Effects of seawater acidification and salinity alterations on metabolic, osmoregulation and oxidative stress markers in Mytilus galloprovincialis

Ecological Indicators 79 (2017) 54–62

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Original Articles

Effects of seawater acidification and salinity alterations on metabolic, osmoregulation and oxidative stress markers in Mytilus galloprovincialis

MARK

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Rosa Freitasa, , Lucia De Marchia, Miguel Bastosa, Anthony Moreiraa, Cátia Veleza, Stefania Chiesaa, Frederick J. Wronab, Etelvina Figueiraa, Amadeu M.V.M. Soaresa a b

Departamento de Biologia & CESAM, Universidade de Aveiro, 3810-193 Aveiro, Portugal Department of Geography David Turpin Building University of Victoria, Victoria, BC, V8P 5C2, Canada

A R T I C L E I N F O

A B S T R A C T

Keywords: Climate change Mussels Osmoregulation and biomineralization capacity Cellular damages Antioxidant mechanisms

The impacts of seawater acidification and salinity shifts on metabolism, energy reserves, and oxidative status of mussels have been largely neglected. With the aim to increase the current knowledge for the mussel Mytilus galloprovincialis a 28-day chronic test was conducted during which mussels were exposed to two pH (7.8 and 7.3; both at control salinity 28) and three salinity (14, 28 and 35, at control pH, 7.8) levels. After exposure to different conditions, mussels electron transport system activity, energy reserves (protein and glycogen content) carbonic anhydrase activity, antioxidant defences and cellular damage were measured. Results obtained showed that mussels exposed to seawater acidification presented decreased metabolic capacity that may have induced lower energy expenditure (observed in higher glycogen, protein and lipids content at this condition). Low pH condition induced the increase of carbonic anhydrase activity that was related to acid-base balance, while no significant activation of antioxidant defence mechanisms was observed resulting in higher LPO. Regarding the impacts of salinity, the present study showed that at the highest salinity (35) mussels presented lower metabolic activity (also related to lower energetic expenditure) and an opposite response was observed at salinity 14. Carbonic anhydrase slightly increased at stressful salinity conditions, a mechanism of homeostasis maintenance. Lower metabolic activity at the highest salinity, probably related to valves closure, helped to mitigate the increase of LPO in this condition. At low salinity (14), despite an increase of antioxidant enzymes activity, LPO increased, probably as a result of ROS overproduction from higher electron transport system activity. The present findings demonstrated that Mytilus galloprovincialis oxidative status and metabolic capacity were negatively affected by low pH and salinity changes, with alterations that may lead to physiological impairments namely on mussels reproductive output, growth performance and resistance to disease, with ecological and economic implications. Indicators: Physiological and biochemical changes in Mytilus galloprovincialis in response to low pH and salinity changes

1. Introduction Due to human activities, atmospheric CO2 partial pressure (pCO2) is increasing, and is predicted to reach 500–1000 μatm by the end of this century (Caldeira and Wickett, 2003, 2005; IPCC, 2013; Orr et al., 2005; Raven et al., 2005). These changes have led to an increase of global mean temperature (a process called global warming) and to a decrease in both pH and the availability of carbonate ions in seawater (a phenomenon known as ocean acidification) (Caldeira and Wickett, 2003; Feely et al., 2009; Orr et al., 2005; Raven et al., 2005). As a result of global warming salinity shifts are also expected to occur since the atmospheric temperature rise is causing changes in the hydrological

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cycle at a global scale, with increases in precipitation at high latitudes and near the tropics, and decreases in the sub-tropical and mid-latitude regions (Fenoglio et al., 2010). As a consequence, many areas may experience frequent flood and drought events which lead to prolonged and more frequent periods of decreased or increased salinity, particularly in estuarine areas (Bussell et al., 2008). Additionally, the continuous release of CO2 into the atmosphere has caused a decrease in ocean pH of approximately 0.1 pH units with respect to pre-industrial levels, and further decrease from 0.3 to 0.5 pH units are predicted to occur up to year 2100 (Caldeira and Wickett, 2003, 2005; IPCC, 2013; Orr et al., 2005; Raven et al., 2005). Besides studies on climate change predictions, a growing body of

Corresponding author at: Departamento de Biologia, Universidade de Aveiro, Campus Universitário de Santiago, 3810-193, Aveiro, Portugal. E-mail address: [email protected] (R. Freitas).

http://dx.doi.org/10.1016/j.ecolind.2017.04.003 Received 26 October 2016; Received in revised form 22 February 2017; Accepted 3 April 2017 1470-160X/ © 2017 Elsevier Ltd. All rights reserved.

