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Responses of the sea anemone, Exaiptasia pallida, to ocean acidification conditions and zinc or nickel exposure
Accepted Manuscript Title: Responses of the sea anemone, Exaiptasia pallida, to ocean acidification conditions and zinc or nickel exposure Author: Christina G. Duckworth Codie R. Picariello Rachel K. Thomason Krina S. Patel Gretchen K. Bielmyer-Fraser PII: DOI: Reference:
S0166-445X(16)30344-7 http://dx.doi.org/doi:10.1016/j.aquatox.2016.11.014 AQTOX 4537
To appear in:
Aquatic Toxicology
Received date: Revised date: Accepted date:
10-10-2016 11-11-2016 16-11-2016
Please cite this article as: Duckworth, Christina G., Picariello, Codie R., Thomason, Rachel K., Patel, Krina S., Bielmyer-Fraser, Gretchen K., Responses of the sea anemone, Exaiptasia pallida, to ocean acidification conditions and zinc or nickel exposure.Aquatic Toxicology http://dx.doi.org/10.1016/j.aquatox.2016.11.014 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Responses of the sea anemone, Exaiptasia pallida, to ocean acidification conditions and zinc or nickel exposure
Christina G. Duckworth1, Codie R. Picariello1, Rachel K. Thomason1, Krina S. Patel1, Gretchen K. Bielmyer-Fraser2
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Department of Biology
Valdosta State University, Valdosta, GA, USA 2
Department of Chemistry
Jacksonville University, Jacksonville, FL, USA
Corresponding Author: Gretchen K. Bielmyer-Fraser Department of Chemistry, Jacksonville University, 2800 University Blvd. North, Jacksonville, FL 32211; Email: [email protected]
Highlights
Ni and Zn exposure to E. pallida resulted in concentration-dependent tissue metal accumulation and the rate of accumulation was increased with exposure of 1000 ppm CO2
Both Ni and Zn were well regulated in the anemone and the effects of CO2 exposure on tissue metal concentration were no longer observed after 96 h
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GR activity was significantly reduced by increased Ni or Zn, and the inhibition was decreased by addition of 1000 ppm CO2.
Abstract Ocean acidification, caused by increasing atmospheric carbon dioxide (CO2), is a growing concern in marine environments. Land-based sources of pollution, such as metals, have also been a noted problem; however, little research has addressed the combined exposure of both pollutants to coral reef organisms. In this study we examined tissue metal accumulation and physiological effects (activity of anti-oxidant enzymes, catalase and glutathione reductase) in the sea anemone, Exaiptasia pallida after exposure to increased CO2, as well as zinc (Zn) or nickel (Ni). After exposure to four concentrations (nominal values = control, 10, 50, 100 µg/L) of Zn or Ni over 7 days, both metals accumulated in the tissues of E. pallida in a concentrationdependent manner. Anemones exposed to elevated CO2 (1000 ppm) accumulated significant tissue burdens of Zn or Ni faster (by 48 h) than those exposed to the same metal concentrations at ambient CO2. No differences were observed in catalase activity due to Zn exposure; however, 50 µg/L Ni caused a significant increase in catalase activity at ambient CO2. No significant effect on catalase activity from CO2 exposure alone was observed. Glutathione reductase activity was affected by increased Zn or Ni exposure and those effects were influenced by increased CO2. Results of this study provide insight into the toxic mechanisms and environmental implications of CO2 and Zn or Ni exposure to the cnidarian E. pallida.
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1. Introduction Coral reefs are among the most diverse and productive ecosystems on Earth, containing a wealth of species complexity and substantial economic importance (Brown and Howard, 1985). Despite their vital role in the global economy and ecology, these valuable marine environments have experienced vast degradation in recent years (Brown and Howard, 1985; Albright and Langdon, 2011). Ocean acidification has become a prevalent environmental stressor to many coral reef species. Current oceanic values of CO2 approximate 380 µatm, and by the years 2100 and 2300 these values are expected to rise to 1,000 and 1,900 µatm, respectively, which would cause a decrease of up to 0.77 pH units with the higher CO2 level (Caldeira and Wickett 2003; Meehl et al., 2007; Fabry et al. 2008; Esbaugh et al., 2012; Albright and Langdon, 2011). The uptake of atmospheric CO2 by the ocean alters the ocean carbonate chemistry, lowering the concentration and saturation states of calcium carbonate (CaCO3) minerals (Gomez et al., 2014). Several studies have demonstrated a negative correlation with higher concentrations of CO2 in coral reef organisms that rely on a saturated state of CO32- for calcification and growth (Gomez et al., 2014; Ateweberhan et al., 2013). Towle et al. (2015) reported a decline of as much as 98% in the dominant reef-building coral A. cervicornis in the Florida Reef Tract since the 1970s. Environmental stressors affecting calcification and the coral-algal symbiosis may induce a phase shift from communities of hard, reef-building corals to more resilient species of soft corals and fleshy macroalgae (Peters et al., 1997; Gomez et al., 2014; Anthony et al., 2011).
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In addition to the global stressors of ocean acidification, coral reef ecosystems are also faced with more localized stressors, such as heavy metals, from land-based anthropogenic sources (Howard and Brown, 1984; Anthony et al., 2011). Many reefs are located near densely populated areas with substantial industrialization and coastal development. Sewage discharge, dredging, coastal petroleum refineries, fossil fuel combustion, sacrificial anodes on boats, leachate from metal-based antifouling paints, marine disposal of municipal solid waste and metallic bulk waste, mining, smelting, refining, and alloy processing, and many other agricultural and industrial activities contribute to increased heavy metal pollution in nearshore marine environments (Howard and Brown, 1984; Guzmán and Jiménez, 1992; Reichelt and Jones, 1994; Gonzalez et al., 1999; Evans et al., 2000; Voulvoulis et al., 2000; Naoum et al., 2001; Stylianous et al., 2007; Jones, 2010). Zinc and nickel are both essential metals to marine organisms and function in multiple roles as a cofactorsfor enzymes and in DNA. As essential metals, they are needed in low concentrations, but may become toxic at concentrations above certain thresholds (Poonkothai and Vijayavathi, 2012; Ferrier-Pages et al., 2005). High concentrations of zinc can affect main physiological functions, such as calcification rates, and can affect photosynthetic efficiency and proper enzyme function in the coral symbionts, zooxanthellae (Houlbrèque et al., 2011; Ferrier-Pages et al., 2005). Heavy metals have been shown to accumulate in coral reef organisms, particularly those containing algal symbionts such as zooxanthellae (Peters et al., 1997; Main et al., 2010; Horwitz et al., 2014). In many cases, corals and anemones exposed to elevated concentrations of metals will release their symbiotic algae as a possible method of detoxification, since algae have been shown to accumulate some metals to a larger extent than their animal symbiont (Bielmyer et al., 2010; Horwitz et al., 2014). This process of ―bleaching‖ is a potentially detrimental condition
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and can occur in conjunction with physiological disruptions in the holobiont (Biemyer et al., 2010; Main et al., 2010; Brock and Bielmyer, 2013; Patel and Bielmyer-Fraser, 2015). Physiological effects of metal exposure include changes in the activity of antioxidant enzymes (Main et al., 2010; Brock and Bielmyer, 2013). When exposed to metals, corals and anemones containing a photosynthetic symbiont are susceptible to increased production of reactive oxygen species (ROS) that can denature proteins, mutate DNA, and cause lipid peroxidation (Yakovleva et al., 2004). Antioxidant defenses utilizing the enzymes glutathione peroxidase (GPx), glutathione reductase (GR), and catalase (CAT), have been shown to reduce these harmful effects (Yakovleva et al., 2004; Brock and Bielmyer, 2013; Patel and Bielmyer-Fraser, 2015; Siddiqui et al., 2015; Siddiqui and Bielmyer-Fraser, 2015). GPx and CAT are produced to catalyze the conversion of hydrogen peroxide into water and oxygen, thus combating the effects of ROS (Higuchi et al., 2010; Forman et al., 1990; Sies, 1999; Sunagawa et al., 2008; Brosnan and Brosnan, 2009). GR is then produced to reduce glutathione so that it may be recycled for the reaction above (Forman et al., 1990; Sies, 1999; Sunagawa et al., 2008; Brosnan and Brosnan, 2009). Though several studies have examined the effects of ocean acidification or metal exposure, few studies have analyzed the effects of both stressors combined (Houlbrèque et al., 2011; Gomez et al., 2014; Peters et al., 1997; Horwitz et al., 2014). The lower pH in marine ecosystems can increase the solubility of some metals thus increasing their bioavailability and/or toxicity to the organism, while also directly inducing physiological stress, such as decreased photosynthesis, respiration, calcification rates, and the rate of nitrogen-fixation (Ateweberhan et al., 2013; Horwitz et al., 2014; Houlbrèque et al., 2011). Exaiptasia pallida (formerly Aiptasia pallida and now synonymous with Aiptasia
