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Metabolism of estrogens and androgens by scleractinian corals
Comparative Biochemistry and Physiology Part B 136 (2003) 473–485
Metabolism of estrogens and androgens by scleractinian corals Ann M. Tarranta,*, C.H. Blomquistb,c, P.H. Limab, M.J. Atkinsond, S. Atkinsone a Department of Oceanography, University of Hawaii at Manoa, Honolulu, USA Department of Obstetrics and Gynecology, HealthPartners Regions Hospital, St Paul, USA c Department of Obstetrics and Gynecology, University of Minnesota Medical School, Minneapolis, USA d Hawaii Institute of Marine Biology, University of Hawaii at Manoa, Kaneohe, USA e Alaska SeaLife Center, University of Alaska Fairbanks, Seward, USA b
Received 9 April 2003; received in revised form 2 August 2003; accepted 3 August 2003
Abstract Estrogens and androgens are steroids that act as reproductive hormones in vertebrates. These compounds have also been detected in reef-building corals and other invertebrates, where they are hypothesized to act as bioregulatory molecules. Experiments were conducted using labeled steroid substrates to evaluate metabolism of estrogens and androgens by coral homogenates. GC-MS analysis of 13C-labeled steroids showed that Montipora capitata coral homogenates or fragments could convert estradiol to estrone and testosterone to androstenedione and androstanedione, evidence that M. capitata contains 17b-hydroxysteroid dehydrogenase and 5a-reductase. When homogenates from three coral species and symbiotic dinoflagellates (zooxanthellae) were incubated with tritiated steroid substrates, metabolites separated by thin-layer chromatography confirmed that 17b-hydroxysteroid dehydrogenase activity occurred in all species tested. NADPq was the preferred cofactor in dehydrogenation reactions with coral homogenates. Reduction of estrone and androstenedione occurred at lower rates and aromatization of androgens was not observed. It is unclear whether estrogens detected previously in coral tissues are produced endogenously or sequestered in coral tissue from dietary or environmental sources. Previous studies have demonstrated that corals can take up estrogens from the water column overlying coral reefs. Considered in total, these observations suggest corals could alter the concentration or form of steroids available to reef organisms. 䊚 2003 Elsevier Inc. All rights reserved. Keywords: Androgen; Coral; Dehydrogenase; Estrogen; Invertebrate; Metabolism; Scleractinia; Steroid
1. Introduction Estrogens and androgens are steroids with broad hormonal activity, studied particularly with respect to regulation of vertebrate reproduction. These steroids have been identified in diverse invertebrates, but the physiological roles of estrogens and *Corresponding author. Present address: Biology Department, Woods Hole Oceanographic Institution MS-32, Woods Hole, MA 02543, USA. Tel.: q1-508-289-3398; fax: q1508-457-2134. E-mail address: [email protected] (A.M. Tarrant).
androgens in invertebrates are poorly understood. Invertebrates, including echinoderms (Schoenmakers and Voogt, 1980; Voogt et al., 1986), mollusks (Lupo di Prisco and Dessi’Fulgheri, 1975; but see also Young et al., 1992; Hines et al., 1996), and cnidarians (Gassman, 1992; but see also Slattery et al., 1997), can synthesize androgens, such as testosterone (T) and androstenedione (D4), from progesterone. Evidence for biosynthesis of estrogens by invertebrates is less widespread. Protochordate (Callard et al., 1984), and mollusk (Gottfried et al., 1967; Bose et al., 1997; Matsu-
1096-4959/03/$ - see front matter 䊚 2003 Elsevier Inc. All rights reserved. doi:10.1016/S1096-4959(03)00253-7
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Fig. 1. Metabolism of androgen and estrogen substrates by coral. Solid arrows indicate metabolic transformations that were observed during experimental incubations. Dotted arrows indicate transformations that were not observed during these experiments but occur in other animals (reviewed in Miller, 1988). Numbers represent enzymes or families of enzymes. (1-2) 17b-hydroxysteroid dehydrogenase, showing dehydrogenation in (1) and reduction in (2), (3) 5a-reductase, (4) aromatase.
