Coral Reefs (2014) 33:523–535 DOI 10.1007/s00338-014-1136-3
REPORT
Natural variations in xenobiotic-metabolizing enzymes: developing tools for coral monitoring L. R. A. Rouge´e • R. H. Richmond A. C. Collier
•
Received: 9 August 2013 / Accepted: 27 February 2014 / Published online: 13 March 2014 Ó Springer-Verlag Berlin Heidelberg 2014
Abstract The continued deterioration of coral reefs worldwide demonstrates the need to develop diagnostic tools for corals that go beyond general ecological monitoring and can identify specific stressors at sublethal levels. Cellular diagnostics present an approach to defining indicators (biomarkers) that have the potential to reflect the impact of stress at the cellular level, allowing for the detection of intracellular changes in corals prior to outright mortality. Detoxification enzymes, which may be readily induced or inhibited by environmental stressors, present such a set of indicators. However, in order to apply these diagnostic tools for the detection of stress, a detailed understanding of their normal, homeostatic levels within healthy corals must first be established. Herein, we present molecular and biochemical evidence for the expression and activity of major Phase I detoxification enzymes cytochrome P450 (CYP450), CYP2E1, and CYP450 reductase, as well as the Phase II enzymes UDP, glucuronosyltransferase (UGT), b-glucuronidase, glutathione-S-transferase (GST), and arylsulfatase C (ASC) in the coral Pocillopora damicornis. Additionally, we characterized enzyme expression and activity variations over a reproductive cycle within a coral’s life history to determine natural
Communicated by Biology Editor Dr. Ruth Gates L. R. A. Rouge´e R. H. Richmond (&) Kewalo Marine Laboratory, Pacific Biosciences Research Center, University of Hawaii at Manoa, 41 Ahui Street, Honolulu, HI 96813, USA e-mail:
[email protected] L. R. A. Rouge´e A. C. Collier Department of Tropical Medicine, Medical Microbiology and Pharmacology, John A. Burns School of Medicine, University of Hawaii at Manoa, 651 Ilalo Street, Honolulu, HI 96813, USA
endogenous changes devoid of stress exposure. Significant changes in enzyme activity over the coral’s natural lunar reproductive cycle were observed for CYP2E1 and CYP450 reductase as well as UGT and GST, while bglucuronidase and ASC did not fluctuate significantly. The data represent a baseline description of ‘health’ for the expression and activity of these enzymes that can be used toward understanding the impact of environmental stressors on corals. Such knowledge can be applied to address causes of coral reef ecosystem decline and to monitor effectiveness of mitigation strategies. Achieving a better understanding of cause-and-effect relationships between putative stressors and biological responses in corals, and other marine invertebrates, can guide and evaluate mitigation and conservation approaches for marine ecosystem protection. Keywords Detoxification Enzymes Coral Xenobiotic metabolism Marine conservation Reproduction
Introduction Coral reef ecosystems are declining at an alarming rate with nearly 20 % of the world’s reefs already considered completely degraded since 1950, and an estimated 35 % of the remaining reefs under imminent risk of being lost within the next few decades (Wilkinson 2008). This is an issue that has gained heightened levels of attention due to increasing episodes of mass coral bleaching and mortality caused by climate change and pollution (Normile 2010). Coral reefs are highly diverse and threatened ecosystems that are ecologically, economically, and culturally valuable through supporting fisheries and ecotourism, subsistence activities, and providing protection from wave damage for
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Fig. 1 Biotransformation of compounds (endogenous and exogenous) into polar metabolites for excretion following Phase I (solid line), Phase II (solid line), or Phase I and II enzyme reactions (dotted line). Conjugated compounds can be regenerated to parent compound through cleavage of large polar group by cleavage enzymes
tropical islands and coastal communities (Richmond 1993; Sebens 1994). Economists estimate the global asset value of coral reefs at 800 billion US dollars, with a net 30 billion US dollars in goods and services provided annually (Cesar et al. 2003). Despite their vital role in our biosphere, our understanding of coral reef ‘health’ is limited. Typically referred to as the state of an organism that is free of injury or illness, this definition of health encompasses a wide range of measurements that can be considered indicators. Current monitoring techniques for coral reefs use population-level measurements such as species diversity, species abundance, percent coral cover, and coral distribution to quantify reef state, with mortality or loss as the key metric of change. Applying a human health analogy using a person’s death as the primary evidence of a health problem would be a poor diagnostic approach. On the other hand, cellular diagnostics present an approach to defining molecular parameters (biomarkers) that can reflect the systematic cellular operation during normal cellular function (homeostasis) for an individual organism (Downs 2005). However, it is only after the homeostatic levels for healthy organisms are established, that theses measures can be applied to understand how particular stressed conditions, such as pollutant exposure or disease, alter the levels. Unfortunately, coral molecular physiology is poorly understood, necessitating studies that identify potential cellular measures of interest, provide evidence for their existence in corals and ultimately determine the levels in nonstressed, healthy corals prior to attempting to establish the measures as biomarkers of health. Continued human interaction with our oceans has resulted in the increased presence and levels of
