Effects of cadmium exposure on expression and activity of P-glycoprotein in eastern oysters, Crassostrea virginica Gmelin

Aquatic Toxicology 88 (2008) 19–28

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Effects of cadmium exposure on expression and activity of P-glycoprotein in eastern oysters, Crassostrea virginica Gmelin Anna V. Ivanina, Inna M. Sokolova ∗ Biology Department, University of North Carolina at Charlotte, 9201 University City Blvd., Charlotte, NC 28223, USA

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Article history: Received 7 December 2007 Received in revised form 21 February 2008 Accepted 26 February 2008 Keywords: Cadmium P-glycoprotein Mitochondria Cell membrane Activity Energy demand Respiration Protein and mRNA expression

a b s t r a c t Heavy metal pollution is a worldwide problem, and cadmium (Cd) is one of the most noxious pollutants in aquatic environments. We studied P-glycoprotein (P-gp) expression and function in control and Cd exposed (50 ␮g L−1 Cd, 30–40 days) oysters Crassostrea virginica as a possible mechanism of cell protection against Cd. Our data show that P-gp is expressed on cell membrane and in mitochondria of oyster gills and hepatopancreas. Inhibitor studies with verapamil, cyclosporine A and JS-2190 suggest that in the gills, mitochondrial P-gp pumps substrates from cytosol into the mitochondria, while cell membrane P-gp pumps substrates from cytosol out of the cell. Cd exposure resulted in a 2–2.5-fold increase in Pgp protein expression in cell membranes and a 3.5–7-fold increase in transport activity measured as the inhibitor-sensitive rhodamine B extrusion rate. In contrast, p-gp mRNA levels were similar in control and Cd-exposed oysters. No difference in P-gp protein expression was observed between mitochondria of control and Cd-exposed oysters but the apparent transport activity was higher in mitochondria from Cd-exposed oysters. Overall, a stronger increase in substrate transport activity in Cd-exposed oysters compared to a relatively weaker change in P-gp protein levels suggests that P-gp activity is post-translationally regulated. Our data show that direct determination of P-gp transport activity may be the best measure of the xenobiotic-resistant phenotype, whereas p-gp mRNA levels are not a good marker due to the likely involvement of multiple post-transcriptional regulatory steps. Cd exposure resulted in a significantly elevated rate of oxygen consumption of isolated oyster gills by 46%. Specific inhibitors of ATPase function of P-gp (cyclosporine A and JS-2190) had no significant effect on tissue oxygen consumption indicating that P-gp contribution to energy budget is negligible and supporting indirect estimates based on the ATP stoichiometry of substrate transport that also suggest low energy demand for P-gp function. © 2008 Elsevier B.V. All rights reserved.

1. Introduction Heavy metal pollution is a worldwide problem, and cadmium is one of the most noxious environmental pollutants responsible for episodes of chronic poisoning in humans and wildlife (Pinot et al., 2000). This harmful heavy metal is common in coastal areas and estuaries and is released into the environment via anthropogenic sources such as mining, smelting, electroplating and through the use of nickel- and cadmium-containing batteries, pigments and plastics, as well as from the natural sources such as volcanic activity, leaching from Cd-rich soils and diatom deposition in marine sediments (GESAMP, 1987; Frew et al., 1997; Lane and Morel, 2000). At high levels Cd causes mortality and morbidity, but even low

∗ Corresponding author at: Biology Department, 381c Woodward Hall, University of North Carolina at Charlotte, 9201 University City Blvd., Charlotte, NC 28223, USA. Tel.: +1 704 687 8532; fax: +1 704 687 3128. E-mail addresses: [email protected], [email protected] (I.M. Sokolova). 0166-445X/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.aquatox.2008.02.014

sublethal Cd concentrations can result in physiological perturbations affecting performance and survival of aquatic organisms. One of the key mechanisms of Cd toxicity involves a disturbance of energy metabolism including a decrease in mitochondrial efficiency, impaired oxygen uptake and distribution and an increase in standard metabolic rate (SMR)—the energy necessary to carry out basal activities required for an organism to survive (Cherkasov et al., 2006; Lannig et al., 2006, Lannig et al., 2008). Increase in basal metabolism or SMR due to metal exposures is thought to reflect the energy cost for detoxification and/or repair of metal-induced damage that can potentially divert energy from key cellular processes such as maintenance, growth and reproduction (Gordon, 2005; Cherkasov et al., 2006; Lannig et al., 2006, 2008). However, the mechanisms of the metal-induced increase in SMR are not well understood and require further investigation. P-glycoproteins (P-gps) belong to the family of ABC (ATP-binding cassette) transporters responsible for multixenobiotic resistance phenotype in aquatic organisms and multidrug resistance phenotype in mammalian tumor cells (Ambudkar et al., 1999; Borst et al.,