moto et al., 1997; Le Curieux-Belfond et al., 2001) tissues can synthesize estrogens from androgen precursors. Biosynthesis of estrogens by echinoderms has not been documented despite repeated efforts (Hines et al., 1992; Voogt et al., 1992; Watts et al., 1994). In the few published studies that address metabolism of androgen precursors by cnidarians (Slattery et al., 1997) and crustaceans (Swevers et al., 1991), biosynthesis of estrogens has not been observed, although biological effects of exogenous estrogens have been found in both groups (Ghosh and Ray, 1992, 1993; Tarrant et al., in review). In several studies of estrogen and androgen metabolism by invertebrates, 17b-hydroxysteroid dehydrogenase (17b-HSD), which interconverts estradiol (E2) and estrone (E1), and T and D4 (Fig. 1), was the enzyme with the highest activity (Swevers et al., 1991; Hines et al., 1994; Watts et al., 1994; Matsumoto et al., 1997). Activity of 17b-HSD and other steroid-metabolizing enzymes helps to determine the concentrations of individual steroid hormones in animal tissues. Exposure to environmental chemicals can disrupt steroid
metabolism and cause reproductive abnormalities in invertebrates, although disruption of steroid metabolism has not been shown to cause impaired reproduction (Oberdorster and Cheek, 2001). Scleractinian corals contain a variety of steroids, including estrogens; however, few studies have directly investigated steroid metabolism in corals or other cnidarians. Scleractinian corals can convert progesterone to a variety of steroid products, including T (Gassman, 1992). Alcyonacean octocorals can also produce metabolites of labeled progesterone and D4, indicating the presence of several enzymes, including 5a-reductase, 3bhydroxysteroid dehydrogenase, 17b-HSD and acyl transferase (Slattery et al., 1997). Annual profiles of E1 and E2 concentrations in the scleractinian coral, Montipora capitata, are consistent with metabolism of E2 to E1 by coral (Tarrant et al., 1999). Therefore, we hypothesized that corals would contain 17b-HSD and would be able to metabolize estrogens and androgens. To our knowledge, no previous studies have described metabolism of both androgens and estrogens by scleractinian corals.
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While the endogenous roles of estrogens and androgens in coral tissues remain to be clarified, it has been shown that exogenous estrogens can impair coral growth and reproduction (Tarrant et al., in review). Estrogens are present in the nearshore marine environment (Atkinson and Atkinson, 1992; Atkinson et al., 2003), and corals can take up dilute estrogens at rates approaching a maximum uptake rate, predicted by mass transfer theory (Tarrant et al., 2001). Rates of steroid metabolism in coral tissue may determine the effects of these ‘environmental’ steroids on corals or other reef biota. In this study, we describe the metabolism of androgens and estrogens by scleractinian corals. 2. Materials and methods Two types of experiments were conducted: (1) incubations of coral homogenates or fragments with stable-isotope-labeled steroids and analysis with gas chromatography coupled with mass spectroscopy (GC-MS), and (2) incubations of coral homogenates with radioisotope-labeled steroids and analysis with thin-layer chromatography (TLC). The stable-isotope experiments were conducted with M. capitata (previously M. verrucosa), (Maragos, 1977, 1995), a reef-building coral we have used in previous studies of coral steroids (Tarrant et al., 1999, 2001). These stable-isotope experiments were conducted to provide conclusive structural characterization of products formed during incubations. The radioisotope experiments were conducted using a variety of coral species and were aimed at providing a better understanding of reaction kinetics and the variability in enzyme activity. Details of these experiments follow in subsequent sections. 2.1. Preparation of homogenates Coral fragments (M. capitata, Pocillopora damicornis, Tubastrea coccinea) were collected by a snorkeller from the reef flat adjoining Coconut Island (Kaneohe Bay, Oahu, Hawaii). M. capitata and P. damicornis are common reef-building corals which contain photosynthetic dinoflagellate endosymbionts (zooxanthellae). T. coccinea is a fully heterotrophic coral, which lacks zooxanthellae. Coral fragments were stored in an outdoor tank with continuously flowing unfiltered seawater for up to one week. Zooxanthellae (‘flat plate’ strain,