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anthropogenic pollutants (xenobiotics) detected in our waters (Jackson 2010). Detoxification enzymes are responsible for altering the chemical structures of such xenobiotics within an organism, usually by the initial addition of a polar group, followed by conjugation with a substrate, and then excretion in order to avoid cellular damage (Fig. 1). Since detoxification enzymes are readily induced or inhibited by environmental and genetic stressors, they present a set of measures that can potentially be used as diagnostic markers (Poly 1997; Buckley and Klaassen 2009). The enzymes performing these reactions are designated Phase I and Phase II (Scott 1999; Guengerich 2006), with the former responsible for hydrolysis, reduction, and oxidation (Jokanovic´ 2001; Parkinson and Ogilvie 2001) and the latter for conjugation with large hydrophilic groups, increasing water solubility, and facilitating excretion (Fig. 1). Among the Phase I enzymes, the cytochromes P450 (CYP) superfamily is responsible for the majority of Phase I biotransformation reactions (Guengerich 2008). Within this superfamily, particular isoforms are responsible for the detoxification of various types of chemical structures including environmental toxicants (Table 1). The Phase II reactions are performed by a variety of enzyme families that differ in the substrate molecule conjugated to various xenobiotics (Table 1). Since these enzymes are also involved in essential biological processes such as digestive metabolism, growth and development, cell–cell communication, sexual reproduction, and hormone metabolism, it is insufficient to merely identify protein presence or activity at a discrete time point. Rather, it is vital to characterize normal expression and activity fluctuations over the coral’s life history stages and cycles in order to
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Table 1 Substrate examples for major Phase I and Phase II enzymes Classification
Enzyme
Substrate examples
Phase I
Cytochrome P450 reductase
Responsible for electron transfer to all cytochromes P450 isoforms
Cytochrome P450 1A1 & 1A2
Aromatic hydrocarbons (e.g., phenanthrene), biogenic aromatics
Cytochrome P450 2E1
Ethanol, benzene, acetaldehyde, trichloroethylene, arachidonic acid
Cytochrome P450 3A4
Tamoxifen, alfatoxin, steroids, prostaglandins
UDP glucuronosyltransferase
Pesticides (e.g., methoxychlor), acetaminophen, propofol, steroid hormones
b-glucuronidase
Regeneration enzyme— removes conjugated glucuronic acid molecule Pesticides (e.g., DDT), acetaminophen, chemotherapeutics, carcinogens
Phase II
Glutathione-S-transferase
Sulfotransferase
Arylsulfatase C
Pesticides (e.g., pentachlorophenol), carcinogens, steroid hormones, catecholamines Regeneration enzyme— removes conjugated sulfate molecule
later consider changes caused by exogenous chemicals and stressors in relation to normal baseline fluctuations of these enzymes. The application of molecular techniques that use protein biomarkers as indicators of cellular alteration has proven successful in corals to assess sublethal stress (Gassman and Kennedy 1992; Downs et al. 2000, 2005, 2006; Morgan et al. 2001; Galloway et al. 2002; Rouge´e et al. 2006; Ramos and Garcı´a 2007). However, concerns have been raised regarding the use of antibodies for coral enzyme detection where the genome did not exist to confirm the specificity of the antibodies generated. Alternatively, others have focused solely on transcriptional or genomic methods. Studies based purely on mRNA expression or gene sequencing may be problematic, since it is widely recognized that many enzymes require a high degree of post-transcriptional and post-translational modification for activity (Lodish et al. 2008). To the best of our knowledge, there are currently no complete studies in corals that demonstrate protein expression and activity levels
(including the correlation between the two) for detoxification enzymes. Additionally, no studies have presented a comprehensive description of natural intra- and inter-colony fluctuations of major Phase I and Phase II metabolizing enzymes over reproductive and life cycles in corals. This study provides molecular and biochemical evidence for the presence and activity of Phase I and Phase II metabolizing enzymes in the pan-Pacific reef coral Pocillopora damicornis. Additionally, we describe fluctuations in expression and activity of these enzymes that correlate with natural gametogenic and reproductive cycles in healthy corals. The species of coral in question is ideal for this investigation because it produces larvae on a lunar cycle each month throughout the year in many locations across its wide range, providing an opportunity to study enzyme activity changes tied to both endogenous cues and exogenous stressors.