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Marin Reef Mix) prepared at salinity 28 ± 1, pH 7.8, temperature maintained at 17 ± 1 °C, 12 light: 12 dark photoperiod and continuous aeration. Animals were fed two times per week with Algamac Protein Plus (150.000 cells/animal). After this period organisms were distributed in different aquaria to test different salinities (14, 28, 35) at control pH (7.8) and different pH levels (7.8 and 7.3) at control salinity (28). The following conditions were used for exposures: salinity 14 (pH 7.8), salinity 28 (control salinity, pH 7.8), salinity 35 (pH 7.8), pH 7.3 (control salinity). For each condition 3 replicates were used, with 5 organisms per replicate (15 organisms per condition). For each replicate 1L of artificial seawater per individual was used. Organisms were exposed to each condition for 28 days. During the experimental period dissolved oxygen concentration was monitored in all aquaria and animals were checked for mortality every day. Aquaria were maintained at constant temperature (17 ± 1 °C), 12 light: 12 dark photoperiod and continuous aeration. Two times per week, animals were fed with Algamac Protein Plus (150.000 cells/animal), seawater was renewed every week, and pH and salinity levels re-established. Dead organisms were removed when identified. After exposure (28 days), surviving organisms were frozen until biochemical and physiological analysis. For pH acclimation, organisms were submitted to a gradual pH decrease (0.2 values per day) to reach pH 7.3 and maintained at salinity 28 ± 1. The remaining organisms were maintained at pH 7.8 and salinity 28 ± 1. To reach the salinity levels of 14 and 35, mussels under pH 7.8 were exposed every 2–3 days to different salinities, which were gradually lowered or increased in 2–3 values to reach the test levels. Mussels exposed to different salinity conditions were maintained under pH 7.8 (considered as control pH). Low pH was obtained by directly diffusing CO2 into aquaria. Individual aquarium pH levels were continuously monitored and controlled using a pH Stat system (Aquamedic AT Controller) and crosschecked with independent probes (Hanna Instruments). pH Stat system probes were calibrated using NIST buffers (NBS scale). Seawater pH was crosschecked against an independent probe (Hanna Instruments) at least twice a week, and the pH Stat computer reset to match the independent probes pH if needed. Lowered pH was set to 7.3, to give a 0.5 pH unit reduction relative to control (pH 7.8). Control pH was considered as the average pH measured at the sampling area (7.7–7.9) and a pH decrease of 0.5 units as predicted for the end of the twenty first century (Raven et al., 2005). Tested pH values are within naturally occurring range in temperate estuarine systems (Ringwood and Keppler, 2002; Cochran and Burnett, 1996; Ramos et al., 2006; Ansari and Gill, 2013). Testing conditions gave pCO2 values ranging from ∼940 (pH 7.8) to ∼3700 μatm pCO2 (pH 7.3), within values that occur in coastal habitats (Cochran and Burnett, 1996; Melzner et al., 2012). Salinity 28 was used as control taking into account the salinity at the sampling area. Salinities tested (14 and 35) were selected to resemble extreme weather events of rain and drought periods, namely in the study area (Dias et al., 1999; Dias and Lopes, 2006; Génio et al., 2008; Vaz and Dias, 2008). Water samples were collected from pH test aquaria (conditions: pH 7.8 and salinity 28; pH 7.3 and salinity 28), prior to water renewal, and used to determine Total alkalinity (TA) by potentiometric titration (Gran, 1952). Calculated TA values and measured parameters (tem-

evidence shows that seawater acidification and salinity alterations can impact aquatic species, with particular attention to calcifying organisms such as bivalves (e.g Beniash et al., 2010; Bressan et al., 2014; Dickinson et al., 2012; Velez et al., 2016). Despite a growing number of studies published on seawater acidification, that show impacts on bivalves physiological performance including feeding and excretion rates, growth and larval development, calcification, respiration and reproduction (e.g., Bressan et al., 2014; Dupont et al., 2010; FernándezReiriz et al., 2011; 2012; Gazeau et al., 2010; Kurihara, 2008; Michaelidis et al., 2005; Navarro et al., 2013; Sun et al., 2016; Wang et al., 2015; Xu et al., 2016), less information is available regarding alterations on bivalves oxidative status and metabolic capacity exposed to such scenarios (Hu et al., 2015; Matoo et al., 2013; Matozzo et al., 2013). Furthermore, few studies evaluated the effects of salinity on bivalves physiological (Dickinson et al., 2012; Kim et al., 2001; Hamer et al., 2008) and biochemical (e.g.: Bussel et al., 2008; Carregosa et al., 2014a, 2014b; Gonçalves et al., 2017; Hamer et al., 2008; Moreira et al., 2016a; Velez et al., 2016) performance. Among marine bivalves, several mussel species are commonly used as bioindicators of environmental stressors since they present physiological and biochemical changes when exposed to inorganic and organic contaminants (among others Apeti et al., 2010; Filimonova et al., 2016; Lavradas et al., 2016; Milun et al., 2016; Signa et al., 2015), including Mytilus galloprovincialis (e.g. Cajaraville et al., 1996; Roberto et al., 2010; Rocha et al., 2016; Vlahogianni et al., 2007). Most recently, M. galloprovincialis has also been used as model organism to study climate change related stressors (Anestis et al., 2007; Bressan et al., 2014; Duarte et al., 2014; Fernández-Reiriz et al., 2012; Matozzo et al., 2013; Michaelidis et al., 2005; Range et al., 2012). However, no studies identified and compared the osmotic, metabolic and oxidative stress alterations induced in M. galloprovincialis under salinity alterations and seawater acidification. To enhance the current knowledge on this species performance under a changing environment, the present study aimed to evaluate the capacity of M. galloprovincialis as a bioindicator of salinity changes and seawater acidification. For this, mussels physiological and biochemical responses were identified, including metabolic capacity (electron transport system activity), energy reserves content (protein, glycogen and lipid), osmoregulation capacity (carbonic anhydrase activity), cellular damage (lipid peroxidation levels) and antioxidant capacity (superoxide dismutase and catalase activity), after mussels chronic exposure (28 days) to different salinity (14, 28, 35) and pH (7.8 and 7.3) levels.