475
cultured from M. capitata) were provided by Dr R. Kinzie, University of Hawaii at Manoa. Coral fragments were weighed and ground in a chilled mortar and pestle. The resulting slurry was scraped into a 200-ml beaker. One of two buffers (100 ml per 8 g coral wet weight) was added to the ground tissue. Artificial seawater buffered with 20 mM HEPES was used in most stable-isotope experiments (HEPES buffer). A potassium phosphate buffer with 20% glycerol was used to prepare homogenates for radioisotope experiments (GKP buffer). In all experiments, buffer pH was adjusted to 7.2 for aromatization and ketosteroid or 5areduction and to 9.0 for hydroxysteroid dehydrogenation. Tissue was homogenized for three 10-s bursts, with 1 min cooling on ice between bursts (to reduce protein degradation and maintain enzyme activity), using the lowest setting of a stand-mounted Ultra-Turrax T25 homogenizer. The mixture was centrifuged for 5 min at 1380=g. For stable-isotope experiments, the supernatant was used immediately. For radioisotope experiments, the supernatant was packed on ice and shipped to the HealthPartners Regions Hospital (St. Paul, MN; typical shipping time was 48 h). 2.2. Stable-isotope experiments with GC-MS detection All solvents were GC-MS grade and obtained from Fisher Scientific (Houston, TX). Glassware was combusted for 5 h at 450 8C or extracted serially with several rinses of methanol, dichloromethane and hexane. Incubations of M. capitata coral homogenate with steroid precursors were conducted during August–September 2001. For each incubation, 70 ml of homogenate was measured into a 500-ml round-bottomed flask. Because radioisotope experiments indicated higher metabolism of steroid precursors with NADP(H) than with NAD(H) as a cofactor, NADP(H) was used as a cofactor in all GC-MS experiments. To each incubation, 0.5 mM of the appropriate cofactor was added (27 mg NADPq for dehydrogenation reactions or 29 mg NADPH for aromatizationy reduction reactions). The stable-isotope label was added as 0.25–0.5 ml of a 0.5 mg mly1 solution of 13C-labeled steroid (E2-3,4-13C2 or T-3,4-13C2, Cambridge Isotope Laboratories Inc., Andover MA) in methanol. In initial experiments 0.25 ml of the stock solution was added (assay concentrations: 5.8 mM E2, 6.2 mM T); the amount of
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substrate added was doubled in subsequent experiments (assay concentrations 11.6 mM E2, 12.4 mM T) to improve assay precision. The sample was incubated with continuous stirring for 4 h and then extracted for 30 min with 90 ml of chloroform: methanol (2:1). The mixture was transferred quantitatively into a 500-ml separatory funnel and shaken vigorously. Aqueous sodium chloride (100 ml of a 5% solution) was added to increase the ionic strength of the polar phase. The lower organic phase was removed quantitatively into a clean flask and centrifuged for 5 min at 1380=g to further separate the chloroform extract from remaining methanol and a fatty layer present at the solvent interphase. After centrifuging, the lower chloroform fraction was dried for )8 h over sodium sulfate, concentrated to -5 ml using a rotary evaporator and dried further through a 5-ml sodium sulfate column. The crude extract was evaporated to dryness under a gentle stream of prepurified nitrogen, reconstituted in 0.5 ml hexane and applied to a silica gel column that was pre-equilibrated with hexane. The sample was fractionated by increasing the polarity of the mobile phase by adding an increasing proportion of ethyl acetate (steroid metabolites eluted in 50–66% ethyl acetate in hexane) to obtain a purified steroid fraction. An n-alkane (n-C23) was added as an internal standard to allow quantitation of steroids. Trimethylsilyl (TMS) derivatives were synthesized according to the manufacturer’s instructions (BSTFA solution, Alltech, Deerfield, IL). The GC-MS system consisted of a Hewlett-Packard 6890 GC system coupled to Hewlett Packard 5973 mass selective detector. A HP 7683 autoinjector was used to apply 0.5 ml of sample directly onto a J and W DB-5 column (60 m, 0.32-mm internal diameter, 0.25-mm film thickness) through a cooled oncolumn inlet. The initial GC oven temperature was 50 8C (1 min hold) followed by a ramp of 20 8C miny1 to 250 8C (0 min hold), and a 4 8C miny1 ramp to 320 8C (20 min hold). Steroids were identified through the use of authentic standards (Steraloids, Newport, RI) andyor by their mass spectra. During two additional experiments, intact M. capitata coral fragments (8 g) were incubated with 13 C-E2 or 13C-T in 100 ml HEPES buffer for 4 h. At the end of the incubation, fragments were removed from the incubation buffer and homoge-