Materials and methods Coral collection and experimental procedures Coral branches (*7 cm tall, 2.5 cm wide) and whole coral colonies (15 cm diameter) of P. damicornis were collected from Coconut Island in Kaneohe Bay, Oahu, under Special Activities Permit 2008-47, granted by the Department of Land and Natural Resources under the Division of Aquatic Resources. Coral colonies were sampled at random, cleaned of any foreign organisms, and quarantined prior to introduction to the tanks at the Kewalo Marine Laboratory facilities. Colonies were maintained in separate seawater tanks and allowed a minimum of 14 days to recuperate from possible stress from collection and to acclimatize to the Kewalo water system (salinity of 36 ± 1 parts per thousand, temperature 27 ± 2 °C, geometric mean ± standard error (SE); measurements taken manually almost daily; n = 25) prior to tissue sampling. Fragments from separate colonies (n = 5) were processed immediately after sampling at a predetermined ‘reference’ time point between peak reproductive events. Individual colonies (n = 5) were repeatedly sampled at selected time points during the reproductive lunar cycle, defined as from one full moon to the next, approximately 29 d (Richmond and Jokiel 1984). Samples were taken in seven-day increments before and after a P. damicornis planulation event, which occurs every third quarter in the lunar cycle (Richmond and Jokiel 1984; Kolinski and Cox 2003). To appropriately represent the reproductive cycle timing of P. damicornis, the data are reported as days from the start of sampling, with day 1 representing the reference time point at the first quarter moon for the coral fragments only. This time point also represents 2 weeks after the
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previous planulation event, and 2 weeks before the following planulation event. The subsequent sampling time points represent the lunar quarters for the coral colonies sampled consecutively throughout the lunar cycle as follows: first quarter moon (2 weeks before planulation event, day 28), full moon (one week before next planulation event, day 35), third quarter moon (planulation event, day 42), and new moon (one week after planulation event, day 49). If the lunar cycle fluctuations are reliable, day 28 sample results should mirror results from day 1. Sampling and preparation of postmitochondrial subcellular fraction of coral protein At the designated time points, coral samples were taken and placed into 50-ml Falcon tubes. The tubes were immediately submerged in liquid nitrogen to snap-freeze protein levels and activities, and to prevent further coral stress reactions. Samples were then stored at -80 °C until further processing. The postmitochondrial subcellular fraction of coral host protein, void of zooxanthellae, mitochondria, plasma membrane, and nuclei was prepared as follows. Coral samples were crushed using sterilized instruments, placed into a 100-ml Erlenmeyer flask on ice, and 15 ml of cold homogenization buffer made of 0.2 lm filtered sea water (FSW) and 1 mM phenylmethylsulfonylfluoride added. Flasks were shaken for 10 min by hand; then, the liquid containing the tissue samples was transferred to a clean vessel and homogenized on ice with an Ultra-Turrax homogenizer for 30 s. The homogenate was centrifuged at 800g for 4 min at 4 °C in an Eppendorf Centrifuge 5810R (Eppendorf, Hauppauge, NY), and the pellet containing coral skeleton fragments was discarded. The supernatant, containing coral tissue, was centrifuged at 10,000g for 20 min at 4 °C, the supernatant decanted, and the tissue pellet resuspended in 1 ml homogenization buffer and then processed on ice using a glass homogenizer for 2 min (approx. 30 strokes). The homogenate was subsequently centrifuged at 10,000g for 10 min at 4 °C in an Eppendorf Microcentrifuge 5415D (Eppendorf, Hauppauge, NY). After this final centrifugation, the supernatant represented the postmitochondrial subcellular fraction of host coral protein containing cytosol and microsomes, subsequently referred to as ‘coral protein,’ and was aliquoted and frozen at -80 °C until use. Before use, all samples were assessed for protein concentration and standardized using the BCA method described by Smith et al. (1985). Kinetic enzyme assays Coral protein (10–150 lg), buffer, and substrate were loaded into wells on 96-well format microplates kept on
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ice. For substrates dissolved in solvents, that volume was never more than 2 % of the reaction volume; hence, solvent carrier properties did not affect enzyme activities (Williams et al. 2008). Microplates were pre-warmed inside a microplate reader (Spectra Max or Gemini XS, Molecular Devices, Sunnyvale, CA) at 37 °C before the addition of the cofactors used to initiate the reaction. Fluorescent assays were performed in black flat-bottomed microplates, colorimetric assays in clear microplates, and UV assays in optically clear microplates. Individual enzyme activities were assessed as outlined below. Cytochrome P450 reductase (CYPR) Cytochrome P450 reductase (CYPR) was measured using a modification of the method of Ardies et al. (1987). Coral protein (30 lg) in 0.1 M potassium phosphate buffer, pH 7.4, and 50 lg cytochrome c were added to each well, preincubated for 3 min at 37 °C, and a reference reading at k = 550 nm taken. Reactions were then initiated by the addition of 1 mM NADPH. Absorbance was monitored continuously at k = 550 nm and activity calculated using Beer’s Law with e = 21 mM-1 cm-1 (Ardies et al. 1987). Diphenyliodonium chloride was used as a specific inhibitor for CYPR at 5 mM to provide further evidence of specific, inhibitable enzyme activity (Tew 1993). Cytochrome P4501A (CYP1A) Substrates ethoxyresorufin (Dutton and Parkinson 1989), 3-cyano-7-ethoxycoumarin (Crespi et al. 1997), 7-benzyloxy-4-trifluoromethylcoumarin (BFC), and methoxyresorufin were used to determine general cytochrome P4501A (CYP1A class) activity. Specific inhibitors a-naphthoflavone (Dubey et al. 2003) and furafylline (Donato et al. 2004), for CYP1A1 and CYP1A2, respectively, were added to separate reactions in order to determine isoform specific contributions to the general CYP1A activity. Cytochrome P450 2E1 (CYP2E1) Activity of cytochrome P450 2E1 (CYP2E1) was determined using a modification of the method of Donato et al. (2004). Briefly, coral protein (150 lg) in 0.1 M potassium phosphate buffer, pH 7.4, with 5 mM MgCl2 and 1 mM 7-methoxy-4-trifluoromethylcoumarin (MFC) were added to each well. Pre-incubation at 37 °C occurred for 3 min; then, reactions were initiated with the addition of 1 mM NADPH. Plates were incubated in the dark at 37 °C for 30 min and fluorescence measured at 410 nm ex/510 nm em. Results were transformed to fmol min-1 mg of protein-1 using a standard curve generated with 7-hydroxy-4trifluoromethylcoumarin. While MFC substrate is
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principally metabolized by CYP2E1, other enzymes contribute to its metabolism; hence, to isolate the CYP2E1specific activity, separate reactions were performed in parallel that included 100 lM sodium diethyldithiocarbamate that specifically inhibits CYP2E1 (Chang et al. 1994; Donato et al. 2004). Total CYP2E1 activity was defined as total MFC turnover minus MFC turnover in the presence of diethyldithiocarbamate.
