 cycles with the following conditions: denaturation for 5 s at 95 °C, annealing for 5 s at 50 °C and extension for 30 s at 72 °C. After this thermal program, dissociation curves were obtained by following the decrease in the SyberGreen fluorescence level during gradual heating of the PCR products from 60 to 95 °C. The hybridization efficiency of two qRAPD primers (OPB7 and OPB11) was monitored. These primers were chosen based on their previous successful use in studies examining responses in zebrafish to various genotoxic stressors (Cambier et al. 2010; Orieux et al. 2011; Geffroy et al. 2012; Lerebours et al. 2013). For each treatment condition, ten DNA samples were analysed. To quantify the number of hybridization sites per qRAPD primer, reference PCR assays were also performed using b-actin 1 gene primers. The sequences for the qRAPD primers were adopted from Cambier et al. (2010), whereas those sequences for the b-actin gene originated from Geffroy et al. (2012). The primer sequences are listed in Online Resource 1 (Table S1). The melting temperature curve analysis was conducted using Light Cycler Software 3.5 (Roche, Switzerland). For every qRAPD-PCR assay, the melting temperature (Tm) peaks were obtained for 12-14 different temperature intervals ranging from 69 to 90 °C. The frequency of occurrence of melting peaks for each interval was compared between the control and treated fish. AFLP The AFLP methodology was based on the principles originally described by Vos et al. (1995), with slight modifications. For each treatment condition, ten DNA samples were assessed. Restriction digestion and adapter ligation were performed simultaneously on genomic DNA at room temperature overnight in a total volume of 11 lL. Five units of EcoRI (Thermo Scientific, USA) and 1 U of MseI (Thermo Scientific, USA) were used for the digestion of 150 ng DNA. One unit of T4 DNA ligase (Thermo Scientific, USA) was used for the ligation of 5 pmol EcoRI and 50 pmol MseI double-stranded nucleotide adapters. Preselective and selective amplifications were performed with a MyCycler thermal cycler (Bio-Rad, USA). The primer pair adapters, preselective primers and primers used for selective amplification were adopted from Bagley et al. (2001) and synthesized by Sigma Aldrich, USA. Primer sequences are presented in Online Resource 1 (Table S1). The restriction-ligation mixture was diluted with nuclease-

free water (Qiagen, Germany) to obtain a concentration of 2.5 ng/lL of DNA. This solution was used as a template in the preamplification reaction with the DNA primers EP1 and MP1, which are complementary to the cores of the EcoRI and MseI adapters, respectively. The 20 lL preamplification mixture contained 4 lL of the diluted restriction-ligation mixture, 0.2 mM of a mixture of all dNTPs (Thermo Scientific, USA), 2 mM MgCl2, 0.6 lM of both EP1 and MP1 primers, 19 Taq DNA polymerase buffer and 0.5 U of Taq DNA polymerase (Thermo Scientific, USA). After 2 min at 72 °C, DNA fragments were amplified through 20 cycles under the following conditions: denaturation for 20 s at 94 °C, annealing for 30 s at 56 °C and extension for 2 min at 72 °C. After the final elongation step for 30 min at 60 °C, the preamplification product was diluted 1:20 with nuclease-free water and used for selective amplification. This procedure was performed using the EcoRI-MseI primer pairs Eco-GTA and MseCAT, Eco-GCA and Mse-CAT, Eco-GGA and Mse-CAC, Eco-GAA and Mse-CAC. The EcoRI primers were endlabeled with the Hex or 6-Fam fluorescent label (SigmaAldrich, USA). The final amplification mixture (20 lL) contained 3 lL of the diluted preamplification mixture, 0.2 mM of a mixture of all dNTPs (Thermo Scientific, USA), 2.5 mM of MgCl2, 0.05 lM of the labeled EcoRI primer, 0.25 lM of the MseI primer, 19 Hot Start DNA polymerase buffer and 0.5 U of Hot Start DNA Polymerase (Thermo Scientific, USA). After 2 min at 94 °C, the DNA fragments were amplified through 30 cycles under the following conditions: denaturation for 20 s at 94 °C; annealing for 30 s at 66 °C for the first cycle, followed by decreasing temperatures (1 °C each cycle) during the next 10 cycles, 56 °C for the remaining 20 cycles and extension for 2 min at 72 °C. A final extension step was performed for 30 min at 60 °C. The resultant DNA fragments were detected using an automatic GeneScan ABI3130 apparatus from Macrogen Inc. (Korea). The presence or absence of fragments was scored on chromatograms using GeneMapper AFLP Genotyping Software 1.5. All fragments between 50 and 400 bp were scored. The obtained peaks were automatically transposed onto a binary matrix. To quantify amplification of each fragment, the fluorescence intensity of stable fragments (i.e., those fragments present in all control replicates) was analysed using the same software. Gene expression analysis RNA isolation and cDNA synthesis Total RNA was extracted using an Absolutely RNA RTPCR Miniprep Kit (Stratagene, USA) according to the manufacturer’s recommendations. First-strand cDNA was

