Phase I and II biotransformation enzymes and polycyclic aromatic hydrocarbons in the Mediterranean mussel (Mytilus galloprovincialis, Lamarck, 1819) collected in front of an oil refinery

Marine Environmental Research 79 (2012) 29e36

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Phase I and II biotransformation enzymes and polycyclic aromatic hydrocarbons in the Mediterranean mussel (Mytilus galloprovincialis, Lamarck, 1819) collected in front of an oil refinery Anna Trisciani a, *, Guido Perra a, Tancredi Caruso b, Silvano Focardi a, Ilaria Corsi a a b

Department of Environmental Sciences “G. Sarfatti”, University of Siena, Via Mattioli 4, 53100 Siena, Italy Freie Universität Berlin, Institut für Biologie e Ökologie der Pflanzen, Berlin, Germany

a r t i c l e i n f o

a b s t r a c t

Article history: Received 12 March 2012 Received in revised form 24 April 2012 Accepted 27 April 2012

The aim of the present study was to investigate the responses of phase I and II biotransformation enzymes and levels of PAHs in the Mediterranean mussel (Mytilus galloprovincialis, Lamarck, 1819) collected from three sites at different distance from an oil refinery. Phase I enzyme activities as NAD(P)Hcyt c red, NADH ferry red, B(a)PMO and phase II as UDPGT, GST were measured in digestive gland while 16 PAHs (US-EPA) in whole soft tissue. An added value to the data obtained in the present study rely on the RDA analysis which showed close correlations between PAHs levels and phase I enzyme activities in mussels collected in front of the refinery. And again a significant spatial correlation between B(a)P levels and NADPH-cyt c red activities was observed using linear models. No differences among sites for B(a) PMO and phase II GST activities were observed, while the application of UDPGT as biomarkers requires further investigation. Ó 2012 Elsevier Ltd. All rights reserved.

Keywords: Hydrocarbons Pollution monitoring Biomarker Cytochrome P450 NADPH cytochrome c reductase GST B(a)P Oil refinery Mussel

1. Introduction The aquatic environment is increasingly threatened by a vast number of hydrocarbon compounds such as petroleum and its derivates, that may enter the marine environment through discharge of industrial and urban effluents, shipping activities, offshore oil production, oil spills, fossil fuel combustion and natural seepage (Medeiros et al., 2005). Polycyclic aromatic hydrocarbons (PAHs) are among the most toxic components of petroleum products (Lima et al., 2007). They released into the marine environment tend to be rapidly absorbed by suspended matter and sediment and become available to fish and other marine organisms through the food chain (Perugini et al., 2007). So PAHs and their metabolites possess the highest toxicity among petroleum products, being mutagenic, cytotoxic and potentially carcinogenic (IARC, 1989). Once released in the water column and sediment they cause a series of harmful effects in both vertebrates and invertebrates species, including genetic damage, immune and endocrine

* Corresponding author. Tel.: þ39 (0) 577 232877; fax: þ39 (0) 577 232806. E-mail address: [email protected] (A. Trisciani). 0141-1136/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.marenvres.2012.04.006

disfunction, malformations, fibrosis and cancer (Aas et al., 2000). Information about their bioaccumulation and the biological responses capable to exert in exposed organisms are therefore essential to assess how dangerous they are in a natural environment and to devise strategies to protect biological resources, including those for human consumption. The exposure of marine organisms to various classes of environmental contaminants including PAHs can be inferred from assays of contaminant-selective induction of enzyme activities. The primary biological system for detoxifying/bioactivating PAHs is the cytochrome P450 (P450) system, which consists of several multigenic families of structurally and functionally related haemproteins (Snyder, 2000). Marine pollution monitoring programs have successfully included biological responses known as biomarkers as tools for assessment of potential toxic effects of pollutants (Den Besten, 1998; Cajaraville et al., 2000). In particular induction of cytochrome P450-dependent monooxygenase enzymes (phase I) has been successfully used as a sensitive biomarker of exposure to organic contaminants including PAHs in marine organisms including molluscs (Livingstone et al., 1997; Porte et al., 2001; Petushock et al., 2002; Serafim et al., 2008). A clear evidence of

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interaction between organic pollutants and the flavoprotein NAD(P)H cytochrome c reductase (NAD(P)H-cyt c red) has been widely described in bivalve molluscs as mussels. In particular increases in digestive gland microsomal NADPH-cyt c red activities have been reported in mussels in both laboratory exposed (diesel oil and B(a)P) (Livingstone, 1987, 1988, 1989; Michel et al., 1993) and in situ studies including other organic contaminants (Michel et al., 1994; Solé et al., 1996, 1998; Peters et al., 1999; Nasci et al., 2002; Lima et al., 2007). Activation of the cytochrome P450 system has been also measured by determining benzo(a)pyrene monooxygenase (B(a) PMO) activity as a biomarker of exposure to PAHs in mussels (Narbonne et al., 1999; Akcha et al., 2000; Solé and Livingstone, 2005; Banni et al., 2010). Michel et al. (1998) reported a positive correlation between B(a)PMO activity in mussels and levels of PAHs in sediment along the north-western Mediterranean coast. Same evidences have been also obtained in other field studies (Peters et al., 1999; Damiens et al., 2007). More often, glutathione Stransferase (GST) activity of the phase II of detoxification system has been widely measured in mussels as a sensitive biomarker of exposure to several classes of pollutant, including PAHs and petrochemical products both in field and laboratory studies (Moreira and Guilhermino, 2005; Rocher et al., 2006; Tim-Tim et al., 2009; Vidal-Liñàn et al., 2010). Therefore, mussels like the Mediterranean Mytilus galloprovincialis (Lamarck, 1781) have been extensively used as bioindicator of marine pollution including monitoring studies of PAHs contamination in marine coastal areas (Goldberg, 1975; D’Adamo et al., 1997; Piccardo et al., 2001; Sureda et al., 2011). Bioaccumulation of organic contaminants by aquatic organisms is a balance between essentially passive processes of uptake and depuration and elimination of xenobiotics via biotransformation pathways (Livingstone, 1991). B(a)P is known to be metabolised by M. galloprovincialis (Michel et al., 1995) but rates of metabolism of PAHs are significantly less than uptake rates in molluscs, resulting in strong bioaccumulation (Livingstone et al., 1998). Mussels take up and concentrate contaminants to levels well above those present in the surrounding seawater and therefore provide information on spatial and temporal pollution trends, enabling identification of ‘hot spots’ in coastal areas (Phillips, 1980). It has been estimated that mussels take up PAHs from suspended particulate five times faster than from water (Ribera et al., 1991). Studies using ‘Prestige’ fuel oil show that mussels accumulate up to 17,033 mg of PAHs per kg dry weight of tissue (Pérez-Cadahìa et al., 2004). Mussels therefore show a series of (sub-lethal) biochemical responses to pollutants (see Livingstone, 1991, for a review) which may be used as early warning signs of exposure (Huggett et al., 1992). The aim of the present field study was to investigate the response of phase I and II biotransformation enzymes and the levels of PAHs in the Mediterranean mussel M. galloprovincialis collected from a marine coastal area located in front of an oil refinery. Enzyme activities, such as NAD(P)H-cyt c red, NADH ferrycianide reductase (NADH ferry red), B(a)PMO, uridine diphosfate glucuronosil transferase (UDPGT) and glutathione-S-transferase (GST), were measured in mussel digestive glands. Levels of the 16 most toxic PAHs (EPA) were measured in whole soft tissue. 2. Materials and methods 2.1. Mussel sampling Specimens of native Mediterranean mussel M. galloprovincialis (Lamarck, 1819) (valve length 5.8  0.2 cm, SD) were collected in November 2007 on a stretch of coast of the north-western Adriatic Sea, extending from 43.65890 N, 13.32977 E to 43.58148 N,