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pulchella) contain photosynethetic dinoflagellate zooxanthellae, and occupy a range of nearshore environments in southeastern United States where metal pollution is more prevalent (Leal et al., 2012). This species has also been shown to be affected by both metals and ocean acidification (Siddiqui and Bielmyer-Fraser, 2015). For all of these reasons, E. pallida serve as useful surrogates for corals in toxicity testing. The objectives of this research were to assess tissue metal accumulation and physiological effects of elevated CO2 with Zn or Ni exposure in the sea anemone Exaiptasia pallida. 2. Materials and methods 2.1. Test organisms E. pallida were shipped from University of Miami (Miami, FL, USA) and maintained in a 30 L holding tank at Valdosta State University filled with synthetic saltwater with continuous filtration and aeration. Synthetic saltwater at a salinity of 30 ppt was prepared by mixing Instant Ocean salt and 18.2 mΩ Milli-Q water 24 h prior to use. Anemones were fed brine shrimp (Artemia sp.) ad libitum daily and acclimated to testing conditions for at least two weeks prior to testing. Salinity was measured using a portable refractometer (Aquatic Ecosystems, Inc.) and temperature and dissolved oxygen (DO) were measured using a YSI 85 Meter (YSI, OH, USA) daily. Measured values (mean ± standard deviations) for salinity, temperature and DO were as follows: 29.9 ± 0.45 ppt, 24.2 ± 0.48 ºC, and 8.28 ± 0.19 mg/L, respectively. Anemone wet and dry weight averaged 0.09 ± 0.05 g and 0.01 ± 0.007 g, respectively. 2.2. Experimental design Sixteen 10-gallon tanks were used in the experiments. Eight of these tanks contained a nominal CO2 value of 1000 ppm and the remaining eight tanks had no added CO2 and were at ambient conditions. The pH/pCO2 stat system used for this experiment is from Loligo systems
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with CAPCTRL software and set up by manual instruction. This system has been used in our laboratory previously (Siddiqui and Bielmyer-Fraser, 2015) and by several other investigators (Heuer et al., 2012). A standard curve for the calibration was prepared using a known CO2/O2 gas mixture of 2250 ppm. The pH of each tank was continuously monitored and regulated using an automated negative feedback system which dispensed pure CO2 gas into the water when needed. Each tank was also continuously aerated. Metal exposure concentrations were prepared from mixing 10 g/L stock solutions of NiCl2 and ZnCl2 (Ricca Chemical Company) with 30 g/L synthetic saltwater 24 h prior to the beginning of the experiment and 24 h prior to each water change. At ambient CO2 or 1000 ppm CO2, anemones were exposed to nominal concentrations of 0 (control), 10, 50, and 100 μg/L Ni or Zn in two sequential experiments and there were two replicate tanks for each metal concentration. At the start of each experiment, eight anemones were transferred from the holding tank to each experimental tank in the Ni experiment; and in the Zn experiment, six anemones were transferred to each control tank and four anemones to each Zn treatment. Additionally, five E. pallida were obtained from the holding tank, anesthetized with Tricaine methanesulfonate (MS222), and cut in half using a scalpel. Half of the anemone was collected for enzyme analysis and immediately frozen at -80 ºC. After dissection, the other halves were dried in an oven at 65 ºC for 24 h, weighed, and acidified using trace metal grade nitric acid for later metal analysis. This procedure was followed on 2, 4, and 7 d with two anemones collected per tank in the Ni experiment. For the Zn experiment, two anemones were collected from the control tank on 2 d, while on 4 and 7 d one anemone was collected. For all other tanks, one anemone was sampled on 2 and 4 d and two anemones were collected on 7 d. This pattern was due to the number of
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anemones available for the experiment. Behavior was monitored and recorded daily by taking pictures of every tank. Water samples were collected from each tank on each sampling day and before each 50 % water change. Temperature and DO were measured daily using a YSI 85 Meter (YSI, OH, USA). Salinity was measured daily using a portable refractometer (Aquatic Eco-Systems, Inc.). Ammonia, nitrate, and nitrite levels were measured at the beginning and end of each experiment using a LaMotte saltwater aqua-culture test kit (model AQ-2). The measured values (mean ± standard deviation) in the Ni experiment were 23.8 ± 0.30 ºC, 6.37 ± 0.10 mg/L DO, 30.0 ± 0.0 g/L salinity, 0.09 ± 0.12 mg/L ammonia, 0.00 ± 0.0 mg/L nitrite, and 0.00 ± 0.0 mg/L nitrate. The measured values (mean ± standard deviation) in the Zn experiment were 23.9 ± 0.52 ºC, 6.34 ± 0.11 mg/L DO, 30.1 ± 0.41 g/L salinity, 0.00 ± 0.0 mg/L ammonia, 0.00 ± 0.0 mg/L nitrite, and 0.00 ± 0.0 mg/L nitrate. Saltwater (50%) from each experimental tank was replaced on 2, 4, and 6 d. The pH was measured manually using a calibrated pH meter and probe (WTW, Xylem Inc., USA) in each of the tanks designated as ambient CO2, while the automated pH/pCO2 stat system measured the pH continuously in the tanks at 1000 ppm CO2 on a daily basis. Measured CO2 and pH values are provided in Table 1. 2.3. Metal analysis Water samples were filtered (0.45 μm Nylon, Fisher Scientific, Pittsburg, PA), acidified using trace metal grade nitric acid (Fisher Scientific, Pittsburg, PA), and diluted using 18.2 mΩ Milli-Q water. The samples were then analyzed for Ni or Zn using atomic absorption spectrophotometry (AAS; Perkin Elmer AAnalyst 800) with graphite furnace detection (detection limits ~ 1 µg/L). Anemone tissue samples were weighed, dried in an oven at 65 °C for 24 h, weighed again and acidified with trace metal grade nitric acid (Fisher Scientific, Pittsburg, PA). Samples were then diluted with 18.2 mΩ Milli-Q water and analyzed for Ni or
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Zn using AAS. Certified Ni and Zn standards from Ricca Chemical Company (Arlington, TX with 3% HCl) were used for each calibration, and recalibration was performed after every 40 samples. The metal accumulation in the tissue is presented as μg metal/g dry weight (dw). 2.4. Enzyme assays Frozen anemone samples were finely ground using a mortar and pestle on ice, and 5 mL 50 mM KH2PO4/K2HPO4 buffer (Sigma-Aldrich, MO, USA) was added. The homogenate was then transferred to polypropylene centrifuge tubes and centrifuged at 5000 rpm for 10 m at 4°C. The supernatant was collected into new tubes and preserved at -80°C for protein quantification. A Bradford assay (Bradford, 1976) was performed for total soluble protein quantification using the Bio-Rad® quick start kit (Bio-Rad, CA, USA) according to the manufacturer‘s instructions. Bovine serum albumin (BSA) standards (0, 0.02, 0.04, 0.08, 0.12, 0.24 mg/mL) were prepared from a 20 mg/mL stock solution and a 50 mM KH2PO4/K2HPO4 buffer. The absorbance was recorded for each sample and standard using PerkinElmer Lamda 35 UV/Vis spectrophotometer at a wavelength of 595 nm. CAT activity was measured following a Sigma method (Sigma, 1994a; EC 1.11.1.6). In short, an 11.9 mM hydrogen peroxide (H2O2) stock solution was prepared via dilution with a 50 mM phosphate monobasic buffer (pH 7.0, 25 °C). Anemone supernatant (500 μL) and 11.9 mM H2O2 (1500 μL) were added together in a quartz cuvette and inverted for 20 s. The absorbency was then measured at a wavelength of 240 nm for 90 s using a PerkinElmer Lamda 35 UV/Vis spectrophotometer. CAT activity was determined by the rate of H2O2 decomposition using the following equation: )(
( (
) )
Where, 9
(
) (
)
GR activity was measured using an assay kit (Sigma Aldrich; EC 1.6.4.2) and following a Sigma protocol (Sigma, 1994b). Using the UV assay method, standards were prepared by adding 500 μL of 2 mM oxidized glutathione, 450 μL of 0.1 M KH2PO4/K2HPO4, and 5 μL of 2 mM®-nicotinamide adenine dinucleotide phosphate, reduced form (NADPH), in a quartz cuvette. Samples were prepared by adding and mixing each of the following reagents in a quartz cuvette: 500 μL of 2 mM oxidized glutathione, 350 μL of Assay buffer, 100 μL of sample (homogenate of sample), and 50 μL of 2 mM NADPH. NADPH initiated the reaction and the change in absorbance was measured for 4 minutes at 340 nm using Perkin Elmer Lambda 35 UV/Vis spectrophotometer. GR activity in the sample was calculated using the following formula: ) (
( (
) )
Where, 2.5. Data analysis Data were tested for equality of variance and normal distribution using a Bartlett's test and Shapiro–Wilk's test, respectively. Statistical differences between treatments at each time point and over time were determined using a one-way analysis of variance (ANOVA) followed by the multiple comparison test of Tukey (p ≤ 0.05) with Sigma Plot 8.0. 3. Results Measured pCO2 and metal concentrations were similar to nominal values, except in the 100 µg/L treatment, where measured values were higher (Table 1). All concentrations were fairly stable throughout the experiments (Table 1). The pCO2 levels were relatively similar in both experiments. The pH values in the 1000 ppm CO2 exposed groups were ~0.2 pH units below those with no added CO2 (Table 1). 10
Ni and Zn accumulated in E. pallida tissues in a concentration-dependent manner; however, temporal differences were observed dependent on CO2 level (Figs. 1,2). Anemones exposed to 15 µg/L Ni at 1000 ppm CO2 for 48 h had significantly elevated tissue Ni, unlike in the ambient CO2 treatment; however, the tissue Ni values returned to control levels by 7 d (Fig. 1). Anemones exposed to 52 µg/L Ni at 1000 ppm CO2 had significantly elevated tissue Ni by 2 d, which persisted until 4 d and then returned to control levels by 7 d; whereas those anemones exposed to 51 µg/L Ni at ambient CO2 only had significantly elevated tissue Ni at 4 d (Fig. 1). The anemones exposed to 131 and 138 µg/L Ni with ambient and 1000 ppm CO2, respectively, had significantly elevated tissue Ni throughout the 7 d. Zn did not significantly accumulate in the tissues of anemones exposed to 13 µg/L Zn at either CO2 level (Fig. 2). Tissue Zn was significantly elevated in anemones exposed for 2 d to 74 µg/L Zn at 1000 ppm CO2 only. Anemones exposed to 141 and 144 µg/L Zn with ambient and 1000 ppm CO2, respectively, had significantly elevated tissue Zn throughout the experiments (Fig. 2). No significant differences in CAT activity were observed due to Ni or Zn exposure; however, there was a trend of increasing CAT activity in the anemones exposed to 51 µg/L Ni at ambient CO2 over the first 4 d (Fig. 3). No differences in GR activity were observed after exposure to 13 µg/L Ni or Zn at either CO2 concentration (Fig. 4). The pattern of GR activity in anemones exposed to 71 and 74 µg/L Zn differed between the two CO2 exposure groups, whereas such difference was not observed in anemones exposed to 51 and 52 µg/L Ni (Fig. 4). Anemones exposed to 144 µg/L Zn or 138 µg/L Ni at elevated CO2 had a significantly reduced GR activity by 7 d as compared to controls; however, no differences were observed in the anemones exposed to similar metal concentrations at ambient CO2 (Figs. 4, 5). The CO2 concentration significantly influenced the GR activity in the anemones exposed to 141 and 144