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synthesized from 5 lg of total RNA using an Affinity Scrip Multiple Temperature cDNA Synthesis Kit (Stratagene, USA) following the manufacturer’s instructions. After the addition of 1 lL of oligo (dT), 1 lL of random primers, 0.8 lL of 100 mM dNTPs, 2 lL of 109 first-strand buffer and 5 lg of total RNA, the reaction was incubated at 65 °C for 5 min. Then, 0.5 lL of RNase inhibitor and 1 lL of StrataScript reverse transcriptase were added. The reaction was incubated for 1 h at 42 °C in an Eppendorf Mastercycler. The cDNA mixture was maintained at -20 °C until subjected to real-time PCR analysis. Real-time PCR and gene expression analysis Real-time PCR amplifications were performed in a qPCR MX3000P thermal cycler (Stratagene, USA) using Brilliant III Ultra-Fast SYBR Green QPCR Master Mix (Stratagene, USA) according to the manufacturer’s recommendations. For each treatment condition, four cDNA samples were analysed. Each 20 lL reaction contained 17 lL of master mix, 2 lL of a 2 lM primer pair mix and 1 lL of the reverse-transcribed product template. After 10 min at 95 °C, the products were amplified through 45 cycles under following conditions: denaturation for 30 s at 95 °C, annealing for 30 s at 52 °C and extension for 45 s at 72 °C. After a final elongation step for 1 min at 95 °C, the reaction specificity was determined for each reaction from the dissociation curve of the PCR product by following the SyberGreen fluorescence level during gradual heating of the PCR products from 60 to 95 °C. Melting curves were examined to verify that only one target was amplified and to assure that no genomic contamination was present in the RNA samples. The relative quantification of the expression level of each gene was normalized according to b-actin gene expression. The expression of seven genes involved in different pathways of DNA repair was monitored in the zebrafish larvae (nucleotide excision repair (NER): xpc, xpd, hr23b; base excision repair (BER): apex1; homologous recombination (HR): rad51; mismatch repair (MMR): msh2, msh6), whereas the expression of four genes was examined in adult zebrafish (xpc, xpd, hr23b, rad51). Additionally, the expression of the sod(Cu/Zn) and cyp1a1 genes was also measured. The primer pairs used for the amplification of the analysed genes were adopted from the literature or designed using LightCycler Probe Design Software 1.0 (Roche, Switzerland). Their sequences, accession numbers and reference sources are presented in Online Resource 1 (Table S2). Statistical analyses The results of the comet assay (% tDNA) were calculated based on the mean of each replicate/individual within a

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treatment group, and the data are presented as the mean and corresponding standard error of the mean (SEM) per group. Statistical analyses was performed with the Mann–Whitney U-test. The following levels of significance were reported: P B 0.05; P B 0.01 and P B 0.001. Both quantitative and qualitative qRAPD results are shown. The quantitative results are presented as the relative hybridization efficiency of each qRAPD primer calculated using the following formula: 2(Ct(b-actin)-Ct(qRAPD primer)?2) (Geffroy et al. 2012), where Ct represents the cycle threshold, and the qRAPD primer is the OPB7 or OPB11 primer. This formula provides the relative number of hybridization sites for the qRAPD primers in the genome. The results are presented as the mean of each treatment group and corresponding SEM. Qualitative results are presented as a frequency of the PCR products obtained with the OPB7 and OPB11 primers, according to the temperature intervals to which their Tm belonged. For both the quantitative and qualitative results, statistical analysis was performed using the Mann–Whitney U-test. The following levels of significance were reported: P B 0.05; 0.01. The AFLP results are also presented quantitatively and qualitatively. The qualitative results are given as the number of lost and introduced fragments for each treatment and each primer pair, and are expressed as the number of fragments lost for all replicates or introduced in at least one replicate within the treatment group compared to the control. The total fragment number per treatment group represents the cumulative number of fragments with different lengths recorded within the analysed group. The quantitative analysis of the amplification efficiency was calculated based on the peak heights of stable fragments, for which the mean (±SEM) peak height is calculated using the values for all replicates. The peak heights intensities were compared between the control and treated groups with the Mann–Whitney U-test (data not shown). The results are presented as the percentage of fragments with significantly decreased or increased intensities, calculated with respect to the total number of analysed fragments. Fragments that were introduced and lost in zebrafish larvae and adults were analysed to reveal specific fragments that were affected in both models upon exposure to a particular genotoxicant. The relative quantification of each gene expression level was normalized according to b-actin gene expression. The results are presented as the relative expression calculated using the following formula: Relative expression = 2(Ct(b-actin)-Ct(gene)). The results are presented as the mean for each treatment group and the corresponding SEM. Statistical analysis was performed using the Mann–Whitney U-test. The following level of significance was reported: P B 0.05.