Fluorescence was continuously monitored at 355 nm ex/ 460 nm em. Results were transformed to pmol min-1 mg protein-1 using a standard curve generated with 4-MU. To confirm the specificity of the reaction, saccharolactone (100 lM) was used to inhibit b-glucuronidase activity (Oleson and Court 2008).
Cytochrome P450 3A4 (CYP3A4)
The activity of sulfotransferase (SULT) 1A1 isoform was measured using the method of Frame et al. (2000), while general SULT (including 1A1, 1A2, 1A3, 1B1, 1E1, 2A1 isoforms) was measured using the method by Tabrett and Coughtrie (2003).
The BFC substrate was used to determine cytochrome P450 3A4 (CYP3A4) activity, and ketoconazole, 100 lM, was used to specifically inhibit CYP3A4 (Crespi et al. 1997; Donato et al. 2004).
Sulfotransferase
Arylsulfatase C Glutathione-S-transferase Activity of glutathione-S-transferase (GST) was determined using a method adapted from Habig et al. (1974) and Gonza´lez et al. (1989). Optically clear microplates on ice (Greiner Bio-One, Monroe, NC) were loaded with 0.5 mM 1-chloro-2,4-dinitrobenzene (in DMSO) and coral protein (10 lg). After pre-incubation (3 min at 37 °C), reactions were initiated through the addition of 1 mM L-glutathione. Absorbance was monitored continuously at k = 340 nm. Total activity was calculated using Beer’s law with e = 9.6 mM-1 cm-1 (Habig et al. 1974). Specific activity of GST-Pi (GST-P) isoform was further confirmed by using triphenyl phosphate (100 lM) to knock out GST-P activity (Wu et al. 2007). UDP–glucuronosyltransferase Total glucuronosyltransferase (UGT) activity was determined using the method of Collier et al. (2000) with the exception that 50 lg ml-1 alamethicin (in DMSO) was used as the UGT activator, 30 lg of coral protein was used, and 5 mM sacchrolactone was included in each reaction, since we identified b-glucuronidase activity. Fluorescence was monitored continuously at 355 nm ex/460 nm em and results transformed to pmol min-1 mg protein-1 using a standard curve generated with 4-methyl umbelliferone sodium salt (4MU). Curcumin (100 lM) was used to confirm specific UGT activity by inhibiting 4-MU metabolism (Volak et al. 2008). b-Glucuronidase b-Glucuronidase activity was determined using the method of Trubetskoy and Shaw (1999). Microplates on ice were loaded with coral protein (10 lg) and buffer; plates were pre-incubated at 37 °C for 5 min, and the reaction initiated with 100 lM 4-methyl umbelliferyl-beta-D-glucuronide.
