Genotoxic Effect and Carcinogenic Potential of a Mixture of As and Cd in Zebrafish at Permissible Maximum Contamination Levels for Drinking Water

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Genotoxic Effect and Carcinogenic Potential of a Mixture

of As and Cd in Zebrafish at Permissible Maximum

Contamination Levels for Drinking Water

Oguzhan Doganlar&Zeynep Banu Doganlar&

Fulya Dilek Gokalp Muranlı&Utku Guner

Received: 19 October 2015 / Accepted: 8 February 2016 / Published online: 18 February 2016

# Springer International Publishing Switzerland 2016

Abstract Currently, the toxic effects and carcinogenic potential of individually treated arsenic (As) or cadmium (Cd) are well documented both in animal and human tissues. However, there are no data focusing on the genotoxicity of these heavy metals as a mixture at the very low concentrations of permissible limits for drinking water. In this study, we examine the genotoxicity and carcinogenic potential of single and combined treatments of As and Cd, as well as attempt to elucidate the mechanism of action of certain cell defense systems such as antioxidants, gene repair, heat shock, cell cycle control, and the apoptosis pathway. Zebrafish (Danio rerio), reared under controlled conditions with artificial diets, were treated with As and Cd, either individually or in combination, at concentrations commonly found in water (10 ppb for As and 5 ppb for Cd) and tenfold higher concentrations for 48 h. Our results indicate that separately, As and Cd treatments at low dose selectively

induce antioxidant enzymes, gene repair, and caspase- independent apoptosis in gill tissue, by targeting the mitochondria, leading to oxidative stress and sub-lethal levels of DNA damage. However, tenfold higher (100 ppb As + 50 ppb Cd) treatment caused significant downregulation of genes involved in double-strand break repair and molecular chaperone genes.

Additionally, the highest BCL2/BAX ratio (1.6) and lowest expression levels of caspase-3 (8.4-fold) in all treated groups were observed in same condition. These results demonstrate that both single and combined exposure to As and Cd at permissible levels is potentially safe and causes repairable genotoxicity in gill tissue.

However, the highest concentration is potentially carcinogenic due to ineffective DNA repair and insufficient apoptosis.

Keywords Arsenic . Cadmium . Genotoxicity . Apoptosis . Carcinogenesis . Gene expression

1 Introduction

Recently, several clinical and epidemiological studies have reported that the increased incidence of a certain type of cancer, in a particularly polluted area, correlated with the presence of carcinogenic agents. At present, a number of studies aim to investigate the relationship between the toxic chemicals and the carcinogenic effect to detect ultimate metabolism before biodegradation, water, air, soil, and food pollution. Environmental pollution with toxic chemicals—such as heavy metals, DOI 10.1007/s11270-016-2779-1

O. Doganlar

:

Z. B. Doganlar (*)

Department of Medical Biology, Faculty of Medicine, Trakya University, 22030 Edirne, Turkey

e-mail: zdoganlar@trakya.edu.tr Z. B. Doganlar

e-mail: zdoganlar@yahoo.com.tr O. Doganlar

Technology Research and Development Center (TUTAGEM), Trakya University, 22030 Edirne, Turkey

F. D. G. Muranlı

:

U. Guner

Department of Biology, Faculty of Science, Trakya University, 22030 Edirne, Turkey

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polycyclic aromatic hydrocarbons, pesticides, and volatile organic compounds—progressively increases with rapid industrialization. Because of the simultaneous release of these substances, thousands of compounds can be present as a mixture in the soil, water, and atmo- sphere. In particular, arsenic (As) and cadmium (Cd) often occur together due to their resistance to biodegradation and persistence in the environment (Fay and Mumtaz 1996; Wang and Fowler 2008). Therefore, humans are potentially exposed to As and Cd simultaneously and/or sequentially. Both As and Cd are classi- fied as genotoxic carcinogens in humans. The International Agency for Research on Cancer (IARC) reported that As and Cd are group 1 (which means that there is sufficient evidence for their carcinogenicity in humans) human carcinogens. It has also been reported that the exposure to As via drinking water causes skin, lung, bladder, and kidney cancer; metabolic diseases (diabetes mellitus, black food disease); and neurological diseases in humans (Çöl et al.1999; Smith et al.2000).

While acute Cd exposure causes nausea, vomiting, di- arrhea, muscle cramps, salivation, sensory disturbances, liver injury, convulsions, shock, and renal failure, chronic exposure has been associated with kidney damage, osteoporosis, and cancer in multiple organs (Jennings et al.1996; Liu et al.2000).

The presence of As and Cd in drinking water directly affects public health. Because of the toxic effects of these heavy metals, governments and agen- cies, such as the American Public Health Association (APHA), the World Health Organization (WHO), the Indian Standard Institution (ISI), the Central Pollution Control Board (CPCB), and the Indian Council of Medical Research (ICMR), enforce some limitations on the levels of heavy metals that are permissible in drinking water. TheBRegulation of Water Intended for Human Consumption,^ published by the Turkish Government in 2005, notes that the permissible concentrations of As and Cd are 10 and 5μg/L, respectively. Although people are exposed to numerous organic and inorganic agents simultaneously, these permissible limits were determined based on single agents. Considering people’s chronic low-dose co- exposure to As and Cd, we thought that investigating the genotoxic effects of As and Cd alone or combined is important.

It was reported that the toxic effects of As and Cd occur via similar mechanisms, such as DNA damage, production of reactive oxygen species

(ROS), induction of stress proteins [heat shock proteins (HSPs)], and metal-binding proteins (metallothioneins) interfering with essential metals (Doğanlar et al. 2014; Liu et al. 2000). DNA damage and misfolded proteins play major roles in carcinogenesis (Bernstein et al. 2005). In the cell, single- or multi-nucleotide mutations of the DNA—such as point mutations, deletions, and in- sertions—can occur in a gene’s promoter and change its expression or can occur in the gene body and change the characteristics or stability of the protein product. Several survival pathways— such as those involved in gene repair, antioxidants, apoptosis, and cell cycle control—directly affect this unstable condition. Inhibition of gene repair mechanisms, insufficient apoptosis, and uncon- trolled cell proliferation result in carcinogenesis (Knudson 2001; Wood et al. 2007; Croce 2008).

The zebrafish (Danio rerio) is a powerful model organism for studying genotoxicity, cancer, DNA damage, and pharmacology due to its genes being highly conserved with human genes (Chen et al.

2014; Dai et al. 2014; Shive 2013). Additionally, the determination of the toxicant response of gill tissue is important due to its direct contact with, and being a primary target of, toxic chemicals (Wang et al. 2015). The effects of As and Cd on the expression of several genes and their toxic potential are well documented (Andrew et al.

2003; Qin et al. 2012; Wieland et al. 2009; Zhou et al. 2012). However, there are no data focusing on the carcinogenic potential of these heavy metals as a mixture at permissible limits for drinking and surface water.

In this paper, we determined the amounts of As and Cd in the gill tissue of zebrafish and evaluated the genotoxic effect on total DNA and genomic template stability. We also demonstrated the role of mixtures of As and Cd at trace levels on heat shock proteins (HSP60, HSP70), antioxidant [man- ganese superoxide dismutase (Mn-SOD), copper- zinc superoxide dismutase (CuZn-SOD), catalase (CAT), glutathione peroxidase (GPX), glutathione synthase (GS)], gene repair (RAD-18, SMUG-1, XRCC3), intrinsic apoptosis (Survivin, BCL 2, XIAP, BAX, APAF1, Cyc-C), and cell cycle control (P21^Cip1, P27^Kip1, CDKN3, Cyclin D1) pathways via target gene expression profiles. The aim of this study was to investigate the carcinogenic

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potential and genotoxic mechanisms of As and Cd mixtures at permissible limits for drinking water.

