Toxic effects of the antihistamine cetirizine in mussel Mytilus galloprovincialis

Water Research 114 (2017) 316e326

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Toxic effects of the antihistamine cetirizine in mussel Mytilus galloprovincialis ^ ^nia Calisto a, Valdemar I. Esteves a, Miguel Teixeira a, Angela Almeida b, Va Rudolf J. Schneider c, Frederick J. Wrona d, Amadeu M.V.M. Soares b, Etelvina Figueira b, Rosa Freitas b, * a

Department of Chemistry & CESAM, University of Aveiro, 3810-193 Aveiro, Portugal Department of Biology & CESAM, University of Aveiro, 3810-193 Aveiro, Portugal BAM Federal Institute for Materials Research and Testing, Richard-Willstaetter-Str. 11, Berlin, Germany d Department of Geography, University of Victoria, National Water Research Institute, STN CSC, Victoria, BC, Canada b c

a r t i c l e i n f o

a b s t r a c t

Article history: Received 28 October 2016 Received in revised form 4 January 2017 Accepted 14 February 2017

Recent studies have become increasingly focused on the assessment of pharmaceuticals occurrence in aquatic ecosystems, however the potential toxicity to non-target organisms is still largely unknown. The antihistamine cetirizine is a commonly used pharmaceutical, already detected in surface waters of marine aquatic systems worldwide. In the present study Mytilus galloprovincialis mussels were exposed to a range of cetirizine concentrations (0.3, 3.0, 6.0 and 12.0 mg/L), resembling moderate to highly contaminated areas, over 28 days. The responses of different biochemical markers were evaluated in mussels whole soft tissue, and included energy-related parameters (glycogen content, GLY; protein content, PROT; electron transport system activity, ETS), and oxidative stress markers (superoxide dismutase activity, SOD; catalase activity, CAT; glutathione S-transferases activity, GSTs; lipid peroxidation levels, LPO; reduced (GSH) and oxidized (GSSG) glutathione content). The results obtained demonstrated that with the increase of exposure concentrations mussels tended to increase their energy reserves and maintain their metabolic potential, which was significantly higher only at the highest concentration. Our findings clearly revealed that cetirizine inhibited the activity of GSTs and although induced the activity of antioxidant enzymes (SOD and CAT) mussels were not able to prevent cellular damages observed through the increase of LPO associated to the increase of exposure concentrations. Thus, this study confirmed that cetirizine induces toxic effects in Mytilus galloprovincialis, which, considering their trophic relevance, wide use as bioindicator and wide spatial distribution of this species, can result in ecological and economic negative impacts at a large scale. © 2017 Elsevier Ltd. All rights reserved.

Keywords: Bivalves Biomarkers Oxidative stress Metabolic capacity Pharmaceuticals

1. Introduction The increasing consumption of pharmaceuticals by an exponentially growing human population has resulted in ubiquity of these compounds in the environment (e.g. Fent et al., 2006; Kümmerer, 2010; Nikolaou et al., 2007; Puckowski et al., 2016). Furthermore, due to their incomplete removal in Wastewater Treatment Plants (WWTPs), which may only reach 10% for some substances, pharmaceuticals are continuously introduced into

* Corresponding author. Departamento de Biologia & CESAM, Universidade de rio de Santiago, 3810-193 Aveiro, Portugal. Aveiro, Campus Universita E-mail address: [email protected] (R. Freitas). http://dx.doi.org/10.1016/j.watres.2017.02.032 0043-1354/© 2017 Elsevier Ltd. All rights reserved.

aquatic environment (Voulvoulis et al., 2016). This fact, associated to their environmental persistence, may explain the detected concentrations of pharmaceuticals in the environment which range from ng L
1 to mg L
1 (for review see, Fent et al., 2006; Kümmerer, 2009; Santos et al., 2010). For these reasons, and because pharmaceuticals may preserve their biological activity in the environment (Huerta et al., 2012) with potential impacts to aquatic wildlife, over the last years increasing attention has been given to understand the impacts of these contaminants in aquatic ecosystems, namely on the inhabiting organisms (among others, AguirreMartínez et al., 2013; Almeida et al., 2014; Canesi et al., 2007; Freitas et al., 2016, 2015a,b; Martin-Diaz et al., 2009a,b; Pires et al., 2016; Quinn et al., 2011). Several studies have demonstrated that different pharmaceuticals accumulate and cause toxic effects