Activity of arylsulfatase C (ASC) isoform was determined using a modification of the method of Roy (1958). Microplates were kept on ice, and coral protein in 0.1 M Tris– HCl buffer, pH 7.4, containing (50 lg) was added to each well. The samples were pre-incubated for 5 min at 37 °C; then, reactions were initiated with the addition of 100 lM para-nitrophenyl sulfate. Absorbance was monitored continuously at k = 400 nm and results generated using a standard curve of para-nitrophenol. ASC activity was significantly reduced through a decrease in the pH of the assay buffer to pH 6.5 (Roy 1976; Chang and Moudgil 1984). SDS-PAGE and Western blot gel electrophoresis SDS-polyacrylamide gel electrophoresis (PAGE) and Western blot analysis were used to separate all target proteins (Laemmli) except for b-glucuronidase, which was electrophoresed under nonreducing conditions without SDS prior to Western blot analysis. Total soluble protein (20 lg) for each sample was electrophoresed in 7–12 % polyacrylamide gels, and the proteins transferred to polyvinylidene difluoride membranes using a Bio-Rad semi-dry transfer system (Hercules, CA, USA) and blocked overnight (*8–10 h) at 4 °C with 10 % nonfat dry milk in phosphate-buffered saline (PBS) containing 0.05 % Tween-20 (PBST). The next day, blots were washed three times for 10 min using PBST, and the membranes incubated for 2 h at room temperature with biotinylated primary antibody (1:50 for b-glucuronidase; 1:100 for SULT1A1, ASC; 1:200 for CYP1A1, CYP1A2, UGT1A1, UGT 2B; 1:500 for GST; 1:1,000 for CYPR; 1:2,000 for CYP2E1, in 20 % nonfat dry milk in PBST). Antibodies were purchased from Abcam (Cambridge, MA, USA) and Santa Cruz Biotechnology Inc. (Santa Cruz, CA, USA). The primary antibody was decanted, and blots were washed three times for 10 min with PBST and then incubated with streptavidin-biotinylated horseradish peroxidase (GE
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Healthcare Bio-sciences, Piscataway, NJ) for 1 h at room temperature. After a final three 10-min washes with PBST and one 5-min wash with PBS, membranes were developed using chemiluminescent substrate. The amount of enzyme proteins was quantified by comparison with an internal standard (human liver protein) using area/density analysis and the program Image J (Wayne Rasband, NIH, Bethesda, MD). Statistical analyses Statistical analyses were performed using the program GraphPad Prism Program version 5.0. Normality of the data was checked through the D’Agostino-Person normality test. Equality of variance of the samples was verified using the Bartlett’s test. A one-way analysis of variance (ANOVA) was used if the data were found to be normally distributed and homogeneous. However, if the data did not meet the requirements for homogeneity or variances, the Kruskal–Wallis one-way ANOVA (KS) on ranks was used. The Bonferroni (for one-way ANOVA) and the Dunns (for KS) methods for multiple comparison post hoc tests were used. Statistical significance was defined as p B 0.05.
Results The Phase I enzymes CYPR and CYP2E1, as well as the Phase II enzymes UGT, b-glucuronidase, GST, and ASC
Fig. 2 The activity and protein expression of cytochrome P450 reductase (CYPR) and cytochrome P450 2E1 (CYP2E1). a CYPR enzyme activity, b CYP2E1 enzyme activity, c CYP2E1 Western Blot protein intensity, and d CYP2E1 correlation of protein intensity with enzyme activity. Dashed line represents 95 % confidence interval with Pearson correlation coefficient (r) and p value (p). Data with different lower case letters differ significantly at a B 0.05. Asterisk denotes significant difference between all other groups at a B 0.05. Bars are means ± SE of n = 5 corals assayed in triplicate
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were present and active in colonies of P. damicornis. In contrast, CYP1A, CYP3A4, and SULT1A were not present. Significant fluctuations of CYPR, CYP2E1, UGT, and GST occurred over the lunar reproductive cycle (29 d) demonstrating that these enzymes fluctuate naturally in this coral, necessitating consideration of natural fluctuations in any subsequent measurements of these coral enzymes where the purpose of such measurements is to quantify changes caused by environmental, biological, or exogenous stressors (Figs. 2, 3, 4). Phase I enzymes Activity was observed biochemically for CYPR and CYP2E1 (Fig. 2). No activity was observed for CYP1A1, CYP1A2, and CYP3A4, and similarly, Western blots for these enzymes yielded no conclusive results under our conditions (data not shown). Variations in the activity rates of CYPR are shown in Fig. 2a, with the geometric mean ± SE enzyme rates for each time point listed in Table 2. These data show variations exist between different coral colonies, as well as within a colony at different time points in the reproductive cycle. This is evidenced by the significant differences (ANOVA, F4,20 = 37.67, p \ 0.0001) in the same monthly time point, 2 weeks after planulation, taken one month apart from different coral colonies (day 1 vs. day 28; Fig. 2a). We hypothesize that the differences in the 2 weeks after planulation time points that were observed indicate inter-colony variation in
a
b
c
d
Coral Reefs (2014) 33:523–535 Fig. 3 The activity and protein expression of UGT and bglucuronidase. a General UGT enzyme activity, b bglucuronidase enzyme activity, c UGT1A1 Western blot protein intensity, d b-glucuronidase Western blot protein intensity, e UGT2B Western blot protein intensity, f b-glucuronidase correlation of protein intensity with enzyme activity, g correlation of UGT1A1 protein intensity with general UGT enzyme activity, and h Correlation of UGT2B protein intensity with general UGT enzyme activity. Dashed line represents 95 % confidence interval with Pearson correlation coefficient (r) and p value (p). Data with different lower case letters differ significantly at a B 0.05. Asterisk represents significance with a p value B 0.05 only between groups under the caped line. Bars are means ± SE for n = 5 corals assayed in triplicate