2 Materials and Methods

2.1 Test Organism

Zebrafish (D. rerio) were obtained from Bogazici University, Experimental Animal Unit, Turkey, and ac- climated for 2 weeks in a glass aquarium (50 × 50 × 100 cm, 100 L) under routine approved animal welfare protocols at the aquarium room. Fish were fed with commercial fish feed. The photoperiod (12:12 light/

dark), water temperature (26 °C), and pH (7–8) were maintained until at the end of experiment. After accli- matization, fish were transferred to smaller propylene aquariums, each containing 1 L heavy metal and control water.

2.2 Experimental Design

The analytical standards of arsenic (CAS 7440-38-2) and cadmium (CAS 7440-43-9) were prepared as a stock solution dissolved in water, and final concentrations in the test aquarium were adjusted to 10 and 100 ppb for As, 5 and 50 ppb for Cd, 10 + 5 ppb and 100 + 50 ppb for mixed solutions of As + Cd, respectively for 48 h. These concentrations were selected based on the permissible limits of As (10 ppb) and Cd (5 ppb) in drinking water. At the end of the experimental period, dissected gill tissues were frozen in liquid nitro- gen and deposited in a −86 °C deep freezer (Daihan Scientific) for the subsequent steps of the chemical and genetic analyses. Both controls and heavy metal-treated experiments were performed with three replicates.

2.3 Determination of As and Cd Accumulation

As and Cd accumulations in gill tissue of both control and treated fish were determined according to Doğanlar et al. (2014). Gill tissues were digested with 65 % ultrapure nitric acid solution (1 mL). Digests were diluted with deionized water, and the final volume of the solution was adjusted to 10 ml. After the dilution of digests, inductively coupled plasma mass spectrometry (ICP-MS, Agilent 7700xx) was done for the analysis.

2.4 Genetic analyses

2.4.1 DNA Extraction and RAPD Procedures

Genomic DNA was isolated from D. rerio using a DNeasy® Blood and Tissue Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions.

The total genomic DNA was diluted with nuclease-free water to a concentration of 25 ng/μL, and this diluted DNA was used as a template DNA for the PCR reaction.

Standard 50 μL PCR reactions were performed using 2 μL (50 ± 10 ng) of template DNA, AmpliTaq Gold PCR Master Mix (4326717, Applied Biosystems®), 1 μL of each primer, and nuclease-free water (10977- 015, Invitrogen™). DNA amplification was performed using the thermal cycler Applied Biosystems®

ProFlex™ PCR System (Table1). After the amplification of DNA, 10μL of the product with 2 μL of loading dye (R0611, Fermentas) was loaded onto a 2 % agarose gel with ethidium bromide in 2× TAE (Tris 1.6 M, acetic acid 0.8 M, EDTA 40 mM, Ambion® 10X TAE) buffer.

The molecular weight standard 100 bp DNA ladder (Geneaid, DL007) was used according to the manufacturer’s instructions. The DNA bands were visualized with a UV transilluminator (Vilber Lourmat), and the sizes of all of the bands were calculated with program software. The percentage of genomic template stabilities (GTS) was calculated using random amplified polymorphic DNA (RAPD) data GTS (%) = (1− [(a + b) / n]) × 100 where a and b indicate appearance of new bands and disappearance of normal bands, respectively;

n is the number of total bands in the control.

2.4.2 Isolation of Total RNA and cDNA Synthesis

At the end of each treatment period, the gill tissues were dissected and total RNA was isolated from each gill specimen using the PureLink® RNA Mini Kit (Life Technologies, USA) according to the manufacturer’s instructions. The extracted RNA concentrations were measured by the OPTIZEN NanoQ micro volume pho- tometer. The concentration of total RNA was adjusted to 50 ng/μL for the synthesis of the first strand of comple- mentary DNA (cDNA) using a High-Capacity cDNA Reverse Transcription Kit (Life Technologies, USA).

cDNA synthesis was performed using the thermal cycler Applied Biosystems® ProFlex™ PCR System (step 1 25 °C, 10 min; step 2 37 °C, 120 min; step 3 85 °C,

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5 min). The cDNA was stored−20 °C for subsequent steps of the analysis.

2.4.3 Quantitative Real-Time PCR Analysis

The relative expression levels of the genes belong to antioxidant system (CuZn-SOD, Mn-SOD, CAT, GPX, GS), heat shock protein family (HSP60 and HSP70), gene repair pathway (RAD-18, SMUG-1, XRCC3),

apoptosis pathway (Survivin, BCL 2, XIAP, BAX, APAF1, Cyc-C, Caspase-3), and cell cycle arrest pathway (P21Cip1, P27Kip1, CDKN3, Cyclin D1) in response to treatment with the As and Cd were analyzed by quantitative real-time PCR (qRT-PCR) using the SYBR® Select Master Mix (Life Technologies, USA) on an ABI Step One Plus Real-Time PCR system with the primer pairs given in Table1. Gene expressions were determined as the relative fold change compared to the Table 1 Genes, primer sequences, and PCR conditions belong to qRT-PCR and RAPD assays

Genes/Primers Primer sequences Genes/Primers Primer sequences PCR conditions qRT-PCR

Mn-SOD F:5′-GTTCGGTGACAACA CCAATG-3′

R:5′-GGAGTCGGTGATGT TGACCT-3′

BAX F:5′-TTCATCCAGGATCG AGCAGA-3′ R:5′-GCAAAGTAGAAG

GCAACG-3′

1 cycle of 2 min at 50 °C and 10 °C min at 95 °C followed by 40 cycles of denaturation at 95 °C for 15 s, annealing, and extension at 60 °C for 1 min CuZn-SOD F: 5′-TCTGAAGAAGGCC

ATCGAGT-3′

R: 5′-GCAGATAGTAGGCG TGCTCC-3′

BCL-2 F:5′-ATGTGTGTGGAGAG CGTCAA-3′

R:5′-ACAGTTCCACAAAG GCATCC-3′

CAT F:5′-TACGAGCAGGCCAA GAAGTT-3′

R:5′-ACCTTGTACGGGCA GTTCAC-3′

XIAP F:5′-GGGGTTCAGTTTCAAG GAC-3′

R:5′-TGCAACCAGAACCTC AAGTG-3′

GS F:5′-TGGGACCAGCAAG TAAAACC-3′ R:5′-TCGCGAATG

TAGAACTCGTG-3′

Cyt-C F:5′-AGTGGCTAGAGTGGTCA TTCATTTACA-3′

R:5′-TCATGATCTGAATTCTG GTGTATGAGA-3′ HSP60 F:5′-GTCGCGCCCCGTTA

GCAC-3′

R:5′-CATCGCGTCCCACCTT CTTCAT-3′

APAF-1 F:5′-GATATGGAATGTCTCAGA TGGCC-3′

R:5′-GGTCTGTGAGGACTCC CCA-3′

HSP70 F:5′-CGAGETCGACGCATTG TTTG-3′

R:5′-GAGTGGATCCGCCGA CGAGTA-3

Caspase-3 F:5′-GGTATTGAGACAGACA GTGG-3′

R:5′-CATGGGATCTGTTTCT TTGC-3′

RAD-18 F:5′-TTCACAAAAGGAAGC CGCTG-3′

R:5′-TTACTGAGGTCATATTA TCTTC-3′

Survivin F:5′-GACGACCCCATAGAG GAACA-3′

R:5′-GACAGAAAGGAAAG CGCAAC-3′

SMUG-1 F:5′-CTCTGTGGCTGAGGGT TGAT-3′

R:5′-TTGTAGATGATGCCCA CAGG-3′

CDKN3 F:5′-GCCGCCTTCTGACT CTTC-3′

R:5′-GATCCGCTTCTGGT TTCTTA-3′

P21^Cip1 F:5′-GGCGTTTGGAGTGG TAGAAA-3′

R:5′-GACTCTCAGGGTCG AAAACG-3′

CCND1 F:5′-AACAGAAGTGCGAG GAGGAG-3′

F:5′-TGAGGCGGTAGTAG GACAGG-3′

P27^Kip1 F:5′-CCGGCTAACTCTGAG GACAC-3′

R:5′-TGGATCCAAGGCTC TAGGTG-3′

GADPH F:5′-TTGGTATCGTGGAAGG ACTCA-3′

R:5′-TGTCATCATATTTGGC AGGTTT-3′

RAPD-PCR

AP5 5′-TCCCGCTGCG-3′ 1253 5′-GTTCCGCCCC-3′ 40 cycles of 95 °C denaturation (30 s), 37 °C annealing (30 s) and 72 °C elongation (90 s) with an initial 94

°C denaturation (3 min) and a final 72 °C extension (30 min)

OPC 11 5′-AAAGCTGCGG-3′ P2 5′-ATGTAACGCC-3′

OPC 15 5′-GACGGATCAG-3′ RAPD4 5′-CTACTGCGCT-3′

OPW10 5′-TCGCATCCCT-3′ OPB 05 5′- TGCGCCCTTC-3′

RAPD1 5′-CCGATATCCC-3′ RAPD3 5′-CCAGCGTATT-3′

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control and normalized with 18S mRNA expressions.