Zebrafish genome instability

Results

The comet assay

General health and survival of exposed zebrafish

Analysis of zebrafish larvae following 6 days of exposure to EMS and B[a]P using the comet assay revealed a significant increase in DNA damage at both tested concentrations, and the damage remained significantly increased after the recovery period (Fig. 1a). EMS exposure induced DNA damage in a dose-dependent manner after 6 days, with 90 and 180 % increases in damage observed following treatment with 0.5 and 1 mM EMS, respectively (P = 0.0008 for both concentrations). B[a]P treatment caused DNA damage, but not in a dose-dependent manner; both concentrations resulted in an approximate 45 % increase in DNA damage (P = 0.016 for 0.001 lM B[a]P; P = 0.002 for 0.01 lM B[a]P) (Fig. 1a). In adult zebrafish, 6 days of EMS exposure caused a significant increase in DNA damage (P = 0.007 for 0.5 mM; P = 0.0009 for 1 mM EMS), which remained significantly increased after the recovery period in the 1 mM EMS treatment group (P = 0.007) (Fig. 1b). B[a]P treatment caused significant DNA damage only at the lower tested concentration, and this damage persisted following recovery (P = 0.01) (Fig. 1b).

High larval mortality (97–100 %) was observed in the treatment groups exposed to 0.1 and 1 lM B[a]P at the time of hatching (3 dpf). Therefore, the B[a]P concentrations had to be lowered to 1 and 10 nM to observe sublethal genotoxic effects. In these treatment groups, we did not record any developmental morphological abnormalities or increased mortality in comparison to the control group. In contrast, EMS treatment triggered significant (Mann–Whitney U-test) developmental abnormalities without causing mortality. In the 1 mM EMS treatment group, after the recovery period, 67 % of the larvae were smaller compared to the controls, 56 % exhibited decreased pigmentation and 58 % displayed curled tails. The lower EMS concentration triggered fewer morphological abnormalities, with 50 % of the larvae showing a smaller size and exhibiting decreased pigmentation compared to control larvae, whereas no curved tail phenotype was noted. In the control group all the larvae had well developed pigmentation and no curled tail phenotype was observed. The exposure of adult zebrafish to the two genotoxicants did not cause any mortality or any noticeable morphological changes.

Fig. 1 Results of the comet assay after 6 days of exposure to EMS and B[a]P, followed by 6 days of recovery (experimental time point 12 days) in a zebrafish larvae and b adult zebrafish. The data are presented as the mean and corresponding SEM. *Statistical significance compared with the respective control (Mann–Whitney U-test: *P B 0.05, **P B 0.01, and ***P B 0.001)

qRAPD qRAPD analysis revealed a significant increase in the number of hybridization sites in zebrafish larvae exposed to the lower B[a]P concentration for both tested qRAPD primers (Fig. 2a). No significant changes were observed following EMS exposure. The frequency of particular PCR products changed significantly following B[a]P treatment using both qRAPD primers (Table 1a). For OPB7, significant changes in the frequency of PCR products as a consequence of B[a]P treatment were noted at three different temperature intervals, whereas for the OPB11 primer, such changes were evident at seven temperature intervals. The lower EMS concentration (0.5 mM) induced a significant change in the frequency of PCR products in only one of the temperature intervals for each of the OPB7 and OPB11 primers, and no significant changes were observed in zebrafish larvae exposed to the higher EMS concentration. The results of the quantitative RAPD analysis in adult zebrafish revealed a general, yet insignificant decrease in the number of hybridization sites for the OPB7 qRAPD primer and, in most cases, a slight increase for the OPB11 primer. This increase was significant only under the 1 lM B[a]P treatment (Fig. 2b). The frequency of PCR products observed following EMS treatment changed significantly in two temperature intervals (69–72, 88–89) for the OPB7 primer and in one interval (72–74) for the OPB11 primer (Table 1b). After B[a]P treatment, significant changes in