2. Materials and methods 2.1. Study organisms and experimental setup Mussels were collected in September 2015 in the Ílhavo channel, the most narrow and short main channel of the Ria de Aveiro lagoon (Northwest coast of Portugal), with a length of ca. 15 km. Previous studies demonstrated low metal contamination levels in this channel (Freitas et al., 2014; Velez et al., 2015). After collection mussels were transferred to the laboratory, where they were acclimated and depurated for 15 days prior to exposure. During this period, mussels were kept in artificial seawater (Tropic

Table 1 Carbonate system physicochemical parameters for pH experiments (mean ± SD). Measured pH, and determined total alkalinity (At) from weekly water sampling (Temperature 17.1 °C ± 0.75 and salinity 28.4 ± 0.5). Partial CO2 pressure (pCO2), bicarbonate (HCO3−) and carbonate ion concentrations (CO32−), and saturation states of calcite (ΩCal) and aragonite (ΩAg), calculated with CO2SYS software ().

pH 7.8 pH 7.3

pH

At (μmol kg−1)

ƿCO2 (μatm)

HCO3− (μmol kg-1)

CO32− (μmol kg−1)

ΩCal

ΩAra

7.86 ± 0.06 7.34 ± 0.02

2271 ± 127 2495 ± 92

940 ± 149 3645 ± 218

2081 ± 126 2429 ± 90

78.3 ± 10.0 27.2 ± 1.8

1.95 ± 0.26 0.68 ± 0.04

1.2 ± 0.14 0.43 ± 0.03

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2.3.2. Energy reserves GLY content was quantified according to the phenol-sulphuric acid method, described by Yoshikawa (1959). Glucose standards were used to obtain a calibration curve (0–5 mg/ mL). Samples were incubated at room temperature for 30 min and absorbance measured at 492 nm. Results were expressed in mg per g FW. PROT content was determined according to the Biuret method (Robinson and Hogden, 1940), using bovine serum albumin (BSA) as standard (mg/mL). Absorbance was measured at 540 nm and the results were expressed in mg per g FW. LIP content was quantified according to Cheng et al. (2011). Absorbance was measured at 540 nm after 1 h of color development, at room temperature. A standard curve was determined using cholesterol standards (0–100%). Results were expressed in percentage per g dry weight (DW).

perature, salinity and pH values, read at the time of water sampling) were plotted in CO2SYS software (Robbins et al., 2010) and carbonate chemistry calculated using dissociation constants K1, K2 (Mehrbach et al., 1973; Dickson and Millero, 1987) and KSO4 (Dickson, 1990) (Table 1).

2.2. Taxonomic identification by molecular markers After collection mussels were randomly selected for molecular taxonomic analyses. Prior to DNA extraction, whole mussels were preserved in absolute ethanol and conserved at −20 °C. Foot tissue was selected for genomic DNA extraction. High weight genomic (HWG) DNAs were obtained by extraction with DNeasy Blood & Tissue Kit (Qiagen) following the manufacturer protocol. The Me15/16 marker developed by Inoue et al. (1995) is considered the most consistent and reliable for Mytilus species identification (Wood et al., 2003). The Me15/Me16 primers were used to amplify a segment of gene coding an adhesive protein of byssus, with diagnostic PCR products length differences among the three European taxa M. edulis, M. trossulus and M. galloprovincialis (Inoue et al., 1995). The reaction mix used was as follows: 25 μL reaction volume containing 1U of Taq Polymerase (Nzytech), 50 mM MgCl2, 10 mM dNTPs, and 10 pmol of each primer. Polymerase chain reaction (PCR) profile for Me15 and Me16 primers was modified from the original one (Inoue et al., 1995) as follows: 3 min of denaturation at 93 °C, then 35 cycles of 25 s at 93 °C, 20 s at 45 °C, and 30 s at 72 °C, followed by a final elongation step of 3 min at 72 °C. Three different DNA fragments, each representing an allele diagnostic for one of the Mytilus taxa, could be observed at the codominant Me-locus (molecular weight 180 bp for M. edulis, 168 bp for M. trossulus and 126 bp for M. galloprovincialis). PCR products were visualized by electrophoresis in a 2.5% agarose gel with a specific 50–1000 and 100–1000 bp DNA markers for fragment weight identification.