The comparative cycle threshold (ΔCt) method (User Bulletin 2, Applied Biosystems, CA, USA) was performed to analyze the expression levels of mRNAs.

2.5 Statistical Analyses

The experiment was performed with three replicates per treatment time for each control and experimental group.

Dose-dependent alterations in GTS % value were normalized with arcsine transformations. Differences in the metal accumulations in tissues, normalized GTS % values, and the degree of the relative fold change resulting from gene expressions under the effect of As, Cd, and their mixture were compared using analysis of variance (ANOVA) with Duncan’s separation of means test using SPSS 18 software at a significance level of P≤ 0.05. Correlations between the relative expressions levels of genes were analyzed by a bivariate correlation test with Pearson correlation coefficient and a two-tailed test of significance using SPSS 18 software at significance levels of P≤ 0.01 and 0.05.

3 Results

3.1 Metal Accumulation

The first step taken to understand the effect of As and Cd in this study was to determine the accumulation of heavy metals in gill tissue following treatment with increasing concentrations of As or Cd. While the highest As accumulation occurred with an As exposure of

100 ppb (9.4-fold, compared to control), the administra- tion of 50 ppb Cd in 100 ppb As (Mix2) caused a decline in As accumulation (3.8-fold, compared to control) (FAs= 9.610; P = 0.0001). Conversely, Mix2 caused more Cd accumulation (11.51 ± 1.09 ppb) than 50 ppb Cd (6.13 ± 1.30 ppb). The highest Cd accumulation was found in gill tissues exposed to Mix2 (FCd= 33.349;

P = 0.0001). Both As and Cd accumulation in the gill tissues exposed to 5 ppb Cd in 10 ppb As (Mix1) do not show any difference to separate exposure to each metal (Fig.1). Our results indicate that while As uptake was reduced by Cd, Cd accumulation was increased approximately twofold by a high concentration mixture of As and Cd.

3.2 Oxidative Stress, DNA Damage, and Repair Mechanisms

In several organisms, As and Cd produce ROS due to oxidative stress. Increasing ROS levels due to an inad- equate antioxidant system can lead to genotoxic effects such as DNA polymorphisms and gene induction/re- pression. The antioxidant genes are rapid and sensitive biomarkers that monitor oxidative stress in tissues and cells. Significant increases in the gene expressions of antioxidant enzymes CuZn-SOD, Mn-SOD, CAT, GPX, and GS were observed in gill tissue after both individual and mixed treatments with As and Cd (Fig. 2) (FCuZn- S O D= 392.39, P = 0.0001; FM n - S O D= 1829.24, P = 0.0001; FCAT= 413.82, P = 0.01). These increases were observed in all ROS scavenger genes (cytosolic CuZn-SOD, CAT, and mitochondrial Mn-SOD) in a range of approximately 15.4- to 59.8-fold and in the

Fig. 1 The detected levels of As and Cd elements in gill tissues of zebrafish treated with As, Cd, Mix1 (10 ppb As + 5 ppb Cd), and Mix 2 (100 ppb As + 50 ppb Cd) for 48 h. Data represent ng/

mg (ppb), mean ± SE, n = 4.

*Means followed by the same letters in same metal group do not differ significantly at P≤ 0.05 (results obtained by one-way ANOVA, separated by Duncan test)

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Fig. 2 Relative fold change determined by quantitative real-time PCR (qRT-PCR) analysis of antioxidant (CuZn-SOD, Mn-SOD, CAT, GPX, GS), gene repair (RAD-18, SMUG-1, XRCC3), and heat shock proteins (HSP60, HSP70) genes in heavy metals-exposed gill

tissues of zebrafish for 48 h. All data were normalized with GADPH expression and given as relative to control (control = 1 not shown in the figure); n = 5. Different letters indicating significantly different values were analyzed by one-way ANOVA and Duncan test (P≤ 0.05)

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detoxifying and lipid peroxidation preventive enzyme genes (GPX and GS) in the range of 4- to 34-fold. In general, the high concentrations of individually treated As and Cd caused significant reductions in the expression levels of antioxidant genes compared to low concentrations, except GPX which increased in a dose- dependent manner. While a similar trend was observed in the ROS scavenger genes upon mixed treatment with As and Cd, GPX and GS levels increased with increasing mixture concentrations (FGPX= 858.18, P = 0.029;

FGS= 18,882.64, P = 0.0129).

Our study indicated that As and Cd treatments at the permissible levels cause oxidative stress in gill tissue.

We used the RAPD assay to assess As- and Cd-induced oxidative damage in total DNA. For the RAPD assay,

ten oligonucleotide primers were evaluated to screen genomic DNA isolated from gill tissue: two primers produced inconsistent results and eight primers generat- ed consistently positive results. Among the eight primers, four informative and stress-specific primers produced bands ranging in molecular size from 151 to 2411 bp and were selected for analyses of RAPD profiles (Fig.3). Arsenic and Cd treatments, individually or as a mixture, resulted in the disappearance of bands, the appearance of new bands, and increases/decreases in the intensity of bands in the gill tissues of zebrafish at all exposure concentrations compared to their respective controls (Fig. 3). In total, 64 bands were observed in the control groups and 132 new bands appeared in the treated groups. The 174 bands lost in the treated groups

Fig. 3 RAPD assay band profiles of total genomic DNA belongs to gill tissues of zebrafish exposed to heavy metals for 48 h. L Gen Ruler 100 bp plus DNA ladder (100–3000 bp), As1 10 ppb, As2

100 ppb, Cd1 5 ppb, Cd2 50 ppb, Mix1 10 ppb As + 5 ppb Cd, Mix2 100 ppb As + 50 ppb Cd. Primers = AP5, OPC11, OPC15, and OPW10

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had a molecular size ranging from 172 (OPW10) to 2411 bp (OPC15); these were amplified by four primers (Table2). The increases in band intensity corresponding to the changes in molecular size were prevalent with the Cd2, Mix1, and Mix2 treatments, but a decrease in band intensity was obvious with the As2 treatment for the OPC11 and OPC15 primers. The maximum number of new bands was found for OPW10 with the Mix1 exposure; the bands ranged in molecular size from 1720 to 188 bp. The genomic template stability (GTS %) value of the gill tissues was calculated by the mean band changes for the four primers tested (Fig.4). Our study indicated that GTS decreased with increasing

concentrations of As, Cd, and their mixture and that the highest damage occurred with the Mix2 treatment (FGTS%= 324.14, P = 0.018).