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2000; Bard, 2000). In mollusks, a single P-glycoprotein gene has been described that corresponds to the vertebrate ABC-B (including ABC-B1 P-glycoprotein) and sister P-glycoprotein (sP-gP) subfamilies of transporters and is likely homologous to an ancestral P-gP gene that gave rise to this branch of ABC transporters in vertebrates (Bard, 2000; Tutundjian and Minier, 2007). The primary function of P-gP transporters is pumping out a broad variety of xenobiotic substances including drugs and natural product toxins in ATP-dependent fashion. Export of xenobiotics out of the cell by Pglycoproteins and other ABC transporters requires hydrolysis of 2–3 ATP molecules per molecule of transported substrate (Ambudkar et al., 1999; Hennessy and Spiers, 2007). Moreover, it has been shown that ABC transporters also hydrolyze ATP when idling due to uncoupled ATPase activity and transport of endogenous substrates thus potentially making them an important contributor to basal metabolic rate (Ambudkar et al., 1999; Hennessy and Spiers, 2007). Recently it has been also shown that exposure to heavy metals such as cadmium can induce elevated P-gp expression in mammalian cell lines (Endo et al., 2002; Huynh-Delerme et al., 2005) and freshwater bivalves Corbicula fluminea (Achard et al., 2004; Legeay et al., 2005). Given ATP dependence of P-gp both in active and idling states, it is possible that this protein may contribute to elevated SMR in Cdexposed animals especially if both expression and function of this protein is elevated. The energy cost to sustain a perpetual state of xenobiotic resistance by elevated P-gp expression may ultimately reduce overall fitness of organisms in chemically stressful environments (Renfro et al., 1993; Epel, 1998). However, to the best of our knowledge no studies have been conducted so far to directly assess energy requirements for P-gp function and its possible contribution to SMR. Although P-gp was originally described from plasmalemma, recent studies have shown that it may be also be expressed in intracellular membranes of organelles in multidrug resistant tumor cell lines, including mitochondria (Munteanu et al., 2006; Solazzo et al., 2006; but see Patterson and Gottesman, 2007), golgi apparatus (Meschini et al., 1987; Gervasoni et al., 1991; Rutherford and Willingham, 1993; Molinari et al., 1994) and nuclear membrane (Maraldi et al., 1999). P-gp involvement in the substrate transport was experimentally confirmed in mitochondria of these cell lines, and the direction of transport (in or out of the mitochondrial matrix) was shown to depend on the cell type (Munteanu et al., 2006; Solazzo et al., 2006). However, there are very few studies of P-gP expression in mitochondria of normal (non-transformed) cells and nothing is known about P-gp expression in mitochondria of organisms other than mammals. It is also not known whether P-gps contribute to energy demand of mitochondria and whether they are involved in mitochondrial substrate transport and/or potential protection of mitochondria from xenobiotics in aquatic organisms. Eastern oysters, Crassostrea virginica Gmelin (Bivalvia: Ostreidae), are common filter-feeders in estuaries and coastal waters of the western Atlantic and useful models for studies of the Cd effect on P-gp expression and function in marine ectotherms. Oysters are exposed to a variety of natural toxins and environmental contaminants including Cd and express several defense mechanisms, including multixenobiotic resistance proteins, in order to tolerate these exposures (Keppler and Ringwood, 2001). Oysters have an ability to accumulate Cd in soft tissues to concentrations exceeding the environmental levels by orders of magnitude thus making them susceptible to toxic effects of this metal (Roesijadi, 1996). Longterm sublethal Cd exposures result in a significant energy cost in oysters leading to an approximately 40–90% increase in SMR at 20 or 24 ◦ C (Cherkasov et al., 2006; Lannig et al., 2006), but the relative contribution of different protection mechanisms to this energy cost are not well understood. Indeed, no significant increase in the amount of energy spent to counteract mitochondrial proton leak or

for non-mitochondrial respiration was found in Cd-exposed oysters (Cherkasov et al., 2006). Elevated energy costs of protein synthesis associated with the expression of cellular protection proteins such as metallothioneins and heat shock proteins were shown to contribute to high SMR in Cd-exposed oysters (Cherkasov et al., 2006; Ivanina et al., 2008). However, protein synthesis accounts only for 10–20% of the total energy budget of the cell (Hand and Hardewig, 1996; Cherkasov et al., 2006), and, thus, elevated costs of the protein synthesis alone cannot fully explain Cd-induced increase in SMR. In the present work we aimed to: (1) determine the effect of Cd exposure on P-gp expression on cell membrane and in mitochondria of eastern oysters C. virginica; (2) assess functional activity of P-gp in whole tissues and isolated mitochondria of control and Cd-exposed oysters; (3) estimate energy demand for P-gp function in oyster gills in order to assess its potential contribution to Cd-induced elevation of SMR. 2. Materials and methods 2.1. Animal collection and maintenance Oysters (C. virginica) were purchased from Taylor Shellfish Farms (Shelton, WA) in April 2007, shipped overnight to the University of North Carolina at Charlotte and placed in recirculated aerated tanks with artificial seawater (Instant Ocean® , Kent Marine, Acworth, USA) at the temperature and salinity close to their habitat (12 ± 1 ◦ C and 30‰). They were fed ad libitum on alternate days with a commercial algal blend (2 mL per oyster) containing Nannochloropsis, Tetraselmis, and Isochrysis spp. with a cell size of 2–15 ␮m (PhytoPlex; Kent Marine, Acworth, GA, USA) or Nannochloropsis oculata, Phaeodactylum tricornutum and Chlorella with a cell size of 2–20 ␮m (DT’s Live Marine Phytoplankton, Premium Reef Blend Sycamore, IL, USA). Oysters were allowed to recover for 5 days in the lab. After the preliminary acclimation, half of the tanks were randomly selected, and Cd (as CdCl2 ) was added to the nominal concentration of 50 ␮g L−1 . The remaining tanks were used as controls. To avoid pseudoreplication, at least three tanks were set for control or Cd exposure, and oysters were randomly sampled from these tanks for each experiment. Mortality during experimental exposures (30–40 days) was <5% and did not significantly differ between control and Cd-exposed oysters. It is worth nothing that Cd concentrations in our experimental exposures (50 ␮g L−1 ) were at the upper end of Cd concentrations found in polluted estuaries (Crompton, 1997). However, our previous studies have shown that 30–60 days of exposure under these conditions result in physiologically relevant tissue burdens of Cd similar to those found in oysters from polluted estuaries (Sokolova et al., 2005; Cherkasov et al., 2006 and references therein). 2.2. Preparation of isolated mitochondria and cell membranes from oyster tissues Gill or hepatopancreas (digestive gland) tissues were collected from 2 to 3 oysters and homogenized in homogenization medium (HM) containing 300 mM sucrose, 50 mM KCl, 50 mM NaCl, 8 mM EGTA, 30 mM HEPES, 0.5 mM dithiothreitol (DTT), 0.1% ␤-mercaptoethanol (ME), aprotinin (0.5 ␮g mL−1 ), phenylmethanesulphonylfluoride (PMSF) (40 ␮g mL−1 ), pH 7.4 in PotterElvenhjem glass-Teflon homogenizer. Mitochondrial fraction was isolated on Percoll gradient using a method modified from Crockett and Sidell (1993). Briefly, tissue homogenates were centrifuged for 10 min at 1500 × g to remove cell debris (this and all subsequent centrifugations were conducted at 4 ◦ C). Supernatants were collected and centrifuged for 8 min at 10,000 × g. Pellet was resus-

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Fig. 1. P-glycoprotein (P-gp) expression in subcellular fractions of control and Cd-exposed C. virginica. (A) Representative Western blot for the expression of mitochondrial markers, cytochrome c oxidase subunit I (mitochondrial complex IV, CCOI), F0 , F1 -ATPase (mitochondrial complex V) and a plasma membrane marker, endothelial growth factor receptor (EGFR) in mitochondrial and cell membrane fractions isolated from gills and hepatopancreas (HP) of oysters. (B) Representative Western blots for P-gp expression in cell membrane and mitochondrial fractions in control and Cd-exposed oysters. (C) and (D) P-gp protein expression in cell membrane (C) and mitochondrial (D) fractions of oyster gills. P-gp levels were measured using immunoblotting and expressed as % of an internal standard run on all gels. Vertical bars represent standard errors. Asterisks denote values significantly different from the respective controls (P < 0.05), N = 5.