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µg/L Zn, but not in the anemones exposed to 131 and 138 µg/L Ni (Fig. 5). At 2 d, anemones exposed to 144 µg/L Zn and 1000 ppm CO2 had significantly lower GR activity and at 7 d significantly higher GR activity than anemones exposed to 141 µg/L Zn at ambient CO2 (Fig. 5). 4. Discussion Zn concentrations in E. pallida were similar to those reported from other studies. The anemones, Anemonia viridis and Actinia equina contained tissue Zn concentrations between 100 and 200 µg/g dw in waters containing up to 200 µg/L Zn in the environment (Harland et al., 1990). Additionally, tissue Zn in A. viridis exposed to 2-1000 µg/L Zn for 5 days in the laboratory accumulated up to 400 µg/g dw (Harland et al., 1990). Likewise, Mitchelmore et al. (2003) reported tissue Zn concentrations in A. viridis ranging between 200 and 400 µg/g dw after exposure to 100 µg/L Zn for 42 d. E. pallida exposed for 7 d to 10-100 µg/L of a metal mixture (Zn, Ni, Cu, Cd) contained tissue Zn concentrations ranging from 100 to 500 µg/g dw (Brock and Bielmyer, 2013). It‘s possible that increased Zn uptake occurs in the presence of other metals because E. pallida exposed to 141 µg/L of solely Zn in our study only contained up to 300 µg/g dw. Zn is a vital constituent for more than 200 enzymes and is generally well regulated in most organisms (Eisler, 1993; Viarengo and Nott, 1993) so it is not surprising that the tissue Zn concentrations are fairly similar between different anemone species. Like Zn, tissue Ni concentrations in E. pallida in this study were similar to those reported in A. viridis after exposure to the same range of concentrations for 42 d (Mitchelmore et al., 2003). Alternatively, when exposed to a Ni, Zn, Cd, and Cu mixture, E. pallida accumulated approximately 3.5 times more Ni than was observed in our study after exposure of E. pallida to solely Ni (Brock and Bielmyer, 2013). Ni is also an essential element to most organisms (Eisler 1998); however the baseline concentrations of Ni are much lower than Zn, indicating that Zn
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may be required at higher levels for normal functioning. Additionally, the anemones likely have better homeostatic regulation of Zn. At both CO2 levels, Ni and Zn tissue burdens decreased slightly from 4 d to 7 d. This was likely due to loss of zooxanthellae, which was not quantified in this study, but has been demonstrated in anemones and corals in other studies with increased exposure to metals (Bielmyer et al., 2010; Main et al., 2010; Brock and Bielmyer, 2013). Anemones exposed to Zn or Ni and 1000 ppm CO2 in the present study accumulated the metal significantly faster than those exposed to the same metal concentrations at ambient CO2, potentially due to increased metal bioavailability. At the higher CO2 level and lower pH, metal solubility increases and metal speciation changes (Millero et al., 2009; Lopez et al., 2010; Millero and DiTrolio, 2010). It should be noted that all the pH values (ambient and high CO2 treatments) in the present study were lower than expected. This could have been due to solubility changes in the reconstituted synthetic seawater, although this was not visibly noticed. In any case, the higher CO2 treatments did have significantly lower pH values than the ambient CO2 treatments. In general, as pH decreases, there is an increased formation of metal species that are more bioavailable; and therefore, an increase in toxicity to aquatic organisms. The results of the present work coincide with similar studies using copper in our laboratory. Siddiqui and Bielmyer-Fraser (2015) demonstrated increased Cu uptake at elevated CO2 concentrations in E. pallida. Likewise, anemones exposed to Cu at lower salinity and consequently lower pH exhibited increased Cu accumulation as compared to those exposed to higher salinity and pH (Patel and Bielmyer-Fraser, 2015). However, unlike Cu, the Zn and Ni tissue concentrations in this study returned to those in the ambient CO2 treatments after 96 h. Horwitz et al. (2014) reported substantial changes in tissue metal concentrations in the anemone, Anemonia viridis, located near a CO2 vent with elevated metal concentrations, as compared to a
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control site. Further, they reported increased concentrations of iron, lead, copper and cobalt; but decreased concentrations of Zn, cadmium, and arsenic in anemones close to the vent; with the pedal disk generally containing higher concentrations than the tentacles (Horwitz et al., 2014). Zn concentrations in the pedal disc ranged from 40-90 µg/g dw. The authors suggested that A. viridis regulates Zn well through compartmentalization and excretion. These field results are consistent to what we observed in our study where even though tissue Zn concentrations did increase faster when CO2 levels were elevated, tissue Zn concentrations quickly returned to levels observed in anemones exposed to Zn at ambient CO2. Physiological responses in the anemones were more varied. Exposure to 51 and 52 µg/L Ni did appear to increase CAT activity in the anemones; however, no other significant differences in CAT activity were observed due to either Ni or Zn exposure (Fig. 2). Alternatively, Cu (Main et al., 2010; Patel and Bielmyer-Fraser, 2015; Siddiqui et al., 2015; Siddiqui and Bielmyer-Fraser, 2015) and combined exposure of Cu, Cd, Ni, and Zn (Brock and Bielmyer, 2013) have been shown to increase CAT activity. Significant differences in CAT activity due to elevated CO2 were not observed in this study, consistent with a previous study (Siddiqui and Bielmyer-Fraser, 2015). Exposure to 131 µg/L Ni or 141 µg/L Zn caused a significant reduction in GR activity by 7 d; however, at elevated CO2, the degree of GR inhibition lessened (Fig. 4). Alternative to Ni and Zn, exposure to Cu alone or a mixture of Cu, Cd, Zn and Ni have been shown to cause an increase in GR activity in past studies (Brock and Bielmyer, 2013; Siddiqui and Bielmyer-Fraser, 2015; Siddiqui et al., 2015). The anti-oxidant enzymes measured in this study are known to be vital defenses against ROS (Livingstone et al., 1992; Regoli et al., 2002). Elevated metal concentrations have been demonstrated to cause oxidative damage to proteins, lipids, and DNA, when oxidative defense mechanisms are
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overwhelmed (Krumschnabel et al., 2005; Lushchak, 2011). In the present study, GR was affected and CAT was unaffected by increasing Zn or Ni concentrations and elevated CO2 concentration, suggesting that GPx or another enzyme is likely used for the conversion of peroxide into water, rather than CAT. Additionally, in past studies with Cu (Patel and BielmyerFraser, 2015; Siddiqui and Bielmyer-Fraser, 2015) and increased CO2, both parameters individually and in combination (Siddiqui and Bielmyer-Fraser, 2015), caused an increase in GR activity. On the contrary, exposure to Zn or Ni caused a decrease in GR activity and exposure to elevated CO2 lessened that inhibition. Therefore, Ni and Zn may cause toxicity and elicit antioxidant responses via other pathways than does Cu. 5. Conclusions Ni and Zn exposure to E. pallida resulted in tissue metal accumulation and the rate of accumulation was increased with exposure of 1000 ppm CO2. Tissue metal concentration remained elevated in the highest exposure concentrations of 144 µg/L Zn or 138 µg/L Ni by 7 d. Both Ni and Zn were well regulated in the anemone and the effects of CO2 exposure on tissue metal concentration were no longer observed after 96 h. Varying physiological responses were observed due to exposure of Zn or Ni and elevated CO2. No significant differences in CAT activity were observed due to elevated Ni, Zn or CO2. GR activity was significantly reduced by 131 µg/L Ni or 141 µg/L Zn, and the inhibition was decreased by addition of 1000 ppm CO2. This response differs from the increased GR activity demonstrated in anemones exposed to Cu. The metal concentrations used in this study are within the range of those reported in polluted aquatic environments. These results provide new data concerning metal accumulation and physiological responses in sea anemones exposed to acidification and Zn or Ni. Furthermore,
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these findings are important for application to environmental pollution scenarios with multiple contaminants. Acknowledgements Financial support was provided by individual contributions to the crowdfunding site CREU (creu.tilt.com). Contributors included the following: Southern Oak Insurance Company, Doris and William Bielmyer, T.J. Fraser, Brad Eavenson, Edwin Lunsford, William Lynch, Bill and Michelle Bielmyer, Wayne Richmond, Joseph Pearce, Jenny and Tom Fraser, Holly Schopp, Ben Harper, Bill Tomson, Bob DeSmedt, Chad Lotocki, Herb LeMoyne, Donald Reoderique, Desmond McElroy, and Larry Holder. We would also like to thank the VSU Biology department and Graduate school for funding; Phil Gillette from the University of Miami for the anemones; Pavan Patel for help in the laboratory; and Dr. T.J. Grove for the use of a PerkinElmer Lambda 35 UV/Vis spectrophotometer that was purchased by National Science Foundation grant funds (IOS-0817805to TJG). References Albright, R., Langdon, C., 2011. Ocean acidification impacts multiple early life history processes of the Caribbean coral Porites astreoides. Glob. Change Biol. 17, 2478–2487. Anthony, K.R.N., Maynard, J.A., Diaz-Pulido, G., Mumby, P.J., Marshalls, P.A., Cao, L., Hoegh-Guldberg, O., 2011. Ocean acidification and warming will lower coral reef resilience. Glob. Change Biol. 17, 1798-1808. Ateweberhan, M., Feary, D.A., Keshavmurthy, S., Chen, A., Schleyer, M.H., Sheppard, C.R.C., 2013. Climate change impacts on coral reefs: Synergies with local effects, possibilities for acclimation, and management implications. Mar. Pollut. Bull. 74, 526-539. Bielmyer, G.K., Grosell, M., Bhagooli, R., Baker, A.C., Langdon, C., Gillette, P., Capo, T., 2010. Differential effects of copper on three species of scleractinian corals and their algal Symbionts (Symbiodinium spp.). Aquat. Toxicol. 97, 125-133. Bradford, M.M., 1976. Rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Anal. Biochem. 72, 248–254.