nized in a fresh 70-ml aliquot of HEPES buffer, extracted and chromatographed as described above. The HEPES buffer used during the incubation was extracted separately. 2.3. Radioisotope experiments using TLC Radiolabeled steroid precursors were as follows: w6,7-3HxE2 (1.5 TBqymmol, 40 Ciymmol), w2,4,6,7-3HxE1 (4.0 TBqymmol, 108 Ciymmol), w1,2,6,7-3HxD4 (4.1 TBqymmol, 110 Ciymmol), Amersham Corp (Arlington Heights, IL); w1,23 x H T (1.9 TBqymmol, 52.5 Ciymmol), w1b3 x H D4 (0.9 TBqymmol, 24.1 Ciymmol) DuPont NEN Products (Boston, MA). Two sets of experiments were conducted: one in June 2001 (summer) and a second in January 2002 (winter). All experiments conducted during a given month used the same tissue extracts. Homogenates were stored at 4 8C and all experiments were completed within three weeks of original collection of tissue fragments. Homogenates were used in incubations, except where cytosol is specified. Cytosol was prepared by centrifuging homogenates at 105 000=g for 60 min and collecting the supernatant. To measure 17b-HSD-like activity, homogenates or cytosol were incubated with cofactor (0.5 mM NAD(H) or NADP(H), as specified) and a tritiated steroid substrate (1.0, 21.0 or 101 mM, as specified). All assays were incubated at room temperature. Reaction mixtures were fractionated into substrate and product by thin-layer chromatography using benzene-acetone (4:1 v:v) as the mobile phase (Blomquist et al., 1994). To test the substrate specificity of the conversion of E2 to E1, the rate of production of radiolabeled E1 from E2 was measured in the presence and absence of a 100fold excess of unlabeled T. E1 was confirmed as product of E2 metabolism through recrystallization. Aromatase was assayed based on the NADPHdependent release of 3H2O from 1b-w3HxD4. Microsomes (105 000=g pellet) were incubated at pH 7.2 in 0.5-ml reaction mixtures prepared by combining 290 ml of 100 mM HEPES, pH 7.2, containing 0.5 mM glucose-6-phosphate, 0.5 unity ml glucose-6-phosphate dehydrogenase and 1.0 mM NADPH with 200 ml of microsomes. The reaction was started by the addition of 10 ml of 10 mM w3HxD4. Reaction was stopped by the addition of 0.5 ml of 20% trichloroacetic acid and 1.0 ml of activated charcoal (5% wyv). After
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centrifugation at 1000=g for 10 min, 1.0 ml of the supernatant was taken for liquid scintillation counting of 3H2O. Microsomes prepared from M. capitata, P. damicornis and T. coccinea were incubated in triplicate for 22.5 and 43.5 h. Human placental microsomes were used as a positive control. 3. Results 3.1. Stable-isotope experiments with GC-MS detection When GKP and HEPES buffers with coral homogenate were extracted as blanks, GKP buffer resulted in a higher baseline. Omitting glycerol from the GKP buffer recipe and solvent–rinsing the potassium phosphate salts reduced the concentration of contaminants. HEPES buffer was used in most incubations to minimize the contaminants co-eluting with steroids and to facilitate comparisons with other studies (i.e. Slattery et al., 1997). Androgens and estrogens were only detected in buffers or coral homogenates when added experimentally; unlabelled (endogenous) androgens or estrogens were not detected. In all buffers, an unknown, non-steroidal (fatty acid) contaminant co-eluted with E1. The concentration of the fatty acid was lower in HEPES buffer than in GKP buffer; this concentration was measured in blanks and was subtracted from peak areas from incubations to determine E1 concentration. Although elution times for E1 and the contaminant overlapped, spectra of E1 could be clearly identified. Steroid substrates and products were identified based on peak elution time and mass spectra (Table 1). Coral homogenate spiked with 13C-T and 13CE2 and immediately extracted showed no evidence of labeled steroid metabolites (Table 2). Similarly, when coral rubble was incubated with E2, labeled steroid metabolites were not detected. However, after a 4-h incubation of 13C-E2 with coral homogenate, approximately 2% of the precursor was converted to E1. Incubations with 13C-T resulted in up to 4% conversion of precursor into D4 and androstanedione (Fig. 2, Table 2). Mass spectra of 13 C-labeled products are shown in Fig. 3. 3.2. Radioisotope experiments using TLC As a first approach to assessing 17b-HSD-like activity, assays were run with E2, E1, T and D4 at