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 ~ oz et al., in vertebrates (mainly fish, Abreu et al., 2016; Alvarez-Mu n 2015; Carlsson et al., 2013; Corcoran et al., 2010; Watanabe et al., 2016), and in invertebrates (mainly bivalves, Aguirre-Martínez  ~ oz et al., 2015; et al., 2013; Almeida et al., 2015; Alvarez-Mu n Canesi et al., 2007; Contardo-Jara et al., 2011; Freitas et al., 2016; 2015a,b; Gonzalez-Rey and Bebianno, 2014; Martin-Diaz et al., 2009a,b; Pires et al., 2016; Quinn et al., 2011), when these organisms are exposed to pharmaceuticals under laboratory conditions or to wastewater effluent and surface waters receiving effluent discharges in the field. The effects of pharmaceuticals exposure on freshwater and marine bivalves include impacts at cellular level with increased oxidative stress, embriotoxicity and immunotoxicity (AguirreMartínez et al., 2013; Binelli et al., 2009; Canesi et al., 2007;  et al., 2006; Contardo-Jara et al., 2011; Fabbri et al., 2014; Gagne Martin-Diaz et al., 2009a,b; Matozzo et al., 2012; Munari et al., 2016; Parolini et al., 2011; Tsiaka et al., 2013). To assess the toxic impacts of pharmaceuticals a wide variety of bivalve species has been used, including mussels, which are useful bioindicator species that combine a wide distribution and long-life cycle with their filtration ability allowing bioaccumulation of contaminants from the surrounding environment (among others, Cajaraville et al.,  et al., 2007; Cravo et al., 2009; Ericson et al., 2010 in 2000; Gagne Gonzalez-Rey and Bebianno, 2014). Martin-Diaz et al. (2009a) showed a reduction in haemocyte lysosome membrane stability and an increase in oxidative stress in mussels (Mytilus galloprovincialis) exposed to the antiepileptic drug carbamazepine. Gonzalez-Rey and Bebianno (2014) assessed the effects of diclofenac (a non-steroidal anti-inflammatory drug) in mussel M. galloprovincialis and showed that this drug induced biochemical responses, including a significant induction of the activity of the enzymes superoxide dismutase and glutathione reductase in mussels gills in addition to high catalase activity and lipid peroxidation levels in their digestive gland. Recently Lacaze et al. (2015) showed that exposure to psychotropic drugs and antibiotics led to genotoxicity, immunotoxicity and cytotoxicity in mussel M. edulis. To our knowledge, the effects of cetirizine in aquatic organisms, namely in bivalves and in particular in mussels, have not been evaluated before, despite the presence of this drug in different aquatic systems, which justifies the need for this study. The antihistamine cetirizine was detected in water bodies, namely in surface waters and influents/effluents of WWTPs with concentrations ranging from 4 ng/L to 1.4 mg/L, respectively (Bahlmann et al., 2012; Kosonen and Kronberg et al., 2009a,b; Larsson et al., 2007). Taking into consideration that different studies demonstrated that regardless of their mode of action pharmaceuticals may induce oxidative stress in aquatic organisms, the present study aimed to evaluate cellular damages, defense mechanisms and the metabolic potential of M. galloprovincialis after exposure to cetirizine for 28 days. A range of cetirizine concentrations, resembling medium to highly contaminated areas was used.

channel in the Ria de Aveiro, a shallow lagoon in northwest Portugal. Previous studies already reported the presence of pharmaceutical residues in this aquatic system, which were limited to wastewater treatment plants effluents and in one surface water sample at the Mira channel (Costa Nova region) with cetirizine concentrations varying between 0.04 and 0.6 mgL
1 (Calisto et al., 2011). For this study, mussels with similar fresh weight (20.7 ± 1.3 g) were selected in order to consider all the individuals as replicates. In order to reduce the possible contamination levels inherent to their natural environment (including metals), specimens were depurated and acclimated to laboratory conditions during 2 weeks in several glass aquaria, with artificial seawater (salinity 25) set up by mixing artificial sea salt (Tropic Marin Reef Mix) with reverse osmosis water, under continuous aeration, at 18 ± 1  C, a photoperiod thereabout 12:12 h (light/dark) and fed twice per week with Algamac protein plus (150 000 cells/L/animal) purchased to Aquafauna Bio-Marine, Inc. During this period water was changed twice per week (before feeding the animals). After this period mussels were exposed during 28 days to different conditions: 0.0 (Control, CTL), 0.3, 3.0, 6.0 and 12.0 mg/L of cetirizine. For each condition 3 containers (plastic 5 L vessels) were used, each one with 5 individuals (5 organisms x 3 containers x 5 conditions). During exposure period the medium was renewed every week and the individuals were fed three times per week with Algamac protein plus (150 000 cells/L/animal) purchased to Aquafauna Bio-Marine, Inc. Weekly, after water renewal, cetirizine concentrations were reestablished. During exposure, experimental conditions were maintained: 3 L of medium per container, salinity 25, continuous aeration, temperature 18 ± 1  C and a 12:12 h photoperiod. Dead organisms were removed from the containers when identified. Organisms were considered dead when their shells gaped and failed to shut again after external stimulus. Weekly water samples were collected immediately before and after the water renewal to evaluate cetirizine concentrations over time. Water samples (ca 2 mL) were collected, using a pipette, and preserved in the freezer at
20  C until cetirizine quantification. After exposure (28 days) individuals were collected and immediately frozen with liquid nitrogen and preserved at
80  C. Before laboratory analyses, frozen organisms were removed from their shells and the soft tissue pulverized in a mill with liquid nitrogen, for cetirizine quantification and biochemical markers measurement. For each organism, pulverized tissue was distributed in 0.5 g aliquots. In parallel to the exposure assay a quality control experiment was conducted to evaluate cetirizine losses (due to degradation or adsorption onto containers) along the 28 days exposure period. For this, containers with the same exposure conditions were prepared without mussels. Water samples were taken weekly immediately before and after water renewals. Water samples (ca 2 mL) were collect using a pipette, and preserved in the freezer at
20  C until cetirizine quantification.