Damage to DNA is one of the major causes of carcinogenesis. Cells have to rapidly repair this damage with own repair mechanisms in order to survive. To further estimate the gene and protein repair efficiency of damaged DNA upon heavy metal exposure, we investigated the role of post-replication repair (RAD-18), base excision repair (SMUG-1), double-strand break repair (XRCC3) and misfolded or damaged protein repair, and molecular chaperone (HSP60 and HSP70) genes in repair mechanisms in gill tissue. Our results revealed Table 2 Changes in the RAPD profiles based on the appearance (A) and disappearance (D) of bands with specific molecular sizes (bp) using four primers in the control and heavy metal-exposed zebrafish

Primers Control As1 As2 Cd1 Cd2 Mix1 Mix2

AP5 1822; 1500; 994; 956; 748;

562; 500; 464; 378; 369;

327; 270; 242; 228

A 1039; 982;

900; 600;

435; 349;

253

1005; 982;

900; 771;

627; 527

1039; 982;

900; 811;

700;627;

358; 0.253

1005; 900;

521; 253

1005; 982;

971; 900;

771; 627;

358; 345

1005; 900;

771; 627;

521; 358;

349 D 1822; 1500;

500; 464;

378; 270;

242

1822; 1500;

994; 748;

500; 369;

270

1822; 1500;

748; 500;

369; 228

1822; 1500;

500; 369;

270; 242

1822; 1500;

994; 748;

500; 464;

369; 228

1822; 1500;

748; 500;

369; 270 OPC 11 1293; 1158; 1000; 959; 643;

533; 520; 428; 357; 331;

285; 272

A 1349; 1188;

810; 700;

664; 554;

446; 339

664; 554 1112; 738;

446

1349; 1239;

1112; 664;

500

1239; 1188;

1062; 664;

600; 339

1500; 1062;

900; 664;

600; 339;

314 D 1293; 1158;

959; 643;

533; 520;

428; 357;

272

1293; 900;

810; 520;

357; 285

1293; 1158;

52; 428;

357; 272

1293; 1158;

643; 520;

428; 285;

272

1293; 1158;

1000; 643;

428; 357;

285

1293; 1000;

643; 533;

537; 331;

300; 285 OPC 15 2411; 1571; 1280; 900; 818;

800; 720; 619; 597; 559;

529; 780; 400; 358; 316;

265; 231; 207; 176

A 1340; 940;

659; 375

1500; 1126;

940; 836;

675; 427;

375; 281

1340; 1008;

659; 427;

375; 247

1500; 1340;

1126; 659;

281

2299; 1500;

1126; 659;

427

1500; 1126;

1108; 659

D 2411; 1280;

900; 800;

597; 556;

265; 231;

207

2411; 1571;

900; 818;

800; 619;

400; 358;

265; 207

2411; 1280;

800; 597;

559; 400;

316; 231;

207

1571; 1280;

818; 800;

619; 358;

316; 265;

310; 207

2411; 1571;

818; 619;

400; 231;

207

1571; 800;

619; 559;

265; 231;

207 OPW10 1555; 1120; 944; 900; 836;

755; 700; 628; 574; 548;

507; 474; 400; 364; 344;

311; 283; 266; 172

A 1213; 1063;

1000;600;

439; 203

1213; 1063;

439; 203;

151

1835; 1489;

1213; 670;

650; 439;

203; 151

1720; 1489;

800

1720; 1489;

1063;

1000;873;

800; 670;

600; 439;

188

2255; 1634;

1489; 873;

670; 439;

203; 160;

144 D 1555; 1120;

944; 574

1555; 1120;

944; 548

1555; 1120;

700; 574;

548; 172

1555; 839;

574; 548;

266; 172

1555; 836;

755; 700;

574; 474;

312; 283;

266; 172

1555; 900;

700; 587;

507; 364;

172

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statistically significant increases in RAD-18 and SMUG-1 mRNA levels in all exposure groups, compared to their respective controls (FRAD18= 730.25, P = 0.0001; FSMUG-1= 1425.95, P = 0.003). However, compared to the groups singly treated with either 10 ppb As or 5 ppb Cd, these genes showed significantly lower expression levels with high concentration treatments. While the highest increase in the mRNA levels of RAD-18 and XRCC3 occurred at the 5 ppb Cd exposure (37.32- and 1.88-fold compared to their controls, respectively), Mix1 and Mix2 caused the highest increases in SMUG-1 gene expression (44.81- and 43.96-fold compared to their controls, respectively) (Fig.2). The expression level of XRCC3 significantly decreased with mixed As and Cd and high-dose Cd treatments (FXRCC3= 21.614, P = 0.001). Low-dose Cd and both Mix1 and Mix2 treatments significantly induced HSP60 expression in gill tissue (FHSP60= 8117.42, P = 0.0101). Both As and high-dose Cd treatments gave similar expression levels and these levels were significantly lower than other groups. While expression of the stress-specific HSP70 genes significantly increased with A s t r e a t m e n t i n a d o s e - d e p e n d e n t m a n n e r (FHSP70= 4651.59, P = 0.003), significantly lower expression levels of this gene were observed in the Cd- and mixture-treated groups (Fig.2).

3.3 Intrinsic Apoptosis Pathway and Cell Cycle Control Genes

The irreparable DNA damage caused by several toxic chemicals, especially heavy metals, has been linked to the activation of apoptosis signal transduction pathways

in different tissues and cells. For this reason, we investigated the apoptotic effect of As and Cd in gill tissue using gene expression profiles of three principle com- ponents of the intrinsic pathway: apoptosis inhibitor, pro-apoptotic, and cell cycle. In gill tissues, both 10 and 100 ppb As treatments caused a significant increase in Survivin expression (52.2- and 25.6-fold, respectively) (Fig.5). A similar induction of BCL2 expression was observed, with 26.1- and 25.4-fold increases with 10 and 100 ppb, respectively. The BCL2/BAX ratio was 1.56 and 1.43 for 10 and 100 ppb As, respectively ( FB C L 2= 3 0 7 . 7 5 , P = 0 . 0 0 0 1 ; FB A X= 3 9 1 . 3 8 , P = 0.0001). Although the Apaf1 gene was significantly induced in response to As treatment, Cyt-c, Caspase-9, and Caspase-3 were unaffected by this increase, and the relative expression level of the cell cycle checkpoint gene P21^Cip1 and cell cycle arrest gene P27^Kip1 was significantly lower compared to other treatment groups (Figs.5and6). For this reason, the apoptotic effect of As on gill tissue remained at a low level. While the low- dose Cd treatment caused significant induction of the Survivin and XIAP genes (37.6- and 90-fold, respectively), we observed significantly lower expression of the same genes (16.4- and 33.6-fold) at high- concentration Cd treatment. However, in the same conditions, the BCL 2/BAX ratio (0.61 and 0.41) was significantly reduced, and this reduction was triggered by the increase in pro-apoptotic genes (BAX, APAF1, Cyc-C, and Caspase-3) belonging to the intrinsic apoptosis pathway, approximately in the range of 12- to 136-fold. Additionally, a significant induction in the P21^Cip1 gene was observed with both Cd treatments.

Our results indicate that the apoptotic effect of Cd and As was particularly strong upon Mix1 treatment. At this condition, while apoptosis inhibitor genes exhibited low expression levels, the apoptosis regulator genes of the intrinsic apoptosis pathway and bax were overexpressed (47.1-fold) and triggered the release of Cyt-C (276.8- fold) from mitochondrial membranes. This gene linked the APAF1 (47.4-fold) and activated caspase-3 (22.2- fold). Additionally, the G1/S transition checkpoint gene P21^Cip1(overexpressed 40-fold) and the cell cycle arrest gene P27^Kip1(225.7-fold) stopped the cycle in the G1 phase (Fig.6).