13.57237 E (20 km). The marine coastal area under investigation is located in front of an oil refinery, occupying an area of 70 ha, that processes 3.9 million tons of crude oil per year and produces 2 million MWh/y of electricity. Storage capacity (about 1,500,000 m3) is among the largest in Europe. Three sites were selected based on previous findings on PAH levels in sediments (data not shown) and on the distance from the oil refinery: (1) the high-impact site (HIS) immediately offshore from the refinery (PAHs in sediment in the range 20e500 ng/g d.w.); (2) the moderate-impact site (MIS) (3e10 ng/g d.w.) 5 km north of HIS and the low-impact site (LIS) 10 km south of HIS (3e10 ng/g d.w.). Fifty mussels from each site (HIS; MIS; LIS) were collected from artificial and natural barriers. Half of samples (25 samples per site) were used for analytical determinations of PAHs in whole mussel tissue and stored at 20  C while half were used for enzymatic determination using digestive glands which were dissected, frozen in liquid nitrogen and stored at 80  C. 2.2. PAH analysis The sixteen most toxic PAHs listed by the EPA were determined in 5 pools of 5 mussels each from each site. Tissue (5 g) was extracted (Dionex mod. 200 accelerated solvent extractor, Sunnyvale, USA) according to US-EPA (1996) method 3545A (modified by Perra et al., 2010) and quantified by high-performance liquid chromatography (HPLC) (Waters mod. 474 scanning fluorescence detector (SFD) and mod. 996 photodiode array (PDA) detector, Milford, Massachusetts). PAHs were identified and measured by HPLC. Acenaphthylene was determined with a Waters 996 PDA detector and all other compounds with a Waters 474 SFD. Chromatographic separation was performed on a SupelcosilÔ LC-PAH HPLC chromatographic column (250  4.6 mm i.d., particle size 5 mm, Supelco) with the following mobile phase: acetonitrile:water 60:40 linear gradient for 40 min, then acetonitrile:water 100:0 for 10 min at a flow rate of 1.5 mL min1. Maximum elution time was 50 min. Quantitative analysis was performed using a three-point linear calibration of a PAH solution obtained by dilution of the certified standard mixture TLC 16-PAH mix (Supelco). Linearity was satisfactory, with values of correlation coefficient r > 0.99. Detection limits of the method for each PAH, estimated as 3s (IUPAC criterion), ranged from 0.01 to 0.5 ng/g. A certified reference material, HS-6 harbour sediments, purchased from NRC (Canada), procedural blanks and replicate samples were used for quality control procedures; their reproducibility and recovery were high (70e80%). Precision, evaluated in terms of reproducibility of experimental results (n ¼ 10) for analysis of a real sample and expressed as relative standard deviation, ranged from 4.3% (DBA) to 18.5% (N) and was below 10% in most cases. Content of PAHs were expressed as ng/g dry weight (d.w.). 2.3. Biochemical assays 2.3.1. Isolation of cytosolic and microsomal fractions Cytosolic and microsomal fractions of digestive gland of mussel were obtained by standard procedures using the method of Corsi et al. (2003). Digestive glands were individually homogenized in a 1:4 weight/volume ratio with sucrose buffer (50 mM K2HPO4, 0.75 mM sucrose, 1 mM EDTA, 0.5 mM DTT, 400 mM PMSF, 1 mg/l Aprotinin, 10 mM Leupeptin and 1 mM Pepsatin; pH 7.5), using a Potter-Elvenjem glass/Teflon homogenizer at 2000 rpm. Microsomes were obtained by differential centrifugation in a Sorvall RC28S Ultracentrifuge. Homogenates were first centrifuged at 9000 g for 20 min to remove the nuclei, mitochondria, lysosomes, and cell debris, and the resulting supernatants (cytosolic fractions) were transferred and centrifuged at 100,000 g for 60 min to isolate