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Brock, J.R., Bielmyer, G.K., 2013. Metal accumulation and sublethal effects in the sea anemone Aiptasia pallida, after waterborne exposure to metal mixtures. Comp. Biochem. Physiol. C. 158, 150-158. Brosnan, J.T., Brosnan, M.E., 2009. Glutathione and sulfur-containing amino acids: An overview, in: Masella,R., Mazza,G. (Eds.), Glutathione and sulfur amino acids in human health and Disease. John Wiley & Sons, Hohoken, New Jersey, pp. 3-15. Brown, B.E., Howard, L.S., 1985. Assessing the effects of ‗stress‘ on reef corals. Adv. Mar. Biol. 22, 1–63. Caldeira, K., Wickett, M.E., 2003. Anthropogenic carbon and ocean pH. Nature 425, 365. Eisler, R., 1993. Zinc hazards to fish, wildlife, and invertebrates. Contaminant Hazard Reviews 26, 1–79. Eisler R., 1998. Nickel hazards to fish, wildlife, and invertebrates: A synoptic review. Contaminant Hazard Reviews, Report 34. Biological Science Report USGS/BRD/BSR1998-0001. U.S. Geological Survey, Washington, DC, 1–95. Esbaugh, A.J., Heuer, R.M., Grosell, M., 2012. Impacts of ocean acidification on respiratory gas exchange and acid base balance in a marine teleost, Opsanus beta. J. Comp. Physiol. B. 182, 921-934. Evans, S.M., Birchenough, A.C., Brancato, M.S., 2000. The TBT ban: out of the frying pan into the fire? Mar. Pollut. Bull. 40, 204–211. Fabry, V.J., Seibel, B.A., Feely, R.A., Orr, J.C., 2008. Impacts of ocean acidification on marine fauna and ecosystem processes. ICES. J. Mar. Sci. 65, 414–432. Ferrier-Pages, C., Houlbreque, F., Wyse, E., Richard, C., Allemand, D., Boisson, F., 2005. Bioaccumulation of zinc in the scleractinian coral Stylophora pistilla. Coral Reefs 24, 636-645. Forman, H.J., Liu, R., Shi, M.M., 1990. Glutathione synthesis in oxidative stress. In: Packer, L., Fuchs, J. (Eds.), Biothiols in Health and Diseases. Marcel Dekker, NY, 189–211. Gomez, C.E., Paul, V.J., Riston-Williams, R., Muehllehner, N., Langdon, C., Sanchez, J.A., 2014. Responses of the tropical gorgonian coral Eunicia fusca to ocean acidification conditions. Coral Reefs 34(2), 451-460. Gonzalez, H., Pomares, M., Ramirez, M. & Torres, I., 1999. Heavy metals in organism and sediments from the discharge zone of the submarine sewage outfall of Havanna city, Cuba. Mar. Pollut. Bull. 38, 1048-1051. Guzmán, H.M., Jiménez, C.E., 1992. Contamination of coral reefs by heavy metals along the 17
Caribbean coast of Central America (Costa Rica and Panama). Mar. Pollut. Bull. 24, 554561. Harland, A.D., Bryan, G.W., Brown, B.E., 1990. Zinc and cadmium absorption in the symbiotic anemone Anemonia viridis and the non-symbiotic anemone Actinia equina. J. Mar. Biol. Assoc. U. K. 70, 789–802. Heuer, R., Esbaugh, A., Grosell, M., 2012. Ocean acidification leads to counterproductive intestinal base loss in the gulf toadfish (Opsanus beta). Physiol. Biochem. Zool. 85(5), 450-459. Higuchi, T., Fujimura, H., Hitomi, Y., Arakaki, T., Oomori, T. and Suzuki, Y., 2010. Photochemical formation of hydroxyl radicals in tissue extracts of the coral Galaxea fascicularis. Photochem. Photobiol. 86(6), 1421-1426. Horwitz, R., Borell, E.M., Fine, M., Shaked, Y., 2014. Trace element profiles of the sea anemone Anemonia viridis living nearby a natural CO2 vent. Peer J. 2:e538; DOI 10.7717/peerj.538. Howard, L.S., Brown, B.E., 1984. Heavy metals and reef corals. Oceanogr. Mar. Biol. Ann. Rev. 22, 195–210. Houlbrèque, F., Rodolfo-Metalpa, R., Jeffree, R., Oberhansli, F., Teyssie, J.L., Boisson, F., AlTrabeen, K., Ferrier-Pages, C., 2012. Effects of increased pCO2 on zinc uptake and calcification in the tropical coral Stylophora pistillata. Coral Reefs 31, 101-109. DOI 10.1007/s00338-011-0819-2. Jones, R., 2010. Environmental contamination associated with a marine landfill (seafill) beside a coral reef. Mar. Pollut. Bull. 60, 1993–2006. Krumschnabel, G., Manzl, C., Berger, C., Hofer, B., 2005. Oxidative stress, mitochondrial permeability transition, and cell death in Cu exposed trout hepatocytes. Toxicol. Appl. Pharmacol. 209 (1), 62-73. Leal, M.C., Nunes, C., Engrola, S., Dinis, M.T., Calado, R., 2012. Optimization of monoclonal production of the glass anemone Aiptasia pallida (Agassiz in Verrill, 1864). Aquaculture 354–355, 91–96. Livingstone, D.R., Lips, F., Garcia, M.P., Pipe, R.K., 1992. Antioxidant enzymes in digestive gland of the common mussel Mytilus edulis. Mar. Biol. 112, 265–276. Lopez, I.R., Kalman, J., Vale, C., Blasco, J.. 2010. Influence of sediment acidification on the bioaccumulation of metals in Ruditapes philippinarum. Environ. Sci. Poll. Res. 17, 1519– 1528.
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Lushchak, V.I., 2011. Environmentally induced oxidative stress in aquatic animals. Aquat. Toxicol. 101, 13-30. Main, W.P.L., Ross, C., Bielmyer, G.K., 2010. Copper accumulation and oxidative stress in the sea anemone, Aiptasia pallida, after waterborne copper exposure. Comp. Biochem. Physiol. C. 151, 216–221. Meehl, G.A., Stocker, T.F., Collins, W.D., Friedlingstein, P., Gaye, A.T., Gregory, J.M., Kitoh, A., 2007. Global Climate Projections. Climate Change 2007: The Physical Science Basis. 10, 749-844. Millero, F.J., Woosley, R., Ditrolio, B., Waters, J., 2009. Effect of ocean acidification on the speciation of metals in seawater. Oceanography 22, 72–85. Millero, F.J., DiTrolio, B.R., 2010. Use of thermodynamics in examining the effects of ocean acidification. Elements. 6, 299–303. Mitchelmore, C.L., Verde, E.A., Ringwood, A.H., Weis, V.M., 2003. Differential accumulation of heavy metals in the sea anemone Anthopleura elegantissima as a function of symbiotic state. Aquat. Toxicol. 64, 317–329. Naoum, C., Fatta, D., Haralambous, K.J., Loizidou, M., 2001. Removal of heavy metals from sewage sludge by acid treatment. J. Environ. Sci. Health, Part A. 36(5), 873-881. Patel, P.P., Bielmyer-Fraser, G.K., 2015. The influence of salinity and copper exposure on copper accumulation and physiological impairment in the sea anemone Exaiptasia pallida. Comp. Biochem. Physiol. C. 168, 39-47. Peters, E.C., Gassman, N.J., Firman, J.C., Richmond, R.H., Power, E.A., 1997. Ecotoxicology of tropical marine ecosystems. Environ. Toxicol. Chem. 16, 12-40. Poonkothai, M.V.B.S., Vijayavathi, B.S., 2012. Nickel as an essential element and a toxicant. Int. J. Environ. Sci. 1, 285-288. Regoli, F., Gorbi, S., Frenzilli, G., Nigro, M., Corsi, I., Focardi, S., Winston, G.W., 2002. Oxidative stress in ecotoxicology: from the analysis of individual antioxidants to a more integrated approach. Mar. Environ. Res. 54, 419–423. Reichelt, A.J., Jones, G.B., 1994. Trace metals as tracers of dredging activity in Cleveland Bay—field and laboratory studies. Mar. Freshw. Res. 45(7), 1237-1257. Siddiqui, S., Goddard, R.H., Bielmyer-Fraser, G.K., 2015. Comparative effects of dissolved copper and copper oxide nanoparticle exposure to the sea anemone, Exaiptasia pallida. Aquat. Toxicol. 160, 205-213. Siddiqui, S., Bielmyer-Fraser, G.K., 2015. Responses of the sea anemone, Exaiptasia pallida, 19
to ocean acidification conditions and copper exposure. Aquat. Toxicol. 167, 228-239. Sies, H., 1999. Glutathione and its role in cellular functions. Free Radic. Biol. Med. 27(9), 916921. Sigma, 1994a. Enzyme Assay of Catalase (EC 1. 11. 1. 6). www.sigmaaldrich.com. Sigma, 1994b. Enzamatic Assay of Glutathione Reductase (EC 1.6.4.2). www.sigmaaldrich.com. Stylianous, M.A., Kolia, D., Haralambous, K.J., Inglezakis, V.J., Moustakas, K.G., Loizidous, M.D., 2007. Effect of acid treatment on the removal of heavy metals from sewage sludge. Desalination. 215, 73–81. Sunagawa, S., Choi, J., Forman, H.J., Meidna, M., 2008. Hyperthermic stress-induced in- crease in the expression of glutamate–cysteine ligase and glutathione levels in the symbiotic sea anemone Aiptasia pallida. Comp. Biochem. Physiol. B. 151, 133–138. Towle, E.K., Enochs, I.C., Langdon, C., 2015. Threatened Caribbean coral is able to mitigate the adverse effects of ocean acidification on calcification by increasing feeding rate. Plos. One. 10, 1-17. Viarengo, A., Nott, J.A., 1993. Mechanisms of heavy metal cation homeostasis in marine invertebrates. Comp. Biochem. Physiol. C. 104, 355–372. Yakovleva, I., Bhagooli, R., Takemura, A., Hidaka, M., 2004. Differential susceptibility to oxidative stress of two scleractinian corals: anti-oxidant functioning of mycosporineglycine. Comp. Biochem. Physiol. B. 139, 721-730. Voulvoulis, N., Scrimshaw, M.D., Lester, J.N., 2000. Occurrence of four biocides utilized in antifouling paints, as alternatives to organotin compounds, in waters and sediments of a commercial estuary in the UK. Mar. Pollut. Bull. 40 (11), 938-946.
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Table 1. Measured pCO2, pH, and dissolved metal concentrations (mean ± standard deviation) in the experimental solutions over the 7 day exposure periods. Nominal Ni pCO2 (ppm) treatment (µg/L) Control 10 50 100 Control 10 50 100
Ambient Ambient Ambient Ambient 963 ± 30 1180 ± 650 1180 ± 30 1020 ± 30
Nominal Zn pCO2 (ppm) treatment (µg/L) Control 10 50 100 Control 10 50 100
Ambient Ambient Ambient Ambient 1130 ± 40 1120 ± 330 1240 ± 60 1050 ± 90
pH 8.05 ± 0.01 7.88 ± 0.04 7.88 ± 0.01 7.86 ± 0.02 7.77 ± 0.08 7.79 ± 0.05 7.76 ± 0.10 7.80 ± 0.06 pH 7.92 ± 0.07 7.90 ± 0.04 7.91 ± 0.08 7.88 ± 0.08 7.74 ± 0.09 7.77 ± 0.05 7.77 ± 0.07 7.73 ± 0.10
Measured [Ni] (µg/L) 2 ± 1.3 13 ± 1.6 51 ± 6.9 131 ± 22.2 4 ± 1.9 15 ± 2.0 52 ± 6.4 138 ± 17.0 Measured [Zn] (µg/L) 4 ± 1.8 13 ± 1.3 71 ± 5.6 141 ± 11.1 4 ± 2.1 13 ± 1.1 74 ± 3.8 144 ± 11.6
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Figure 1. Nickel accumulation (mean ± standard error; µg/g dw) in Exaiptasia pallida exposed to nickel at (A) ambient CO2 and (B) 1000 ppm CO2. * indicate significant differences. Refer to Table 1 for measured nickel concentrations.
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Figure 2. Zinc accumulation (mean ± standard error; µg/g dw) in Exaiptasia pallida exposed to zinc at (A) ambient CO2 and (B) 1000 ppm CO2. * indicate significant differences. Refer to Table 1 for measured zinc concentrations.
A. Zn
C. Ni
B. Zn
D. Ni
Figure 3. Catalase (CAT) activity (mean ± standard error; units/mg protein) in Exaiptasia pallida exposed to A. zinc at ambient CO2, B. zinc at 1000 ppm CO2, C. nickel at ambient CO2, D. nickel at 1000 ppm CO2. Refer to Table 1 for measured metal concentrations.
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A. Zn
C. Ni
B. Zn
D. Ni
Figure 4. Glutathione reductase (GR) activity (mean ± standard error; units/mg protein) in Exaiptasia pallida exposed to A. zinc at ambient CO2, B. zinc at 1000 ppm CO2, C. nickel at ambient CO2, D. nickel at 1000 ppm CO2. * indicate significant differences. Refer to Table 1 for measured metal concentrations.
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A. Zn
B. Ni
Figure 5. Glutathione reductase (GR) activity (mean ± standard error; units/mg protein) in Exaiptasia pallida exposed to control or 100 µg/L of A. zinc and B. nickel at ambient CO2 and 1000 ppm CO2. Circles represent control groups and squares represent metal treatments. Filled symbols represent ambient CO2 and open symbols represent 1000 ppm CO2. * indicate significant differences between ambient and 1000 ppm CO2 at a specified metal concentration. Refer to Table 1 for measured metal concentrations.