2. Material and methods

2.2. Laboratory measurements

2.1. Experimental conditions

2.2.1. Cetirizine quantification 2.2.1.1. Reagents. The polyclonal antibody against mouse (IgG F(c) domain, from goat, lot 20 185) and the anti-cetirizine monoclonal antibody (mouse IgG1, clone B3212M, lot 5 K32007) were purchased from Acris Antibodies (Germany) and BIODESIGN International (Meridian Life Science Inc., USA), respectively. The tracer was produced and characterized as described in Bahlmann et al. (2009). 3,30 ,5,50 -Tetramethylbenzidine (99%, CAS number: 54827-17-7); tetrabutylammonium borohydride (>97%, CAS number: 33725-745); sodium phosphate dibasic dihydrate (>99%, CAS Number

For the present study the mussel Mytilus galloprovincialis was selected as it represents one of the most widely used bioindicator species of environmental pollution (among others, Martin-Diaz et al., 2009a). Furthermore, since this species is consumed worldwide, contamination by pharmaceuticals may put at risk not only human health but also the economic status of both producers and harvesters. M. galloprovincialis individuals were collected at the Ilhavo

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10028-24-7); sodium phosphate monobasic dihydrate (>99%, CAS number: 13472-35-0); potassium sorbate (>99%, CAS number: 24634-61-5); potassium dihydrogen citrate (>99%, CAS number: 866-83-1); hydrogen peroxide (30%, CAS number: 7722-84-1) and Tween™20 (CAS number: 005-64-5) were purchased from Fluka. Ethylenediamine tetraacetic acid disodium salt dihydrate (>99%, CAS number: 60-00-4) and sodium chloride (99.5%, CAS number: 7647-14-5) were from Panreac-Applichem. Dimethylacetamide (CAS number: 127-19-5) and tris(hydroxymethyl) aminomethane (TRIS, p.a.) were purchased from VWR Prolabo. Cetirizine dihydrochloride (p.a.) was purchased from Sigma-Aldrich. 2.2.1.2. Immunoassay procedure. Cetirizine was quantified based on the direct competitive ELISA (Enzyme-Linked ImmunoSorbent Assay) following the method developed by Bahlmann et al. (2011) and optimized by Calisto et al. (2011). From the exposure assay, samples from 15 individuals per condition (5 organisms  3 containers) and 3 water samples per condition (1 sample  3 containers) were used. From the quality control assay 2 water samples per condition (1 sample  2 containers) were used. For cetirizine quantification in mussels, supernatants were obtained by extraction of pulverized tissue (0.5 g) with deionized water (1:2, w/v). For this process, samples were sonicated (Branson Sonifier) for 15 s at 4  C and centrifuged (Beckman) for 10 min at 10 000 g at 4  C. The sample buffer was prepared with 1 M TRIS, 2% (w/v) EDTA and 3 M NaCl (pH 7.6). For the analysis of cetirizine in supernatants from tissue, cetirizine standards were prepared, in ultrapure water, by diluting a 10 mg/L stock solution of cetirizine (also prepared in ultrapure water). For the analysis of the medium exposure and blank samples, cetirizine standards were prepared in seawater (25 g/L NaCl) by diluting a stock solution of cetirizine with the same concentration in order to eliminate the effects of water salinity in quantification. All samples and standards (0e100 mg/L) were determined in triplicate for each determination. A fourparametric logistic equation (4PL) (Findlay and Dillard, 2007) was fit to the mean absorbance values obtained for each standard concentration. The absorbance was read on a microplate spectrophotometer at 450 nm and referenced to 650 nm. After cetirizine quantification in mussels tissues and in the exposure water, the Bioconcentration factor (BCF) was determined for each condition dividing the concentration of cetirizine present in mussels tissues by the cetirizine concentration in the exposure medium (Gobas and Morrison, 2000). 2.2.2. Biochemical markers 2.2.2.1. Reagents. Trichloroacetic acid (>99.5%, CAS number: 7603-9), potassium dihydrogen phosphate (>99.5%; CAS number: 7778-77-0) and oxidized glutathione (98.0%, CAS number: 2702541-8) were purchased from Panreac-Applichem. Potassium phosphate dibasic (>99%, CAS number: 7758-11-4); Triton X.100 (CAS number: 9036-19-5) and magnesium sulfate (99.5%, CAS number: 10034-99-8) were purchased from Merck Millipore. Polyvinylpyrrolidone (CAS number: 9003-39-8); bovine serum albumin (98%, CAS number: 9048-46-8); reduced glutathione (98.0%, CAS number: 70-18-8); superoxide dismutase (S8409, CAS number: 9054-89-1); formaldehyde solution (for molecular biology, CAS number: 50-00-0); sulfosalicylic acid (>99%, CAS number: 596583-3) and glucose (97%, CAS number: 492-61-5) were purchase from Sigma-Aldrich. Nitroblue tetrazolium chloride (ultra pure grade, CAS number: 298-83-9) was purchased from AMRESCO. Dithiothreitol (molecular biology grade, CAS number: 3483-12-3) was purchased from VWR Prolabo. 2.2.2.2. Biochemical procedures. Pulverized tissue of each organism (0.5 g aliquots) was used for supernatants extractions with specific