In this study, the relative expression of Survivin and XIAP (28.8- and 50.1-fold, respectively) was significantly higher in the Mix2-treated groups than in the Mix1-treated groups (FSurvivin= 1289.10, P = 0.0001;

FXIAP= 4032.47, P = 0.001). Additionally, the bcl2/bax Fig. 4 Dose-dependent alterations in GTS % in heavy metal-

exposed gill tissues of zebrafish. As1 10 ppb, As2 100 ppb, Cd1 5 ppb, Cd2 50 ppb, Mix1 10 ppb As + 5 ppb Cd, Mix2 100 ppb As + 50 ppb Cd. Different letters indicating significantly different values among concentrations were analyzed by one-way ANOVA and Duncan test (P≤ 0.05)

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ratio in the Mix2-treated group (1.5) was significantly higher than in the Mix1-treated group. In contrast to other treated groups, the relative expression levels of the apoptotic genes Cyt-C, Caspase-9, and Caspase-3 (18.2-, 4.4-, and 8.4-fold, respectively) were significantly lower, as was the case for the cell cycle control genes P21^Cip1and P27^Kip1(19.2- and 20.5-fold, respectively).

4 Discussion

Arsenic and Cd induce several disruptions in many physiological and metabolic pathways, particularly af- fecting oxidative stress. They also cause a degeneration process in many of the cell organelles, tissues, and organs due to having strong DNA damage capacity, Fig. 5 Relative fold change determined by qRT-PCR analysis of

Survivin, XIAP, BCL2, BAX, APAF-1, Cyc-C, Caspase 9, and Caspase 3 gene expressions belong to apoptosis pathway in heavy metals-exposed gill tissues of zebrafish for 48 h. All data were

normalized with GADPH expression and given as relative to control (control = 1 not shown in figure); n = 5. Different letters indicating significantly different values were analyzed by one-way ANOVA and Duncan test (P≤ 0.05)

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persistent characteristic, and carcinogenic potential. In this report we investigate for the first time to our knowl- edge the genotoxic properties of single and combined treatments with As and Cd at levels permissible for drinking water and also assess the carcinogenic potential on the gill tissue of zebrafish. To this end, we firstly determined the amount of As and Cd in gill tissues exposed either individually or as a mixture. Our results indicate that metal accumulation of As and Cd shows significant increases in a dose-dependent manner upon individual exposure (Fig. 1). As reported by previous studies, the accumulation of As and Cd showed concentration-dependent increases in the individually exposed zebrafish (Wicklund-Glynn et al. 1994;

Vellinger et al.2012b). However, in the case of mixed exposure, the accumulated amount of a metal may be increased, decreased, or unchanged by other metals in the mixture (Phillips1976; Casini and Depledge1997;

Bat et al.1998; Shuhaimi-Othman and Pascoe2007). In this study, compared with the single As and Cd treatment while the As accumulation significantly decreased, we observed an approximately twofold increase in the amount of Cd in gill tissue with the Mix2 treatment. It could be said that the marked change in both As and Cd accumulation can be explained by a disturbance in the balance of cellular mechanisms due to the toxic effect of

heavy metals and competition of metals for metal-binding proteins (Holwerda1991; Vellinger et al.2012a).

Oxidative stress to the gill tissue is a common occur- rence during exposure to various types of toxic chemicals. This stress to the zebrafish results in the release of large amounts of ROS into the cell cytosol and triggers the antioxidant systems, which are the first defense mechanisms of cells. Recent studies have shown that during stressful conditions, there is a differ- ential regulation of genes encoding antioxidant enzymes (SOD, CAT, GS) (Doğanlar et al.2014; Doganlar et al.

2015). SOD is the leading and the most important enzyme of the antioxidant system, catalyzing the dismutation of superoxide anions to hydrogen peroxide (H2O2) and water to scavenge ROS. In the second step, CAT catalyzes the decomposition of H2O2to water and oxygen (Chelikani et al. 2004). Glutathione mechanisms depollute toxic xenobiotic and endobiotic electro- philes so that the secondary products can be metabolized and excreted (Doganlar and Doganlar 2015). In this study, we found that all ROS scavenger genes (cytosolic CuZn-SOD, CAT, and mitochondrial Mn-SOD) and the detoxifying and lipid peroxidation preventive enzyme genes (GPX and GS) were induced in gill tissue, as quantified by qRT-PCR. However, the expression profiles of these genes, except GPX, did not vary in a dose- Fig. 6 Relative fold change determined by qRT-PCR analysis of

p21^Cip1, P27^Kip1, CDKN3, Cyclin D1 gene expressions belong to cyclin-dependent kinases proteins in heavy metals-exposed gill tissues of zebrafish for 48 h. All data were normalized with

GADPH expression and given as relative to control (control = 1 not shown in figure); n = 5. Different letters indicating significantly different values were analyzed by one-way ANOVA and Dun- can test (P≤ 0.05)

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dependent manner: in general, high doses caused slight or significant reductions in antioxidant gene expression.

It could be said that the marked decreases of antioxidant gene expression can be explained by disturbances of the expressional balance by exposures to high concentrations of As and Cd. However, in this study, significantly higher expression levels of antioxidant genes were pre- cursors to oxidative damage. It is known that As and Cd can cause increases in ROS via uncoupling oxidative phosphorylation, thus resulting in increases in the enzymatic and non-enzymatic antioxidant system elements.

Recent studies have demonstrated that high expression levels of genes involved in antioxidant system were biomarkers of cell oxidative stress (Silins and Högberg 2011).

Increased ROS and hydroxyl radical levels in cells are potent activators of lipid peroxidation and DNA damage (Møller et al. 2010) and high concentrations are known to affect cellular homeostasis, which ulti- mately leads to apoptotic/necrotic cell death (Winyard et al.2005). The RAPD assay indicated that As and Cd accumulation in gill tissue treated with all the concentrations tested significantly altered the electrophoretic band pattern of total DNA and caused a significant decrease in GTS (%) due mainly to increased ROS levels. The RAPD band polymorphism may be due to the changes in the priming sites of the oligonucleotides caused by genome rearrangement. Newly occurring bands may also be due to DNA damage induced by stressors (Conte et al.1998; Atienzar et al.1999; Liu et al.2005). Moreover, new bands are maybe the result of genomic template instability. The genetic stability is related to the level of DNA damage and the efficiency of DNA repair and replication mechanisms in the cell (Silins and Högberg2011). Liu et al. (2005) reported that the disappearance of PCR products mainly affected the high-molecular-weight bands because the probabil- ity of sustaining DNA damage increased with the length of the amplified fragment. In this study, we observed increases in band intensity in gill tissue with the Cd2, Mix1, and Mix2 treatments, but a decrease in band intensity was obvious with the As2 treatment for the OPC11 and OPC15 primers. Changes observed in the band intensity especially depend on genome rearrangement actions by enzymatic reactions such as Taq polymerase and efficiency of DNA repair mechanisms, par- t i c u l a r l y o n s i n g l e 8 - o x o - 7′,8′-dihydro-2′- deoxyadenosine lesions. We suggest that differences in band patterns and intensity, i.e., DNA damage, related to

insufficient ROS scavenging and gene repair mechanisms are maybe due to disrupted chemical composition and redox potential in the cell due to As and Cd accumulation.

Oxidative stress, resulting from increasing ROS caused by As and Cd toxicity, mediates the genotoxic effects of these metals (Filipič 2012; Adeyemi et al.

2015). In our study, increased expression of antioxidant enzyme genes showed that As, Cd, and their mixtures caused oxidative stress due to accumulation of ROS. As seen in Fig. 4, compared to low concentrations of As and Cd, the expression levels of antioxidant enzymes are generally reduced by high concentrations. The reason for these decreases in mRNA levels of enzymes may be the toxic effects of ROS on DNA, RNA, lipids, proteins, and nucleotides (Slupphaug et al.2003). This was confirmed by the changes in the expressions of HSP (HSP60, HSP70) and DNA repair genes (RAD-18, SMUG-1, and XRCC3). The increasing expression l e v e l s o f H S P 6 0 a n d H S P 7 0 s h o w t h a t t h e cytoprotective mechanism involves the degradation of the misfolded proteins after exposure. While increased HSP70 expression is used as a biomarker for general cell stress, upregulation of HSP60 expression levels is used as an indicator of mitochondrial stress and injury (Martinus et al.1996). In our study, HSP60 expression levels were higher than HSP70 with the Cd and mixed treatments, but single As treatment caused the opposite (Fig.2). Therefore, we can conclude that both As and Cd caused damage to cellular proteins but the Cd and mixed treatments mainly targeted mitochondria.