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S0166-445X(16)30344-7 http://dx.doi.org/doi:10.1016/j.aquatox.2016.11.014 AQTOX 4537
To appear in:
Aquatic Toxicology
Received date: Revised date: Accepted date:
10-10-2016 11-11-2016 16-11-2016
Please cite this article as: Duckworth, Christina G., Picariello, Codie R., Thomason, Rachel K., Patel, Krina S., Bielmyer-Fraser, Gretchen K., Responses of the sea anemone, Exaiptasia pallida, to ocean acidification conditions and zinc or nickel exposure.Aquatic Toxicology http://dx.doi.org/10.1016/j.aquatox.2016.11.014 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Responses of the sea anemone, Exaiptasia pallida, to ocean acidification conditions and zinc or nickel exposure
Christina G. Duckworth1, Codie R. Picariello1, Rachel K. Thomason1, Krina S. Patel1, Gretchen K. Bielmyer-Fraser2
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Department of Biology
Valdosta State University, Valdosta, GA, USA 2
Department of Chemistry
Jacksonville University, Jacksonville, FL, USA
Corresponding Author: Gretchen K. Bielmyer-Fraser Department of Chemistry, Jacksonville University, 2800 University Blvd. North, Jacksonville, FL 32211; Email: [email protected]
Highlights
Ni and Zn exposure to E. pallida resulted in concentration-dependent tissue metal accumulation and the rate of accumulation was increased with exposure of 1000 ppm CO2
Both Ni and Zn were well regulated in the anemone and the effects of CO2 exposure on tissue metal concentration were no longer observed after 96 h
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GR activity was significantly reduced by increased Ni or Zn, and the inhibition was decreased by addition of 1000 ppm CO2.
Abstract Ocean acidification, caused by increasing atmospheric carbon dioxide (CO2), is a growing concern in marine environments. Land-based sources of pollution, such as metals, have also been a noted problem; however, little research has addressed the combined exposure of both pollutants to coral reef organisms. In this study we examined tissue metal accumulation and physiological effects (activity of anti-oxidant enzymes, catalase and glutathione reductase) in the sea anemone, Exaiptasia pallida after exposure to increased CO2, as well as zinc (Zn) or nickel (Ni). After exposure to four concentrations (nominal values = control, 10, 50, 100 µg/L) of Zn or Ni over 7 days, both metals accumulated in the tissues of E. pallida in a concentrationdependent manner. Anemones exposed to elevated CO2 (1000 ppm) accumulated significant tissue burdens of Zn or Ni faster (by 48 h) than those exposed to the same metal concentrations at ambient CO2. No differences were observed in catalase activity due to Zn exposure; however, 50 µg/L Ni caused a significant increase in catalase activity at ambient CO2. No significant effect on catalase activity from CO2 exposure alone was observed. Glutathione reductase activity was affected by increased Zn or Ni exposure and those effects were influenced by increased CO2. Results of this study provide insight into the toxic mechanisms and environmental implications of CO2 and Zn or Ni exposure to the cnidarian E. pallida.
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1. Introduction Coral reefs are among the most diverse and productive ecosystems on Earth, containing a wealth of species complexity and substantial economic importance (Brown and Howard, 1985). Despite their vital role in the global economy and ecology, these valuable marine environments have experienced vast degradation in recent years (Brown and Howard, 1985; Albright and Langdon, 2011). Ocean acidification has become a prevalent environmental stressor to many coral reef species. Current oceanic values of CO2 approximate 380 µatm, and by the years 2100 and 2300 these values are expected to rise to 1,000 and 1,900 µatm, respectively, which would cause a decrease of up to 0.77 pH units with the higher CO2 level (Caldeira and Wickett 2003; Meehl et al., 2007; Fabry et al. 2008; Esbaugh et al., 2012; Albright and Langdon, 2011). The uptake of atmospheric CO2 by the ocean alters the ocean carbonate chemistry, lowering the concentration and saturation states of calcium carbonate (CaCO3) minerals (Gomez et al., 2014). Several studies have demonstrated a negative correlation with higher concentrations of CO2 in coral reef organisms that rely on a saturated state of CO32- for calcification and growth (Gomez et al., 2014; Ateweberhan et al., 2013). Towle et al. (2015) reported a decline of as much as 98% in the dominant reef-building coral A. cervicornis in the Florida Reef Tract since the 1970s. Environmental stressors affecting calcification and the coral-algal symbiosis may induce a phase shift from communities of hard, reef-building corals to more resilient species of soft corals and fleshy macroalgae (Peters et al., 1997; Gomez et al., 2014; Anthony et al., 2011).
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In addition to the global stressors of ocean acidification, coral reef ecosystems are also faced with more localized stressors, such as heavy metals, from land-based anthropogenic sources (Howard and Brown, 1984; Anthony et al., 2011). Many reefs are located near densely populated areas with substantial industrialization and coastal development. Sewage discharge, dredging, coastal petroleum refineries, fossil fuel combustion, sacrificial anodes on boats, leachate from metal-based antifouling paints, marine disposal of municipal solid waste and metallic bulk waste, mining, smelting, refining, and alloy processing, and many other agricultural and industrial activities contribute to increased heavy metal pollution in nearshore marine environments (Howard and Brown, 1984; Guzmán and Jiménez, 1992; Reichelt and Jones, 1994; Gonzalez et al., 1999; Evans et al., 2000; Voulvoulis et al., 2000; Naoum et al., 2001; Stylianous et al., 2007; Jones, 2010). Zinc and nickel are both essential metals to marine organisms and function in multiple roles as a cofactorsfor enzymes and in DNA. As essential metals, they are needed in low concentrations, but may become toxic at concentrations above certain thresholds (Poonkothai and Vijayavathi, 2012; Ferrier-Pages et al., 2005). High concentrations of zinc can affect main physiological functions, such as calcification rates, and can affect photosynthetic efficiency and proper enzyme function in the coral symbionts, zooxanthellae (Houlbrèque et al., 2011; Ferrier-Pages et al., 2005). Heavy metals have been shown to accumulate in coral reef organisms, particularly those containing algal symbionts such as zooxanthellae (Peters et al., 1997; Main et al., 2010; Horwitz et al., 2014). In many cases, corals and anemones exposed to elevated concentrations of metals will release their symbiotic algae as a possible method of detoxification, since algae have been shown to accumulate some metals to a larger extent than their animal symbiont (Bielmyer et al., 2010; Horwitz et al., 2014). This process of ―bleaching‖ is a potentially detrimental condition
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and can occur in conjunction with physiological disruptions in the holobiont (Biemyer et al., 2010; Main et al., 2010; Brock and Bielmyer, 2013; Patel and Bielmyer-Fraser, 2015). Physiological effects of metal exposure include changes in the activity of antioxidant enzymes (Main et al., 2010; Brock and Bielmyer, 2013). When exposed to metals, corals and anemones containing a photosynthetic symbiont are susceptible to increased production of reactive oxygen species (ROS) that can denature proteins, mutate DNA, and cause lipid peroxidation (Yakovleva et al., 2004). Antioxidant defenses utilizing the enzymes glutathione peroxidase (GPx), glutathione reductase (GR), and catalase (CAT), have been shown to reduce these harmful effects (Yakovleva et al., 2004; Brock and Bielmyer, 2013; Patel and Bielmyer-Fraser, 2015; Siddiqui et al., 2015; Siddiqui and Bielmyer-Fraser, 2015). GPx and CAT are produced to catalyze the conversion of hydrogen peroxide into water and oxygen, thus combating the effects of ROS (Higuchi et al., 2010; Forman et al., 1990; Sies, 1999; Sunagawa et al., 2008; Brosnan and Brosnan, 2009). GR is then produced to reduce glutathione so that it may be recycled for the reaction above (Forman et al., 1990; Sies, 1999; Sunagawa et al., 2008; Brosnan and Brosnan, 2009). Though several studies have examined the effects of ocean acidification or metal exposure, few studies have analyzed the effects of both stressors combined (Houlbrèque et al., 2011; Gomez et al., 2014; Peters et al., 1997; Horwitz et al., 2014). The lower pH in marine ecosystems can increase the solubility of some metals thus increasing their bioavailability and/or toxicity to the organism, while also directly inducing physiological stress, such as decreased photosynthesis, respiration, calcification rates, and the rate of nitrogen-fixation (Ateweberhan et al., 2013; Horwitz et al., 2014; Houlbrèque et al., 2011). Exaiptasia pallida (formerly Aiptasia pallida and now synonymous with Aiptasia
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pulchella) contain photosynethetic dinoflagellate zooxanthellae, and occupy a range of nearshore environments in southeastern United States where metal pollution is more prevalent (Leal et al., 2012). This species has also been shown to be affected by both metals and ocean acidification (Siddiqui and Bielmyer-Fraser, 2015). For all of these reasons, E. pallida serve as useful surrogates for corals in toxicity testing. The objectives of this research were to assess tissue metal accumulation and physiological effects of elevated CO2 with Zn or Ni exposure in the sea anemone Exaiptasia pallida. 2. Materials and methods 2.1. Test organisms E. pallida were shipped from University of Miami (Miami, FL, USA) and maintained in a 30 L holding tank at Valdosta State University filled with synthetic saltwater with continuous filtration and aeration. Synthetic saltwater at a salinity of 30 ppt was prepared by mixing Instant Ocean salt and 18.2 mΩ Milli-Q water 24 h prior to use. Anemones were fed brine shrimp (Artemia sp.) ad libitum daily and acclimated to testing conditions for at least two weeks prior to testing. Salinity was measured using a portable refractometer (Aquatic Ecosystems, Inc.) and temperature and dissolved oxygen (DO) were measured using a YSI 85 Meter (YSI, OH, USA) daily. Measured values (mean ± standard deviations) for salinity, temperature and DO were as follows: 29.9 ± 0.45 ppt, 24.2 ± 0.48 ºC, and 8.28 ± 0.19 mg/L, respectively. Anemone wet and dry weight averaged 0.09 ± 0.05 g and 0.01 ± 0.007 g, respectively. 2.2. Experimental design Sixteen 10-gallon tanks were used in the experiments. Eight of these tanks contained a nominal CO2 value of 1000 ppm and the remaining eight tanks had no added CO2 and were at ambient conditions. The pH/pCO2 stat system used for this experiment is from Loligo systems