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M. Sˇrut et al. Table 1 Qualitative assessment of PCR products frequency in different temperature intervals obtained with OPB7 and OPB11 qRAPD probes after 6 days exposure of (a) zebrafish larvae and (b) adult zebrafish to a concentration range of EMS and B[a]P followed by 6 days recovery Control

EMS (mM)

B[a]P (lM)

0.5

1

0.001

0.01

(a) OPB7

Fig. 2 Quantitative assessment of hybridization sites detected via qRAPD analysis using the OPB7 and OPB11 primers after 6 days of recovery following 6 days of exposure of a zebrafish larvae and b adult zebrafish to a range of EMS and B[a]P concentrations. The data are presented as the mean and corresponding SEM. *Statistical significance compared with the control (Mann–Whitney U-test: *P B 0.05 and **P B 0.01). The number of hybridization sites for the OPB11 primer is multiplied by 103 to facilitate reading

Aflp

123

0.5

0.6

0.3

0.5

0.5

72–74

0.4

0.5

0.3

0.3

0*

74–76

0.2

0.1

0.3

0

0.7*

76–78

0.3

0.4

0.2

0.4

0.1

79–80

0

0.1

0

0

0

80–81

0.1

0

0.3

0.3

0.1

81–82

0.9

1

0.7

0.7

0.9

84–85 85–86

1 0

1 0

0.9 0.1

1 0

1 0

86–87

0.9

0.5

0.9

1

1

88–89

0.5

0*

0.4

1*

0.6

90–92

0.8

0.6

0.7

1

0.9

69–72

0

0.4*

0.2

0.2

0

72–74

0

0.2

0.3

0.5*

0.1

74–76

0.1

0.1

0

0

0.4

76–78

0

0.1

0.1

0.2

0.1

79–80

0.2

0.3

0.3

0.7*

1*

80–81

0.1

0.1

0.1

0

0

82–83

0

0

0

0.6*

0.1

83–84

1

1

1

0.4*

0.9

85–86

0.3

0.5

0.5

0.5

0.9*

86–87 87–88

0.5 0.2

0.5 0

0.4 0.1

0.5 0

0* 0.1

89–90

0

0.1

0.2

0

0

90–92

0

0

0.1

0.6*

0.8*

Control

EMS (mM)

B[a]P (lM)

0.5

1

0.1

1

OPB11

the frequency of PCR products were observed in one temperature interval (69–72) for the OPB7 primer and in three temperature intervals (79–80, 85–86, 86–87) for the OPB11 primer (Table 1b).

Both qualitative and quantitative fragment changes were observed in zebrafish larvae after exposure to B[a]P and EMS. Qualitative changes in the obtained AFLP patterns were observed for all treatment conditions compared with the control. The loss of fragments present in all controls and the introduction of new fragments not observed in the control replicates were registered for all treatments and for all primer pair combinations (Table 2a). The greatest number of lost fragments was observed for the 0.01 lM B[a]P treatment, whereas the greatest number of introduced fragments was evident under the 0.001 lM B[a]P treatment. Significant quantitative alterations in stable fragment intensities were found for all treatment conditions. EMS treatment primarily resulted in increased fragment intensities, whereas B[a]P treatment resulted in decreased intensities for the majority of the observed fragments (Fig. 3a). The AFLP profiles of adult zebrafish treated with 0.01 lM B[a]P during embryonic and larval development

69–72

(b) OPB7 69–72

0.9

0.7

0.2*

0.5

0.4*

72–74

0.6

0.5

0.7

0.5

0.3

74–76

0.5

0.4

0.4

0.4

0.3

76–78

0.5

0.4

0.2

0.3

0.5

78–79

0

0.1

0.2

0

0.1

80–81

0

0

0.1

0.1

0.1

81–82

1

1

0.9

0.9

0.9

84–85

0.9

1

0.8

0.9

0.7

85–86

0.1

0

0.2

0.1

0.3

86–87

0.5

0.5

0.5

0.9

0.3

87–88

0.1

0.3

0

0.1

0.1

Zebrafish genome instability Table 1 continued Control

EMS (mM)

B[a]P (lM)