A. Trisciani et al. / Marine Environmental Research 79 (2012) 29e36

the microsomes. The microsomal pellets were subsequently transferred and resuspended in a 1:2.6 weight/volume ratio with Tris-base buffer (10 mM Tris [base]), 20% weight/volume glycerol, 0.5 mM DTT, 400 AM PMSF, pH 7.5. All procedures were carried out at 4  C. 2.3.2. Phase I enzymes NAD(P)H-cyt c red activity was assayed in microsomal fraction and measured by the spectrophotometric method of Livingstone and Farrar (1984), using a Shimadzu UV-160A visible recording spectrophotometer. Assay conditions in the reaction mixture (final volume 1 ml) were as follows: pH 7.6, 25  C, in a spectrophotometer cuvette containing 100 mM TriseHCl, 20 mM KCN, 1.2 mM cytochrome c (cyt c) (extinction coefficient of 19.6 mM1 cm1), distilled water, 6 mM NAD(P)H and 25e100 ml microsomal fraction. The reaction was started by adding 50 ml NAD(P)H and the progressive increase in absorbance of the reaction product was recorded at l ¼ 550 nm. Activities were expressed as nmol min1 mg prot1. NADH ferry red activity was assayed in microsomal fraction and measured by the spectrophotometric method of Livingstone and Farrar (1984), using a Shimadzu UV-160A visible recording spectrophotometer. Assay conditions in the reaction mixture (final volume 1 ml) were as follows: pH 7.6, 25  C, in a spectrophotometer cuvette containing 100 mM TriseHCl, 20 mM KCN, 10 mM potassium ferrycianide (extinction coefficient 1.02 mM1 cm1), distilled water, 6 mM NADH and 25 ml microsomal fraction. The reaction was started by adding 50 ml NADH and the progressive increase in absorbance of the reaction product was recorded at l ¼ 420 nm. Activities were expressed as nmol min1 mg prot1. B(a)PMO activity was measured on 12 pools of microsomes (4 per site, 5 individuals each) by the fluorimetric method of Kurelec et al. (1977) using a PerkineElmer LS50B luminescence spectrofluorimeter. Assay conditions in the reaction mixture (final volume 2.25) were as follows: pH 7.5, 30  C, in a fluorimeter cuvette containing 110 mM TriseHCl, 15 mM MgCl2$6H2O, 1.8 mM NADPH and 100 ml pooled microsomal fraction. B(a)P (2 mM) was used as substrate in a 1-h reaction, stopped with cool acetone. The amount of 3 OH-B(a)P produced was read at lEX ¼ 396 nm/lEM ¼ 522 nm, with H2SO4 1 M and quinine sulphate 1 mg/ml as standards. Activity was determined in duplicate and compared to a blank treated with acetone prior to incubation; activity was expressed as units of fluorescence (UF) min1 mg prot1. 2.3.3. Phase II enzymes GST activity was assayed in cytosolic fraction and measured by the spectrophotometric method of Habig et al. (1974), using a Wallac Victor 3 Multilabel Counter mod. 1420 spectrophotometer. Assay conditions in the reaction mixture (final volume 220 ml) were as follows: pH 7.4, 18  C in a microplate containing 1.5 mM 1-chloro-2,4-dinitrobenzene (CDNB, extinction coefficient of 9.6 mM1 cm1) in 0.1 M NaH2PO4/Na2HPO4 buffer, 1.5 mM reduced gluthathione (GSH) and 20 ml diluted cytosolic fractions. The increase in absorbance of the reaction product was measured at 340 nm before and after 2 min of incubation at 18  C. Activities were expressed as nmol mg prot1 min1. UDPGT activity was assayed in microsomal fraction and measured by the fluorimetric method of Collier et al. (2000), using a Wallac Victor 3 Multilabel Counter mod. 1420 spectrofluorimeter at lEX ¼ 355 nm, lEM ¼ 460 nm. Assay conditions in the reaction mixture (final volume 150 ml) were as follows: pH 7.4, 30  C in a microplate containing 0.1 mM 4-methyl-umbelliferon (4-MU) in 0.1 M TriseHCl buffer (containing 5 mM MgCl2 and 0.05% BSA), 2 mM 50 -diphospho-glucuronic acid (UDPGA) and 15 ml microsomal fractions. The progressive decrease in fluorescence was recorded

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after 10 min of incubation at 30  C. A calibration curve with 4-MU (125 mMe1.25 mM) was used as reference for quantification. Activities were expressed as nmol min1 mg prot1. Total proteins were measured according to Bradford (1976) using a Shimadzu UV-160A visible recording spectrometer and bovine serum albumin as standard, reading at 595 nm.

2.4. Statistical analysis The results are presented as means (SD). The comparison of the biochemical parameters among the three sites was performed by analysis of variance (ANOVA, Test of Bonferroni). The normality and homogeneity of variance of data was verified and data transformation was applied as required to fulfil ANOVA assumptions. In addition, the ordination technique of redundancy analysis (RDA) was performed to order samples as a function of biomarkers and using PAHs as constraining, explanatory factors (Legendre and Legendre, 1998). RDA was originally intended to detect relationships between environmental data and species distribution but the method can be used to describe multivariate correlation between any data sets (Legendre and Legendre, 1998). In the present study, biomarkers were here used as “species”, i.e. descriptor variables of samples to be ordered. The permutation procedure (based on 9999 cycles) was used to test the significance of constraints in RDA for all eigenvalues, and additive linear models were used for the multivariate regression (Ter Braak, 1986; Quinn and Keough, 2002; Oksanen et al., 2006). Thus, the model we use was Biomarkers w PAHs. Statistical analyses of data were performed using the software Statistica 7.0 (StatSoft Inc., 2006, USA), with the exception of the RDA that was performed using the software R (version 2.8.1) (http://www.r-project.org).

3. Results Levels of the 16 PAHs measured in whole mussel tissue from the three investigated sites are shown in Table 1. The highest levels were measured in specimens from HIS (56.05  29.37 ng/g d.w.) followed by MIS (32.72  24.64 ng/g d.w.) and LIS (14.05  10.80 ng/g d.w.), though huge variations were observed in all three sites.

Table 1 Levels of 16 most toxic PAHs indicated by EPA (ng/g dry weight) in whole soft tissue from the three sites HIS, MIS and LIS. Results are mean (SD) of 5 pools of five mussels P P from each site. Minus sign indicates below detection limit. 16 PAHs and P450inducers PAHs are also reported. PAHs

HIS

Naphthalene Acenaphthylene Acenaphthene Fluorene Phenanthrene Anthracene Fluoranthene Pyrene Benzo(g,h,i)perylene Benzo(a)anthracene Chrysene Benzo(b)fluoranthene Benzo(k)fluoranthene Benzo(a)pyrene Dibenzo(a,h)anthracene Indeno(c,d)pyrene P 16 PAHs P P450-inducers PAHs

1.92 1.39 3.58 0.80 1.61 0.46 1.72 5.12 7.29 1.56 2.8 4.88 3.31 8.80 4.4 6.75 56.05 39.73

(1.12) (1.72) (3.75) (0.35) (0.75) (0.61) (1.72) (3.04) (4.30) (0.85) (1.47) (3.33) (1.92) (4.84) (4.38) (5.79) (29.37) (22.58)

MIS

LIS

3.48 (5.59) 5.68 (9.40) 3.94 (5.96) <0.05 1.32 (1.62) 0.69 (0.44) 2.85 (3.60) 2.93 (3.04) <0.04 0.27 (0.41) 9.50 (14.57) <0.01 0.96 (1.64) <0.03 0.93 (1.44) <0.03 32.72 (24.64) 11.56 (13.93)

2.3 (3.37) <0.25 1.45 (2.33) 0.27 (0.55) 0.64 (0.48) 0.39 (0.21) 0.60 (0.88) 0.86 (1.26) 3.91 (8.31) 0.26 (0.35) 2.13 (3.39) 0.05 (0.08) 0.36 (0.55) 0.03 (0.01) <0.10 0.45 (1.02) 14.05 (10.80) 7.07 (8.82)

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Table 2 NAD(P)H-cyt c red, NADH ferry red, UDPGT, GST (nmol min1 mg prot1) and B(a) PMO (UF min1 mg prot1) activities in digestive gland of mussel (M. galloprovincialis) (n ¼ 25). Values are means (SD). Significant differences (p < 0.05) between sites are denoted by different superscript letters. Biomarkers

HIS

NADPH-cyt c red NADH-cyt c red NADH ferry red B(a)PMO UDPGT GST

11.82 50.81 435 122 7.46 30.77

MIS (3.57)a (11.99)a (92)a (15) (1.51)a,b (6.21)

6.48 36.74 335 108 9.01 38.18

LIS (0.50)b (2.25)a,b (8)a,b (38) (0.28)a (2.14)