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a
b
c
d
e
f
g
h
activity for CYPR. This is similar to the significant interindividual variation observed for CYPR in humans and other animal species (Hart et al. 2008). Despite strong activity data, including the utility of specific CYPR inhibitor substrates, we were unable to confirm the presence of CYPR proteins with Western blots since nonspecific protein bands occurred in the correct molecular weight region for our CYPR antibody. CYP2E1 enzyme activities were significantly lower (*twofold decrease; ANOVA, F4,20 = 8.450, p = 0.0004) at the planulation (larval release) time point compared with
all other time points in the lunar cycle (Fig. 2b; Table 2). These data are further supported by Western blot (ANOVA, F4,20 = 6.818, p = 0.0012), where relative levels of the CYP2E1 protein expression showed a significant, strong, positive correlation with enzyme activity (Pearson, r = 0.4390, p = 0.0281; Fig. 2c, d). Phase II enzymes The activity of the UGT superfamily of enzymes was significantly higher at the 2 weeks after planulation time
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530 Fig. 4 The activity and protein expression of glutathione-Stransferase (GST) and arylsulfatase C (ASC) enzymes. a GST enzyme activity, b ASC enzyme activity, c GST Western blot protein intensity, d ASC Western blot protein intensity; e GST correlation of protein intensity with enzyme activity; f ASC correlation of protein intensity with enzyme activity. Dashed line represents 95 % confidence interval with Pearson correlation coefficient (r) and p value (p). Data with different lower case letters differ significantly at a B 0.05. Bars are means ± SE for n = 5 corals assayed in triplicate
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a
b
c
d
e
f
Table 2 Enzyme activity rates for Phase I and Phase II enzymes detected over the lunar reproductive cycle given as means ± SE of n = 5 colonies Enzyme
Day 1
Day 28
Day 35
Day 42
Day 49
Units
Cytochrome P450 reductase
366.5 ± 28.83
694.8 ± 39.28
993.7 ± 61.68
922.1 ± 41.22
632.0 ± 19.62
pmol/min/mg protein
Cytochrome P450 2E1
479.6 ± 54.09
398.3 ± 36.2
371.9 ± 32.24
208.1 ± 15.01
454.0 ± 34.64
fmol/min/mg protein
UDP glucuronosyltransferase
744.8 ± 201.1
714.2 ± 125.3
190.2 ± 11.74
247.4 ± 11.13
372.2 ± 33.46
pmol/min/mg protein
b-glucuronidase
6.65 ± 0.78
7.26 ± 1.06
10.14 ± 1.62
10.13 ± 1.45
9.04 ± 1.39
pmol/min/mg protein
Glutathione-S-transferase
2.16 ± 0.05
2.41 ± 0.02
2.59 ± 0.15
2.49 ± 0.07
2.39 ± 0.06
nmol/min/mg protein
84.38 ± 18.81
145.7 ± 23.42
145.8 ± 30.34
121.9 ± 11.65
181.3 ± 24.11
pmol/min/mg protein
Arylsulfatase C
points for both months (day 1 and day 28; KS, p = 0.0004). Hence, it is likely that the changes in enzyme activity observed are due to the lunar/physiologically derived changes in UGT, and not inter-individual variation (Fig. 3a; Table 2). The substrate used in the general UGT activity assay (4MU) is metabolized by all known UGT isoforms, from both the 1A and 2B subfamilies except UGT1A4 (Uchaipichat et al. 2004). Antibodies to both
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UGT1A1 and a conserved region of the UGT2B subfamily (encompassing all 7 isoforms) were positive for protein bands (Fig. 3c, e). No significant differences across the lunar cycle were observed for UGT1A1 expression (Fig. 3c). Only the 2 weeks after planulation (day 1, coral fragments) and one week before planulation time points (day 35, coral colonies) differed significantly (ANOVA, F4,20 = 3.837, p = 0.0180; Fig. 3e). No correlation was
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found between enzyme activity and relative protein intensity for UGT1A1 (Fig. 2g). However, UGT2B was found to have a strong, significant, inverse correlation, with a higher activity observed with lower amounts of protein (Pearson, r = -0.4788, p = 0.0155; Fig. 2h). For b-glucuronidase, an enzyme performing the inverse metabolic reaction to UGT by hydrolyzing b-glucuronides, no significant differences, or correlations were observed over the lunar time cycle for enzyme activities or protein levels (Table 2; Fig. 2b, d, f). The activity of GST enzymes was mainly consistent across points measured (Table 2) with the exception of the 2 weeks after planulation (day 1, coral fragments) and one week before planulation time points (day 35, coral colonies), which differed significantly (KS, p = 0.0347; Fig. 4a). Relative protein expression mirrored activity levels, except for the 2 weeks after planulation (day 28, coral colonies) where protein expression was significantly lower compared to the one week before (day 35) and planulation (day 42) time points (Fig. 4c). These data indicate a potential cyclical difference in GST expression, not variability in the coral population (inter-colony variation). A strong significant correlation was observed (Pearson, r = 0.4052, p = 0.0445), suggesting that enzyme activity is correlated with amount of protein produced during the lunar cycle (Fig. 4e). No activity was observed in the SULT1A1-specific assay (Frame et al. 2000). Similarly, Western blots for SULT1A1 yielded no visible positive results. A general SULT activity assay using para-nitrophenol was likewise inconclusive. Finally, activity of the ASC enzyme, which performs the inverse reaction to SULT specifically on steroid hormones and phenolic molecules, was observed, but no significant differences or correlations across the lunar reproductive cycle were observed (Fig. 4b, d, f).