0.5

1

0.1

1

0.3

0.2

0.8*

0.4

0.5

69–72

0.2

0.3

0.4

0.2

0.1

72–74

0

0.2

0.4*

0.2

0.1

74–76

0.2

0.1

0.2

0.1

0.1

76–78

0.2

0.2

0.3

0.2

0.1

78–79

0

0

0

0.1

0

79–80

0.1

0.2

0.3

0

0.6*

80–81

0.6

0.3

0.7

0.7

0.2

82–83

0.3

0.5

0.3

0.3

0.3

83–84

0.7

0.7

0.7

0.7

0.7

84–85

0

0.1

0.1

0.3

0.1

85–86

0

0

0.2

0.4*

0

86–87 87–88

1 0

1 0

0.8 0

0.4* 0.3

0.9 0.1

89–90

0.3

0.1

0.3

0

0

88–89 OPB11

* Statistical significance compared to the control

showed that out of the 47 introduced fragments observed in exposed larvae after a 6-day recovery period, 25 persisted in adulthood, whereas out of the 127 lost fragments, 124 were restored in the adults (Table 2a). Both treatments induced measurable qualitative and quantitative fragment changes in adult zebrafish but to a much lower extent than in zebrafish larvae. Qualitative modifications such as the loss of some fragments and the introduction of new fragments not observed in the control replicates were registered for all treatments and for all primer pair combinations (Table 2b). Significant alterations in stable fragment intensities were observed primarily as a decrease in the fragment amplification intensity (Fig. 3b). The comparison of the AFLP profiles between zebrafish larvae and adults revealed that a certain number of the same fragments were either lost or introduced in both models (Fig. 4). We found a much higher relative number of identical fragments that were affected in both models for newly introduced compared with lost fragments upon B[a]P treatment. Gene expression analysis Zebrafish larvae exposure to either genotoxicant for 6 days did not alter the expression of any of the analysed DNA repair genes (Fig. 5a, c, e). After the recovery period, we observed significant down-regulation of some NER, BER and MMR genes (xpd, hr23b, apex1 and msh6) in the

0.5 mM EMS treatment group and of some NER genes (xpd and hr23b) in the 0.001 lM B[a]P treatment group (Fig. 5b, d, f). Exposure to 0.01 lM B[a]P resulted in a significant increase in the expression of the sod(Cu/Zn) and cyp1a1 genes, whereas after the recovery period significant down-regulation of cyp1a1 gene was observed in 0.001 lM B[a]P treatment group (Fig. 5g, h). Interestingly, when the 12-day-old control zebrafish larvae were compared with the 6-day-old larvae, a 2- to 3-fold increase in the expression of all of the analysed genes was demonstrated, and this difference reached statistical significance for the hr23b, rad51, msh2, msh6, sod(Cu/Zn) and cyp1a1 genes. Adult zebrafish exposure to EMS resulted in downregulation of some NER genes and of the sod(Cu/Zn) and cyp1a1 genes, whereas B[a]P treatment caused up-regulation of the rad51 and cyp1a1 genes in the gills (Fig. 6a, c, e). After the recovery period, most of the values returned to the control level (Fig. 6b, d, f). Analysis of the same genes in the digestive tract revealed less pronounced effects, with significant alterations in the expression of some NER genes being observed upon B[a]P and EMS treatment (data not shown).

Discussion The comet assay is a commonly used genotoxicity test that can reveal various aspects of DNA damage but can only detect certain lesions produced by B[a]P or EMS, such as strand breaks, alkali-labile sites and breaks induced by the DNA repair process. However, stable B[a]P-7,8-diol-9,10 epoxide (BPDE) bulky adducts at the N2 position of the guanine bases in the DNA (Tarantini et al. 2009), EMS alkylation of nucleotides or mutations that arise as a result of such adducts or alkylated bases cannot be directly revealed by this method (Sˇtambuk et al. 2008). In this study, significant DNA damage was recorded in both zebrafish larvae and adult specimens upon exposure to model genotoxicants, and in some cases, this damage remained significantly increased after the recovery period. In general EMS induced lesions decreased after the recovery period whereas B[a]P induced lesions were either at the same level or even higher in comparison to the level observed directly after exposure. Possible explanation for this discrepancy is that EMS-induced AP sites (caused either by spontaneous release of ethylated nucleotides or by DNA repair processes), are more effectively repaired over the course of time than B[a]P induced lesions. B[a]P metabolism leads to the formation of BPDE bulky adducts which can persist for several days after the end of exposure period (Akcha et al. 2000). It is plausible that even after 6 days of recovery comet assay is detecting lesions caused by ongoing DNA repair of B[a]P-induced DNA adducts. The