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Table 1 Steroid standards (E1, E2), substrates (13C-E2, 13C-T), and products (13C-E1, 13C-D4, 13C-Androstanedione) detected by GC-MS Compound
Time
MW
TMS
E1 13 C-E1 E2 (mono-silane) 13 C-E2 (mono-silane) 13 C-D4 E2 13 C-E2 13 C-T 13 C-Androstanedione
22.10 22.10 22.32 22.34 22.50 22.61–22.66 22.64 22.98–23 22.76–22.79
342 344 344 346 288 416 418 362 290
1 1 1 1 0 2 2 1 0
Time is peak elution time, MW is molecular mass (measured as the mass to charge ratio), TMS is the number of trimethylsilyl groups attached during derivatization. Note that both mono- and bis-TMS forms of E2 were detected. Unlabeled and 13 C-steroids coeluted; however, endogenous unlabeled estrogens and androgens were not detectable in coral homogenates.
a low concentration (1.0 mM) and a high concentration (21.0 mM E2 or E1, the solubility limit and 101 mM T or D4) of steroid substrate. These conditions have been shown to be optimal for detecting various isoforms of 17b-HSD in human tissues and subcellular fractions (Blomquist, 1995). During the summer experiments, coral homogenates converted E2 to E1, indicating the presence of a 17b-HSD-like enzyme (Table 3). Approximately 10-fold greater conversion occurred when NADPq was used as a cofactor than when NADq was used. When homogenates were incubated for 3 h with 1.0 mM E2 and 0.5 mM NADPq, conversion to E1 ranged from 12% for T. coccinea to 31% for M. capitata. Protein concentrations varied among samples, so the specific activity of conversion (pmol (mg protein)y1 3 hy1) of E2 to E1 was lowest for T. coccinea and highest for P. damicornis. Reduction of E1 to E2 by coral homogenates ranged from below detection to 9.7% conversion. Coral homogenates also converted T to D4 (Table 3). As with the estrogens, NADPq was preferred as a cofactor over NADq. When homogenates were incubated for 3 h with 1.0 mM testosterone and 0.5 mM NADPq, conversion to D4 ranged from 1.7% for P. damicornis to 10% for T. coccinea (Table 3). With 101 mM T, the specific activity increased, but only 0.6–6.7% of the precursor was converted. Reduction of D4 to T by coral homogenates ranged from below detection to 4.7%. Coral microsomes incubated with
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478 Table 2 Conversion of Time (h)
pH
13
C-labeled steroid substrates by M. capitata
Sample
Wet weight (g)
Substrate (mg)
6.5 5.5 5.8 5.4 5.6
110 220 110 220 220
9
220
Recovery E2
E1
84 117 55 197 136
ND ND 0.99 4.8 2.5 0.03 (0.5)
Conversion (%)
Specific activity pmol gy1
pmolymg proteiny1
– – 1.8 2.4 1.8
– – 500 2600 1300
– – 290 1500 760
1.2
870
510
– 10.5 3.1 1.6 3.8
– 7800 200 3200 1200
– 4600 120 1900 740
1.7
1600
970
13
C-Estradiol substrate 0 9 Homogenate 4 9 Rubble 4 9 Homogenate 4 9 Homogenate 4 9 Homogenate* Fragment 4 8.2 (buffer)
13
C-Testosterone substrate 0 9 Homogenate 4 9 Homogenate 4 9 Homogenate 4 7.2 Homogenate 4 7.5 Homogenate Fragment 4 8.2 (buffer)
6.5 5.4 5.7 5.5 5.6
125 252 252 252 252
8.4
252
12 (32) T
D4
A
20 125 13 29 76 0.44 (4.3)
ND ND 0.4 6.1 2.4 ND (0.07)
ND 14.6 ND 0.2 0.7 ND
Sample types: homogenatesM. capitata homogenized in buffer, rubblesM. capitata skeletal fragment homogenized in HEPES buffer, fragmentsM. capitata live coral fragment incubated in artificial seawater with 20 mM HEPES (fragment and buffer incubated together for 4 h with a labeled steroid substrate and extracted separately). One sample homogenized in GKP buffer is marked with an asterisk. Substrate added and steroids recovered are given in milligram; AsAndrostanedione. Percent conversion is given as a percent of labeled steroids recovered. ND indicates no detectable conversion. Specific activity (per 0- or 4-h incubation) is given in pmol (gram wet weight)y1 and pmol (milligram protein)y1. Protein concentration was estimated as 1.7 mg protein per gram wet weight (Tarrant et al., 1999). Activity is normalized to estimated protein concentration to allow comparison with radioisotope results.