The data from the qRT-PCR of antioxidant systems genes, RAPD assay, and GTS % experiment indicated that a significant amount of total DNA underwent damage, especially at higher concentrations of single and mixed treatments, which was further validated by relative gene expression levels of DNA repair pathway genes of control and As-Cd-exposed gill tissues. In our study, expression of the post-replication repair gene RAD-18 and base excision repair gene SMUG-1 significantly increased upon both single and combined As and Cd exposure (Fig. 2). But expression of the DNA double-strand break repair gene XRCC3 increased only upon both As concentrations and 5 ppb Cd exposure (Fig.2). The high concentration of Cd and both mixture treatments resulted in significantly lower expression levels of XRCC3 genes compared to other treatment groups. Previous studies have reported that mismatch repair may be important in regulating recombination

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events between distinct DNA templates that could result in different types of genetic instability (Rayssiguier et al.

1989; Selva et al.1995; Datta et al.1996). In addition, several genes belong to nucleotide excision repair, which is responsible for reintegration of physical and chemical damage to DNA (Karran and Marinus1982;

Fram et al.1985; Feng et al.1991; Mellon and Champe 1996) and contributes to a cell-cycle control pathway by interacting with different types of DNA damage and triggering cell-cycle arrest or other responses to DNA damage (Hawn et al.1995; Anthoney et al.1996). We thought that the induction of both antioxidant enzyme and DNA repair gene expression demonstrated that both mechanisms act together to overcome genotoxic damage. Additionally, according to this study in gill tissue, after receiving especially sub-lethal DNA damage, cell cycle checkpoints are activated to permit extra time for repair of the damaged DNA and ensure genomic stability (Kanaar et al. 1998; Zona et al. 2014). Cell cycle checkpoints and apoptosis are closely linked because following cell cycle arrest, the cell determines its fate:

the cell cycle resumes if the DNA is repaired, and cellular apoptosis occurs when the damage is severe and irreparable (King and Cidlowski1998).

Apoptosis is a process involving interactions with anti-apoptotic and pro-apoptotic genes within a cell. In our study, we showed that both anti-apoptotic and pro- apoptotic genes are upregulated upon heavy metal exposure (Fig.5). While the upregulation of anti-apoptotic genes can be explained as a means to ensure the cell’s survival in the case of sub-lethal damage (Roos and Kaina2006; Fulda et al.2010), the increased expression of pro-apoptotic genes may trigger the activation of apoptotic pathways due to the severe damage (Fernández et al.2003). It was reported that while the anti-apoptotic BCL2 blocks, BAX promotes the release of Cyc-C from mitochondria (Clark et al. 1997;

Abdelkader et al.2013). In our study, we determined that the increasing mRNA levels of anti-apoptotic genes were not enough to inhibit apoptosis. Additionally, the upregulation of Cyc-C due to increasing BAX expression and activated Caspase-3 means triggering the mitochondrial apoptotic pathway (Bernardi et al. 2001).

We determined that both As and Cd and their low-dose mixture even at permissible limits in drinking water caused a high BCL2/BAX ratio and upregulation of Cyc-C and Caspase-3 genes, which indicates activation of the mitochondrial intrinsic pathway (Senapati et al.

2015). Therefore, we thought that the As, Cd, and Mix1

caused apoptosis in gill tissue due to severe DNA damage that cannot be repaired by antioxidant and gene repair mechanisms.

Our results indicate that the Mix2 treatment, unlike other treatment groups, caused greater heavy metal accumulation and higher genetic template instability. We observed significant downregulation of the XRCC3 and HSP70 genes with the Mix2 treatment compared to other treatment groups. Insufficient DNA repair mechanisms can cause more DNA damage to accumulate and increase the risk of cancer. In several cancer types, a DNA repair deficiency is sporadically due to damage of a DNA repair gene. Generally, however, downregulated expression of DNA repair genes is due to methylation, mutation, and suppression that decrease or silences gene expression.

Previous studies of As- and Cd-induced carcinogenesis report that such heavy metals cause reduced expression of DNA repair enzymes. While As inhibits poly(ADP-ribose) polymerase (PARP), X-ray repair cross-complementing protein (XRCC), DNA excision repair protein (ERCC), and the ligase group of DNA repair genes (Andrew et al 2003; Ebert et al. 2011; Qin et al. 2012), cadmium inhibits the DNA repair genes MutS protein homologue (MSH), ERCC, and XRCC (Wieland et al.2009; Schwerdtle et al. 2010; Zhou et al. 2012). Moreover, Bernstein et al. (2002) proposed that DNA repair inhibition and its effect on pro-apoptotic molecules is to be a predominant mechanism in carcinogenesis. In the present study, the highest BCL2/BAX ratio (1.6) and lowest relative expression levels of Cyt-C (19.2-fold) and Caspase-3 (8.4-fold) among all treated groups occurred with the Mix2 treatment.

Additionally, with the Mix2 treatment, we observed significantly lower expression levels of the cell cycle arrest gene P27^Kip1and the cell cycle checkpoint gene P21^Cip1compared to other groups (Fig.6). We thought that this treatment suppressed apoptosis in gill tissue damaged by the toxic effect of As and Cd.

The process of apoptosis, i.e., programmed cell death, is closely related to the different types of caspase activations, and these mechanisms regulate gene expression, including antioxidant, apoptosis inhibitor, apoptotic, and cell cycle arrest genes. Caspases are one of the major players in apoptosis and act as common apoptotic molecules in a number of different substrates in the cytoplasm in various forms of cell death (Degterev et al. 2003). Apoptosis occurs as a

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defense mechanism when cells are damaged by toxic agents (Norbury and Hickson2001). It is vital for the cell and tissue to prevent sub-lethal DNA damage f r o m b e i n g t r a n s f e r r e d t o n e w g e n e r a t i o n s . Inappropriate apoptosis (either too little or too much) plays a major role in neurodegenerative and autoim- mune disorders, ischemic damage, and many types of cancer (Elmore2007).

5 Conclusions

The results of this study indicate that individual and mixed As and Cd found in drinking water at the very low levels permitted by several countries may cause a genotoxic effect on the gill tissue of zebrafish, as indicated by DNA damage due to genomic template instability and altered expression profiles of genes in the following categories: antioxidant, HSP, gene repair, cell cycle control, and apoptosis. However, sub-lethal DNA damage and disrupted cell function due to single and low-dose mixture treatments may be repaired and restored by effectively activated antioxidant, gene repair, and heat shock systems. In single and low-dose mixture treatments, we observed sufficient apoptosis in irreparable cells of gill tissue. Therefore, we suggest that permissible levels and tenfold higher concentrations of As and Cd had a clear genotoxic effect, but this toxicity may be tolerated by cell defense mechanisms working together.

In this study, we report that exposure to high concentrations of mixed As and Cd was insufficient in inducing expression of gene repair, apoptosis, and cell cycle control genes in gill tissue. This indicates that mixed As and Cd (present in tenfold higher concentrations than permissible limits for drinking water) could act synergistically to damage the DNA template and several cell mechanisms such as gene repair and apoptosis pathway at very low accumulation in gill tissue, unlike single exposures. These findings highlight that the mixture of high concentrations of As and Cd may have a carcinogenic potential.

Consequently, although As and Cd exposure alone at the permissible limits caused a repairable genotoxic effect, much attention should be given to chronic low- dose exposure that can threaten human health due to the bioaccumulation and non-biodegradable properties of heavy metals. We propose that these limits

should be revised with further studies on the genotoxicity and carcinogenic potential of As and Cd mixtures in different organisms.