123

M. Sˇrut et al. Table 2 Qualitative assessment of AFLP data Primer pair

Treatment

Total fragment number

Retained fragments

Lost fragments

Introduced fragments

Fragments retained in the adulthooda

(a) Eco-GTA ? Mse-CAT

Eco-GCA ? Mse-CAT

Eco-GGA ? Mse-CAC

Control

128

0.001 lM B[a]P

128

110

18

0.01 lM B[a]P

60

55

73

5

0.5 mM EMS

125

111

17

14

1 mM EMS

105

98

30

7

Control

125

0.001 lM B[a]P

126

115

10

11

0.01 lM B[a]P

103

94

31

9

0.5 mM EMS

120

112

13

8

1 mM EMS Control

121 89

112

13

9

0.001 lM B[a]P

89

81

8

8

0.01 lM B[a]P

95

83

6

12

0.5 mM EMS

71

69

20

2

82

76

13

6

1 mM EMS Eco-GAA ? Mse-CAC

All primers

Primer pair

18

Control

6 introduced, 1 lost

4 introduced, 1 lost

121

0.001 lM B[a]P

125

110

11

15

0.01 lM B[a]P

125

104

17

21

0.5 mM EMS

104

95

26

9

1 mM EMS

113

99

22

14

Control

463

0.001 lM B[a]P

468

416

47

52

0.01 lM B[a]P

383

336

127

47

0.5 mM EMS

420

387

76

33

1 mM EMS

421

385

78

36

Treatment

4 introduced, 0 lost

Total fragment number

Retained fragments

Lost fragments

11 introduced, 1 lost

25 introduced, 3 lost

Introduced fragments

(b) Eco-GTA ? Mse-CAT

Eco-GCA ? Mse-CAT

Eco-GGA ? Mse-CAC

Control

142

0.1 lM B[a]P

143

134

8

9

1 lM B[a]P

150

138

4

12

0.5 mM EMS

130

125

17

5

1 mM EMS

137

132

10

5

Control 0.1 lM B[a]P

130 135

129

1

6

1 lM B[a]P

136

128

2

8

0.5 mM EMS

128

123

7

5

1 mM EMS

128

126

4

2

96

87

2

9

101

87

2

14

Control 0.1 lM B[a]P 1 lM B[a]P

Eco-GAA ? Mse-CAC

123

89

0.5 mM EMS

86

83

6

3

1 mM EMS

89

86

3

3

Control

135

0.1 lM B[a]P

143

127

8

16

1 lM B[a]P

143

128

7

15

Zebrafish genome instability Table 2 continued Primer pair

All primers

Treatment

Total fragment number

Retained fragments

Lost fragments

Introduced fragments

0.5 mM EMS

121

120

15

1

1 mM EMS

131

126

9

5

Control

496

0.1 lM B[a]P

517

477

19

40

1 lM B[a]P

530

481

15

49

0.5 mM EMS

465

451

45

14

1 mM EMS

485

470

26

15

Total number of generated, lost and introduced fragments per primer pair in (a) zebrafish larvae and (b) adult zebrafish after 6 days exposure to EMS and B[a]P followed by 6 days recovery period compared to the control a

Fragments analysed in 1 year old adult zebrafish exposed during embryonic and early larval stages to 0.01 lM B[a]P. Number of introduced or lost fragments retained in the adulthood is recorded

results of the comet assay gave us an indication of the DNA damage occurring under B[a]P and EMS exposure. However, qRAPD and AFLP were applied to further investigate and explain the DNA damage observed after recovery and to assess more permanent DNA alterations, such as DNA adducts and mutations. These methods were able to detect DNA alterations even in cases when the comet assay did not indicate any genotoxic effect. This was particularly evident in adult zebrafish treated with 1 lM B[a]P, in which the comet assay did not detect any significant genotoxic effects, whereas qRAPD revealed an increase in the number of OPB11 primer hybridization sites, and AFLP revealed both quantitative and qualitative alterations. These results reaffirmed the usefulness of qRAPD and AFLP for supplementing comet assay data, as previously noted (Orieux et al. 2011; Sˇrut et al. 2013). Alterations that are evident in qRAPD and AFLP analyses (both loss and appearance of hybridization sites/ fragments and decrease and increase in their amplification intensity) can be a consequence of an entire range of different factors and can be explained by various mechanisms, as discussed by Sˇrut et al. (2013). Briefly, these alterations could be a consequence of DNA adducts or of mutations occurring at qRAPD and AFLP primerannealing sites or at AFLP enzyme restriction sites, or they could be due to structural DNA changes occurring as a consequence of mutations outside the primer binding sites (Atienzar and Jha 2004, 2006; Bowditch et al. 1993; Pietrasanta et al. 2000; Qi et al. 2006). Furthermore, epigenetic events (e.g. changes in DNA methylation patterns) can lead to structural changes that may affect qRAPD and AFLP profiles. Carcinogens such as B[a]P can alter DNA methylation patterns by modifying the target DNA or by inactivating DNA methyl transferase (Atienzar and Jha 2004). Specifically, B[a]P can induce global and gene-specific DNA hypomethylation, which can alter gene expression, increase mutation rates and