buffers (1:2, w/v) for each biochemical parameter. After adding specific buffers to pulverized tissue, samples were sonicated (Branson Sonifier) at 4  C during 15 s and centrifuged (Beckman) for 10 min at 10 000 g and 4  C. The supernatants were frozen or immediately used to determine: energy-related markers (total protein content, PROT; glycogen content, GLY; and electron transport system activity, ETS), oxidative stress markers (superoxide dismutase activity, SOD; catalase activity, CAT; glutathione Stransferases activity, GSTs; lipid peroxidation levels, LPO; reduced (GSH) and oxidized (GSSG) glutathione content). All biochemical parameters were performed in duplicate. For all biomarkers, absorbances were measured using a Microplate reader (BioTek Synergy). Energy-related parameters For PROT and GLY quantification extraction was performed using potassium phosphate buffer (50 mM potassium dihydrogen phosphate; 50 mM potassium phosphate dibasic; 1 mM ethylenediamine tetraacetic acid disodium salt dihydrate (EDTA); 1% (w/v) Triton X-100; 1% (w/v) polyvinylpyrrolidone (PVP); 1 mM dithiothreitol (DTT); pH 7.0). The supernatants for ETS were extracted in 0.1 M Tris-HCl (pH 8.5) with 15% (w/v) PVP, 153 mM magnesium sulfate (MgSO4) and 0.2% (v/v) Triton X-100) (Almeida et al., 2014). For GLY quantification the sulphuric acid method was used, as described by Dubois et al. (1956). Glucose standards were used (0e10 mg/mL). Absorbance was measured at 492 nm. The results were expressed in mg per g fresh weight (FW). PROT content was determined according to the spectrophotometric method of Biuret (Robinson and Hogden, 1940). Bovine serum albumin (BSA) was used as standard (0e40 mg/mL). Absorbance was measured at 540 nm. The results were expressed in mg per g FW. ETS activity was measured based on King and Packard (1975) and modifications performed by De Coen and Janssen (1997). Absorbance was measured at 490 nm during 10 min in 25 s intervals. The amount of formazan formed was calculated using Ɛ ¼ 15 900M
1cm
1 and the results expressed in nmol per min per g FW. Oxidative stress markers Antioxidant (SOD and CAT) and biotransformation (GSTs) enzymes were quantified in supernatants extracted with potassium phosphate buffer (50 mM potassium dihydrogen phosphate; 50 mM potassium phosphate dibasic; 1 mM ethylenediamine tetraacetic acid disodium salt dihydrate (EDTA); 1% (v/v) Triton X-100; 1% (w/v) polyvinylpyrrolidone (PVP); 1 mM dithiothreitol (DTT); pH 7.0). LPO supernatants were extracted with 20% (w/v) trichloroacetic acid (TCA). GSH and GSSG were determined using 0.6% (w/v) sulfosalicylic acid in potassium phosphate buffer (0.1 M dipotassium phosphate; 0.1 M potassium dihydrogen phosphate; 5 mM EDTA; 0.1% (v/v) Triton X-100; pH 7.5) (Almeida et al., 2014). SOD was determined following the method described in Beauchamp and Fridovich (1971) and adaptations performed by Carregosa et al. (2014). The standard curve was determined using SOD standards (0e60 U/mL). Absorbance was measured at 560 nm. The enzymatic activity was expressed in U per g FW, where U corresponds to a reduction of 50% of nitroblue tetrazolium (NBT). CAT was quantified according to Johansson and Borg (1988) and modifications performed by Carregosa et al. (2014). The standard curve was determined using formaldehyde standards (0e150 mM). Absorbance was determined at 540 nm. The enzymatic activity was expressed in U per g FW, where U represents the amount of enzyme that catalyzes the formation of 1 nmol formaldehyde. GSTs were quantified following Habig et al. (1974) protocol with some adaptations performed by Carregosa et al. (2014). GSTs activity was determined spectrophotometrically at 340 nm (ε ¼ 9.6 mM
1 cm
1). The enzymatic activity was expressed in U

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per g FW where U is defined as the amount of enzyme that catalyzes the formation of 1 mmol of dinitrophenyl thioether per min. LPO determination was performed following the method described by Ohkawa et al. (1979). LPO levels were measured trough the quantification of malondialdehyde (MDA), a by-product of lipid peroxidation. Absorbance was determined at 535 nm (ε ¼ 156 mM
1cm
1). LPO levels were expressed in nmol of MDA formed per g FW. The quantification of GSH and GSSG was performed following the method described in Rahman et al. (2007), using GSH and GSSG as standards (0e90 mmol/L). Absorbance was measured at 412 nm and the results expressed in mmol per g FW.

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0.79e1.02 for 6.0 mg/L; 0.92e1.02 for 12.0 mg/L (for concentrations obtained immediately after water contamination) and between 0.61 and 0.79 for 0.3 mg/L; 0.88e0.96 for 3.0 mg/L; 0.96e1.06 for 6.0 mg/L; 0.95e1.10 for 12.0 mg/L (for concentrations obtained immediately before water renewal). Cetirizine was present in M. galloprovincialis tissues at concentrations that increased significantly with the increase of the exposure concentration (Fig. 1E), in samples obtained after 28 days of exposure. The BCF values showed no significant differences among concentrations of 0.3 (BCF: 1.3 ± 0.2), 3.0 (BCF: 1.2 ± 0.3) and 6.0 mg/L (BCF: 1.4 ± 0.2) and significantly lower values were found at the highest exposure concentration (12.0 mg/L, BCF: 1.0 ± 0.1) (Fig. 1F).