Acknowledgments The authors are grateful to Prof. Dr. Yener Yoruk and the Technology Research and Application Centre (TUTAGEM), which is funded by the T.R. State Planning Orga- nization (Project Number 2011K120390), for providing the labo- ratory equipment.

References

Abdelkader, T. S., Chang, S. N., Kim, T. H., Song, J., Kim, D. S.,

& Park, J. H. (2013). Exposure time to caffeine affects heartbeat and cell damage‐related gene expression of zebrafish Danio rerio embryos at early developmental stages.

Journal of Applied Toxicology, 33, 1277–1283.

Adeyemi, J. A., da Cunha Martins-Junior, A., & Barbosa, F.

(2015). Teratogenicity, genotoxicity and oxidative stress in zebrafish embryos (Danio rerio) co-exposed to arsenic and atrazine. Comparative Biochemistry and Physiology, Part C:

Toxicology & Pharmacology, 172, 7–12.

Andrew, A. S., Karagas, M. R., & Hamilton, J. W. (2003).

Decreased DNA repair gene expression among individuals exposed to arsenic in United States drinking water.

International Journal of Cancer, 104, 263–268.

Anthoney, D. A., McIlwrath, A. J., Gallagher, W. M., Edlin, A. R.,

& Brown, R. (1996). Microsatellite instability, apoptosis, and loss of p53 function in drug-resistant tumor cells. Cancer Research, 56, 1374–1381.

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.

Environmental Toxicology and Chemistry, 18, 2275–2282.

Bat, L., Raffaelli, D., & Marr, I. L. (1998). The accumulation of copper, zinc and cadmium by the amphipod Corophium volutator (Pallas). Journal of Experimental Marine Biology and Ecology, 223, 167–184.

Bernardi, P., Petronilli, V., Di Lisa, F., & Forte, M. (2001). A mitochondrial perspective on cell death. Trends in Biochemical Sciences, 26, 112–117.

Bernstein, C., Bernstein, H., Payne, C. M., & Garewal, H. (2002).

DNA repair/pro-apoptotic dual-role proteins in five major DNA repair pathways: fail-safe protection against carcinogenesis. Mutation Research, 511(2), 145–178.

Bernstein, H., Bernstein, C., Payne, C. M., Dvorakova, K., &

Garewal, H. (2005). Bile acids as carcinogens in human gastrointestinal cancers. Mutation Research, Reviews in Mutation Research, 589(1), 47–65.

Casini, S., & Depledge, H. (1997). Influence of copper, zinc, and iron on cadmium accumulation in the talitrid amphipod, Platorchestia platensis. Bulletin of Environmental Contamination and Toxicology, 59, 500–506.

(15)

Chelikani, P., Fita, I., & Loewen, P. C. (2004). Diversity of structures and properties among catalases. Cellular and Molecular Life Sciences, 61(2), 192–208.

Chen, Y. Y., Zhu, J. Y., & Chan, K. M. (2014). Effects of cadmium on cell proliferation, apoptosis, and proto-oncogene expression in zebrafish liver cells. Aquatic Toxicology, 157, 196– 206.

Clark, R. S., Chen, J., Watkins, S. C., Kochanek, P. M., Chen, M., Stetler, R. A., & Graham, S. H. (1997). Apoptosis-suppressor gene bcl-2 expression after traumatic brain injury in rats. The Journal of Neuroscience, 17(23), 9172–9182.

Çöl, M., Çöl, C., Soran, A., Sayli, B. S., & Oztürk, S. (1999).

Arsenic-related Bowen's disease, palmar keratosis, and skin cancer. Environmental Health Perspectives, 107, 687.

Conte, C., Mutti, I., Puglisi, P., Ferrarini, A., Regina, G., Maestri, E. A., & Manmiroli, N. (1998). DNA fingerprinting analysis by a PCR based method for monitoring the genotoxic effects of heavy metals pollution. Chemosphere, 37, 2739–2749.

Croce, C. M. (2008). Oncogenes and cancer. New England Journal of Medicine, 358, 502–511.

Dai, Y. J., Jia, Y. F., Chen, N., Bian, W. P., Li, Q. K., Ma, Y. B., Chen, Y. L., & Pei, D. S. (2014). Zebrafish as a model system to study toxicology. Environmental Toxicology and Chemistry, 33, 11–17.

Datta, A., Adjiri, A., New, L., Crouse, G. F., & Robertson, S. J.

(1996). Mitotic crossovers between diverged sequences are regulated by mismatch repair proteins in Saccaromyces cerevisiae. Molecular and Cellular Biology, 16, 1085–1093.

Degterev, A., Boyce, M., & Yuan, J. (2003). A decade of caspases.

Oncogene, 22, 8543–8567.

Doğanlar, Z. B., Doğanlar, O., & Tabakçıoğlu, K. (2014).

Genotoxic effects of heavy metal mixture in Drosophila melanogaster: expressions of heat shock proteins, RAPD profiles and mitochondrial DNA sequence. Water, Air, &

Soil Pollution, 225, 1–14.

Doganlar, O. & Doganlar, Z. B. (2015). Effects of a mixture of volatile organic compounds on total DNA and gene expression of heat shock proteins in Drosophila melanogaster.

Archives of Environmental Contamination and Toxicology, 68(2), 395–404.

Doganlar, O., Doganlar, Z. B., & Tabakcioglu, K. (2015). Effects of permissible maximum-contamination levels of VOC mixture in water on total DNA, antioxidant gene expression, and sequences of ribosomal DNA of Drosophila melanogaster.

Environmental Science and Pollution Research, 22(20), 15610–15620.

Ebert, F., Weiss, A., Bultemeyer, M., Hamann, I., Hartwig, A., &

Schwerdtle, T. (2011). Arsenicals affect base excision repair by several mechanisms. Mutation Research, 715, 32–41.

Elmore, S. (2007). Apoptosis: a review of programmed cell death.

Toxicologic Pathology, 35, 495–516.

Fay, R., & Mumtaz, M. (1996). Development of a priority list of chemical mixtures occurring at 1188 hazardous waste sites, using the HazDat database. Food and Chemical Toxicology, 34, 1163–1165.

Feng, W.-Y., Lee, E., & Hays, J. (1991). Recombinagenic process- ing of UV-light photoproducts in nonreplicating phage DNA by the Escherichia coli methyl-directed mismatch repair system. Genetics, 129, 1007–1020.

Fernández, E. L., Gustafson, A.-L., Andersson, M., Hellman, B.,

& Dencker, L. (2003). Cadmium-induced changes in

apoptotic gene expression levels and DNA damage in mouse embryos are blocked by zinc. Toxicological Sciences, 76, 162–170.

Filipič, M. (2012). Mechanisms of cadmium induced genomic instability. Mutation Research, Fundamental and Molecular Mechanisms of Mutagenesis, 733, 69–77.

Fram, R. J., Cusick, P. S., Wilson, J. M., & Marinus, M. G. (1985).

Mismatch repair of cis-diamminedichloroplatinum (II)-induced DNA damage. Molecular Pharmacology, 28, 51–55.

Fulda, S., Gorman, A. M., Hori, O., & Samali, A. (2010). Cellular stress responses: cell survival and cell death. International Journal of Cell Biology, 2010, 1–23.

Hawn, M. T., Umar, A., Carethers, J. M., Marra, G., Kunkel, T. A., Boland, C. R., & Koi, M. (1995). Evidence for a connection between the mismatch repair system and the G2 cell cycle checkpoint. Cancer Research, 55, 3721–3725.

Holwerda, D. A. (1991). Cadmium kinetics in freshwater clams. V.

Cadmium–copper interaction in metal accumulation by Anodonta cygnea and characterization of the metal-binding protein. Archieves of Environmental Contamination and Toxicology, 21, 432–437.

Jennings, G., Sneed, R. & Clair, M. (1996). St. Metals in drinking water. North Carolina Cooperative Extension Service Publication no AG-473-1. Electronic version 3. http://

infohouse.p2ric.org/ref/01/00073.htm. Accessed 19 Oct 2015.