2.3.3. Osmorregulation capacity CA activity was measured spectrophotometrically, following an adaptation of the titrimetric method described by Warrier et al. (2014) with modifications described in Moreira et al. (2016b). The absorbance was measured at 436 nm on a microplate reader for 1 min, and the variation of absorbance per min (ΔABS min−1) determined in triplicate for each sample. Non-enzymatic reaction rate was also determined in triplicate, following the same procedure, after denaturing samples at 100 °C for 15 min. Mean non-enzymatic reaction rate from each sample was subtracted from each sample mean enzymatic reaction. A set of samples were analyzed through a standard pH assay described by Weis and Reynolds (1999). The same samples were also analyzed spectrophotometrically and a calibration curve between ΔpH min−1 and ΔABS min−1 was established (r2 = 0.998). Results were expressed in ΔpH min−1 per g FW. 2.3.4. Antioxidant defences SOD activity was quantified following Beauchamp and Fridovich (1971), using SOD standards 0.25–60 U/mL. Enzyme activity was measured at 560 nm after adding xanthine oxidase at 5mU, diluted in phosphate buffer 50 mM (pH 8.0). Absorbance was read after 20 min incubation at 22 °C, and the rate of nitroblue tetrazolium (NBT) reduction calculated. SOD was expressed in U per g FW where U corresponds to a reduction of 50% of nitroblue tetrazolium (NBT). CAT activity was determined according to Johansson and Borg (1988) at room temperature (22 °C). A standard curve was determined using formaldehyde standards (0–150 μM). Absorbance was read at 540 nm and the enzyme activity was expressed in U per g FW. One unit of enzyme (U) is defined as the amount of enzyme that caused the formation of 1.0 nmol of formaldehyde per min.

2.3. Biochemical descriptors After 28 days of exposure, two mussels per replicate were used for Lipid content determination, and three mussels per replicate were used for the remaining biomarkers determination. For Lipid content determination, mussels were opened and placed inside an oven at 45 °C for 48 h. After this period dried tissue was homogenized using a mortar and pestle, and stored for further analysis. For the remaining biochemical analysis each mussel was opened, tissue was immediately fixed using liquid nitrogen, and samples stored at −80 °C. Latter, samples were manually homogenized in liquid nitrogen, and each specimens homogenate separated in 0.5 g aliquots (except for lipid determination furtherly discussed). Extractions were performed with specific buffers (see Moreira et al., 2016a) for each analysis and samples centrifuged for 10 min at 10,000g or 3,000g(depending on the biomarker) (4 °C). Supernatants were stored at −80 °C or immediately used to determine mussels: metabolic capacity (assessed by electron transport system (ETS) activity); energy reserves (glycogen (GLY) content, total soluble protein (PROT) content, lipids (LIP) content); osmoregulation (carbonic anhydrase, CA) and antioxidant (superoxide dismutase, SOD; catalase, CAT) capacity; and cellular damage (measured by lipid peroxidation (LPO) level).

2.3.5. Cellular damage LPO was measured by the quantification of malondialdehyde (MDA), according to the method described by Ohkawa et al. (1979). Absorbance was read at 535 nm (ε = 156 mM−1 cm−1). LPO levels were expressed in nmol of MDA per g FW. 2.4. Data analysis The biochemical descriptors (LPO, CAT, SOD, CA, ETS, GLY, PROT, LIP) were submitted to hypothesis testing using permutational analysis of variance, employing the PERMANOVA+ add-on in PRIMER v6 (Anderson et al., 2008). The effects of salinity (for pH 7.8) as well as the impacts of pH (for salinity 28) were evaluated separately. When the main test revealed statistical differences (p ≤ 0.05), pairwise comparisons among conditions were performed, and differences among salinity conditions (14, 28, 35; pH 7.8) and between pH conditions (7.8, 7.3; salinity 28) were evaluated separately. Biochemical descriptor data were analyzed following a one-way hierarchical design, with salinity (pH 7.8) or pH (salinity 28) as the main fixed factors. The null

2.3.1. Metabolic capacity ETS activity was measured based on King and Packard (1975) and modifications by Coen and Janssen (1997). Absorbance was measured at 490 nm every 25 s during 10 min. The amount of formazan formed was calculated using the extinction coefficient 15,900 M−1 cm−1and the results expressed in nmol min per g fresh weight (FW). 56

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Fig. 1. Results of amplification of the non-repetitive region of the adhesive protein gene. M1, M2, M3: original mussel samples from this study. L100 & L50 = molecular markers (100–1000 bp; 50–1000 bp).

hypotheses tested were: i) no significant differences existed among salinities (14, 28, 35; pH 7.8); ii) no significant differences existed between pH levels (7.8, 7.3; salinity 28). Significant differences (p ≤ 0.05) among salinities were presented with different letters, while significant differences between pH levels were represented with an asterisk. 3. Results 3.1. Taxonomic identification by molecular markers The analysis of Me 15–16 marker showed that all samples exhibited a single band, indicating that they were all homozygotes. The electrophoresis separation identified a single band of 126 bp molecular weight in each analyzed sample, as showed in Fig. 1. Therefore, all analyzed mussels presented the typical genotype profile of Mytilus galloprovincialis. 3.2. Biochemical responses 3.2.1. Metabolic capacity Results on the ETS activity in mussels exposed to different salinities (14, 28 and 35, pH 7.8) showed the highest values in mussels exposed to salinities 28 and 14, with significant differences compared to mussels exposed to salinity 35 (Fig. 2). Comparing the ETS activity in mussels exposed to different pH levels (7.8 and 7.3, salinity 28) significantly Fig. 3. Energy reserves: A- Glycogen (GLY) content, B- Protein (PROT) content, C- Lipid (LIP) content in Mytilus galloprovincialis exposed to different salinity levels (14, 35, 28, pH 7.8) and low pH (7.3, salinity 28). Significant differences (p ≤ 0.05) among salinities (14, 28, 35, pH 7.8) are represented with letters. Significant differences (p ≤ 0.05) between pH levels (7.8 and 7.3, salinity 28) are represented with an asterisk.