Discussion Herein, the protein expressions and/or activities for CYPR, CYP2E1, UGT, b-glucuronidase, GST, and ASC are confirmed in the coral P. damicornis. This study adds to the literature regarding metabolizing enzymes in corals by addressing concerns regarding the use of biochemical substrates (alone) to confirm the presence/functionality of xenobiotic-metabolizing enzymes (XMEs). Specifically, results from enzyme activity assays with specific substrates were further confirmed using enzyme-specific inhibitors. Where substrates were biochemically metabolized, enzyme activity was not accepted unless the specific inhibitor similarly caused the elimination of substrate biotransformation. Additionally, commercial polyclonal antibodies, raised to conserved sequences in enzymes with multiple
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species (both vertebrate and invertebrate) application, provided further evidence for the expression of proteins for these detoxification enzymes. In the past, using antibodies raised to a species, for example in cnidarians, for which the genome sequence is unavailable, has been controversial. The approach taken here was that if an antibody is polyclonal, has been raised to a highly conserved sequence, and has been successfully used across multiple vertebrate and invertebrate species to define protein presence, then this antibody can provide strong supporting evidence of protein presence in the coral P. damicornis. These data raise interesting questions regarding the natural, cyclical variations of detoxification and metabolizing enzymes in corals and other marine invertebrates. Clearly, a natural variation of some enzymes was observed. Specifically, in the case of the UGT enzymes, this appeared to follow reproductive cycles, which is unsurprising given the importance of UGT enzymes for sex steroid and thyroid hormone biotransformation and elimination (Tukey and Strassburg 2000). Conversely, the cleavage enzymes bglucuronidase and ASC that play significant roles in the intracellular transport, conservation, and homeostasis of endogenous compounds did not fluctuate (Beyler and Szego 1954). Based on the known physiological roles of UGT and its apparent fluctuations within the reproductive cycle, this enzyme can be useful for investigating cellular responses to exogenous factors as long as natural cycles are considered in the analyses. The presence of steroid hormones in corals and the glucuronidated forms of these hormones (namely estradiol and testosterone) in surrounding waters during mass spawning events have been extensively documented, suggesting that UGT enzymes have value for better understanding key life history stages critical to the persistence of coral populations (Tarrant et al. 1999, 2003; Pernet and Anctil 2002; Twan et al. 2003, 2006a, b). On the other hand, for enzymes with no known reproductive roles, such as CYP2E1 and CYPR, which still change over the lunar cycle, physiological ‘regulation’ of these enzymes within the cells with other downstream effects may be inferred. The up- or down-regulation of these enzymes may occur in response to the endogenous variations in other cellular pathways as well as in response to exogenous stressors. These types of responses were recently observed by Ramos et al. (2011) while investigating enzyme biomarkers on the coral Siderastrea siderea during nonreproductive and reproductive seasons in two national parks: one afflicted by anthropogenic stressors and one relative reference site. The study observed that the levels of CYP450 and CYP420 were undetectable during the nonreproductive session and that activity of NADPH c reductase and GST differed significantly between the nonreproductive and reproductive seasons (Ramos et al. 2011). Their results demonstrated that the enzymes were
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not only affected by habitat quality, but that the reproductive status of the coral, without exogenous stress, could influence significant changes in the enzymes. Exposure to environmental contaminants is known to effect the regulation of CYP enzymes, resulting in a cascade effect onto other downstream signaling and biotransformation roles. This was a concern highlighted by the recent oil spill in the Gulf of Mexico, which released million of gallons of hydrocarbons that may persist for decades. The results of the exposure may be sublethal to many species of invertebrates, but if these hydrocarbons interfere with critical functions including reproduction and responses to other stressors, the persistence of coral species may be compromised. Since longitudinal reproduction in any species takes several years to study, early detection approaches before populations decline may be facilitated by closer examination of enzymes linked to reproduction and to detoxification. Since our study describes a relatively stable range of activity levels for the b-glucuronidase, ASC, and GST enzymes over the reproductive cycle of ‘healthy’ P. damicornis, the findings suggest that these enzymes may be useful to apply as biomarkers for investigating cellular responses to environmental contamination, particularly acute toxic events, given that the activity resulting from exposure is significantly higher or lower than the inter-colony variation. It is interesting that we did not detect activity for several major enzymes associated with xenobiotic metabolism and exposure to xenobiotics in higher organisms, including CYP3A4, CYP1A1, and CYP1A2, particularly in light of our activity evidence for CYPR, the obligate partner for all CYP isoforms. While sequence evidence suggests that the CYP3 subfamily was derived from a single ancestor gene some 680 million years ago (Nelson 1998), more current phylogenetic data have revealed strong positive selection on human CYP3A4 occurring after the divergence of chimpanzee and human lineages (Qiu et al. 2008). Coupled with the large interspecies variation in the vertebrate CYP3A family (McArthur et al. 2003; Nelson 2003), this suggests that the evolution of the CYP3A4 enzyme may be a result of selective pressure resulting from exposure to particular xenobiotic compounds. In contrast, our lack of identification of CYP1A enzymes is less troubling. In mammals and fish, CYP1A expression is correlated with environmental exposure to compounds such as polyaromatic hydrocarbons, and they are commonly not constitutively expressed (Ortiz de Montellano 2005; Olsvik et al. 2007; Monostory et al. 2009). Hence, our inability to measure CYP1A1 and CYP1A2 in ‘naı¨ve’ corals likely results from the lack of induction by toxicant exposure in Hawaiian waters, which are understood to be comparatively free of many toxicants found elsewhere. Although not constitutively expressed, it is uncertain whether