pended in 15 mL wash buffer (30 mM HEPES, 500 mM sucrose, 0.5 mM DTT, 0.1% ␤-ME, pH 7.5) and centrifuged again for 8 min at 10,000 × g to obtain crude mitochondrial fraction. Supernatants from both centrifugations were collected and used to isolate cell membrane fraction (see below). Mitochondrial fractions were purified on Percoll (Crockett and Sidell, 1993; Rajapakse et al., 2001); briefly, they were resuspended in 1 mL of wash buffer, mixed with 20 mL of wash buffer containing 30% Percoll and centrifuged for 30 min at 25,000 rpm. Pellets were resuspended in 0.5 mL of wash buffer to obtain purified mitochondrial fraction. Cell membrane fractions were isolated on two-step sucrose gradient using methods modified from Schimmel et al. (1973) and Crockett (1999). Buffer containing 2.4 M sucrose and 30 mM HEPES, pH 7.5 was added to the combined supernatants from the abovedescribed centrifugations to obtain the final sucrose concentration of 1.45 M. A two-layer step gradient was prepared in ultracentrifuge tubes that consisted of 5/6 volumes of supernatant adjusted to 1.45 M sucrose and overlaid with approximately 1/6 volume of 0.25 M sucrose, enough to fill the tube. The tubes were centrifuged at 70,000 × g for 60 min at 4 ◦ C. Interphase fraction of the two-step gradient was collected and centrifuged again for 100,000 × g for 60 min at 4 ◦ C to collect purified cell membrane fraction. The pellet was resuspended in 2 mL of wash buffer. Five to six separate mitochondrial and cell membrane preparations were obtained from control and Cd-exposed oysters, each containing tissues pooled from 2 to 3 individuals. To confirm purity of the collected fractions, marker proteins were detected by immunoblotting as described below. Epidermal

growth factor receptor (EGFR) was used as a cell membrane-specific marker and mitochondrial complexes IV (cytochrome c oxidase subunit I, CCOI) and V (F0 , F1-ATPase) were used as mitochondrial markers. Cell membrane fractions showed a strong EGRF signal and no CCOI or F0 , F1-ATPase signals while mitochondrial fractions had only CCOI and F0 , F1-ATPase signals and no EGRF signal (Fig. 1A) indicating sufficient purification of the two fractions. 2.3. Immunoblotting of cell membrane and mitochondrial marker proteins and P-gp Detection and analysis of expression of P-gp, cell membrane and mitochondrial marker proteins were performed by standard immunoblotting in membrane and mitochondrial fractions from gills and hepatopancreas of control and Cd-exposed oysters. Protein content was measured in cell membrane and mitochondrial suspensions using Bio-Rad Protein Assay kit according to the manufacturer’s instructions (Bio-Rad, Hercules, CA, USA). Bovine serum albumin (BSA) was used as a standard. Samples (30 ␮g of protein per lane) were loaded onto 8% polyacrylamide gels and run at 100 mA for 2 h at room temperature. The resolved proteins were transferred onto a nitrocellulose membrane in 96 mM glycine, 12 mM Tris and 20% methanol (v/v) using a Trans-Blot semi-dry cell (Thermo Fisher Scientific Inc., Portsmouth, NH, USA). To verify equal protein loading, membranes were stained with Amido Black Stain Solution (1 g L−1 Amido Black in 10% methanol, 10% glacial acetic acid) for 30 s. The membranes were then destained by washing in several changes of water and blocked overnight in 5% non-fat

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milk in Tris-buffered saline, pH 7.6 (TBST). Blots were probed with primary monoclonal antibodies for P-glycoprotein (P-gp) (C219, Signet, Dedham, MA), a cell membrane marker EGFR (EGFR(29.1), NB120-10414, Novus Biological Inc., Littleton, CO) or mitochondrial markers complex IV subunit I (␣-COX-I, MitoSciences, Eugene, OR) and complex V subunit ␣ (CV-␣, MitoSciences, Eugene, OR). After washing off the primary antibody, membranes were probed with the polyclonal secondary antibodies conjugated with horseradish peroxidase (Jackson Immunoresearch, West Grove, PA, USA) and proteins detected by enhanced chemiluminescence according to the manufacturer’s instructions (Pierce, Rockford, IL, USA). All antibodies used in this study produced a single band of expected size on immunoblots indicating specific reaction (see “Section 3”). Densitometric analysis of the signal was performed by GelDoc 2000TM System with Quantity One 1-D Analysis Software (Bio-Rad Laboratories Inc., Hercules, CA, USA). To allow for cross-gel comparisons of P-gp expression, the same sample of a cell membrane fraction from control oysters was run on all gels as an internal standard, and P-gp expression in other samples was expressed as per cent of this internal standard. 2.4. RNA extraction and RT-PCR amplification Expression of P-glycoprotein mRNA (p-gp gene expression) was measured using quantitative real-time PCR (QRT-PCR) and semiquantitative RT-PCR. RNA was extracted by homogenization in Tri Reagent (Sigma–Aldrich, Saint Louis, Missouri, USA) according to the manufacturer’s protocol. Tissue to TRI reagent ratio was kept below 1:10 (weight:volume). RNA concentration and quality were verified by UV spectroscopy. This method yielded high purity total RNA with 280/260 absorbance ratio > 1.9. For real-time PCR analysis, reverse transcription was performed using 200 U ␮L−1 SuperScript III Reverse Transcriptase (Invitrogen, Carlsbad, CA, USA), 50 ␮M of oligo(dT)18 primers and 5 ␮g total RNA to obtain single stranded cDNA. Specific primers were designed to amplify cDNA for Pglycoprotein and ␤-actin using C. virginica sequences published in GenBank (NCBI accession numbers AAP92331.1 and X75894.1). Expression of ␤-actin was used as a housekeeping gene to normalize expression of P-glycoprotein. Our earlier studies showed that ␤-actin is an appropriate housekeeping gene for standardization of mRNA expression in control and Cd-exposed oysters (Sanni et al., 2008). The following primers were used: for p-gp gene: • Pgp-C.v. F 5 -AAA CAA ATC GGC ATC GTT TC-3 • Pgp-C.v.R 5 -ATG ATC TCT GCC ATC GGA AC-3 for ␤-actin gene: • Act-Cv-F437 5 -CAC AGC CGC TTC CTC ATC CTC C-3 • Act-Cv-R571 5 -CCG GCG GAT TCC ATA CCA AGG-3 . Quantitative RT-PCR was performed using a LightCycler® 2.0 Real Time PCR System (Roche Applied Science, Indianapolis, IN) and QuantiTect SYBR Green PCR kit (Qiagen, Valencia, CA) according to the manufacturers’ instructions. The reaction mixture consisted of 5 ␮l of 2× QuantiTect SYBR Green master mix, 0.3 ␮M of each forward and reverse primers, 1 ␮l of 10× diluted cDNA template and water to adjust to 10 ␮l. Ten microliters of reaction mixture were loaded into LightCycler 20 ␮l capillaries (Roche Applied Science, Indianapolis, IN) and subjected to the following cycling: 15 min at 95 ◦ C to denature DNA and activate Taq polymerase; 40–55 cycles of 15 s at 94 ◦ C, 20 s at 55 ◦ C and 15 s at 72 ◦ C. SYBR Green fluorescence (acquisition wavelength 530 nm) was measured at the end