lower values were found in mussels under low pH conditions (Fig. 2). 3.2.2. Energy reserves GLY content (Fig. 3A) in mussels under control pH (7.8) was significantly higher in mussels exposed to the highest salinity (35) compared to mussels exposed to salinities 14 and 28, with no significant differences between mussels exposed to salinity 14 and 28 (Fig. 3A). Significantly higher GLY content was observed in mussels exposed to pH 7.3 (salinity 28) in comparison to mussels exposed to pH 7.8 (salinity 28) (Fig. 3A). PROT content (Fig. 3B) was significantly higher in mussels exposed to salinity 35 (pH 7.8) in comparison to mussels exposed to salinities 14 and 28 (pH 7.8), with significantly lower values in mussels exposed to salinity 28 (Fig. 3B). Comparing mussels exposed to different pH levels (salinity 28) significantly higher PROT content was observed at low pH conditions (Fig. 3B). The lowest LIP content was observed in mussels under the lowest

Fig. 2. Metabolic capacity: Electron transport system (ETS) activity in Mytilus galloprovincialis exposed to different salinity levels (14, 35, 28, pH 7.8) and low pH (7.3, salinity 28). Significant differences (p ≤ 0.05) among salinities (14, 28, 35, pH 7.8) are represented with letters. Significant differences (p ≤ 0.05) between pH levels (7.8 and 7.3, salinity 28) are represented with an asterisk.

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Fig. 4. Osmotic regulation: Carbonic Anhydrase (CA) activity in Mytilus galloprovincialis exposed to different salinity levels (14, 35, 28, pH 7.8) and low pH (7.3, salinity 28). Significant differences (p ≤ 0.05) among salinities (14, 28, 35, pH 7.8) are represented with letters. Significant differences (p ≤ 0.05) between pH levels (7.8 and 7.3, salinity 28) are represented with an asterisk.

salinity (14, pH 7.8), with significant differences to mussels exposed to salinities 28 and 35 (pH 7.8). No significant differences were observed between mussels exposed to salinity 28 (pH 7.8) and 35 (pH 7.8) (Fig. 3C). Comparing the LIP content in mussels exposed to different pH conditions, significantly higher values were found at pH 7.3 (Fig. 3C). 3.2.3. Osmorregulation capacity CA activity showed no significant differences among organisms exposed to salinities 14, 35 and 28 (pH 7.8). Significantly higher CA activity was observed in mussels exposed to low pH (salinity 28) in comparison to organisms exposed to pH 7.8 (salinity 28) (Fig. 4).

Fig. 5. Antioxidant defences: A- Superoxide Dismutase (SOD) activity, B- Catalase (CAT) activity in Mytilus galloprovincialis exposed to different salinity levels (14, 35, 28, pH 7.8) and low pH (7.3, salinity 28). Significant differences (p ≤ 0.05) among salinities (14, 28, 35, pH 7.8) are represented with letters. Significant differences (p ≤ 0.05) between pH levels (7.8 and 7.3, salinity 28) are represented with an asterisk.

3.2.4. Antioxidant defences The activity of SOD was significantly higher in organisms exposed to salinity 14 (pH 7.8) in comparison to values observed in mussels exposed to salinities 28 and 35 (pH 7.8), with no significant differences between mussels exposed to these two salinities (Fig. 5A). Comparing results obtained at both pH levels (salinity 28), although SOD activity increased at low pH conditions no significant differences were found between values obtained at pH 7.8 and 7.3 (Fig. 5A). The activity of CAT was significantly higher in mussels exposed to the lowest salinity (14, pH 7.8) in comparison to mussels exposed to higher salinities (28 and 35, pH 7.8) (Fig. 5B). Although mussels exposed low pH (salinity 28) showed lower CAT activity than organisms exposed to pH 7.8 at the same salinity no significant differences were observed between these two conditions (Fig. 5B). 3.2.5. Cellular damage LPO levels observed at different salinities (pH 7.8) showed significantly higher values in mussels exposed to the lowest salinity (14), with no significant differences between salinities 28 and 35 (Fig. 6). Comparing LPO levels in mussels exposed to different pH conditions (at salinity 28), significantly higher values were observed in mussels at low pH (Fig. 6).

Fig. 6. Cellular damage: Lipid peroxidation (LPO) levels in Mytilus galloprovincialis exposed to different salinity levels (14, 35, 28, pH 7.8) and low pH (7.3, salinity 28). Significant differences (p ≤ 0.05) among salinities (14, 28, 35, pH 7.8) are represented with letters. Significant differences (p ≤ 0.05) between pH levels (7.8 and 7.3, salinity 28) are represented with an asterisk.