4.70 33.40 283 104 5.73 35.71

(0.75)b (4.21)b (30)b (60) (1.53)b (3.63)

In HIS, the total concentration of eight P450-inducer PAHs (benzo(g,h,i)perylene (B(g,h,i)P), benzo(a)anthracene (B(a)A), benzo(b)fluoranthene (B(b)F), benzo(k)fluoranthene (B(k)F), benzo(a) pyrene (B(a)P), dibenzo(a,h)anthracene (DBA), chrysene (Chry), indeno(c,d)pyrene (IP)) was 3.4 fold than that recorded in MIS and 5.6 fold than LIS. Their range in specimens from HIS was 1.6e8.8 ng/g d.w. with B(a)P > B(g,h,i)P > IP > B(b)F > D(b) A > DBA > B(k)F > Chry > B(a)A. All P450-inducer PAHs were below 1 ng/g d.w. in the other two sites except Chry in MIS and LIS and B(g,h,i)P in LIS. In HIS, high molecular weight PAHs with 4e6 rings were 70% of all PAHs and B(a)P showed the highest levels. Regarding phase I enzymes, significant differences in NADPHcyt c red activities (p < 0.05) were observed in mussels from the three sites, being higher in specimens from HIS than in those from MIS and LIS which resulted very similar (Table 2). The same trend was observed for NADH-cyt c red and NADH ferry red activities (Table 2) with significant differences (p < 0.05) between HIS and LIS but not with MIS. On the contrary, B(a)PMO

activities did not shown differences among the three sites (Table 2). Concerning phase II enzymes, a decreasing trend in UDPGT activities was observed with higher values from MIS than HIS and LIS, but significant differences were found only between MIS and LIS (p < 0.05) (Table 2). No differences were observed in the GST activities among the three sites (Table 2). Regarding RDA, the statistical analysis of mussel content of PAHs and phase I enzyme activities (Fig. 1) showed that the first and second axes explained 36% and 0.5% of the variance, respecP tively. The vectors describing PAHs and B(a)P levels were positively correlated with all three reductases activities (NAD(P)H-cyt c red and NADH ferry red) which were in turn correlated with each other. As shown by the linear models of the spatial distribution of B(a) P level (Fig. 2a) and NADPH-cyt c red activity (Fig. 2b), about 60% of variance was described by the two variables. B(a)P levels in mussels and NADPH-cyt c red activities decreased exponentially with increasing distance from the oil refinery. A slight but significant quadratic effect was observed for NADPH-cyt c red activity which showed an outliner at distances of around 15 km from HIS. The distribution of measured activities on the first axis showed that mussels were discriminated mainly by a negative correlation between reductase activities (phase I; positive score on RDA 1) and the phase II enzyme activities, UDPGT and GST (phase II; negative score on RDA 1) (Fig. 1). Specimens from HIS clustered on the positive part of RDA 1 (higher reductase and lower UDPGT and GST activities) while those from MIS and LIS clustered on the negative part of RDA 1. The permutation test indicated that all the overall RDA model (all eigenvalues) was significant at p < 0.05 as well as the effect of PAHs and B(a)P.

Fig. 1. RDA analysis of phase I and II enzyme activities and the mussel content of

P PAHs and BaP.

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Fig. 2. General linear model for B(a)P. The interpolated line indicates values expected from the model (a). General linear model for NAD(P)H-cyt c red. The interpolated line indicates values expected from the model (b). Legend: DR, distance from refinery (m); B(a)P, levels of benzo(a)pyrene (ng g1 d.w.) in whole mussel tissue; NAD(P)H-cyt c red, enzyme activities in mussel digestive gland (nmol min1 mg prot1).

4. Discussion The aim of the present study was to investigate the responses of phase I and II biotransformation enzymes and levels of PAHs in mussels collected from a petroleum product-contaminated site as an oil refinery on north-western Adriatic Sea. Specimens collected from the closest oil-refinery site (HIS) showed significant higher levels of PAHs than those from the two more distant sites, MIS and LIS (Table 1). PAHs levels detected in native mussels reflected the gradient of petroleum products contamination from HIS to MIS and LIS according to previous findings reported in other studies (Della Torre et al., 2010; Trisciani et al., 2011). The two calculated PAHs distribution indexes as phenanthrene/anthracene (<10) and fluoranthene/pyrene (<1) ratios on mussel’s levels detected suggest that mussels have been exposed to petrogenic and pyrogenic sources. In HIS there was a prevalence (70%) of high molecular weight PAHs (4e6 rings) which are known to be more toxic to aquatic organisms (Solé et al., 1996). Since they are fairly stable and mostly of pyrogenic origin, it is likely that they might reflect the chronic contamination related to the presence of the oil refinery. Looking at B(a)P levels those detected in mussels from HIS resulted the highest compared to those in specimens from MIS and LIS which both were below the detection limit. The highest levels observed in HIS seem in agreement with the assumption that despite in marine invertebrates B(a)P is rapidly metabolized, in molluscs rates of metabolism are generally less than those of uptake. This could explain what observed in the present study in a chronic exposure condition as in front of an oil refinery (Michel et al., 1995; Livingstone et al., 1998). Similar results in terms of PAH levels body burden have been reported both in the same species as in the blue mussel Mytilus edulis from other polluted marine coastal areas: the Galician coast near “La Coruña” (NW Spain) polluted by hydrocarbons (Solé et al., 1996; Porte et al., 2001) and the Gulf of Rijeka (Croatia) characterized by heavy industrial loads (Bihari et al., 2007). In our study, the pattern observed in levels of PAHs in wild mussels in all three sites was significantly reflected by phase I enzymes; in particular by NADPH-cyt c red activities and less for

NADH-cyt c red and NADH ferry activities (Table 2). Nevertheless, to date most of field studies provide contradictory results despite the sensitivity of NADPH-cyt c red to PAHs and in particular B(a)P exposure has been already reported in several laboratory studies (Livingstone, 1988; Michel et al., 1993; Okay et al., 2000). Linking the exposure to PAHs with biomarker responses in the field is related to multi-factorial interaction of different environmental and physiological variables (Hagger et al., 2010). No changes in NADPH-cyt c red activity were observed in specimens of M. edulis collected along a PAH-gradient caused by an oil spill off the Galician coast (Solé et al., 1996). Significant induction of NADPHcyt c red activity was observed by other authors in marine coastal areas heavily contaminated with PAHs and PCBs (Nasci et al., 2002; Serafim et al., 2008). In general different trends of NAD(P)H-cyt c red and NADH ferry red were reported in field and laboratory studies (Livingstone et al., 1985; Petushock et al., 2002): fewer and less marked changes were observed in NADH ferry red and NADH-cyt c red activities compared to NADPH-cyt c red (Livingstone, 1987). In the present study the sensitivity of all three reductases (NAD(P)H-cyt c red, NADH ferry red) to PAHs exposure was explained by RDA (Fig. 1). A positive correlation with total PAHs and even with only B(a)P levels was in fact observed not only with all three reductases but also among them (Fig. 1). In particular a significant spatial correlations between phase I enzymes (NADPHcyt c) and B(a)P levels is evident by linear models: both decreased exponentially with increasing distance from the refinery (Fig. 2). The slight but significant quadratic effect observed for NADPH-cyt c red activity which rose around 15 km might be explained by several factors both abiotic affecting bioavailability of the pollutant and biotic driving different biological responses in the exposed organism (Fig. 2). The observed outlier at 15 km of distance might probably reflect the biological variability of responses to environmental stressors of natural populations (native mussels) as in the present study. The potential exposure to other classes of environmental pollutants as PCBs or other alogenated compounds able to affect NADPH-cyt c red activity in mussels could not be also excluded (Nasci et al., 2002; Hagger et al., 2010).