of each cycle for 2 s at the read temperature of 78 ◦ C. The read temperatures of the amplified fragments were determined in the pilot experiments and set at the value to melt all primer dimers but not the amplified gene product. At the end of each run, melting temperature profiles were run between 50 and 99 ◦ C with continuous fluorescence acquisition to confirm the expected melting temperatures for the amplified fragments. In each run, serial dilutions of a cDNA standard were amplified to determine amplification efficiency (Pfaffl, 2001), and an internal standard was included to test for amplification variability between the runs. Dilutions of the experimental cDNA samples were selected so that their crosspoints for fluorescence fell within the range of the crosspoints for cDNA standards. Amplification efficiencies (E) were 1.71 ± 0.02 (N = 10) and 1.72 ± 0.03 (N = 10) for p-gp and ␤-actin, respectively. Expression of p-gp gene was calculated relative to the expression of ␤-actin and normalized against the internal standard as proposed by Pfaffl (2001): R=

EtCPt CPref Eref

where Et and Eref are amplification efficiencies for target (p-gp) and the reference (␤-actin) genes, respectively, and CPt and CPref are differences between crosspoints for fluorescence of the sample and the internal standard for p-gp gene and ␤-actin, respectively. For semi-quantitative PCR, 0.5 ␮L of undiluted cDNA template was added to a PCR reaction mixture containing 10 pmol of each forward and reverse primer for p-gp or ␤-actin genes, 12.5 ␮L of Go Taq mix (Promega, Madison, WI, USA) and water to adjust to 24.5 ␮L. Amplification was performed in a Mastercycler Gradient thermal cycler (Brinkmann, Westbury, NY, USA) using 35 or 30 cycles (for p-gp and ␤-actin, respectively) of 95 ◦ C for 2 min, 94 ◦ C for 30 s, 55 ◦ C for 30 s and 72 ◦ C for 40s. Cycle numbers were selected for each gene in pilot experiments in order to allow amplification in all samples while avoiding signal saturation. The amplified products were resolved on 1% agarose gel and visualized by ethidium bromide staining. Signal densitometry was performed by GelDoc 2000TM System as described for immunoblotting. Levels of the target gene expression were calculated as ratios of p-gp to ␤-actin signal in the respective samples. 2.5. MXR activity in isolated gill tissue MXR activity was determined in isolated gill tissues of control and Cd-exposed oysters using efflux assay modified from Smital et al. (2000), which measures the rate of efflux of a fluorescent substrate, rhodamine B (RB) from the tissues in the presence or absence of model MXR inhibitors. Briefly, gills fragments (80–120 mg) from control or Cd-exposed oysters were placed into glass containers with 8 mL of filtered artificial seawater (ASW) supplemented with 7.5 ␮M RB for 30 min in the dark. After the exposure, gills were washed three times in 8 mL of clean ASW to remove surfaceassociated RB and transferred to another glass container with 8 mL of filtered seawater for a 5 min “pre-efflux” period. This was necessary to allow the tissue to release the captured volume of loading medium containing high concentration of RB to avoid high background noise at later measurements of the efflux rates. After the “pre-efflux” period gill fragments were individually transferred to new glass containers with 16 mL of filtered seawater without (control) or with addition of known P-gP inhibitors verapamil, cyclosporine A or JS-2190. Verapamil is a high-affinity substrate and a model competitive inhibitor of multixenobiotic resistance (MXR) transporters including P-gp; it is pumped by P-gP and therefore, stimulates its ATPase activity (Litman et al., 2001). In contrast, cyclosporine A and JS-2190 inhibit MXR trans-

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port by blocking ATPase activity of MXR transporters; cyclosporine A has a broader specificity interacting with P-gP as well as other membrane ATPases whereas JS-2190 specifically inhibits P-gp and MRP1 (Watanabe et al., 1997; Litman et al., 2001; Kimura et al., 2005). Pilot dose–response experiments were performed for each P-gp inhibitor in the range from 0 to 1000 ␮M, and the lowest effective concentrations causing maximum inhibition of the fluorescent dye transport were selected for subsequent experiments (13 ␮M cyclosporine A, 66.7 ␮M verapamil or 16.7 ␮M JS-2190). These concentrations are within the range of concentrations used in other MXR studies (Smital and Kurelec, 1998; Kurelec et al., 2000). Glass vials containing gill fragments in seawater with or without inhibitors were placed on a shaker, and RB efflux was measured over a 60 min exposure period at room temperature (20–22 ◦ C). Two replicate aliquots (200 ␮L each) of the media from each vial were collected every 10 min during a 60-min exposure period and transferred into a 96-well dark microplate. After the last collection fluorescence of the effluxed dye was immediately measured using a fluorescence multi-well plate reader (Cyto Fluor Series 4000, Framingham, MA, USA) at excitation wavelength of 535 nm and emission wavelength of 590 nm. Pilot experiments showed that RB efflux was linear over the 60-min exposure period (data not shown). To calculate RB extrusion rates, a calibration curve was made using serial dilutions of known concentrations of RB. P-gp activity was calculated as a difference between the net rates of RB efflux in the absence and presence of P-gp inhibitors and expressed as pmol RB min−1 g−1 wet mass. 2.6. MXR activity in isolated mitochondria Mitochondria were isolated from gills of control and Cd-exposed (50 ␮g L−1 ) Cd oysters after 40 days of exposure at 12 ◦ C. Isolation was performed as described in Sokolova (2004). To check for the quality of mitochondrial isolations, mitochondrial oxygen uptake was measured in 1 mL water-jacketed chambers using Clarketype oxygen electrodes (Qubit Systems, Kingston, ON, Canada) at 20 ◦ C. Mitochondrial assay conditions were as described in Sokolova (2004), and calibration of oxygen electrodes, data acquisition and MO2 calculations were performed as described in Sokolova et al. (2005). Succinate was used as a substrate at saturating amounts (10–15 mM) in the presence of 5 ␮M of rotenone. Maximal respiration rates (state 3) were achieved by addition of 200–300 nmol ADP, and state 4 respiration was determined in ADP-conditioned mitochondria as described by Chance and Williams (1955). Respiration rates were corrected for the electrode drift and non-mitochondrial respiration as described in Sokolova (2004), and respiratory control ratio (RCR) was determined as a ratio of state 3 over state 4 respiration according to Estabrook (1967). Average RCR of mitochondrial isolates used in this study was 2.5 which is typical for molluscan mitochondria (Sokolova, 2004 and references therein) and indicates good coupling. Six to seven mitochondrial isolates, each obtained from gill tissues of 2–3 oysters, were analyzed for the control and Cd-exposed groups. Mitochondrial MXR activity was measured in state 4 (resting mitochondria) by the difference in uptake of a fluorescent dye rhodamine 123, into isolated mitochondria in the presence and absence of MXR inhibitors verapamil (100 ␮M) or JS-2190 (20 ␮M) at 20 ◦ C. Our earlier studies showed that this temperature is close to the optimum for oyster mitochondria (Cherkasov et al., 2006). Briefly, mitochondrial suspensions were diluted to 3–5 mg protein mL−1 with assay buffer containing 150 mM sucrose, 250 mM KCl, 10 mM glucose, 10 mM KH2 PO4 and 30 mM HEPES, pH 7.2. Rhodamine 123 was added to the final concentration of 2.5 ␮M, mixed by inver-