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CYP1A enzymes may be induced, or indeed, whether novel CYP1A isoforms or orthologs are present that do not share substrate identity or protein structure with the forms currently described. Our findings are also supported by Firman (1995) who was unsuccessful in detecting CYP1A activity using the ethoxyresorufin substrate in individual or pooled samples from the coral Montastraea faveolata, a complete lack of published reports for CYP1A in corals, and investigation of the cnidarian Nematostella vectensis genome where CYP1-like genes were absent (Goldstone 2008). Evidence of ethoxyresorufin activity in a cnidarian has been presented for the sea anemones Anthopleura elegantissima and Anthopleura xanthogrammica, although the activity levels reported were sparingly low and the authors presenting the data speculated on possible confounding due to the presence of algal symbionts in their samples (Heffernan and Winston 1998). Previous investigations have documented the presence of XMEs, including CYP 1-class enzymes, in the coral holobiont (Downs et al. 2006; Rouge´e et al. 2006). However, the fact that our study yielded no activity or definitive expression results for this class of enzymes in the coral host tissue alone suggests that if present, CYP1A enzymes may reside within the zooxanthellae and might begin to unravel some aspects of the coral/zooxanthellae symbiosis. After exposure to benzo[a]pyrene, Phase I metabolites have been observed in zooxanthellae fractions of the reef corals Favia fragum and Montastrea annularis (Kennedy et al. 1992). Additionally, Rosic et al. (2010) discovered novel CYP450 genes in dinoflagellate symbionts that were not found in the coral host tissue that shares metazoan origins and close sequence relationships with other known CYP450 hemoproteins. Further investigation is needed to elucidate the presence and activity of XMEs in the zooxanthellae, may reveal their role in detoxification within the coral, and further illuminate the symbiotic relationship between the two organisms. The absence of the phase II family of enzymes SULT1, which plays a particular role in xenobiotic and drug metabolism, is curious due to the presence of the regeneration enzyme ASC. However, the cnidarian Nematostella vectensis genome revealed that while SULT genes were present, they related more to membrane-bound SULT enzymes responsible for energy metabolism, rather than cytosolic SULTs known to be involved in detoxification reactions in vertebrates (Goldstone 2008). Moreover, SULT1A isoforms have developed relatively recently in evolutionary terms (Yoshinari et al. 2001; Bradley and Benner 2005). Therefore, further investigation of SULT at the individual isoform level is needed to elucidate sulfotransferase pathways that may exist in corals. Previous evidence for the presence of CYP enzymes in cnidarians (Sole´ and Livingstone 2005), as well as UGT
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and GST, has been presented (Gassman and Kennedy 1992). However, this study is the first to demonstrate both protein presence and activity, and describe changes over time in the lunar reproductive cycle for both Phase I (CYPR, CYP2E1) and Phase II (UGT, GST, b-glucuronidase, and ASC) enzymes. Additionally, while the enzymes CYP2E1, b-glucuronidase, and ASC have been discovered in other marine invertebrates (Leon et al. 1960; Janer et al. 2005), this is the first description in adult corals, in the phylum Cnidaria. Although diagnostic tests are available for humans and other mammals, few have been developed and applied to the majority of the earth’s vast biodiversity. The current study has value toward the development of biomarkers of coral health by establishing the presence and describing the natural changes in levels of the cellular parameters in healthy organisms. This baseline knowledge can now be applied to understand the effect of environmental stressors on these cellular pathways during the coral’s reproductive life cycle, allowing for detection of changes outside of normal ‘healthy’ variation, prior to outright mortality when intervention has the greatest potential for success. Additionally, quantification of proteins that are up-regulated or down-regulated (as opposed to current qualitative, presence/absence approaches) can guide the application of financial, institutional, and human resources for management responses to pollution events by providing numerical measures of the effectiveness and comprehensiveness of mitigation approaches. The deterioration of marine ecosystems worldwide and accompanying economic, ecological, and cultural losses is an essential focus area for researchers. The development of tools that can allow us to understand cause-and-effect relationships between stressors and organismal health can lead to early interventions and prevent loss. Due to the sessile nature of corals, they are a good sentinel species for assessing and monitoring contamination, and as with human health, it is critical to have tools to detect problems before mortality occurs. Acknowledgments This research was supported by grants to Dr. Robert Richmond from The Pew Environmental Group, NOAA CSCOR Grant NA09NOS4780178 and the Hawaii Coral Reef Initiative. We thank the Hawaii Human Organ and Tissue Bank and staff of the Organ Donor Center of Hawaii for providing the human tissues used as positive controls in these experiments and Grant G12 MD007601.
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