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On the opposite, still regarding phase I enzymes, B(a)PMO activity did not change among sites except in specimens from HIS which resulted higher. Generally, in in vivo studies, B(a)PMO in mussels has been described as the most sensitive marker to PAHs exposure and in particular to B(a)P (Akcha et al., 2000; Banni et al., 2010). Also in situ, our previous findings showed that DNA adduct levels resulted high correlated with B(a)PMO activity in digestive gland of mussels from a high PAHs contaminated marine coastal area confirming the existence of activation pathways in the Mediterranean mussel by the involvement of the P450 system (Pisoni et al., 2004). Other studies showed contradictory results, for instance, no changes in B(a)PMO activity were observed in Mytilus sp. related to exposure to aliphatic compounds and/or PAHs and along a PAH-gradient caused by an oil spill (Solé et al., 1996; Peters et al., 1999; Porte et al., 2001). In some studies it’s postulated that despite the proven capacity of M. galloprovincialis to metabolize B(a)P, part of this metabolism is constitutive, explaining why the enzyme B(a)PMO is difficult to induce; intraspecies variations and interactions with environmental parameters can also be added (Michel et al., 1995; Livingstone et al., 1997). Regarding phase II enzymes, in agreement with previous findings from field studies, GST activities show lower sensitivity to PAHs exposure compared to phase I (Table 2) (Canesi et al., 1999; Gowland et al., 2000; Bebianno et al., 2007; Tsangaris et al., 2010). No significant changes in GST activity were found in digestive gland of M. edulis collected along a PAH-gradient caused by an oil spill (Solé et al., 1996) and in M. galloprovincialis exposed to petrochemical contamination off the north-western coast of Portugal (Lima et al., 2007). The latter authors hypothesized that toxic intermediates produced in the digestive gland during metabolism may inactivate the enzyme, resulting in reduced GST activity, as found by Cheung et al. (2001) in Perna viridis. No correlation was found between PAHs and GST activities in whole soft tissues of mussels (M. galloprovincialis) transplanted in the Bay of Cannes (NW Mediterranean Sea) (Damiens et al., 2007) or between PAHs and GST activities in gills of mussels of the same species collected along the Mediterranean coast of Spain (Fernàndez et al., 2010). The lack of significant GST responses could be due to the use of the substrate CDNB, which gives global GST activity, whereas activities determined with other substrates may evoke isoforms induced in response to different pollutants (Fitzpatrick et al., 1995; Hoarau et al., 2001). Recent studies carried out with M. edulis showed that several GST isoforms with various binding properties may be induced by different types of pollutants (Fitzpatrick et al., 1997; Vidal-Liñàn et al., 2010). In addition, counteractive GST responses to prooxidant challenge by environmental pollutants in Mediterranean caged mussels have been reported by Bocchetti et al. (2008a). They hypothesised that GST responses can be overwhelmed by longer periods (chronic exposure) or higher intensity of chemical exposure when perturbation exceeds the efficiency of specific antioxidant defences as probably occur in chronic exposure conditions (oil-refinery areas). Finally, the natural influence of biotic and abiotic factors may have a role in the low sensitivity to PAHs in the field (Livingstone, 1988; Regoli, 1998; Bocchetti et al., 2008b; Gorbi et al., 2008). Availability of nutrients, temperature and reproductive status are the most common factors influencing natural fluctuations of antioxidant defences in Mediterranean mussels (Bocchetti and Regoli, 2006). Concerning UDPGT, a different trend was observed with significantly higher activities in specimens from MIS than LIS and HIS (p < 0.05) (Table 2). The only data available is from Livingstone et al. (1995) who reported similar UDPGT activities as those reported in LIS in specimens from Venice Lagoon (3.8 nmol min1 mg prot1). A potential involvement of UDPGTactivity in PAHs metabolism might be hypothesized but need further investigations both in vivo and in vitro.

5. Conclusions A significant spatial correlation between a biological response and a pollutant in a marine coastal area has been observed in the present study based on RDA analysis and linear models. The significant correlations between the three reductase activities and levels of total PAHs and in particular B(a)P evidence for the first time a strong relationship P450 enzymes and PAHs exposure in mussels in the field; B(a)P levels and NADPH-cyt c red activities decreased exponentially with increasing distance from the refinery reflecting a decreasing pattern of PAHs contamination (HIS). The lack of significant induction of B(a)PMO activity and GST seems to reveal the unsuitability of both markers for PAHs monitoring with mussels. Finally, the different trend observed for UDPGT activities requires further investigation in vivo and in situ to fully assess their suitability for monitoring petroleum hydrocarbon contamination in the field. Acknowledgements The authors are grateful to Dr. Camilla Della Torre for technical assistance during the laboratory analysis and Dr. Valerio Volpi, Dr. Wanda De Stefano, Dr. Angela Pastore and Costanza Burroni and Alessio Moroni for sampling campaign.