2.3. Data analyses Data obtained from cetirizine determination and biochemical analyses were submitted to hypothesis testing using permutational analysis of variance with the PERMANOVA þ add-on in PRIMER v6 (Anderson et al., 2008). The pseudo-F values in the PERMANOVA main tests were evaluated in terms of significance. When the main test revealed statistical significant differences (p  0.05), pairwise comparisons were performed. The t-statistics in the pairwise comparisons were evaluated in terms of significance. Values lower than 0.05 were considered as significantly different. A one-way hierarchical design was followed, with cetirizine concentrations as the main fixed factor. The null hypothesis tested was: for each parameter no significant differences were found among exposure conditions. Significance levels among conditions were presented with different letters. 3. Results 3.1. Cetirizine quantification For water samples from exposures as well as from blanks, data on cetirizine quantification, acquired at the experiment onset (immediately after contamination and immediately before water renewal) and at the last week of exposure onset (immediately after contamination and immediately before water renewal), were selected as representative of the entire exposure period. The concentrations of cetirizine quantified in the exposure media (Fig. 1A and B) demonstrated that regardless of the week of exposure, concentrations are significantly different among exposure concentrations and increased along the increasing exposure gradient. Also, independently of the week of analysis, cetirizine concentrations measured immediately after water contamination were not significantly different from the cetirizine concentrations measured after seven days of exposure (immediately before water renewal) demonstrating that the concentration levels were constant between water renewals (1 week) and during the entire course of the experiment. Measured concentrations were also similar to the nominal concentrations, with relative differences ranging between 0.83 and 0.99 for 0.3 mg/L; 0.98e1.23 for 3.0 mg/L; 1.04e1.34 for 6.0 mg/L; 0.93e1.25 for 12.0 mg/L (for concentrations obtained immediately after water contamination) and between 1.28 and 1.35 for 0.3 mg/L; 1.16e1.25 for 3.0 mg/L; 1.22e1.38 for 6.0 mg/L; 1.04e1.31 for 12 mg/L (for concentrations obtained immediately before water renewal). In water samples from blanks (Fig. 1C and D) results obtained showed that cetirizine concentrations at each condition were not affected by biodegradation, photodegradation or adsorption to experiment vessels during the seven days of exposure, the period between each water renewal. Measured concentrations were also similar to the nominal concentrations, with relative differences ranging between 0.62 and 0.90 for 0.3 mg/L; 0.77e1.00 for 3.0 mg/L;

3.2. Biochemical markers 3.2.1. Energy-related parameters After exposure, mussels exposed to the lowest cetirizine concentrations (0.3 and 3.0 mg/L) showed no significant differences in PROT content compared to individuals from the control (Fig. 2A). However, individuals in the highest concentrations (6.0 and 12.0 mg/L) showed significantly higher PROT content than individuals from the control and the lowest exposure concentration (0.3 mg/L) (Fig. 2A). GLY content increased along the increasing exposure gradient, with the highest values at the highest exposure concentration (Fig. 2B). Nevertheless, no significant differences were found among individuals exposed to the respective concentrations (Fig. 2B). Mussels presented significantly higher ETS activity at the highest exposure concentration (12.0 mg/L) but no significant differences were found among tested concentrations (Fig. 2C). 3.2.2. Oxidative stress related markers Along the increasing cetirizine exposure gradient mussels increased the activity of SOD but no significant differences were found among individuals exposed to different concentrations (Fig. 3A). As for SOD activity, mussels showed increasing CAT activity with the increase of cetirizine concentration, with the highest values at the highest exposure concentration (Fig. 3B). However, in this case, significant differences were found between individuals exposed to the highest concentration (12.0 mg/L) and individuals in the control and the lowest concentration (0.3 mg/L), as well as between individuals in the 6.0 mg/L and mussels exposed to the lowest cetirizine concentration (0.3 mg/L) (Fig. 3B). GSTs presented a decreasing trend in enzymatic activity with the increase of exposure concentrations (Fig. 3C), but no significant differences were found between exposed and non-exposed organisms. LPO levels increased along the increasing exposure gradient (Fig. 4A), with the highest values at the highest cetirizine concentration (12.0 mg/L). The results obtained showed significantly higher values in the concentrations 3.0, 6.0, 12.0 mg/L compared to the control and 0.3 mg/L condition. No significant differences in LPO levels were found between individuals exposed to 3.0, 6.0 and 12.0 mg/L (Fig. 4A). As for LPO, mussels presented increasing GSH content with the increase of cetirizine concentrations, but no significant differences were found among conditions (Fig. 4B). GSSG content was higher in mussels exposed to 3.0, 6.0 and 12.0 mg/L compared to the remaining concentrations (CTL and 0.3 mg/L) but no significant differences were found among concentrations (Fig. 4C).

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Fig. 1. Cetirizine (CTZ) concentrations (mg/L) in A: water samples from the experiment at the first week of exposure; B: water samples from the experiment at the fourth week of exposure; C: water samples from blanks at the first week of exposure; D: water samples from blanks at the fourth week of exposure. For each concentration and for each week, dark grey bars represent samples collected immediately after water contamination, while light grey bars represent samples collected immediately before water renewal. E: CTZ concentrations (ng/g FW) in Mytilus galloprovincialis at the fourth week of exposure; and F: Bioconcentration Factor (BCF). Values are the mean (STDEV) of 15 individuals. Significant differences (p  0.05) among concentrations are presented with different letters.

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Fig. 2. A: Protein (PROT) content; B: Glycogen (GLY) content; C: Electron Transport System (ETS) activity, in Mytilus galloprovincialis after 28 days of exposure to increasing concentrations of Cetirizine (CTZ) (0.0, 0.3, 3.0, 6.0 and 12.0 mg/L). Values are the mean (STDEV) of 15 individuals. Significant differences (p  0.05) among concentrations are presented with different letters.