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sion and incubated for 30 min on a shaker at room temperature (20–22 ◦ C) in the dark. Mitochondria were collected by centrifugation for 5 min at 10,000 × g and 4 ◦ C, the supernatant was discarded and the mitochondrial pellet was washed in 1 mL of the assay buffer to remove surface-associated rhodamine 123. Mitochondria were again collected by centrifugation for 5 min at 10,000 × g and 4 ◦ C, resuspended in 1 mL of assay buffer and lysed by addition of 1% Triton-X (final concentration) to release the dye. Suspension was centrifuged for 5 min at 10,000 × g and 4 ◦ C to remove membrane debris, and rhodamine 123 fluorescence was measured in the supernatant at excitation wavelength of 485 nm and emission wavelength of 530 nm using a fluorescence spectrophotometer (Hitachi Ltd., Tokyo, Japan). Protein concentrations in mitochondrial suspensions were measured using a modified Biuret method with 1% Triton-X (Bergmeyer, 1985). Data were expressed as fluorescence of rhodamine 123 (RFU) mg−1 mitochondrial protein. 2.7. Tissue respiration rates Small gill fragments (0.1–0.14 g) were cut out from control or Cd-exposed oysters after 30 days of exposure, and incubated for 30 min at 20–22 ◦ C in tissue support media (TSM, 24.72 g L−1 NaCl, 0.68 g L−1 KCl, 1.36 g L−1 CaCl2 × 2H2 O, 0.18 g L−1 NaHCO3 , 4.66 g L−1 MgCl2 × 6H2 O, 6.29 g L−1 MgSO4 × 7H2 O, 30 mM glucose and 30 mM HEPES at pH 7.5) in the presence or absence of cyclosporine A (13.3 ␮M) and JS-2190 (16.67 ␮M) to block ATPase activity of MXR transporters. Antibiotics (10,000 U mL−1 of penicillin and 10,000 U mL−1 streptomycin) were added to the incubation media to prevent bacterial respiration. Pilot studies showed that there was no change in tissue oxygen consumption rate for at least 2 h of incubation at 20–22 ◦ C in the absence of inhibitors (longer incubation times were not tested). Tissue fragments were then mounted on plastic frames to ensure maximum contact between the tissue surface and respiration media and placed in 3 mL of antibiotics- and glucose-supplemented TSM in water-jacketed chambers with Clarke-type oxygen electrodes (Qubit Systems, Kingston ON, Canada) at 20 ◦ C. Inhibitors were added as appropriate to maintain the same concentration as in the incubation media, and the rate of oxygen consumption was recorded for 12–15 min. All respiration rates were corrected for the electrode drift. Tissue oxygen consumption rate (MO2 ) was expressed in ␮mol O2 h−1 g−1 wet tissue mass, and oxygen demand for MXR function was determined as the difference between tissue MO2 without inhibitors and in the presence of inhibitors. Rhodamine 123 was purchased from Molecular Probes (Invitrogen, Carlsbad, CA, USA), JS-2190 was from Kamiya Biomedical Company (Seattle, WA, USA). All other chemicals were purchased from Sigma–Aldrich (St. Louis, MO, USA) or Fisher Scientific (Suwanee, GA, USA) and were of analytical grade or higher. 2.8. Statistics Effects of cadmium exposure, tissue type and presence/absence of inhibitors were analyzed using split-plot repeated measures ANOVA after testing the assumptions of normality of data distribution and homogeneity of variances. Individual tissue or mitochondrial isolate was used as a repeated measures variable. Least squared difference (LSD) tests were used for planned comparisons of sample means. For data expressed as % of control, t-tests were used to test the hypothesis that sample mean significantly deviates from 100%. Statistical analyses were performed using SAS 9.1.3 software (SAS Institute, Cary, NC, USA), and differences were considered significant if the probability for Type II error was less than 0.05.