References Aas, E., Baussant, T., Balk, L., Liewenborg, B., Andersen, O.K., 2000. PAH metabolites in bile, cytochrome P4501A and DNA adducts as environmental risk parameters for chronic oil exposure: a laboratory experiment with Atlantic cod. Aquat. Toxicol. 51, 241e258. Akcha, F., Izuel, C., Venier, P., Budzinski, H., Burgeot, T., Narbonne, J.-F., 2000. Enzymatic biomarker measurement and study of DNA adduct formation in benzo[a]pyrene-contaminated mussels, Mytilus galloprovincialis. Aquat. Toxicol. 49, 269e287. Banni, M., Negri, A., Dagnino, A., Jebali, J., Ameur, S., Boussetta, H., 2010. Acute effects of benzo[a]pyrene on digestive gland enzymatic biomarkers and DNA damage on mussel Mytilus galloprovincialis. Ecotoxicol. Environ. Saf. 73, 842e848. Bebianno, M.J., Lopes, B., Guerra, L., Hoarau, P., Ferreira, A.M., 2007. Glutathione Stransferases and cytochrome P450 activities in Mytilus galloprovincialis from the South coast of Portugal: effect of abiotic factors. Environ. Int. 33, 550e558. Bihari, N., Fafandel, M., Piskur, V., 2007. Polyciclic aromatic hydrocarbons and ecotoxicological characterization of seawater, sediment, and mussel M. galloprovincialis from the gulf of Rijeka, the Adriatic Sea, Croatia. Arch. Environ. Contam. Toxicol. 52, 379e387. Bocchetti, R., Regoli, F., 2006. Seasonal variability of oxidative biomarkers, lysosomal parameters, metallothioneins and peroxisomal enzymes in the Mediterranean mussel Mytilus galloprovincialis from Adriatic Sea. Chemosphere 65, 913e921. Bocchetti, R., Fattorini, D., Pisanelli, B., Macchia, S., Oliviero, L., Pilato, F., Pellegrini, D., Regoli, F., 2008a. Contaminant accumulation and biomarker responses in caged mussels, Mytilus galloprovincialis, to evaluate bioavailability and toxicological effects of remobilized chemicals during dredging and disposal operations in harbour areas. Aquat. Toxicol. 89, 257e266. Bocchetti, R., Lamberti, C.V., Pisanelli, B., Razzetti, E.M., Maggi, C., Catalano, B., Sesta, G., Martuccio, G., Gabellini, M., Regoli, F., 2008b. Seasonal variations of exposure biomarkers, oxidative stress responses and cell damage in the clams, Tapes philippinarum, and mussels, Mytilus galloprovincialis, from Adriatic sea. Mar. Environ. Res. 66, 24e26. Bradford, M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248e254. Cajaraville, M.P., Bebianno, M.J., Blasco, J., Porte, C., Sarasquete, C., Viarengo, A., 2000. The use of biomarkers to assess the impact of pollution in coastal environmental on the Iberian Peninsula: a practical approach. Sci. Total Environ. 247, 295e311. Canesi, L., Viarengo, A., Leonzio, C., Filippelli, M., Gallo, G., 1999. Heavy metals and glutathine metabolism in mussel tissues. Aquat. Toxicol. 46, 67e76. Cheung, C.C.C., Zheng, G.J., Li, A.M.Y., Richardson, B.J., Lam, P.K.S., 2001. Relationships between tissue concentrations of polycyclic aromatic hydrocarbons and antioxidative responses of marine mussels, Perna viridis. Aquat. Toxicol. 52, 189e203. Collier, A.C., Tingle, M.D., Keelan, J.A., Paxton, J.W., Mitchell, M.D., 2000. A highly sensitive fluorescent microplate method for the determination of

A. Trisciani et al. / Marine Environmental Research 79 (2012) 29e36 UDP-glucoronosyl transferase activity in tissues and placental cell lines. Drug Metab. Dispos. 28, 1184e1186. Corsi, I., Mariottini, M., Sensini, C., Lancini, L., Focardi, S., 2003. Cytochrome P450, acetylcholinesterase and gonadal histology for evaluating contaminant exposure levels in fishes from a highly eutrophic brackish ecosystem: Orbetello Lagoon, Italy. Mar. Pollut. Bull. 46, 203e212. D’Adamo, R., Pelosi, S., Trotta, P., Sansone, G., 1997. Bioaccumulation and biomagnification of polycyclic aromatic hydrocarbons in aquatic organisms. Mar. Chem. 56, 45e49. Damiens, G., Gnassia-Barelli, M., Loquès, F., Roméo, M., Salbert, V., 2007. Integrated biomarker response index as a useful tool for environmental assessment evaluated using transplanted mussels. Chemosphere 66, 574e583. Della Torre, C., Corsi, I., Nardi, F., Perra, G., Tomasino, M.P., Focardi, S., 2010. Transcriptional and post-tanscriptional response of drug-metabolizing enzymes to PAHs contamination in red mullet (Mullus barbatus, Linnaeus 1758): a field study. Mar. Environ. Res. 70, 95e101. Den Besten, P.J., 1998. Concepts of the implementation of biomarkers in environmental monitoring. Mar. Environ. Res. 46, 253e256. Fernàndez, B., Campillo, J.A., Martìnez-Gòmez, C., Benedicto, J., 2010. Antioxidant responses in gills of mussel (Mytilus galloprovincialis) as biomarkers of environmental stress along the Spanish Mediterranean coast. Aquat. Toxicol. 99, 186e197. Fitzpatrick, P.J., Sheehan, D., Livingstone, D.R., 1995. Studies on isoenzymes of glutathione S-transferase in the digestive gland of Mytilus galloprovincialis with exposure to pollution. Mar. Environ. Res. 39, 241e244. Fitzpatrick, P.J., O’Halloran, J., Sheehan, D., Walsh, A.R., 1997. Assessment of a glutathione S-transferase and related proteins in the gill and digestive gland of Mytilus edulis, as a potential organic pollution biomarkers. Biomarkers 2, 51e56. Goldberg, E.D., 1975. The mussel watch: a first step in global marine monitoring. Mar. Pollut. Bull. 6, 111e132. Gorbi, S., Lamberti, C.V., Notti, A., Benedetti, M., Fattorini, D., Moltedo, G., Regoli, F., 2008. An ecotoxicological protocol with caged mussels, Mytilus galloprovinicialis, for monitoring the impact of an offshore platform in the Adriatic Sea. Mar. Environ. Res. 65, 34e49. Gowland, B.T.G., McIntosh, A.D., Davies, I.M., Moffat, C.F., Webster, L., 2000. Implication from field study regarding the relationship between polycyclic aromatic hydrocarbons and glutathione S-transferase activity in mussels. Mar. Environ. Res. 54, 231e235. Habig, W.H., Pabst, M.J., Jakoby, W.B., 1974. Glutathione S-transferases. The first enzymatic step in mercapturic acid formation. J. Biol. Chem. 249, 7130e7139. Hagger, J.A., Lowe, D., Dissanayake, A., Jones, M.B., Galloway, T.S., 2010. The influence of seasonality on biomarker responses in Mytilus edulis. Ecotoxicology 19, 953e962. Hoarau, P., Gnassia-Barelli, M., Roméo, M., Girard, J.P., 2001. Differential induction of GST in the clams Ruditapes decussatus. exposed to organic compounds. Environ. Toxicol. Chem. 20, 523e529. Huggett, R.J., Kimerle, K.A., Mehrle, J.P.M., Bergman, H.L. (Eds.), 1992. Biomarkers: Biochemical, Physiological and Histological Markers of Anthropogenic Stress. Lewis Publishers, Boca Raton, FL. IARC, 1989. IARC Monographs on the Evaluation of Carcinogenic Risk of Chemicals to Humans. Diesel and Gasoline Engine Exhausts and Some Nitroarenes, vol. 46. Lyon, International Agency for Research of Cancer, France. Kurelec, B., Britvic, S., Rijavec, M., Muller, W.E.G., Zahn, R.K., 1977. Benzo(a)pyrene monooxygenase induction in marine fish. Molecular response to oil pollution. Mar. Biol. 44, 211e216. Legendre, P., Legendre, L.L., 1998. Numerical Ecology, second english ed. Elsevier Sciences B.V., Amsterdam, 853 pp. Lima, I., Moreira, S.M., Rendòn-Von Osten, J., Soares, A.M.V.M., Guilhermino, L., 2007. Biochemical responses of the marine mussel Mytilus galloprovincialis to petrochemical environmental contamination along the North-western coast of Portugal. Chemosphere 66, 1230e1242. Livingstone, D.R., Farrar, S.V., 1984. Tissue and subcellular distribution of enzyme activities of mixed-function oxygenase and benzo[a]pyrene metabolism in the common mussel Mytilus edulis L. Sci. Total Environ. 39, 209e235. Livingstone, D.R., Moore, M.N., Lowe, D.M., Nasci, C., Farrar, S., 1985. Responses of the cytochrome P-450 monooxygenase system to diesel oil in the common mussel, Mytilus edulis l., and the periwinkle, Littorina littorea L. Aquat. Toxicol. 7, 79e91. Livingstone, D.R., Kirchin, M.A., Wiseman, A., 1989. Cytochrome P450 and oxidative metabolism in molluscs. Xenobiotica 19, 1041e1062. Livingstone, D.R., Lemaire, P., Matthewes, A., Peters, L.D., Porte, C., Fitzpatrick, P.J., Forlin, L., Nasci, C., Fossato, V., Wootton, N., Goldfarb, P., 1995. Assessment of the impact of organic pollutants on goby (Zosterisessor ophiocephalus) and mussel (Mytilus galloprovincialis) from the Venice Lagoon, Italy: biochemical studies. Mar. Environ. Res. 39, 235e240. Livingstone, D.R., Nasci, C., Solé, M., Da Ros, L., O’Hara, S.C.M., Peters, L.D., Fossato, V., Wootton, A.N., Goldfarb, P.S., 1997. Apparent induction of a cytochrome P450 with immunochemical similarities to CYP1A in digestive gland of the common mussel (Mytilus galloprovincialis L.) with exposure to 2,20 3,4,40 ,50 -hexachlorobiphenyl and Aroclor 1254. Aquat. Toxicol. 38, 205e224. Livingstone, D.R., Chipman, J.K., Lowe, D.M., Minier, C., Mitchelmore, C.L., Moore, M.N., Peters, L.D., Pipe, R.K., 1998. Development of biomarkers to detect the effects of organic pollution on aquatic invertebrates: recent molecular,