4. Discussion Several studies have shown that pharmaceuticals like nonsteroidal anti-inflammatory drugs (ibuprofen and diclofenac), antihypertensive (propranolol), antiepileptics (carbamazepine), and antibiotics (fluoxetine, paroxetine and erythromycin) induce distinct impacts in bivalves, including mussels (Almeida et al., 2014, 2015; Ericson et al., 2010; Gonzalez-Rey and Bebianno, 2011, 2014; Lacaze et al., 2015). Nevertheless, scarce information is available on

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the effects of antihistaminic drugs in bivalves (Meinertz et al., 2012). Since antihistamines are considered one of the most used drugs worldwide (e.g. Minguez et al., 2016; Petrie et al., 2015), it is of most relevance to evaluate the impacts that this type of drugs may exert in the aquatic environment. The present study demonstrated increased accumulation of cetirizine in mussels tissues with the increase of exposure concentration, although lower bioconcentration rate was observed at the highest tested cetirizine concentration. Furthermore, this study clearly revealed that cetirizine accumulation observed in M. galloprovincialis was correlated to the toxicity effects induced in mussels, with alterations on energy-related and oxidative stress related markers. The balance between mitochondrial electron transport system activity and energy reserves has been reported as a relevant marker to predict changes in invertebrates due to stressful conditions (De  et al., Coen and Janssen, 1997; Smolders et al., 2004; Gagne 2006). Also Smolders et al. (2004) pointed out that the use of energy-related parameters, like PROT and GLY content, allow to measure the energy costs associated to the activation of organisms defense mechanisms under stressful conditions. In the present study M. galloprovincialis presented higher GLY and PROT content with the increase of cetirizine exposure concentration which may indicate that mussels were able to avoid the expenditure of energy even under stress conditions. Since mussels only presented increased metabolic potential (measured by the ETS activity) at the highest exposure concentration this may indicate that up to the highest exposure concentration organisms were able to maintain their metabolism at control level, avoiding energy expenditure. Higher energy reserves in exposed mussels may be achieved by closing their valves, which was a protecting behavior identified by Gosling (2003) in bivalves exposed to pollutants. Duquesne et al. (2004) also demonstrated that the clam Ruditapes philippinarum increased the GLY content along the increase of carbamazepine exposure concentrations which was associated to the reduction of the metabolic activity based in the decreasing of clams clearance rate. At the highest exposure concentration, although the metabolic potential was significantly higher compared to the remaining concentrations, probably metabolic activation was only triggered at the end of the assay and did not occur along the entire exposure period. Therefore mussels were not able to use promptly their energy reserves and/or did not have enough time to use their energy reserves, which were the highest at this condition. Similarly, Cruz et al. (2016) showed that PROT content increased in R. philippinarum with the increasing caffeine concentration but GLY content was lower in exposed than in non-exposed organisms. In this case authors discussed that the increase on clams ETS activity with the increase of caffeine exposure concentrations induced the use of GLY and organisms were able to preserve PROT content. In the present study the increase of PROT content in mussels may also be associated to the increase on enzymes production, including enzymes involved in oxidative stress defense mechanisms (e.g. antioxidant enzymes) which is in line with Smolders et al. (2003) findings showing that pollution triggers the increase of the synthesis of proteins related to defense mechanisms in zebrafish individuals. In fact, the antioxidant enzymes activity increased along the increasing exposure gradient that could result not only from the increase of enzymes activity per se but also from the increase of enzymes production. The present study clearly demonstrated an increase on SOD and CAT activities with the increase of cetirizine concentration as a defense mechanism against cetirizine impacts, namely overproduction of reactive oxygen species (ROS) responsible for LPO (among others, Regoli and Giuliani, 2014). SOD and CAT are ROS naturally occurring scavengers (Regoli and Giuliani, 2014; Santovito

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Fig. 3. A: Superoxide dismutase (SOD) activity; B: Catalase (CAT) activity; C: Glutathione S-transferases (GSTs) activity, in Mytilus galloprovincialis after 28 days of exposure to increasing concentrations of Cetirizine (CTZ) (0.0, 0.3, 3.0, 6.0 and 12.0 mg/L). Values are the mean (STDEV) of 15 individuals. Significant differences (p  0.05) among concentrations are presented with different letters.

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Fig. 4. A: Lipid peroxidation (LPO) levels; B: Reduced glutathione (GSH) content; C: Oxidized glutathione (GSSG) content, in Mytilus galloprovincialis after 28 days of exposure to increasing concentrations of Cetirizine (CTZ) (0.0, 0.3, 3.0, 6.0 and 12.0 mg/L). Values are the mean (STDEV) of 15 individuals. Significant differences (p  0.05) among concentrations are presented with different letters.