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3. Results 3.1. Expression of P-gp protein and mRNA Immunoblotting showed that a ∼170 kDa protein specifically reacting with a P-gp antibody was expressed in both cell membrane and mitochondrial fractions of gills and hepatopancreas from control and Cd-exposed oysters (Fig. 1B), and expression levels (on per unit protein basis) were similar in these two organelles. In control oysters, P-gp expression in cell membrane fraction was twofold higher in gills compared to hepatopancreas, whereas in mitochondria similar levels of P-gp protein were found in the two studied tissues (Fig. 1). Effect of tissue type on P-gp expression was significant for cell membrane fraction (F1, 16 = 6.45, P = 0.022) but not for mitochondrial fraction (F1, 16 = 0.52, P = 0.481). Cadmium exposure resulted in 2–2.5-fold increase in P-gp protein expression in cell membranes of gills and hepatopancreas but had no effect on mitochondrial P-gp expression (Fig. 1C and D) (F1, 16 = 13.01, P = 0.002 and F1, 16 = 0.56, P = 0.464 for cell membrane and mitochondria, respectively). Factor interactions between tissue type and Cd exposure were not significant in either of the two studied fractions (F1, 16 = 0.03, P = 0.8582 and F1, 16 = 1.23, P = 0.2842 for cell membrane and mitochondrial fractions, respectively). Analysis of mRNA expression for p-gp gene (Fig. 2) showed no significant differences in p-gp mRNA levels between control and Cd-exposed oysters in gills and hepatopancreas (F1, 36 = 1.10, P = 0.301). Differences in mRNA expression between gill and hepatopancreas were also non-significant (F1, 36 = 0.66, P = 0.420). No significant interactions were found between the effects of tissue type and Cd exposure on p-gp mRNA levels (F1, 36 = 0.63, P = 0.432). Semi-quantitative PCR showed essentially the same pattern indicating a good correspondence between the two methods to assess expression of this gene (data not shown). 3.2. MXR activity in whole tissue and mitochondrial fraction of oyster gills In Cd-exposed oysters, both the total efflux rate of a fluorescent dye (rhodamine B, RB) (Fig. 3A) and the contribution of MXR activity to this efflux were considerably higher than in the controls. In control oysters, 27% (according to the estimates using verapamil and JS-2190 as inhibitors) to 51% (using inhibition with cyclosporine A) of RB efflux was due to MXR activity. In contrast, in Cd-exposed oysters 71–83% of RB efflux was due to MXR pumping. Non-MXR mediated efflux rate of RB was similar in control and Cd-exposed oysters (28–42 pmol RB g−1 min−1 and 22–39 pmol RB g−1 min−1 , respectively). In contrast, MXR-dependent efflux was 3.5–7-fold

Fig. 3. MXR activity in whole gills and isolated mitochondria from control and Cdexposed C. virginica. (A) (insert) Total net efflux of rhodamine B (RB) from the gills of control and Cd-exposed oysters in the absence of P-gp inhibitors. Differences in total net RB efflux between control and Cd-exposed oyster gills were marginally significant (P = 0.06). Vertical bars represent standard errors, N = 5. (B) Putative P-gpmediated RB efflux determined as a difference between the total net efflux and efflux in the presence of P-gp inhibitors (verapamil, cyclosporine A or JS-2190) in gills from control and Cd-exposed oysters. Vertical bars represent standard errors. Asterisks denote values significantly different from the respective controls (P < 0.05), N = 5. (C) Rhodamine 123 uptake in mitochondria from control and Cd-exposed oysters with or without P-gp inhibitors (JS-2190 or verapamil). Asterisks denote values significantly different from the respective values in the absence of inhibitors (P < 0.05). Vertical bars represent standard errors, N = 7–8.

higher in Cd-exposed oysters compared to their control counterparts (Fig. 3B). ANOVA showed significant effect of Cd exposure on MXR activity (when estimated with JS-2190: F1, 8 = 7.30, P = 0.027; for cyclosporine A: F1, 8 = 5.44, P = 0.048; for verapamil: F1, 8 = 5.61, P = 0.045). The net uptake rates of a fluorescent dye rhodamine 123 tended to be higher in gill mitochondria from Cd-exposed oysters compared to the controls (Fig. 3C); however, this effect was not statistically significant (ANOVA: F1, 13 = 3.46, P = 0.085). ANOVA showed no significant overall effect of the inhibitors (F2, 25 = 1.76, P = 0.193) or factor interactions (Cd exposure × inhibitors: F2, 25 = 1.92, P = 0.167) on rhodamine 123 uptake in mitochondria. However, post hoc comparisons showed that incubation with the inhibitors resulted in a decrease in rhodamine 123 accumulation in mitochondria from Cd-exposed oysters (P = 0.024 and 0.046 for JS-2190 and verapamil, respectively) but not in their control counterparts (P > 0.80). MXR inhibitors had no effect on oxygen consumption rates of oyster mitochondria (data not shown). 3.3. Tissue respiration rates

Fig. 2. P-glycoprotein mRNA expression in gills and HP in control and Cd-exposed oysters measured by quantitative real-time RT-PCR. Levels of p-gp mRNA were normalized to the ␤-actin mRNA expression. Effects of Cd exposure on P-gp mRNA levels were not significant (P > 0.05). Vertical bars represent standard errors, N = 10.

Cd exposure resulted in a significantly elevated (by 46%) rate of oxygen consumption (MO2 ) of isolated oyster gills (Fig. 4) (ANOVA: F1, 10 = 8.65, P = 0.015). No decrease in MO2 was observed in response to the inhibitors of P-gp ATPase activity, cyclosporine A and JS-2190 in gills from control oysters (Fig. 4). In Cd-exposed oys-

A.V. Ivanina, I.M. Sokolova / Aquatic Toxicology 88 (2008) 19–28

Fig. 4. Effect of P-gp inhibitors (cyclosporine A and JS-2190) on tissue oxygen consumption rate (MO2 ) of isolated gills from control and Cd-exposed oysters. Total respiration rate was measured in the absence of inhibitors. An asterisk denotes the MO2 value significantly different from the respective control (no Cd) (P < 0.05). Change in tissue MO2 in response to the inhibitors of ATPase activity of P-gps (cyclosporine A and JS-2190) was not significant (P > 0.05). Vertical bars represent standard errors, N = 6.