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with CAPCTRL software and set up by manual instruction. This system has been used in our laboratory previously (Siddiqui and Bielmyer-Fraser, 2015) and by several other investigators (Heuer et al., 2012). A standard curve for the calibration was prepared using a known CO2/O2 gas mixture of 2250 ppm. The pH of each tank was continuously monitored and regulated using an automated negative feedback system which dispensed pure CO2 gas into the water when needed. Each tank was also continuously aerated. Metal exposure concentrations were prepared from mixing 10 g/L stock solutions of NiCl2 and ZnCl2 (Ricca Chemical Company) with 30 g/L synthetic saltwater 24 h prior to the beginning of the experiment and 24 h prior to each water change. At ambient CO2 or 1000 ppm CO2, anemones were exposed to nominal concentrations of 0 (control), 10, 50, and 100 μg/L Ni or Zn in two sequential experiments and there were two replicate tanks for each metal concentration. At the start of each experiment, eight anemones were transferred from the holding tank to each experimental tank in the Ni experiment; and in the Zn experiment, six anemones were transferred to each control tank and four anemones to each Zn treatment. Additionally, five E. pallida were obtained from the holding tank, anesthetized with Tricaine methanesulfonate (MS222), and cut in half using a scalpel. Half of the anemone was collected for enzyme analysis and immediately frozen at -80 ºC. After dissection, the other halves were dried in an oven at 65 ºC for 24 h, weighed, and acidified using trace metal grade nitric acid for later metal analysis. This procedure was followed on 2, 4, and 7 d with two anemones collected per tank in the Ni experiment. For the Zn experiment, two anemones were collected from the control tank on 2 d, while on 4 and 7 d one anemone was collected. For all other tanks, one anemone was sampled on 2 and 4 d and two anemones were collected on 7 d. This pattern was due to the number of
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anemones available for the experiment. Behavior was monitored and recorded daily by taking pictures of every tank. Water samples were collected from each tank on each sampling day and before each 50 % water change. Temperature and DO were measured daily using a YSI 85 Meter (YSI, OH, USA). Salinity was measured daily using a portable refractometer (Aquatic Eco-Systems, Inc.). Ammonia, nitrate, and nitrite levels were measured at the beginning and end of each experiment using a LaMotte saltwater aqua-culture test kit (model AQ-2). The measured values (mean ± standard deviation) in the Ni experiment were 23.8 ± 0.30 ºC, 6.37 ± 0.10 mg/L DO, 30.0 ± 0.0 g/L salinity, 0.09 ± 0.12 mg/L ammonia, 0.00 ± 0.0 mg/L nitrite, and 0.00 ± 0.0 mg/L nitrate. The measured values (mean ± standard deviation) in the Zn experiment were 23.9 ± 0.52 ºC, 6.34 ± 0.11 mg/L DO, 30.1 ± 0.41 g/L salinity, 0.00 ± 0.0 mg/L ammonia, 0.00 ± 0.0 mg/L nitrite, and 0.00 ± 0.0 mg/L nitrate. Saltwater (50%) from each experimental tank was replaced on 2, 4, and 6 d. The pH was measured manually using a calibrated pH meter and probe (WTW, Xylem Inc., USA) in each of the tanks designated as ambient CO2, while the automated pH/pCO2 stat system measured the pH continuously in the tanks at 1000 ppm CO2 on a daily basis. Measured CO2 and pH values are provided in Table 1. 2.3. Metal analysis Water samples were filtered (0.45 μm Nylon, Fisher Scientific, Pittsburg, PA), acidified using trace metal grade nitric acid (Fisher Scientific, Pittsburg, PA), and diluted using 18.2 mΩ Milli-Q water. The samples were then analyzed for Ni or Zn using atomic absorption spectrophotometry (AAS; Perkin Elmer AAnalyst 800) with graphite furnace detection (detection limits ~ 1 µg/L). Anemone tissue samples were weighed, dried in an oven at 65 °C for 24 h, weighed again and acidified with trace metal grade nitric acid (Fisher Scientific, Pittsburg, PA). Samples were then diluted with 18.2 mΩ Milli-Q water and analyzed for Ni or
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Zn using AAS. Certified Ni and Zn standards from Ricca Chemical Company (Arlington, TX with 3% HCl) were used for each calibration, and recalibration was performed after every 40 samples. The metal accumulation in the tissue is presented as μg metal/g dry weight (dw). 2.4. Enzyme assays Frozen anemone samples were finely ground using a mortar and pestle on ice, and 5 mL 50 mM KH2PO4/K2HPO4 buffer (Sigma-Aldrich, MO, USA) was added. The homogenate was then transferred to polypropylene centrifuge tubes and centrifuged at 5000 rpm for 10 m at 4°C. The supernatant was collected into new tubes and preserved at -80°C for protein quantification. A Bradford assay (Bradford, 1976) was performed for total soluble protein quantification using the Bio-Rad® quick start kit (Bio-Rad, CA, USA) according to the manufacturer‘s instructions. Bovine serum albumin (BSA) standards (0, 0.02, 0.04, 0.08, 0.12, 0.24 mg/mL) were prepared from a 20 mg/mL stock solution and a 50 mM KH2PO4/K2HPO4 buffer. The absorbance was recorded for each sample and standard using PerkinElmer Lamda 35 UV/Vis spectrophotometer at a wavelength of 595 nm. CAT activity was measured following a Sigma method (Sigma, 1994a; EC 1.11.1.6). In short, an 11.9 mM hydrogen peroxide (H2O2) stock solution was prepared via dilution with a 50 mM phosphate monobasic buffer (pH 7.0, 25 °C). Anemone supernatant (500 μL) and 11.9 mM H2O2 (1500 μL) were added together in a quartz cuvette and inverted for 20 s. The absorbency was then measured at a wavelength of 240 nm for 90 s using a PerkinElmer Lamda 35 UV/Vis spectrophotometer. CAT activity was determined by the rate of H2O2 decomposition using the following equation: )(
( (
) )
Where, 9
(
) (
)
GR activity was measured using an assay kit (Sigma Aldrich; EC 1.6.4.2) and following a Sigma protocol (Sigma, 1994b). Using the UV assay method, standards were prepared by adding 500 μL of 2 mM oxidized glutathione, 450 μL of 0.1 M KH2PO4/K2HPO4, and 5 μL of 2 mM®-nicotinamide adenine dinucleotide phosphate, reduced form (NADPH), in a quartz cuvette. Samples were prepared by adding and mixing each of the following reagents in a quartz cuvette: 500 μL of 2 mM oxidized glutathione, 350 μL of Assay buffer, 100 μL of sample (homogenate of sample), and 50 μL of 2 mM NADPH. NADPH initiated the reaction and the change in absorbance was measured for 4 minutes at 340 nm using Perkin Elmer Lambda 35 UV/Vis spectrophotometer. GR activity in the sample was calculated using the following formula: ) (
( (
) )
Where, 2.5. Data analysis Data were tested for equality of variance and normal distribution using a Bartlett's test and Shapiro–Wilk's test, respectively. Statistical differences between treatments at each time point and over time were determined using a one-way analysis of variance (ANOVA) followed by the multiple comparison test of Tukey (p ≤ 0.05) with Sigma Plot 8.0. 3. Results Measured pCO2 and metal concentrations were similar to nominal values, except in the 100 µg/L treatment, where measured values were higher (Table 1). All concentrations were fairly stable throughout the experiments (Table 1). The pCO2 levels were relatively similar in both experiments. The pH values in the 1000 ppm CO2 exposed groups were ~0.2 pH units below those with no added CO2 (Table 1). 10
Ni and Zn accumulated in E. pallida tissues in a concentration-dependent manner; however, temporal differences were observed dependent on CO2 level (Figs. 1,2). Anemones exposed to 15 µg/L Ni at 1000 ppm CO2 for 48 h had significantly elevated tissue Ni, unlike in the ambient CO2 treatment; however, the tissue Ni values returned to control levels by 7 d (Fig. 1). Anemones exposed to 52 µg/L Ni at 1000 ppm CO2 had significantly elevated tissue Ni by 2 d, which persisted until 4 d and then returned to control levels by 7 d; whereas those anemones exposed to 51 µg/L Ni at ambient CO2 only had significantly elevated tissue Ni at 4 d (Fig. 1). The anemones exposed to 131 and 138 µg/L Ni with ambient and 1000 ppm CO2, respectively, had significantly elevated tissue Ni throughout the 7 d. Zn did not significantly accumulate in the tissues of anemones exposed to 13 µg/L Zn at either CO2 level (Fig. 2). Tissue Zn was significantly elevated in anemones exposed for 2 d to 74 µg/L Zn at 1000 ppm CO2 only. Anemones exposed to 141 and 144 µg/L Zn with ambient and 1000 ppm CO2, respectively, had significantly elevated tissue Zn throughout the experiments (Fig. 2). No significant differences in CAT activity were observed due to Ni or Zn exposure; however, there was a trend of increasing CAT activity in the anemones exposed to 51 µg/L Ni at ambient CO2 over the first 4 d (Fig. 3). No differences in GR activity were observed after exposure to 13 µg/L Ni or Zn at either CO2 concentration (Fig. 4). The pattern of GR activity in anemones exposed to 71 and 74 µg/L Zn differed between the two CO2 exposure groups, whereas such difference was not observed in anemones exposed to 51 and 52 µg/L Ni (Fig. 4). Anemones exposed to 144 µg/L Zn or 138 µg/L Ni at elevated CO2 had a significantly reduced GR activity by 7 d as compared to controls; however, no differences were observed in the anemones exposed to similar metal concentrations at ambient CO2 (Figs. 4, 5). The CO2 concentration significantly influenced the GR activity in the anemones exposed to 141 and 144
11
µg/L Zn, but not in the anemones exposed to 131 and 138 µg/L Ni (Fig. 5). At 2 d, anemones exposed to 144 µg/L Zn and 1000 ppm CO2 had significantly lower GR activity and at 7 d significantly higher GR activity than anemones exposed to 141 µg/L Zn at ambient CO2 (Fig. 5). 4. Discussion Zn concentrations in E. pallida were similar to those reported from other studies. The anemones, Anemonia viridis and Actinia equina contained tissue Zn concentrations between 100 and 200 µg/g dw in waters containing up to 200 µg/L Zn in the environment (Harland et al., 1990). Additionally, tissue Zn in A. viridis exposed to 2-1000 µg/L Zn for 5 days in the laboratory accumulated up to 400 µg/g dw (Harland et al., 1990). Likewise, Mitchelmore et al. (2003) reported tissue Zn concentrations in A. viridis ranging between 200 and 400 µg/g dw after exposure to 100 µg/L Zn for 42 d. E. pallida exposed for 7 d to 10-100 µg/L of a metal mixture (Zn, Ni, Cu, Cd) contained tissue Zn concentrations ranging from 100 to 500 µg/g dw (Brock and Bielmyer, 2013). It‘s possible that increased Zn uptake occurs in the presence of other metals because E. pallida exposed to 141 µg/L of solely Zn in our study only contained up to 300 µg/g dw. Zn is a vital constituent for more than 200 enzymes and is generally well regulated in most organisms (Eisler, 1993; Viarengo and Nott, 1993) so it is not surprising that the tissue Zn concentrations are fairly similar between different anemone species. Like Zn, tissue Ni concentrations in E. pallida in this study were similar to those reported in A. viridis after exposure to the same range of concentrations for 42 d (Mitchelmore et al., 2003). Alternatively, when exposed to a Ni, Zn, Cd, and Cu mixture, E. pallida accumulated approximately 3.5 times more Ni than was observed in our study after exposure of E. pallida to solely Ni (Brock and Bielmyer, 2013). Ni is also an essential element to most organisms (Eisler 1998); however the baseline concentrations of Ni are much lower than Zn, indicating that Zn
12