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et al., 2005) and it has been shown that when under stressful conditions (namely exposure to pharmaceuticals) bivalves, including mussels, show increased activity of these enzymes to prevent peroxidation of membrane lipids. Freitas et al. (2015a) showed an increase of SOD activity in clam Scrobicularia plana up to 3.0 mg/L of carbamazepine but at higher concentrations (6.0 and 9.0 mg/L) the activity of this enzyme decreased to control levels. These authors also demonstrated that CAT activity was maintained for 28 days regardless of carbamazepine concentration. Assessing the impacts of caffeine in R. philippinarum Cruz et al. (2016) demonstrated that SOD and CAT activity increased with increasing exposure concentration (0.5, 3.0 and 18.0 mg/L), after 28 days of exposure. Nevertheless, several other studies revealed opposite trends in bivalves exposed to drugs, from maintenance to decrease of enzymes activity. Martin-Diaz et al. (2009a) revealed that Mytilus galloprovincialis exposed to carbamazepine (0.1 and 10.0 mg/L) for 7 days showed increased activity of CAT in the digestive glands but no activation was noticed in gills and mantle/ gonads except at the highest concentration in mantle/gonads tissues. Mezzelani et al. (2016) exposed M. galloprovincialis during 24 days to different pharmaceuticals and showed different response of CAT towards each drug, with no changes on enzyme activity when mussels were exposed to acetaminophen, diclofenac and ibuprofen and significant decreases when organisms were exposed to ketoprofen and nimesulide. Previous studies by Gonzalez-Rey and Bebianno (2011), exposing M. galloprovincialis to ibuprofen (250 ng/L), revealed that SOD activity significantly increased in the presence of this contaminant up to 7 days of exposure after which (15 days of exposure) this defense mechanism fails. Furthermore, these authors showed that CAT was inhibited by this drug, with the lowest values at the longer exposure period (15 days). GonzalezRey and Bebianno (2014) demonstrated that M. galloprovincialis exposed to an environmentally relevant nominal concentration of diclofenac (250 ng/L) presented decreased activity of CAT and SOD in gills and digestive gland after 15 days of exposure compared to control individuals. The biotransformation GSTs enzymes are known to perform dual protective actions in order to minimize the drug impacts: they are involved in the formation of thiol metabolites from oxidation of contaminants (including pharmaceuticals) through the conjugation with GSH; and are also capable to inactivate lipoperoxidation products, such as lipid hydroperoxides, by the use of GSH as reducing agent (Sturve et al., 2008). Vernouillet et al. (2010) demonstrated that these enzymes have an important role fighting oxidative stress caused by pharmaceuticals contamination in aquatic organisms. However, in the present study, although with no significant differences, the activity of GSTs was lower in organisms exposed to cetirizine compared to non-exposed organisms, which indicates the low capacity of these enzymes to detoxify cetirizine in M. galloprovincialis. Mezzelani et al. (2016) also demonstrated that GSTs activity was lower in mussels (M. galloprovincialis) exposed to ibuprofen but no significant changes were found in GSTs activity in mussels exposed to acetaminophen, diclofenac, ketoprofen and nimesulide compared to control individuals. Freitas et al. (2015a) observed that the exposure of S. plana to carbamazepine resulted in the maintenance or even decrease of GSTs activity compare to control organisms. Also Gonzalez-Rey and Bebianno (2011) showed that M. galloprovincialis presented decreased GSTs activity when exposed to ibuprofen for 15 days but Gonzalez-Rey and Bebianno (2014) revealed that when this species was exposed to diclofenac for 15 days no significant increase was noticed in the activity of GSTs (measured in gills and digestive gland) compared to control individuals. Nevertheless Aguirre-Martínez et al. (2016) demonstrated that this group of enzymes, in general, increased the activity in R. philippinarum clams exposed for 15 days to different drugs (0.1,

5.0, 15.0, 50.0 mg/L caffeine, carbamazepine, ibuprofen, novobiocin). Also Cruz et al. (2016) showed that GSTs activity increased in R. philippinarum exposed to caffeine. Such findings may indicate that the defense mechanisms developed by bivalves against pharmaceuticals are only triggered at given levels of cellular damage which are close related to contaminant concentrations. Also, it seems that defense mechanisms although activated, may present limitations to eliminate ROS if their production surpasses certain limits and cellular damages occur. Finally the results obtained by different authors may also indicate that the activation of antioxidant and biotransformation enzymes might be species and drug dependent. We may further hypothesize that even between the same species but from different populations (coming from different geographical areas) the responses may differ, which may be related to adaptation of organisms to certain conditions. Previous studies by Freitas et al. (2015b) demonstrated that clams coming from different areas (with different contamination levels) respond differently to carbamazepine exposure. Our findings further revealed that despite the increase on antioxidant enzymes activity in mussels exposed to cetirizine, especially at the highest concentration, this defense strategy was not able to eliminate effectively the excess of ROS and prevent mussels from LPO formation. In the present study the increase of LPO may not only result from the excess of ROS due to the presence of cetirizine but also due to the increase on ETS activity at the highest exposure concentration. It is known that an increase in ETS activity is frequently related with an increase of ROS generation, since mitochondria are the principal site of their production, phenomenon that is reflected in LPO levels. Several other studies also demonstrated that when bivalves are exposed to environmental realistic concentrations of pharmaceuticals, although their defense mechanisms are activated (including antioxidant and biotransformation enzymes), LPO may still occur. Gonzalez-Rey and Bebianno (2011) showed that M. galloprovincialis mussels contaminated with ibuprofen showed increased LPO levels, especially after 7 days of exposure. Similar results were found in the same species after 15 days exposure to diclofenac (Gonzalez-Rey and Bebianno, 2014). The exposure to gemfibrozil and diclofenac (1 and 1000 mg/L) induced a significant increase of LPO in mussel Dreissena polymorpha (Quinn et al., 2011) and diclofenac (1 mg/L and 1000 mg/L) significantly induced LPO in Mytilus spp. after 96 h of exposure (Schmidt et al., 2011). Also Aguirre-Martínez et al. (2016) showed that R. philippinarum presented higher LPO when exposed to different drugs (caffeine, carbamazepine, ibuprofen, novobiocin) although defense mechanisms were activated. Higher LPO levels were also found in the same species exposed to caffeine for 28 days (Cruz et al., 2016). A similar pattern was found by Freitas et al. (2015a) in S. plana exposed to carbamazepine for 28 days. To antagonize the adverse effects of ROS, besides enzymatic mechanisms, organisms possess scavengers like the cytosolic reduced glutathione (GSH) that neutralize ROS by direct reaction with them. The ROS neutralization occurs through the oxidation of GSH into oxidized glutathione (GSSG), which posteriorly is reconverted in GSH by specific reductases. In the present study mussels showed an increase on total glutathione (GSHt) content along the increasing exposure gradient (data not shown). GSH and GSSG concentrations were higher in exposed organisms, with 2-fold higher GSH content than GSSG. These results may indicate that mussels were not able to convert the amount of GSH produced into GSSG and that this strategy was not efficient in the elimination of ROS in M. galloprovincialis or that mussels do not use this strategy when contaminated by cetirizine. Few studies have assessed the impacts of pharmaceutical drugs in bivalves GSHt content but Mezzelani et al. (2016) showed that in M. galloprovincialis exposed