35

genotoxic, cellular and immunological studies on the common mussel (Mytilus edulis L.) and other mytilids. Int. J. Environ. Pollut. 13, 1e6. Livingstone, D.R., 1987. Seasonal responses to diesel oil and subsequent recovery of the cytochrome P450 monooxygenase system in the common mussel, Mytilus edulis L., and the periwinkle, Littorina littorea L. Sci. Total Environ. 65, 3e20. Livingstone, D.R., 1988. Responses of microsomal NADPH-cytochrome c reductase activity and cytochrome P-450 in digestive glands of Mytilus edulis and Littorina littorea to environmental and experimental exposure to pollutants. Mar. Ecol. Prog. Ser. 46, 37e43. Livingstone, D.R., 1991. Organic xenobiotic metabolism in marine invertebrates. Adv. Comp. Environ. Phys. 7, 45e185. Medeiros, P.M., Bìcego, M.C., Castelao, R.M., Del Rosso, C., Fillmann, G., Zamboni, A.J., 2005. Natural and anthropogenic hydrocarbon inputs to sediments of Patos Lagoon Estuary. Brazil. Environ. Int. 31, 77e87. Michel, X.R., Suteau, P., Robertson, L.W., Narbonne, J.F., 1993. Effects of benzo(a) pyrene, 3,30 ,4,40 -tetrachlorobiphenyl and 2,20 4,40 ,5,50 -hexachlorobiphenyl on the xenobiotic-metabolizing enzymes in the mussel (Mytilus galloprovincialis). Aquat. Toxicol. 27, 335e344. Michel, X.R., Salaun, J.P., Galgani, F., Narbonne, J.F., 1994. Benzo[a]pyrene hydroxylase activity in the marine mussel Mytilus galloprovincialis: a potential marker of contamination by polycyclic aromatic hydrocarbon type compounds. Mar. Environ. Res. 38, 257e273. Michel, X.R., Beasse, C., Narbonne, J.F., 1995. In vivo metabolism of benzo(a)pyrene in the mussel, Mytilus galloprovincialis. Arch. Environ. Contam. Toxicol. 28, 215e222. Michel, M., Narbonne, J.F., Mora, P., Daubèze, M., Ribera, D., Lafaurie, M., Budzinski, H., Garrigues, P., 1998. Indicateurs biochimiques de pollution des écosystèmes marins (GICBEM). In: Lagadic, L., Caquet, T., Amiard, J.-C., Ramade, F. (Eds.), Utilisation de biomarqueurs pour la surveillance de la qualité de l’environnement. Lavoisier TEC&DOC, Paris, pp. 9e32. Moreira, S.M., Guilhermino, L., 2005. The use of Mytilus galloprovincialis acetylcholinesterase and glutathione S-transferases activities as biomarkers of environmental contamination along the northwest portuguese coast. Environ. Monit. Assess. 105, 309e325. Narbonne, J.F., Daubeze, M., Clerandeau, C., Garrigues, P., 1999. Scale of classification based on biochemical markers in mussels: application to pollution monitoring in European coasts. Biomarkers 4, 415e424. Nasci, C., Nesto, N., Monteduro, R.A., Da Ros, L., 2002. Field application of biochemical markers and a physiological index in the mussel, Mytilus galloprovincialis: transplantation and biomonitoring studies in the lagoon of venice (NE Italy). Mar. Environ. Res. 54, 811e816. Okay, O.S., Donkin, P., Peters, L.D., Livingstone, D.R., 2000. The role of algae (Isochrysis galbana) enrichment on the bioaccumulation of benzo(a)pyrene and its effects on the blue mussel Mytilus edulis. Environ. Pollut. 110, 103e113. Oksanen, J., Kindt, R., Legendre, P., O’Hara, R.B., 2006. Vegan: Community Ecology Package Version 1.8-2. http://cran.r-project.org/. Pérez-Cadahìa, B., Laffon, B., Pàsaro, E., Méndez, J., 2004. Evaluation of PAH bioaccumulation and DNA damage in mussels (Mytilus galloprovincialis) exposed to spill Prestige crude oil. Comp. Biochem. Physiol. 138C, 453e460. Perra, G., Renzi, M., Guerranti, C., Focardi, S.E., 2010. Polycyclic aromatic hydrocarbon pollution in sediments: distribution and sources in a lagoon system (Orbetello, Central Italy). Transit. Waters Bull. 38, 45e58. Perugini, M., Visciano, P., Giammarino, A., Manera, M., Di Nardo, W., Amorena, M., 2007. Polycyclic aromatic hydrocarbons in marine organism from the Adriatic Sea, Italy. Chemosphere 66, 1904e1910. Peters, L.D., Shaw, J.P., Nott, M., O’hara, S.C.M., Livingstone, D.R., 1999. Development of cytochrome P450 as a biomarker of organic pollution in Mytilus sp.: field studies in United Kingdom (‘Sea Empress’ oil spill) and the Mediterranean Sea. Biomarkers 4, 425e441. Petushock, N., Gabryelak, T., Palecz, D., Zavodnik, L., Varga, S., Deér, K.A., 2002. Comparative study of the xenobiotic metabolising system in the digestive gland of the bivalve molluscs in different aquatic ecosystems and in aquaria experiments. Aquat. Toxicol. 61, 65e72. Phillips, D.J.H., 1980. Quantitative Aquatic Biological Indicators. Applied Science Publishers, London. Piccardo, M.T., Coradeghini, R., Valerio, F., 2001. Polycyclic aromatic hydrocarbon pollution in native and caged mussels. Mar. Pollut. Bull. 42, 951e956. Pisoni, M., Cogotzi, L., Frigeri, A., Corsi, I., Bonacci, S., Iacocca, A., Lancini, L., Mastrototaro, F., Focardi, S., Svelto, M., 2004. DNA adducts, benzo(a)pyrene monooxygenase activity, and lysosomal membrane stability in Mytilus galloprovincialis from different areas in Taranto coastal waters (Italy). Environ. Res. 96, 163e175. Porte, C., Biosca, X., Solè, M., Albaigés, J., 2001. The integrated use of chemical analysis, cytochrome P450 and stress proteins in mussels to assess pollution along the Galician coast (NW Spain). Environ. Pollut. 112, 261e268. Quinn, G.P., Keough, M.J., 2002. Experimental Design and Data Analysis for Biologists. Cambridge University Press, Cambridge, UK, 537 pp. Regoli, F., 1998. Trace metals and antioxidant enzymes in gills and digestive gland of the Mediterranean mussel Mytilus galloprovincialis. Arch. Environ. Contam. Toxicol. 34, 48e63. Ribera, D., Narbonne, J.F., Michel, X.R., Livingstone, D.R., O’Hara, S., 1991. Responses of antioxidants and lipid peroxidation in mussels to oxidative damage exposure. Comp. Biochem. Physiol. 100C, 177e181. Rocher, B., Goff, J.L., Peluhet, L., Briand, M., Manduzio, H., Gallois, J., Devier, M.H., Geffard, O., Gricourt, L., Augagner, S., Budzinski, H., Pottier, D., André, V.,