may be required at higher levels for normal functioning. Additionally, the anemones likely have better homeostatic regulation of Zn. At both CO2 levels, Ni and Zn tissue burdens decreased slightly from 4 d to 7 d. This was likely due to loss of zooxanthellae, which was not quantified in this study, but has been demonstrated in anemones and corals in other studies with increased exposure to metals (Bielmyer et al., 2010; Main et al., 2010; Brock and Bielmyer, 2013). Anemones exposed to Zn or Ni and 1000 ppm CO2 in the present study accumulated the metal significantly faster than those exposed to the same metal concentrations at ambient CO2, potentially due to increased metal bioavailability. At the higher CO2 level and lower pH, metal solubility increases and metal speciation changes (Millero et al., 2009; Lopez et al., 2010; Millero and DiTrolio, 2010). It should be noted that all the pH values (ambient and high CO2 treatments) in the present study were lower than expected. This could have been due to solubility changes in the reconstituted synthetic seawater, although this was not visibly noticed. In any case, the higher CO2 treatments did have significantly lower pH values than the ambient CO2 treatments. In general, as pH decreases, there is an increased formation of metal species that are more bioavailable; and therefore, an increase in toxicity to aquatic organisms. The results of the present work coincide with similar studies using copper in our laboratory. Siddiqui and Bielmyer-Fraser (2015) demonstrated increased Cu uptake at elevated CO2 concentrations in E. pallida. Likewise, anemones exposed to Cu at lower salinity and consequently lower pH exhibited increased Cu accumulation as compared to those exposed to higher salinity and pH (Patel and Bielmyer-Fraser, 2015). However, unlike Cu, the Zn and Ni tissue concentrations in this study returned to those in the ambient CO2 treatments after 96 h. Horwitz et al. (2014) reported substantial changes in tissue metal concentrations in the anemone, Anemonia viridis, located near a CO2 vent with elevated metal concentrations, as compared to a
13
control site. Further, they reported increased concentrations of iron, lead, copper and cobalt; but decreased concentrations of Zn, cadmium, and arsenic in anemones close to the vent; with the pedal disk generally containing higher concentrations than the tentacles (Horwitz et al., 2014). Zn concentrations in the pedal disc ranged from 40-90 µg/g dw. The authors suggested that A. viridis regulates Zn well through compartmentalization and excretion. These field results are consistent to what we observed in our study where even though tissue Zn concentrations did increase faster when CO2 levels were elevated, tissue Zn concentrations quickly returned to levels observed in anemones exposed to Zn at ambient CO2. Physiological responses in the anemones were more varied. Exposure to 51 and 52 µg/L Ni did appear to increase CAT activity in the anemones; however, no other significant differences in CAT activity were observed due to either Ni or Zn exposure (Fig. 2). Alternatively, Cu (Main et al., 2010; Patel and Bielmyer-Fraser, 2015; Siddiqui et al., 2015; Siddiqui and Bielmyer-Fraser, 2015) and combined exposure of Cu, Cd, Ni, and Zn (Brock and Bielmyer, 2013) have been shown to increase CAT activity. Significant differences in CAT activity due to elevated CO2 were not observed in this study, consistent with a previous study (Siddiqui and Bielmyer-Fraser, 2015). Exposure to 131 µg/L Ni or 141 µg/L Zn caused a significant reduction in GR activity by 7 d; however, at elevated CO2, the degree of GR inhibition lessened (Fig. 4). Alternative to Ni and Zn, exposure to Cu alone or a mixture of Cu, Cd, Zn and Ni have been shown to cause an increase in GR activity in past studies (Brock and Bielmyer, 2013; Siddiqui and Bielmyer-Fraser, 2015; Siddiqui et al., 2015). The anti-oxidant enzymes measured in this study are known to be vital defenses against ROS (Livingstone et al., 1992; Regoli et al., 2002). Elevated metal concentrations have been demonstrated to cause oxidative damage to proteins, lipids, and DNA, when oxidative defense mechanisms are
14
overwhelmed (Krumschnabel et al., 2005; Lushchak, 2011). In the present study, GR was affected and CAT was unaffected by increasing Zn or Ni concentrations and elevated CO2 concentration, suggesting that GPx or another enzyme is likely used for the conversion of peroxide into water, rather than CAT. Additionally, in past studies with Cu (Patel and BielmyerFraser, 2015; Siddiqui and Bielmyer-Fraser, 2015) and increased CO2, both parameters individually and in combination (Siddiqui and Bielmyer-Fraser, 2015), caused an increase in GR activity. On the contrary, exposure to Zn or Ni caused a decrease in GR activity and exposure to elevated CO2 lessened that inhibition. Therefore, Ni and Zn may cause toxicity and elicit antioxidant responses via other pathways than does Cu. 5. Conclusions Ni and Zn exposure to E. pallida resulted in tissue metal accumulation and the rate of accumulation was increased with exposure of 1000 ppm CO2. Tissue metal concentration remained elevated in the highest exposure concentrations of 144 µg/L Zn or 138 µg/L Ni by 7 d. Both Ni and Zn were well regulated in the anemone and the effects of CO2 exposure on tissue metal concentration were no longer observed after 96 h. Varying physiological responses were observed due to exposure of Zn or Ni and elevated CO2. No significant differences in CAT activity were observed due to elevated Ni, Zn or CO2. GR activity was significantly reduced by 131 µg/L Ni or 141 µg/L Zn, and the inhibition was decreased by addition of 1000 ppm CO2. This response differs from the increased GR activity demonstrated in anemones exposed to Cu. The metal concentrations used in this study are within the range of those reported in polluted aquatic environments. These results provide new data concerning metal accumulation and physiological responses in sea anemones exposed to acidification and Zn or Ni. Furthermore,
15
these findings are important for application to environmental pollution scenarios with multiple contaminants. Acknowledgements Financial support was provided by individual contributions to the crowdfunding site CREU (creu.tilt.com). Contributors included the following: Southern Oak Insurance Company, Doris and William Bielmyer, T.J. Fraser, Brad Eavenson, Edwin Lunsford, William Lynch, Bill and Michelle Bielmyer, Wayne Richmond, Joseph Pearce, Jenny and Tom Fraser, Holly Schopp, Ben Harper, Bill Tomson, Bob DeSmedt, Chad Lotocki, Herb LeMoyne, Donald Reoderique, Desmond McElroy, and Larry Holder. We would also like to thank the VSU Biology department and Graduate school for funding; Phil Gillette from the University of Miami for the anemones; Pavan Patel for help in the laboratory; and Dr. T.J. Grove for the use of a PerkinElmer Lambda 35 UV/Vis spectrophotometer that was purchased by National Science Foundation grant funds (IOS-0817805to TJG). References Albright, R., Langdon, C., 2011. Ocean acidification impacts multiple early life history processes of the Caribbean coral Porites astreoides. Glob. Change Biol. 17, 2478–2487. Anthony, K.R.N., Maynard, J.A., Diaz-Pulido, G., Mumby, P.J., Marshalls, P.A., Cao, L., Hoegh-Guldberg, O., 2011. Ocean acidification and warming will lower coral reef resilience. Glob. Change Biol. 17, 1798-1808. Ateweberhan, M., Feary, D.A., Keshavmurthy, S., Chen, A., Schleyer, M.H., Sheppard, C.R.C., 2013. Climate change impacts on coral reefs: Synergies with local effects, possibilities for acclimation, and management implications. Mar. Pollut. Bull. 74, 526-539. Bielmyer, G.K., Grosell, M., Bhagooli, R., Baker, A.C., Langdon, C., Gillette, P., Capo, T., 2010. Differential effects of copper on three species of scleractinian corals and their algal Symbionts (Symbiodinium spp.). Aquat. Toxicol. 97, 125-133. Bradford, M.M., 1976. Rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Anal. Biochem. 72, 248–254.
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Brock, J.R., Bielmyer, G.K., 2013. Metal accumulation and sublethal effects in the sea anemone Aiptasia pallida, after waterborne exposure to metal mixtures. Comp. Biochem. Physiol. C. 158, 150-158. Brosnan, J.T., Brosnan, M.E., 2009. Glutathione and sulfur-containing amino acids: An overview, in: Masella,R., Mazza,G. (Eds.), Glutathione and sulfur amino acids in human health and Disease. John Wiley & Sons, Hohoken, New Jersey, pp. 3-15. Brown, B.E., Howard, L.S., 1985. Assessing the effects of ‗stress‘ on reef corals. Adv. Mar. Biol. 22, 1–63. Caldeira, K., Wickett, M.E., 2003. Anthropogenic carbon and ocean pH. Nature 425, 365. Eisler, R., 1993. Zinc hazards to fish, wildlife, and invertebrates. Contaminant Hazard Reviews 26, 1–79. Eisler R., 1998. Nickel hazards to fish, wildlife, and invertebrates: A synoptic review. Contaminant Hazard Reviews, Report 34. Biological Science Report USGS/BRD/BSR1998-0001. U.S. Geological Survey, Washington, DC, 1–95. Esbaugh, A.J., Heuer, R.M., Grosell, M., 2012. Impacts of ocean acidification on respiratory gas exchange and acid base balance in a marine teleost, Opsanus beta. J. Comp. Physiol. B. 182, 921-934. Evans, S.M., Birchenough, A.C., Brancato, M.S., 2000. The TBT ban: out of the frying pan into the fire? Mar. Pollut. Bull. 40, 204–211. Fabry, V.J., Seibel, B.A., Feely, R.A., Orr, J.C., 2008. Impacts of ocean acidification on marine fauna and ecosystem processes. ICES. J. Mar. Sci. 65, 414–432. Ferrier-Pages, C., Houlbreque, F., Wyse, E., Richard, C., Allemand, D., Boisson, F., 2005. Bioaccumulation of zinc in the scleractinian coral Stylophora pistilla. Coral Reefs 24, 636-645. Forman, H.J., Liu, R., Shi, M.M., 1990. Glutathione synthesis in oxidative stress. In: Packer, L., Fuchs, J. (Eds.), Biothiols in Health and Diseases. Marcel Dekker, NY, 189–211. Gomez, C.E., Paul, V.J., Riston-Williams, R., Muehllehner, N., Langdon, C., Sanchez, J.A., 2014. Responses of the tropical gorgonian coral Eunicia fusca to ocean acidification conditions. Coral Reefs 34(2), 451-460. Gonzalez, H., Pomares, M., Ramirez, M. & Torres, I., 1999. Heavy metals in organism and sediments from the discharge zone of the submarine sewage outfall of Havanna city, Cuba. Mar. Pollut. Bull. 38, 1048-1051. Guzmán, H.M., Jiménez, C.E., 1992. Contamination of coral reefs by heavy metals along the 17
Caribbean coast of Central America (Costa Rica and Panama). Mar. Pollut. Bull. 24, 554561. Harland, A.D., Bryan, G.W., Brown, B.E., 1990. Zinc and cadmium absorption in the symbiotic anemone Anemonia viridis and the non-symbiotic anemone Actinia equina. J. Mar. Biol. Assoc. U. K. 70, 789–802. Heuer, R., Esbaugh, A., Grosell, M., 2012. Ocean acidification leads to counterproductive intestinal base loss in the gulf toadfish (Opsanus beta). Physiol. Biochem. Zool. 85(5), 450-459. Higuchi, T., Fujimura, H., Hitomi, Y., Arakaki, T., Oomori, T. and Suzuki, Y., 2010. Photochemical formation of hydroxyl radicals in tissue extracts of the coral Galaxea fascicularis. Photochem. Photobiol. 86(6), 1421-1426. Horwitz, R., Borell, E.M., Fine, M., Shaked, Y., 2014. Trace element profiles of the sea anemone Anemonia viridis living nearby a natural CO2 vent. Peer J. 2:e538; DOI 10.7717/peerj.538. Howard, L.S., Brown, B.E., 1984. Heavy metals and reef corals. Oceanogr. Mar. Biol. Ann. Rev. 22, 195–210. Houlbrèque, F., Rodolfo-Metalpa, R., Jeffree, R., Oberhansli, F., Teyssie, J.L., Boisson, F., AlTrabeen, K., Ferrier-Pages, C., 2012. Effects of increased pCO2 on zinc uptake and calcification in the tropical coral Stylophora pistillata. Coral Reefs 31, 101-109. DOI 10.1007/s00338-011-0819-2. Jones, R., 2010. Environmental contamination associated with a marine landfill (seafill) beside a coral reef. Mar. Pollut. Bull. 60, 1993–2006. Krumschnabel, G., Manzl, C., Berger, C., Hofer, B., 2005. Oxidative stress, mitochondrial permeability transition, and cell death in Cu exposed trout hepatocytes. Toxicol. Appl. Pharmacol. 209 (1), 62-73. Leal, M.C., Nunes, C., Engrola, S., Dinis, M.T., Calado, R., 2012. Optimization of monoclonal production of the glass anemone Aiptasia pallida (Agassiz in Verrill, 1864). Aquaculture 354–355, 91–96. Livingstone, D.R., Lips, F., Garcia, M.P., Pipe, R.K., 1992. Antioxidant enzymes in digestive gland of the common mussel Mytilus edulis. Mar. Biol. 112, 265–276. Lopez, I.R., Kalman, J., Vale, C., Blasco, J.. 2010. Influence of sediment acidification on the bioaccumulation of metals in Ruditapes philippinarum. Environ. Sci. Poll. Res. 17, 1519– 1528.