trigger genome instability (Kisseljova and Kisseljov 2005; Fang et al. 2013). Although we observed genotoxic effects upon EMS exposure using the comet assay, these observations were not followed by quantitative, but only qualitative qRAPD alterations in both zebrafish models. The number of hybridization sites detected by qRAPD is a function of both their loss and creation. If both loss and appearance of hybridization sites occur at the similar frequency, the final result would not be significantly different from the control. This is a slight drawback of the method that can be supplemented by the qualitative analysis of the PCR product frequency. This issue has been previously discussed in the literature (Cambier et al. 2010; Orieux et al. 2011). Therefore, by getting insight into both qualitative and quantitative aspects of qRAPD method, we assure better insight into the genotoxicant-induced alterations. EMS-induced alterations were evident in both zebrafish models using AFLP. This method thus proved to be sensitive enough to provide evidence of DNA alterations occurring at various sites within the genome. Alterations detected via qRAPD and AFLP analyses could be an indication of more persistent DNA changes that may remain in the genome. We were able to detect the persistence of DNA alterations in adult zebrafish 1 year after the end of B[a]P exposure. More than 50 % of the introduced fragments observed in the larvae 6 days after the end of exposure remained in 1-year-old adult zebrafish, indicating the formation of mutations (point mutations, large rearrangements) that persisted in the genome and that were not repaired long after exposure ended. Interestingly, almost all of the fragments that were lost in the larvae reappeared in the adults. This can be explained by the formation of DNA BPDE bulky adducts, that could have persisted in larval zebrafish shortly after the end of exposure but that were effectively removed by the adult stage. DNA adducts can block or reduce the effectiveness of Taq

123

M. Sˇrut et al.

Fig. 3 Quantitative assessment of AFLP data presented as a percentage of the stable AFLP fragments showing an unchanged, decreased or increased intensity compared with the control in a zebrafish larvae and b adult zebrafish after 6 days of recovery following 6 days of exposure to EMS and B[a]P. Only significant increases and decreases were considered

Fig. 4 Comparison of introduced and lost fragments between zebrafish larvae and adults. The numbers in circles and the sizes of the circles represent the overall number of introduced or lost fragments in either zebrafish larvae or adults exposed to genotoxicants. The numbers and sizes of overlapping sections represent the number of specific fragments where DNA alterations occurred in both models

DNA polymerase and alter enzyme restriction efficiency in the first step of the AFLP procedure (Atienzar et al. 2002; Van der Veen et al. 2005) and therefore can contribute to the loss of certain fragments. If the observed persistent alterations would be passed onto the next generation, they could lead to an increased baseline mutation frequency in the descendants of the exposed individuals, a phenomenon known as transgenerational genomic instability (O’Brien et al. 2013). Understanding of such persistent, delayed and even transgenerational effects of toxicant exposure is necessary as it might have direct and practical implications for ecological risk assessment (Segner 2011). Interestingly, comparison of the AFLP profiles of zebrafish larvae and adults revealed several alteration-sensitive sites in the genome. For instance, B[a]P treatment caused many identical fragments to be newly introduced in both zebrafish models, revealing potential mutational hot spots in the genome. BPDE adduct hot spots and B[a]P mutational hot spots have been previously described in the literature (Smith et al. 2000; Lewis and Parry 2004). The results obtained in this study show that DNA marker technology has the power to detect potentially permanent DNA alterations and thus can complement the standard battery of genotoxicity tests. However, the choice of the optimal method to detect genotoxic effects should take into account all the positive and negative aspects of the particular method. Comet assay is still most cost

effective tool to get an indication of genotoxic effects, although the data can not easily be used to extrapolate more persistant effects. On the other hand qRAPD and AFLP can detect various forms of DNA damage that can potentially persist in the genome. However, the success of qRAPD depends on the choice of adequate primers that can target potential alteration prone sites in the genome. AFLP on the other hand scans the entire genome, and although it has been used for the detection of genotoxic events in plant and animal systems (Labra et al. 2004; Fraser et al. 2008; Liu et al. 2010), it might only be appropriate when highly inbred and genetically homogenous stocks or cell lines are utilized (Sˇrut et al. 2013, 2015). This study provides an indication of the applicability of the AFLP method in inbred zebrafish strains for evaluating DNA alterations. However, more detailed data concerning the genetic diversity of the individuals and populations included in the analysis would be required to further validate the method and to more reliably exclude the scoring of natural genetic variations. The gene expression data for zebrafish larvae showed basal levels of expression for most of the analysed genes after exposure to both model toxicants whereas after the recovery, repression of several DNA repair genes was observed. These data indicate a limited DNA repair activity in zebrafish larvae which could be one of the explanations for the high amount of DNA alterations recorded upon