4. Discussion

Considering M. edulis and M. galloprovincialis contact zone, the hybridization between both subspecies has been observed along the Atlantic coasts from Spain to Ireland (Gosling et al., 2008), specifically with introgression of M. edulis mtDNA in Atlantic M. galloprovincialis individuals (Quesada et al., 1998). The Me15/Me16 nuclear marker is reliable for taxonomic identification of EuropeanMytilus species, and confirmed the species taxonomy (M. galloprovincialis).

4.1. Taxonomic identification by molecular marker The taxonomic identification of European Mytilus species is often difficult, especially if based only on shell characteristics, due to the adaptation of shell morphology to environmental conditions and to hybridization among congeneric taxa (Kijewski et al., 2009, 2011). 58

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a protecting behavior that has been identified by Gosling et al. (2008) in bivalves exposed to pollutants. Low energetic expenditure in response to seawater acidification or salinity alterations in other bivalve species have been described, and are in accordance with our results. Timmins-Schiffman et al. (2014) studied the impacts of ocean acidification on C. gigas and demonstrated that oysters showed no changes in GLY content under different scenarios (pCO2: 400 μatm (pH 8.0), 800 μatm (pH 7.7), 1000 μatm (pH 7.6) and 2800 μatm (pH 7.3)). Freitas et al. (2016) demonstrated that the clam S. plana collected from two different areas presented no change or increased PROT content under pH 7.1 for 96 h. Our study also revealed that organisms exposed to salinity 35 presented lower ETS activity in comparison to organisms under control salinity (28), a response that reflected in higher energetic reserve in mussels at the highest tested salinity. As mentioned above, mussels exposed to stressful conditions may close their valves, which reduces their metabolism (identified by lower ETS activity) that in turn, resulted in higher energetic reserves. Similar results were obtained by Dickinson et al. (2012) for the oyster C. virginica exposed to stressful salinity (15).

4.2. Metabolic capacity and energy reserves Previous studies have already demonstrated the impact of environmental factors in bivalves metabolic status (Freitas et al., 2016; Moreira et al., 2016a; Velez et al., 2016). The metabolic capacity of marine organisms has commonly been assessed by measuring the ETS activity, which represents a proxy of the cellular respiratory potential of a given organism (García-Martín et al., 2014) and therefore provides information on organisms physiological status (Choi et al., 2001; Nahrgang et al., 2013). ETS activity assay has been successfully used to assess the effect of climate change related factors in bivalves (Le Moullac et al., 2007; Moreira et al., 2016a). Overall, the present study demonstrated that M. galloprovincialis metabolic capacity may represent a good biomarker of ocean acidification conditions and salinity shifts. The present study clearly demonstrated that low pH conditions resulted in a decrease of mussels ETS activity in comparison to organisms under control pH (7.8). Several studies on different species of marine invertebrates have described metabolic depression under high seawater pCO2 levels, suggesting that the uncompensated extracellular pH might be the cause for this reduction. Mechanisms like oxygen supply and respiration can strongly respond to alterations in surrounding CO2 conditions (Pörtner et al., 2007; Gazeau et al., 2013). According to Fernández-Reiriz et al. (2011), Ruditapes decussatus clams exposed to acidification conditions is able to lower the standard metabolic rate, by decreasing clearance, ingestion, and respiration rates. Recently, Velez et al. (2016) showed a decrease in ETS activity in clams R. philippinarum exposed to low pH (7.3), suggesting a decrease of metabolic activity. Similar results were observed by Freitas et al. (2016) in Scrobicularia plana clams exposed to low pH (7.1). Michaelidis et al. (2005) demonstrated that elevated CO2 levels caused a decrease in the rate of oxygen consumption in M. galloprovincialis juveniles and adults, indicating a drastic reduction of the metabolic rate (pH 7.3). Navarro et al. (2013) showed a significant reduction of clearance rate after exposing M. chilensis to acidified seawater (pCO2 380, 750 and 1200 ppm) for 35 days. Also food absorption rate and efficiency were lower at high pCO2 levels. These physiological responses resulted in a significant reduction of energy available for growth in mussels. Wang et al. (2015) observed negative effects on respiration rates of M. coruscus exposed to low pH conditions (pH 7.3). Sun et al. (2016) demonstrated that, in mussel M. edulis, mortality increased while condition index and filtering rates decreased steadily with the decrease of pH (7.1; 6.5). Regarding salinity, our results demonstrated that organisms exposed to salinity 35 presented lower ETS activity in comparison to control salinity (28). Velez et al. (2016) showed that the clam R. decussatus exposed to salinity 14 presented increased metabolic activity (ETS). Similarly, the oyster Crassostrea angulata presented increased ETS activity at low salinities (10and 20), and results were attributed to higher metabolic rates associated to hyposmotic stress (Moreira et al., 2016a). Hamer et al. (2008) demonstrated that oxygen consumption rate of M. galloprovincialis acclimated to decreased salinities was a concentration-dependent process and increased considerably to about 51 and 65% in lower salinities (28 and 18) compared to control mussels (37). Also Kim et al. (2001) observed increased oxygen consumption in R. philippinarum clams exposed to salinity stress (15), also indicating metabolic adjustment to hyposmotic stress. The present study demonstrated that results obtained on mussels ETS activity could be related to mussels energetic reserve levels, which increased in mussels with lower ETS activity. In particular our findings showed that in comparison to organisms under control pH, mussels under acidification conditions tended to present lower metabolic potential which resulted in a lower energetic expenditure, reflected by higher GLY, PROT and LIP concentrations observed at pH 7.3. Higher energy reserves in mussels exposed to low pH may result from valve closure, which could have induced lower metabolic activity and could justify lower ETS activity observed in mussels exposed to low pH,