ters, addition of the P-gp inhibitors CA and JS-2190 resulted in an average MO2 decrease by 13–14% (Fig. 4); however, this effect was not significant in either control or Cd-exposed group (F2, 20 = 0.40, P = 0.678). The interaction between the effects of Cd exposure and P-gp inhibitors on gill MO2 was also not significant (F2, 20 = 0.83, P = 0.449). 4. Discussion Our study clearly shows that P-gp is expressed on cell membrane and in mitochondria of the eastern oysters. While earlier studies have shown the presence of P-gp-like proteins in whole cell extracts of C. virginica (Minier et al., 1993; Keppler and Ringwood, 2001) and other bivalves (Eufemia and Epel, 2000; Achard et al., 2004), their subcellular localization was not examined. This study for the first time demonstrates that P-gp is expressed in oyster mitochondria at the levels comparable to those in cell membrane when measured per unit protein mass. These findings are in line with earlier studies on multidrug-resistant mammalian cell lines that demonstrated P-gp expression in intracellular membranes (including mitochondria) (Maraldi et al., 1999; Gong et al., 2003; Rajagopal and Simon, 2003; Munteanu et al., 2006; Solazzo et al., 2006). The function of P-gp in oyster mitochondria remains to be elucidated. In mammalian models, mitochondrial localization of P-gP and its function is still debated. It appears to be predominantly expressed in mitochondria of some multidrug resistant cell lines but not in non-transformed ones (Munteanu et al., 2006; Solazzo et al., 2006; but see Patterson and Gottesman, 2007) and well as in endoplasmic reticulum and Golgi apparatus of multidrug-resistant cells where it has been proposed to pump drugs from cytoplasm into these organelles thus protecting critical nuclear targets from exposure to xenobiotics (Maraldi et al., 1999; Gong et al., 2003; Rajagopal and Simon, 2003; Munteanu et al., 2006; Solazzo et al., 2006). In this study, addition of MXR inhibitors (JS-2190 and verapamil) tended to reduce accumulation of a MXR substrate rhodamine 123 in mitochondria from Cd-exposed oysters consistent with the direction of P-gp transport in multidrug-resistant cells’ mitochondria. It is worth noting that in mammalian models, JS-2190 is known to inhibit both P-gp and MRP1 transporters which opens a possibility that in addition to P-gp, MRP-like proteins may contribute to the observed MXR activity in oyster mitochondria. Currently, there is no evidence that MRP-like transporters are present in mollusks at the gene or transcript levels (Bard, 2000; Tutundjian and Minier, 2007) suggesting that the observed JS-2190-inhibitable transport

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in oyster mitochondria (or whole-tissue preparations, described below) is most likely due to P-gp activity. In contrast, MXR inhibitors had no effect on rhodamine 123 accumulation in control mitochondria possibly indicating lower P-pg activity. Notably, P-gp protein levels were similar in gill mitochondria from control and Cd-exposed oysters indicating that the apparent increase in P-gp activity in Cd-exposed mitochondria may be due to post-translational mechanisms. However, further investigations (e.g. with more sensitive assays and/or P-gp substrates that translocate into mitochondria irrespective of the mitochondrial membrane potential) are needed to confirm the presence of mitochondrial P-gp activity and the direction of their pumping in oysters, especially under the control, non-stressed conditions. Studies of mitochondrial P-gp transport in aquatic organisms under different physiological and environmental conditions could represent an interesting and fruitful avenue for future studies. In contrast to mitochondria, cell membrane P-gp of C. virginica pumps substrates out of the cell as indicated by decreased rates of rhodamine B dye efflux from oyster gills in the presence of MXR inhibitors. This is consistent with earlier studies on P-gp and related ABC transporters in other organisms (Smital et al., 2000; Hennessy and Spiers, 2007). Notably, P-gp protein levels in oyster cell membranes significantly increased (by 2–2.5-fold) in response to Cd exposure accompanied by 3.5–7-fold elevated P-gp activity. This increase in P-gp protein expression may partially contribute (along with other protective proteins such as metallothioneins and heat shock proteins) to an overall increase in protein synthesis rates observed in Cd-exposed oysters (Cherkasov et al., 2006; Ivanina et al., 2008). However, mRNA expression level of p-gp gene did not change in response to Cd exposure suggesting that P-gp regulation in oysters occurs on post-transcriptional level. Other studies have also noted the absence of strict correlation between mRNA and protein levels of P-gp (Shirasaka et al., 2007 and references therein) and suggested that post-transcriptional control of P-gp synthesis may occur via mechanisms such as stabilization of mRNA pool, extension of mRNA half-life and regulation of the transcriptional block of existing mRNA (Randle et al., 2007). Earlier studies have also shown that P-glycoproteins undergo considerable post-translational modification including phosphorylation and glycosylation (Bard, 2000). P-gp phosphorylation by protein kinases A and C have been shown to enhance transport activities and ATP hydrolysis by P-gps (Bard, 2000 and references therein) whereas sphingol (a lysosphingolipid protein kinase C inhibitor) reduced P-gp-mediated substrate transport (Sachs et al., 1995). Glycosylation of P-gp was also shown to affect substrate transport efficiency (Schinkel et al., 1993). Post-translational regulation of P-gp activity may explain why in our study the substrate transport activity of P-gp increased 3.5–7 times after Cd exposure while the protein levels only increased 2–2.5 times. This may also explain a trend to higher activity of P-gps in isolated mitochondria from Cd-exposed oysters despite the absence of elevated P-gp protein levels compared to the controls (c.f. Figs. 1 and 3). Overall, the complexity of post-transcriptional and post-translational regulation of P-gps suggests that direct determination of P-gp transport activity may be the best measure of xenobiotic-resistant phenotype, while P-gp protein expression can be used as an indication of the relative levels (and thus costs) of P-gp protein synthesis in exposed organisms. In contrast, p-gp mRNA levels are not a good marker of the P-gp phenotype due to the likely involvement of multiple post-transcriptional regulatory steps that finally determine activity levels of this protein in the cell. What is the functional significance of elevated P-gp expression in Cd-exposed organisms? Earlier studies have shown that Cd induces elevated levels of P-gp transporter proteins in mammalian