36

A. Trisciani et al. / Marine Environmental Research 79 (2012) 29e36

Labailly, P., Cachot, J., 2006. Genotoxicant accumulation and cellular defence activation in bivalves chronically exposed to waterborne contaminants from the Seine River. Aquat. Toxicol. 79, 65e77. Serafim, A., Lopes, B., Company, R., Ferreira, A.M., Bebianno, M.J., 2008. Comparative petroleum hydrocarbons levels and biochemical responses in mussels from hydrothermal vents (Bathymodiolus azoricus) and coastal environments (Mytilus galloprovincialis). Mar. Pollut. Bull. 57, 529e537. Snyder, M.J., 2000. Cytochrome P450 enzymes in aquatic invertebrates recent advances and future directions. Aquat. Toxicol. 48, 529e547. Solé, M., Livingstone, D.R., 2005. Components of the cytochrome P450-dependent monoooxygenase and ‘NADPH-independent benzo[a]pyrene hydroxylase’ activity in a wide range of marine invertebrates species. Comp. Biochem. Physiol. 141C, 20e31. Solé, M., Porte, C., Biosca, X., Mitchelmore, C.L., Chipman, J.K., Livingstone, D.R., Albaigés, J., 1996. Effects of the “Aegean Sea” oil spill on biotransformation enzymes, oxidative stress and DNA-adducts in digestive gland of the mussel (Mytilus edulis L.). Comp. Biochem. Physiol. 113C, 257e265. Solé, M., Peters, L.D., Magnusson, K., Sjolin, A., Granmo, A., Livingstone, D.R., 1998. Responses of the cytochrome P450-dependent monooxygenase and other protective enzyme systems in digestive gland of transplanted common mussel (Mytilus edulis L.) to organic contaminants in the Skagerrak and Kattegat. Biomarkers 3, 49e62.

Sureda, A., Box, A., Tejada, S., Blanco, A., Caixach, J., Deudero, S., 2011. Biochemical responses of Mytilus galloprovincialis as biomarkers of acute environmental pollution caused by the Don Pedro oil spill (Eivissa Island, Spain). Aquat. Toxicol. 101, 540e549. Ter Braak, C.J.F., 1986. Canonical Correspondence Analysis: a new eigenvector technique for multivariate direct gradient analysis. Ecology 67, 1167e1179. Tim-Tim, A.L.S., Morgado, F., Moreira, S., Rangel, R., Nogueira, A.J.A., Soares, S.M.V.M., Guilhermino, L., 2009. Cholinesterase and glutathione S-transferase activities of three mollusc species from the NW Portuguese coast in relation to the ‘Prestige’ oil spill. Chemosphere 77, 1465e1475. Trisciani, A., Corsi, I., Della Torre, C., Perra, G., Focardi, F., 2011. Hepatic biotransformation genes and enzymes and PAH metabolites in bile of common sole (Solea solea, Linnaeus 1758) from an oil-contaminated site in the Mediterranean Sea. A field study. Mar. Pollut. Bull. 62, 806e814. Tsangaris, C., Kormas, K., Strogyloudi, E., Hatzianestis, I., Neofitou, C., Andral, B., Galgani, F., 2010. Multiple biomarkers of pollution effects in caged mussels on the Greek coastline. Comp. Biochem. Physiol. 151C, 369e378. US-EPA, 1996. US EPA Method 3545B. Pressurized Fluid Extraction (PFE). Office of Water, Washington, D.C. Vidal-Liñàn, L., Bellas, J., Campillo, J.A., Beiras, R., 2010. Integrated use of antioxidant enzymes in mussels, Mytilus galloprovincialis, for monitoring pollution in highly productive coastal areas of Galicia (NW Spain). Chemosphere 78, 265e272.