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to different drugs, similar or even lower GSHt was obtained compared to control individuals. Assessing the impacts of carbamazepine in S. plana, Freitas et al. (2015a) showed that the ratio GSH/GSSG decreased with the increase of exposure concentrations indicating that the amount of GSH decreased and GSSG increased along the exposure gradient and that GSH was used as scavenger. Cruz et al. (2016) further demonstrated that although GSSG increased with the increase of caffeine concentrations (0.5 and 3.0 mg/L) at the highest exposure concentration (18.0 mg/L) the concentration of GSSG decreased indicating that R. philippinarum was no longer able to use this strategy to eliminate ROS. 5. Conclusions Overall, we can conclude that the present study clearly demonstrated that cetirizine induced biochemical alterations in M. galloprovincialis mussels, not only in their metabolic activity and energy budget but also, and especially, on oxidative stress related markers, with alterations on antioxidant and biotransformation enzymes activity and induction of LPO. Acknowledgments ^nia Calisto benefited from post-doc grants Rosa Freitas and Va (SFRH/BPD/92258/2013 and SFRH/BPD/78645/2011, respectively) ^ and Angela Almeida benefited from a PhD grant (SFRH/BD/110218/ 2015) from National Funds through the Portuguese Science Foundation (FCT), supported by FSE, Programa Operacional Capital Humano (POCH) and the European Union. Thanks are due, for the financial support, to CESAM (UID/AMB/50017), to FCT/MEC through national funds, and the co-funding by FEDER, within the PT2020 Partnership Agreement and Compete 2020. References Abreu, M.S., Cristina, A., Giacomini, V., Gusso, D., Rosa, J.G.S., Koakoski, G., ^ncio, R., Oliveira, T.A., Barcellos, H.H.A., Barcellos, L.J.G., Kalichak, F., Idale Bonan, C.D., 2016. Acute exposure to waterborne psychoactive drugs attract zebra fish. Comp. Biochem. Physiol., Part C 179, 37e43. http://dx.doi.org/ 10.1016/j.cbpc.2015.08.009. Aguirre-Martínez, G.V., Del Valls, T.A., Martín-Díaz, M.L., 2013. Early responses measured in the brachyuran crab Carcinus maenas exposed to carbamazepine and novobiocin: application of a 2-tier approach. Ecotoxicol. Environ. Saf. 97, 47e58. http://dx.doi.org/10.1016/j.ecoenv.2013.07.002. Aguirre-Martínez, G.V., DelValls, T.A., Martín-Díaz, M.L., 2016. General stress, detoxification pathways, neurotoxicity and genotoxicity evaluated in Ruditapes philippinarum exposed to human pharmaceuticals. Ecotoxicol. Environ. Saf. 124, 18e31. http://dx.doi.org/10.1016/j.ecoenv.2015.09.031. ^ Calisto, V., Esteves, V.I., Schneider, R.J., Soares, A.M.V.M., Figueira, E., Almeida, A., Freitas, R., 2014. Presence of the pharmaceutical drug carbamazepine in coastal systems: effects on bivalves. Aquat. Toxicol. 156, 74e87. http://dx.doi.org/ 10.1016/j.aquatox.2014.08.002. ^ Freitas, R., Calisto, V., Esteves, V.I., Schneider, R.J., Soares, A.M.V.M., Almeida, A., Figueira, E., 2015. Chronic toxicity of the antiepileptic carbamazepine on the clam Ruditapes philippinarum. Comp. Biochem. Physiol. Part e C Toxicol. Pharmacol. 172e173, 26e35. http://dx.doi.org/10.1016/j.cbpc.2015.04.004.  ~ oz, D., Rodríguez-Mozaz, S., Maulvault, A.L., Tediosi, A., Ferna ndezAlvarez-Mu n  , D., Kotterman, M., Marques, A., 2015. Tejedor, M., Van den Heuvel, F., Barcelo Occurrence of pharmaceuticals and endocrine disrupting compounds in macroalgaes, bivalves, and fish from coastal areas in Europe. Environ. Res. 143 (Part B), 56e64. http://dx.doi.org/10.1016/j.envres.2015.09.018. Anderson, M., Gorley, R.N., Clarke, R.K., 2008. Permanovaþ for Primer: Guide to Software and Statistical Methods. University of Auckland-PRIMER-E, Plymouth. Bahlmann, A., Falkenhagen, J., Weller, M.G., Panne, U., Schneider, R.J., 2011. Cetirizine as pH-dependent cross-reactant in a carbamazepine-specific immunoassay. Analyst 136, 1357e1364. Bahlmann, A., Carvalho, J.J., Weller, M.G., Panne, U., Schneider, R.J., 2012. Immunoassays as high-throughput tools: monitoring spatial and temporal variations of carbamazepine, caffeine and cetirizine in surface and wastewaters. Chemosphere 89 (11), 1278e1286. http://dx.doi.org/10.1016/j.chemosphere.2012. 05.020. Bahlmann, A., Weller, M.G., Panne, U., Schneider, R.J., 2009. Monitoring carbamazepine in surface and wastewaters by an immunoassay based on a monoclonal antibody. Anal. Bioanal. Chem. 395 (6), 1809e1820. http://dx.doi.org/10.1007/

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