Kanaar, R., Hoeijmakers, J. H., & van Gent, D. C. (1998).

Molecular mechanisms of DNA double-strand break repair.

Trends in Cell Biology, 8, 483–489.

Karran, P., & Marinus, M. G. (1982). Mismatch correction at O6- methylguanine residues in E. coli DNA. Nature, 296, 868– 869.

King, K., & Cidlowski, J. (1998). Cell cycle regulation and apoptosis 1. Annual Review of Physiology, 60, 601–617.

Knudson, A. G. (2001). Two genetic hits (more or less) to cancer.

Nature Reviews Cancer, 1, 157–162.

Liu, J., Liu, Y., Habeebu, S. M., Waalkes, M. P., & Klaassen, C. D.

(2000). Chronic combined exposure to cadmium and arsenic exacerbates nephrotoxicity, particularly in metallothionein-I/

II null mice. Toxicology, 147, 157–166.

Liu, W., Li, P., Qi, X., Zhou, Q., Zheng, L., Sun, T., & Yang, Y.

(2005). DNA changes in barley (Hordeum vulgare) seedlings induced by cadmium pollution using RAPD analysis.

Chemosphere, 61, 158–167.

Martinus, R. D., Garth, G. P., Webster, T. L., Cartwright, P., Naylor, D. J., Høj, P. B., & Hoogenraad, N. J. (1996).

Selective induction of mitochondrial chaperones in response to loss of the mitochondrial genome. European Journal of Biochemistry, 240, 98–103.

Mellon, I., & Champe, G. N. (1996). Products of DNA mismatch repair genes mutS and mutL are required for transcription- coupled nucleotide-excision repair of the lactose operon in Escherichia coli. Proceedings of the National Academy of Sciences, 93, 1292–1297.

Møller, P., Jacobsen, N. R., Folkmann, J. K., Danielsen, P. H., Mikkelsen, L., Hemmingsen, J. G., Vesterdal, L. K., Forchhammer, L., Wallin, H., & Loft, S. (2010). Role of oxidative damage in toxicity of particulates. Free Radical Research, 44, 1–46.

Norbury, C. J., & Hickson, I. D. (2001). Cellular responses to DNA damage. Annual Review of Pharmacology and Toxicology, 41, 367–401.

(16)

Phillips, D. J. H. (1976). The common mussel Mytilus edulis as an indicator of pollution by zinc, cadmium, lead and copper. I.

Effects of environmental variables on uptake of metals.

Marine Biology, 38, 59–69.

Qin, X.-J., Liu, W., Li, Y.-N., Sun, X., Hai, C.-X., Hudson, L. G.,

& Liu, K. J. (2012). Poly (ADP-ribose) polymerase-1 inhibition by arsenite promotes the survival of cells with unrepaired DNA lesions induced by UV exposure. Toxicological Sciences, 127, 120–129.

Rayssiguier, C., Thaler, D. S., & Radman, M. (1989). The barrier to recombination between Escherichia coli and Salmonella typhimurium is disrupted in mismatch-repair mutants.

Nature, 342, 396–401.

Roos, W. P., & Kaina, B. (2006). DNA damage-induced cell death by apoptosis. Trends in Molecular Medicine, 12, 440–450.

Schwerdtle, T., Ebert, F., Thuy, C., Richter, C., Mullenders, L. H.,

& Hartwig, A. (2010). Genotoxicity of soluble and particu- late cadmium compounds: impact on oxidative DNA damage and nucleotide excision repair. Chemical Research in Toxicology, 23, 432–442.

Selva, E. M., New, L., Crouse, G. F., & Lahue, R. S. (1995).

Mismatch correction acts as a barrier to homeologous recombination in Saccharomyces cerevisiae. Genetics, 139, 1175– 1188.

Senapati, V. A., Jain, A. K., Gupta, G. S., Pandey, A. K., &

Dhawan, A. (2015). Chromium oxide nanoparticle‐induced genotoxicity and p53‐dependent apoptosis in human lung alveolar cells. Journal of Applied Toxicology, 35, 1179–1188.

Shive, H. (2013). Zebrafish models for human cancer. Veterinary Pathology Online, 50, 468–482.

Shuhaimi-Othman, M., & Pascoe, D. (2007). Bioconcentration and depuration of copper, cadmium, and zinc mixtures by the freshwater amphipod Hyalella azteca. Ecotoxicology and Environmental Safety, 66, 29–35.

Silins, I., & Högberg, J. (2011). Combined toxic exposures and human health: biomarkers of exposure and effect.

International Journal of Environmental Research and Public Health, 8, 629–647.

Slupphaug, G., Kavli, B., & Krokan, H. E. (2003). The interacting pathways for prevention and repair of oxidative DNA damage. Mutation Research, Fundamental and Molecular Mechanisms of Mutagenesis, 531, 231–251.

Smith, A. H., Arroyo, A. P., Mazumder, D., Kosnett, M. J., Hernandez, A. L., Beeris, M., Smith, M. M., & Moore, L.

E. (2000). Arsenic-induced skin lesions among Atacameno people in Northern Chile despite good nutrition and centuries of exposure. Environmental Health Perspectives, 108, 617.

Vellinger, C., Parant, M., Rousselle, P., Immel, F., Wagner, P., &

Usseglio-Polatera, P. (2012a). Comparison of arsenate and cadmium toxicity in a freshwater amphipod (Gammarus pulex). Environmental Pollution, 160, 66–73.

Vellinger, C., Parant, M., Rousselle, P., & Usseglio-Polatera, P.

(2012b). Antagonistic toxicity of arsenate and cadmium in a freshwater amphipod (Gammarus pulex). Ecotoxicology, 21(7), 1817–1827.

Wang, G., & Fowler, B. A. (2008). Roles of biomarkers in eval- uating interactions among mixtures of lead, cadmium and arsenic. Toxicology and Applied Pharmacology, 233, 92–99.

Wang, B., Feng, L., Jiang, W.-D., Wu, P., Kuang, S.-Y., Jiang, J., Tang, L., Tang, W.-N., Zhang, Y.-A., & Liu, Y. (2015).

Copper-induced tight junction mRNA expression changes, apoptosis and antioxidant responses via NF-κB, TOR and Nrf2 signaling molecules in the gills of fish: preventive role of arginine. Aquatic Toxicology, 158, 125–137.

Wicklund-Glynn, A., Norrgrenb, L., & Mussenera, A. (1994).

Differences in uptake of inorganic mercury and cadmium in the gills of the zebrafish, Brachydanio rerio. Aquatic Toxicology, 30, 13–26.

Wieland, M., Levin, M. K., Hingorani, K. S., Biro, F. N., &

Hingorani, M. M. (2009). Mechanism of cadmium- mediated inhibition of Msh2-Msh6 function in DNA mismatch repair. Biochemistry, 48, 9492–9502.

Winyard, P. G., Moody, C. J., & Jacob, C. (2005). Oxidative activation of antioxidant defence. Trends in Biochemical Sciences, 30, 453–461.

Wood, L. D., Parsons, D. W., Jones, S., Lin, J., Sjöblom, T., Leary, R. J., Shen, D., Boca, S. M., Barber, T., & Ptak, J. (2007). The genomic landscapes of human breast and colorectal cancers.

Science, 318, 1108–1113.

Zhou, Z., Lei, Y., & Wang, C. (2012). Analysis of aberrant methylation in DNA repair genes during malignant transformation of human bronchial epithelial cells induced by cadmium.

Toxicological Sciences, 125, 412–417.

Zona, S., Bella, L., Burton, M. J., de Moraes, G. N., & Lam, E. W.

F. (2014). FOXM1: an emerging master regulator of DNA damage response and genotoxic agent resistance. Biochimica et Biophysica Acta, Gene Regulatory Mechanisms, 1839, 1316–1322.