123

Zebrafish genome instability Fig. 5 Relative expression of genes involved in a, b nucleotide excision repair (xpc, xpd, hr23b); c, d homologous recombination (rad51) and base excision repair (apex1); e, f mismatch repair (msh2, msh6); and g, h the oxidative stress response (sod (Cu/Zn)) and xenobiotic metabolism (cyp1a1) in zebrafish larvae after 6 days of exposure to EMS and B[a]P, followed by a 6-day recovery period (experimental time point, 12 days). The data are presented as the mean and corresponding SEM. *Statistical significance compared with the control (Mann–Whitney U-test: *P B 0.05)

exposure and recovery. In contrast, it has been reported that zebrafish larvae possess a very effective DNA repair system for coping with robust cell division and DNA replication, particularly during the early stages of development. Therefore, they have been described as questionable models for genotoxicity evaluation (Sussman 2007). However, this phenomenon may be limited to the early stages of zebrafish development, when the expression of DNA repair genes is at its highest peak. For instance, the NER gene (xpd) has an expression peak during the earliest stages of embryonic development (1 cell–5 hpf), followed by a dramatic decrease in expression during later developmental stages (Silva et al. 2012). MMR genes (msh2 and msh6) were also found to be up-regulated in early embryos (1–2 hpf) (Hsu et al. 2010). In the present study, 6- and 12-day-old zebrafish larvae were found to be more sensitive to the applied genotoxic stress than adult zebrafish, as demonstrated by the fact that the damage detected by the

comet assay persisted after 6 days of recovery in all of the treatment groups and by the greater extent of alterations detected through qRAPD and AFLP analyses compared with those alterations observed in exposed adults. Further evidence of larval sensitivity is provided by the high mortality observed at the time of hatching, under exposure to identical concentrations of B[a]P as adult zebrafish and by the fact that, the applied concentrations had to be decreased one hundred times to observe sublethal genotoxic effects. Therefore, zebrafish larvae represent a low-cost, high-throughput model organism for testing genotoxic compounds. However, further research is required for exploring the responses of zebrafish larvae to various genotoxic agents and for assessing the changes in their gene expression profiles during different life stages. Furthermore, a larger experimental setup would be required to assess the repeatability of the biological phenomena observed in this study.

123

M. Sˇrut et al. Fig. 6 Relative expression of genes involved in a, b nucleotide excision repair (xpc, xpd, hr23b); c, d homologous recombination (rad51); and e, f the oxidative stress response (sod (Cu/Zn)) and xenobiotic metabolism (cyp1a1) in adult zebrafish gills after 6 days of exposure to EMS and B[a]P followed by a 6-day recovery period (experimental time point, 12 days). The data are presented as the mean and corresponding SEM. *Statistical significance compared with the control (Mann–Whitney U-test: *P B 0.05)

Conclusion As qRAPD and AFLP analyses detect a wider range of DNA damage and alterations than the comet assay, these methods enabled the detection of DNA modifications even in cases when DNA damage was not detectable with the comet assay, or when the comet assay indicated complete repair. These findings imply the formation of DNA alterations such as DNA adducts or mutations that cannot be detected by the comet assay and that may lead to the impairment of DNA structure and function. Such alterations were evident through AFLP analysis, even 1 year after the end of the exposure period in adult zebrafish treated with B[a]P during embryonic and larval development, indicating the longevity of the detected DNA alterations. Additionally, AFLP analysis appears to reveal potential mutational hot spots in the genome. Zebrafish larvae proved to be an extremely sensitive model for detecting genotoxicity and DNA alterations, as demonstrated using the comet assay and qRAPD and AFLP methods. Thus, they can be recommended as a sensitive model for in vivo genotoxicity testing as well as a good model for monitoring the persistence of genotoxic effects. Acknowledgments The authors are grateful to the students Antonio Karaga and Aleksandar Lazic´ for help with the preparation and analysis of comet assay slides and to Adam Maguire for English revision of the

123

manuscript. This study was part of a Ph.D. thesis and was conducted within the framework of Project No. 119-0982934-3110 supported by the Ministry of Science, Education and Sports of the Republic of Croatia and of program Tractifs No. ANR-07-SEST-02301 supported by the French National Research Agency. Maja Sˇrut was awarded a scholarship by the Croatian Science Foundation to perform a portion of the research at the University of Bordeaux, France. Ethical standards Experiments performed in this study comply with the current laws of Croatia. Conflict of interest of interest.

The authors declare that they have no conflict

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