4.3. Osmorregulation capacity CA is a major player in osmoregulation processes and acid-base balance of estuarine organisms (Dickinson et al., 2012; Monserrat et al., 2007; Sattin et al., 2010), and is also involved in processes such as gas exchange, nitrogen metabolism and biomineralization (Lionetto et al., 2000; Le Roy et al., 2014). Our results showed that CA activity was significantly higher in mussels exposed to low pH in comparison to control individuals (pH 7.8, salinity 28), which could indicate that mussels responded to maintain acid-base balance. As pointed out by David et al. (2005) while studying the impacts of hypoxia on oysters, the increase of CA activity in organisms exposed to low pH may be explained due to higher oxygen demand and gas exchange efficiency in that particular condition. Also Beniash et al. (2010) indicated the possibility of increased CA activity in oyster C. virginica juveniles in response to hypercapnia due to increased CA mRNA expression. Recently Velez et al. (2016) observed similar results, in R. philippinarum exposed to pH 7.3 that presented higher CA activity under seawater acidification, supporting the role of this enzyme in acid-base balance. Regarding the impacts of salinity on M. galloprocincialis CA activity, higher values were observed in organisms exposed to salinity conditions (14 and 35) different from control salinity (28). Previous studies conducted by Olsowski et al. (1995) demonstrated the role of CA in Na+ and Cl− uptake in organisms under hyposaline stress, and could explain our results showing CA activity increase in M. galloprovincialis exposed to stressful salinity conditions (14 and 35). Higher CA activity was also observed in R. philippinarum clams exposed to salinities 14 and 35, supporting the involvement of this enzyme in the balance of ion homeostasis (Velez et al., 2016). Similarly, CA in C. angulata was higher in oysters maintained at low salinity (10) (Moreira et al., 2016a). 4.4. Antioxidant defences and cellular damages It is known that marine bivalves exposed to stressful conditions increase reactive oxygen species (ROS) production and to prevent lipid, DNA and protein damages organisms increase detoxification mechanisms, namely those related to antioxidant defence (e.g.: SOD and CAT enzymes) (Amiard-Triquet et al., 2012; Batley and Simpson, 2016). Antioxidant enzymes such as CAT and SOD are known for their capacity to eliminate ROS produced in excess when organisms are under stressful conditions (Regoli and Giuliani, 2014). To prevent cellular damages caused by ROS the organisms use SOD to convert the superoxide anion into dioxygen and hydrogen peroxide, while hydrogen peroxide is removed by CAT or glutathione peroxidase (Regoli and Giuliani, 2014). The present findings demonstrated that mussels were able to increase the activity of antioxidant enzymes (SOD and CAT), 59

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enzymes to eliminate the excess of ROS produced. However, Matozzo et al. (2013) revealed that regardless of antioxidant enzymes responses LPO was maintained in M. galloprovincialis exposed to different pH conditions (8.1, 7.7, 7.4). Our study further revealed that although antioxidant defences were enhanced at low salinity conditions (14) mussels presented significantly higher LPO levels at this condition. The electron transport chain is an important source of ROS production (such as superoxide anions) and therefore higher metabolic rates can induce high ROS production (Guzy and Schumacker, 2006; Murphy, 2009), a phenomenon that has been proposed for oysters acclimated to low salinity (Moreira et al., 2016a). This may explain the increase of antioxidant enzymes activity but also higher LPO levels in mussels exposed to salinity 14, where ETS was significantly higher in comparison to control salinity. Furthermore, although antioxidant mechanisms were not activated in mussels exposed to the highest salinity (35) LPO levels in mussels under this condition were similar to the ones observed under control salinity. This situation may result from lower ETS activity observed in mussels exposed to the highest salinity which may not only be associated to mussels valves closure (limiting the stress from higher salinity) but may also limited the production of ROS since no increase on organisms metabolism (measured by ETS activity) was observed. Velez et al. (2016) found similar results, showing that although antioxidant enzymes were induced in R. philippinarum clams exposed to salinity 14 these mechanisms were overwhelmed by ROS, resulting in LPO. Also Moreira et al. (2016a) observed higher LPO levels in C. angulata oysters maintained at low salinity (10), despite the increase of SOD and CAT activity. In conclusion, the present study revealed new data on M. galloprovincialis metabolic, energetic and oxidative status changes when mussels were exposed to climate change related factors (different salinities and different pH levels). The changes observed may alter organisms physiological fitness, which can therefore have consequences on mussels reproductive output, growth performance and resistance to disease, thus affecting species resilience in a changing environment. The present study demonstrated that under climate change predicted scenarios M. galloprovincilais can be recognized as a good bioindicator to pH and salinity changes, and the set of biomarkers used, namely related to metabolic activity, osmoregulation and cellular damages, showed to be efficient to identify the early impacts induced in mussels.