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Lushchak, V.I., 2011. Environmentally induced oxidative stress in aquatic animals. Aquat. Toxicol. 101, 13-30. Main, W.P.L., Ross, C., Bielmyer, G.K., 2010. Copper accumulation and oxidative stress in the sea anemone, Aiptasia pallida, after waterborne copper exposure. Comp. Biochem. Physiol. C. 151, 216–221. Meehl, G.A., Stocker, T.F., Collins, W.D., Friedlingstein, P., Gaye, A.T., Gregory, J.M., Kitoh, A., 2007. Global Climate Projections. Climate Change 2007: The Physical Science Basis. 10, 749-844. Millero, F.J., Woosley, R., Ditrolio, B., Waters, J., 2009. Effect of ocean acidification on the speciation of metals in seawater. Oceanography 22, 72–85. Millero, F.J., DiTrolio, B.R., 2010. Use of thermodynamics in examining the effects of ocean acidification. Elements. 6, 299–303. Mitchelmore, C.L., Verde, E.A., Ringwood, A.H., Weis, V.M., 2003. Differential accumulation of heavy metals in the sea anemone Anthopleura elegantissima as a function of symbiotic state. Aquat. Toxicol. 64, 317–329. Naoum, C., Fatta, D., Haralambous, K.J., Loizidou, M., 2001. Removal of heavy metals from sewage sludge by acid treatment. J. Environ. Sci. Health, Part A. 36(5), 873-881. Patel, P.P., Bielmyer-Fraser, G.K., 2015. The influence of salinity and copper exposure on copper accumulation and physiological impairment in the sea anemone Exaiptasia pallida. Comp. Biochem. Physiol. C. 168, 39-47. Peters, E.C., Gassman, N.J., Firman, J.C., Richmond, R.H., Power, E.A., 1997. Ecotoxicology of tropical marine ecosystems. Environ. Toxicol. Chem. 16, 12-40. Poonkothai, M.V.B.S., Vijayavathi, B.S., 2012. Nickel as an essential element and a toxicant. Int. J. Environ. Sci. 1, 285-288. Regoli, F., Gorbi, S., Frenzilli, G., Nigro, M., Corsi, I., Focardi, S., Winston, G.W., 2002. Oxidative stress in ecotoxicology: from the analysis of individual antioxidants to a more integrated approach. Mar. Environ. Res. 54, 419–423. Reichelt, A.J., Jones, G.B., 1994. Trace metals as tracers of dredging activity in Cleveland Bay—field and laboratory studies. Mar. Freshw. Res. 45(7), 1237-1257. Siddiqui, S., Goddard, R.H., Bielmyer-Fraser, G.K., 2015. Comparative effects of dissolved copper and copper oxide nanoparticle exposure to the sea anemone, Exaiptasia pallida. Aquat. Toxicol. 160, 205-213. Siddiqui, S., Bielmyer-Fraser, G.K., 2015. Responses of the sea anemone, Exaiptasia pallida, 19
to ocean acidification conditions and copper exposure. Aquat. Toxicol. 167, 228-239. Sies, H., 1999. Glutathione and its role in cellular functions. Free Radic. Biol. Med. 27(9), 916921. Sigma, 1994a. Enzyme Assay of Catalase (EC 1. 11. 1. 6). www.sigmaaldrich.com. Sigma, 1994b. Enzamatic Assay of Glutathione Reductase (EC 1.6.4.2). www.sigmaaldrich.com. Stylianous, M.A., Kolia, D., Haralambous, K.J., Inglezakis, V.J., Moustakas, K.G., Loizidous, M.D., 2007. Effect of acid treatment on the removal of heavy metals from sewage sludge. Desalination. 215, 73–81. Sunagawa, S., Choi, J., Forman, H.J., Meidna, M., 2008. Hyperthermic stress-induced in- crease in the expression of glutamate–cysteine ligase and glutathione levels in the symbiotic sea anemone Aiptasia pallida. Comp. Biochem. Physiol. B. 151, 133–138. Towle, E.K., Enochs, I.C., Langdon, C., 2015. Threatened Caribbean coral is able to mitigate the adverse effects of ocean acidification on calcification by increasing feeding rate. Plos. One. 10, 1-17. Viarengo, A., Nott, J.A., 1993. Mechanisms of heavy metal cation homeostasis in marine invertebrates. Comp. Biochem. Physiol. C. 104, 355–372. Yakovleva, I., Bhagooli, R., Takemura, A., Hidaka, M., 2004. Differential susceptibility to oxidative stress of two scleractinian corals: anti-oxidant functioning of mycosporineglycine. Comp. Biochem. Physiol. B. 139, 721-730. Voulvoulis, N., Scrimshaw, M.D., Lester, J.N., 2000. Occurrence of four biocides utilized in antifouling paints, as alternatives to organotin compounds, in waters and sediments of a commercial estuary in the UK. Mar. Pollut. Bull. 40 (11), 938-946.
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Table 1. Measured pCO2, pH, and dissolved metal concentrations (mean ± standard deviation) in the experimental solutions over the 7 day exposure periods. Nominal Ni pCO2 (ppm) treatment (µg/L) Control 10 50 100 Control 10 50 100
Ambient Ambient Ambient Ambient 963 ± 30 1180 ± 650 1180 ± 30 1020 ± 30
Nominal Zn pCO2 (ppm) treatment (µg/L) Control 10 50 100 Control 10 50 100
Ambient Ambient Ambient Ambient 1130 ± 40 1120 ± 330 1240 ± 60 1050 ± 90
pH 8.05 ± 0.01 7.88 ± 0.04 7.88 ± 0.01 7.86 ± 0.02 7.77 ± 0.08 7.79 ± 0.05 7.76 ± 0.10 7.80 ± 0.06 pH 7.92 ± 0.07 7.90 ± 0.04 7.91 ± 0.08 7.88 ± 0.08 7.74 ± 0.09 7.77 ± 0.05 7.77 ± 0.07 7.73 ± 0.10
Measured [Ni] (µg/L) 2 ± 1.3 13 ± 1.6 51 ± 6.9 131 ± 22.2 4 ± 1.9 15 ± 2.0 52 ± 6.4 138 ± 17.0 Measured [Zn] (µg/L) 4 ± 1.8 13 ± 1.3 71 ± 5.6 141 ± 11.1 4 ± 2.1 13 ± 1.1 74 ± 3.8 144 ± 11.6
21
22
Figure 1. Nickel accumulation (mean ± standard error; µg/g dw) in Exaiptasia pallida exposed to nickel at (A) ambient CO2 and (B) 1000 ppm CO2. * indicate significant differences. Refer to Table 1 for measured nickel concentrations.
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Figure 2. Zinc accumulation (mean ± standard error; µg/g dw) in Exaiptasia pallida exposed to zinc at (A) ambient CO2 and (B) 1000 ppm CO2. * indicate significant differences. Refer to Table 1 for measured zinc concentrations.
A. Zn
C. Ni
B. Zn
D. Ni
Figure 3. Catalase (CAT) activity (mean ± standard error; units/mg protein) in Exaiptasia pallida exposed to A. zinc at ambient CO2, B. zinc at 1000 ppm CO2, C. nickel at ambient CO2, D. nickel at 1000 ppm CO2. Refer to Table 1 for measured metal concentrations.
24
A. Zn
C. Ni
B. Zn
D. Ni
Figure 4. Glutathione reductase (GR) activity (mean ± standard error; units/mg protein) in Exaiptasia pallida exposed to A. zinc at ambient CO2, B. zinc at 1000 ppm CO2, C. nickel at ambient CO2, D. nickel at 1000 ppm CO2. * indicate significant differences. Refer to Table 1 for measured metal concentrations.
25
A. Zn
B. Ni
Figure 5. Glutathione reductase (GR) activity (mean ± standard error; units/mg protein) in Exaiptasia pallida exposed to control or 100 µg/L of A. zinc and B. nickel at ambient CO2 and 1000 ppm CO2. Circles represent control groups and squares represent metal treatments. Filled symbols represent ambient CO2 and open symbols represent 1000 ppm CO2. * indicate significant differences between ambient and 1000 ppm CO2 at a specified metal concentration. Refer to Table 1 for measured metal concentrations.
26