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cell lines (Chin et al., 1990; Endo et al., 2002; Huynh-Delerme et al., 2005), a freshwater bivalve Corbicula fluminea (Achard et al., 2004; Legeay et al., 2005) and in a marine mollusk C. virginica (this study). It is tempting to assume that this elevation is a functionally adaptive response elicited by Cd exposure which aims to limit the metal accumulation in exposed cells. Although divalent metals are usually not considered classical P-gp substrates, recent studies suggest that P-gp and related proteins (MDR1, and a distantly related MRP1) may play an important role in the outward Cd transport in some cell types limiting Cd accumulation (Broeks et al., 1996; Endo et al., 2002; Callaghan and Denny, 2002; Kimura et al., 2005). The mechanisms of this transport are not fully understood and may involve extrusion of Cd conjugated to glutathione, ATP or other organic ligands (Tommasini et al., 1996; Li et al., 1997; Borst et al., 2000; Thevenod et al., 2000). In freshwater amphipods, addition of sublethal amounts of Cd (0.22–2.2 ␮M) to the ambient water resulted in a decreased efflux of rhodamine B resembling the effects of verapamil and other chemosensitizers (Timofeyev et al., 2007) and suggesting that Cd (or some endogenous Cd-induced molecules) may serve as MXR substrates. P-gps were also found to contribute to Cd resistance in a variety of systems including nematodes, insects, fungi and mammalian cell lines (Broeks et al., 1996; Thevenod et al., 2000; Callaghan and Denny, 2002). In Cdresistant Caenorhabditis elegance, targeted inactivation of a P-gp family gene, mdr1, resulted in increased sensitivity to Cd (Broeks et al., 1996), and functional inhibition of P-gps with verapamil led to a similar effect in Drosophila melanogaster (Callaghan and Denny, 2002). Mammalian MDR1 was implicated in protection against Cd-induced apoptosis in kidney proximal tubule cells, and pharmacological P-gp inhibitors reversed this protective effect (Thevenod et al., 2000). A yeast mutant with a disrupted p-gp-like gene also had reduced tolerance to Cd exposure and was rescued by heterologous expression of the human multidrug resistance-associated protein (Tommasini et al., 1996). Generally, these data indicate that elevated P-gp expression may be a protective response against Cd exposure in a variety of organisms. Other metals (notably arsenic in the form of arsenate) can also induce elevated P-gp expression suggesting a similar protective action (Chin et al., 1990; Eufemia and Epel, 2000; Yague et al., 2003). In this context it is worth noting that P-gp and related transporters (e.g. MDR1) can also be non-specifically induced by other stressors including heat shock, low salinity, anaerobiosis and UV radiation (Chin et al., 1990; Minier et al., 2000; Yague et al., 2003; ¨ ¨ Ludeking and Kohler, 2004). In ectotherms such as oysters, P-gp expression was shown to vary seasonally and to positively correlate with the habitat temperature (Keppler and Ringwood, 2001). The functional significance of P-gp expression in UV-, heat- or metabolically stressed cells, or the nature of endogenous substrates (if any) that may need to be transported out of the cell under these conditions are currently not known. It has been proposed that elevated P-gp expression in cells exposed to metals (such as Cd), UV, ROS and heat stress may be due to increased production of ceramides (generated by sphingomyelase in response to cellular stress) which serve as substrates for P-gps (Thevenod et al., 2000 and references therein). In any case, it is clear that P-gp expression is an important constituent of cellular protection arsenal and may not only limit uptake of the xenobiotic substrates as traditionally proposed but also provide protection against a variety of other stressors through a mechanism that requires further elucidation. Thus, the current data suggest that P-gp expression may not be a useful biomarker of exposure to specific contaminants (such as metals and/or organic substrates) but should rather be considered as a general biomarker of environmental stress. The energy costs of the transport activity of P-gps were estimated in control and Cd-exposed oysters by two comple-

mentary approaches: directly, by measuring MO2 of isolated gills in the presence and absence of P-gp inhibitors and indirectly, by estimating ATP requirements for the transport of a fluorescent substrate, rhodamine B (RB). Based on the P-gpdependent RB extrusion rates and assuming two ATP molecules consumed for each substrate molecule transported (Ambudkar et al., 1999; Hennessy and Spiers, 2007), we estimate that control oysters consume 5.03–9.54 pmol O2 min−1 g−1 (if respiring on glycogen) or 6.64–12.59 pmol O2 min−1 g−1 (if oxidizing proteins) to fuel the outward P-gp-dependent RB transport; for lipid oxidation, intermediate values of oxygen demand are expected. Translating this into respiratory equivalents, this amount of ATP represents 0.005–0.010% of total respiration activity of control oyster gills. On the other hand, Cd-exposed oysters consume 30.83–36.19 pmol O2 min−1 g−1 (based on glycogen oxidation) or 40.68–47.74 pmol O2 min−1 g −1 (based on protein oxidation) for P-gp substrate transport, which equals to 0.023–0.037% of total respiration. These values are in general agreement with the estimated costs of P-gp efflux transporter activity in developing sea urchin embryos (0.023% of total respiration) (Epel et al., 2006). However, these calculations do not take into account the basal ATP hydrolysis of idling P-gps. Direct determinations of MO2 of oyster gills in the presence and absence of cyclosporine A or JS-2190 both of which inhibit ATPase activity of P-gps and some other MXR transporters (e.g. putative MRP-like proteins) should therefore provide a better insight into energy costs of MXR phenotype in control and Cdexposed oysters. This study indicates that MXR inhibitors have no significant effect on oxygen consumption rates by oyster gills supporting a notion that these pumps are no major players in cellular energy costs. It also indicates that the MXR activity does not significantly contribute to the observed increase in standard metabolic rates in isolated cells (Cherkasov et al., 2006), tissues (this study) and whole oysters (Lannig et al., 2006) in response to Cd exposure and suggests that other, yet not fully understood mechanisms must be invoked to explain significant elevation of SMR in Cd-exposed oysters. Acknowledgements This work was supported by funds provided the National Science Foundation CAREER award (IBN-0347238), Undergraduate Summer Research Fellowship of the American Physiological Society and Johnson C. Smith University MBRS-RISE Program NIGMS 58042. The authors thank Kristen Reynolds for conducting dose–response experiments for P-gp inhibitors and assisting in optimization of MO2 assay. All experiments complied with the current laws of the country (USA) where they were performed. References Achard, M., Baudrimont, M., Boudou, A., Bourdineaud, J.P., 2004. Induction of a multixenobiotic resistance protein (MXR) in the Asiatic clam Corbicula fluminea after heavy metals exposure. Aquat. Toxicol. 67 (4), 347–357. Ambudkar, S.V., Dey, S., Hrycyna, C.A., Ramachandra, M., Pastan, I., Gottesman, M.M., 1999. Biochemical, cellular, and pharmacological aspects of the multidrug transporter 1. Annu. Rev. Pharmacol. 39, 361–398. Bard, S.M., 2000. Multixenobiotic resistance as a cellular defense mechanism in aquatic organisms. Aquat. Toxicol. 48 (4), 357–389. Bergmeyer, H.U., 1985. Methods of enzymatic analysis. Metabolites 3: Lipids. Amino Acids and Related Compounds. VCH Verlagsgesellschaft, Weinheim. Borst, P., Evers, R., Kool, M., Wijnholds, J., 2000. A family of drug transporters: the multidrug resistance-associated proteins. J. Natl. Cancer Inst. 92, 1295–1302. Broeks, A., Gerrard, B., Allikmets, R., Dean, M., Plasterk, R.H.A., 1996. Homologues of the human multidrug resistance genes MRP and MDR contribute to heavy metal resistance in the soil nematode Caenorhabditis elegans. EMBO J. 15, 6132– 6143. Callaghan, A., Denny, N., 2002. Evidence for an interaction between P-glycoprotein and cadmium toxicity in cadmium-resistant and -susceptible strains of Drosophila melanogaster. Ecotoxicol. Environ. Saf. 52, 211–213.

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