especially when exposed to low salinity levels (with enhanced activity of both enzymes) but also when under low pH conditions (in this case only SOD presented increased activity). Regarding the impacts of ocean acidification on the antioxidant defence mechanisms of M. galloprovincilais the present findings showed that although SOD activity was higher at low pH conditions the activity of CAT decreased, but in both cases no significant differences were observed between control (7.8) and low (7.3) pH conditions. These results may indicate that mussels were able to activate SOD and CAT under stressful salinity levels but at low pH conditions the activity of these enzymes was not enhanced which may indicate that the stress induced was not enough to activate these enzymes or other antioxidant defences were activated (such as glutathione peroxidase). Opposite responses were found by other authors, including Hu et al. (2015) that showed significantly higher antioxidant enzymes (SOD, CAT and GPx) activity in M. coruscus gills and digestive gland exposed for 14 days to pH 7.3 compared to control pH (8.1). Matozzo et al. (2013) observed a general upward trend (although not always linear) of SOD and CAT activity in the digestive gland of M. galloprovincialis that were kept for 7 days at decreased pH (7.4) while in the gills both enzymes tended to maintained their activity at values similar to control. Also Velez et al. (2016) reported the induction of SOD and CAT activity in R. philippinarum exposed to low pH (7.3) for 28 days. The results obtained further demonstrated that mussels presented significantly higher SOD and CAT activities when exposed to salinity 14 compared to control (salinity 28), indicating that the activation of these enzymes only occurred at low salinities. These results suggest that both CAT and SOD activities were only induced above a certain limit of stress; i.e., above a certain ROS production. In the present study the increase of antioxidant enzymes activity at salinity 14 may be related to higher metabolic capacity and lower energetic reserves level recorded in mussels under this condition as the increase of ETS may be related to the activation of antioxidant defences which requires energy to fuel such mechanisms. Also Li et al. (2008) reported a raise of SOD activity in the white shrimp Litopenaeus vannamei exposed to lower salinity conditions relating it with an increase of ROS. An increase of antioxidant defences (SOD and especially CAT) was observed by Matozzo et al. (2013) in the gills and digestive gland of M. galloprovincialis exposed to salinities 34 and 40 in comparison to control. Velez et al. (2016) and Moreira et al. (2016a) also reported the induction of SOD and CAT activity in R. philippinarum and C. angulata exposed to low salinity conditions (14 and 10), respectively. However, Zanette et al. (2011) did not observe alterations in CAT activity in oysters C. gigas exposed to a salinity gradient. Also Gonçalves et al. (2017) showed that SOD activity presented a significant inhibition in Cerastoderma edule exposed to salinities higher (30) and lower (10) than the control salinity (20) with an increase of SOD activity only recorded at salinity 35 in relation to control values. If antioxidant defence systems are not able to eliminate the excess of ROS lipid peroxidation (LPO) occurs (Amiard-Triquet et al., 2012; Catalá, 2009). The present study showed that organisms exposed to low pH conditions were not able to significantly increase the activity of SOD and CAT which justifies the increased LPO levels observed at pH 7.3. Furthermore, our findings demonstrated that although antioxidant enzymes were activated in mussels under stressful salinity conditions, this defence mechanism was not enough to eliminate the excess of ROS produced by mussels under salinity 14 and increased LPO was observed in mussels exposed to this condition. These results may indicate the inefficiency of M. galloprovincialis antioxidant mechanisms in organisms exposed to predicted climate change conditions. In particular, the inefficient activation of antioxidant defences in mussels exposed to low pH conditions resulted in higher LPO values in these organisms compared to those under control (pH 7.8). A similar response was observed by Velez et al. (2016) that showed that although R. philippinarum presented higher activity of antioxidant enzymes at low pH conditions, LPO increased, suggesting the inefficiency of these

Acknowledgments Cátia Velez benefited from a PhD grant (SFRH/BD/86356/2012), Anthony Moreira benefited from a PhD grant (SFRH/BD/93107/2013), Lucia De Marchi benefited from PhD grant (SFRH/BD/101273/2014), Rosa Freitas and Stefania Chiesa benefited from post-doc grants (SFRH/ BPD/92258/2013 and SFRH/BPD/91923/2012, respectively) given by the National Funds through the Portuguese Science Foundation (FCT), supported by FSE and Programa Operacional Capital Humano (POCH) e da União Europeia. This work was supported by the Integrated Programme of SR & TD “Smart Valorization of Endogenous Marine Biological Resources Under a Changing Climate” (reference Centro01-0145-FEDER-000018), co-funded by Centro 2020 program, Portugal 2020, European Union, through the European Regional Development Fund. Thanks are due, for the financial support to CESAM (UID/AMB/ 50017), to FCT/MEC through national funds, and the co-funding by the FEDER, within the PT2020 Partnership Agreement and Compete 2020. References Amiard-Triquet, C., Amiard, J.-C., Rainbow, P.S., 2012. Ecological Biomarkers: Indicators of Ecotoxicological Effects. CRC Press. Anderson, M., Gorley, R.N., Clarke, R.K., 2008. Permanova+ for Primer: Guide to Software and Statisticl Methods .

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