1b-w3HxD4 did not produce significant quantities of 3H2O; thus, aromatase activity was not detected in these experiments (data not shown). When extracts were fractionated into cytosol and microsomes by centrifugation at 105 000 g for 60 min, most of the activity with E2 as a substrate was recovered in the cytosol, NADPq was the preferred cofactor, and T in 100-fold excess had only a slight inhibitory effect (Fig. 4). To confirm the requirement for a pyridine nucleotide cofactor, samples of M. capitata cytosol were dialyzed overnight against a 100-fold excess of 20% GKP buffer and then assayed with and without NADPq. No activity was observed in the absence of NADPq. Dialysis with a 3500-mw cutoff membrane had little effect on activity in the presence of NADPq; thus, the removal of small molecules had little effect on the activity, which is most likely mediated through an enzyme (Fig. 5). Because T. coccinea homogenates showed the greatest capacity for androgen metabolism in preliminary experiments (Table 3), T. coccinea homogenate and cytosol were incubated with low (1.0
mM) and high (101 mM) concentrations of T or D4 for up to 21 h. Activity with T and D4 was also recovered largely in the cytosolic fraction. With both a low (1.0 mM) and a high (101 mM) concentration of T or D4, the reaction was essentially complete by 5 h with no further product formation during an overnight incubation (Fig. 6). Winter experiments quantified conversion of E2 to E1 in the presence of NADPq by extracts of M. capitata, T. coccinea and the zooxanthellae culture (Fig. 7). In marked contrast with the results of the summer experiments, activity in T. coccinea cytosol greatly exceeded that in the M. capitata sample. Activity in both coral species, particularly M. capitata, was lower in winter than in summer. The highest specific activity was found in the zooxanthellae. The ratio of specific activity measured with NADPq as cofactor to the specific activity with NADq as cofactor (NADPq yNADq ratio) can be used to identify different enzymes that act on the same steroid substrate (Blomquist, 1995). In converting E2 to E1, the T. coccinea tissue homogenate
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Fig. 2. Chromatographs of extracted M. capitata homogenates. Labels represent elution times for 13C-labeled substrates and products. (a) E2 and T added and immediately extracted, 22.64s13C E2, 23s13C T (b) 4-h incubation of M. capitata rubble homogenate with E2, 22.33s13C E2 (mono-silane), 22.65s13C E2 (c) 4-h incubation of M. capitata homogenate with E2, showing E1 as a product, 22.1s13C E1, 22.3s13C E2 (mono-silane), 22.67s13C E2 (d) 4-h incubation of M. capitata homogenate with T, showing androstanedione as a product, 22.77s13C androstanedione, 23.02s13C T.
had similar NADPq yNADq ratios in winter and summer (Table 4). The corresponding NADPq y NADq ratios for M. capitata ranged from approximately 10 in the first summer experiment to -2 in the winter. The ratio for the zooxanthellae sonicate was 1.1. 4. Discussion These experiments clearly demonstrate that corals (zooxanthellate and azooxanthellate) and their dinoflagellate symbionts contain a functional 17bHSD and a 5a-reductase. Reduction (E1™E2 and D4™T) occurred at a lower rate than dehydrogenation (E2™E1 and T™D4). NADPq was the preferred cofactor in dehydrogenation reactions and dehydrogenation of E2 was slightly inhibited by an excess of T (Fig. 4). These results are consistent with metabolism by an enzyme with
both a steroid recognition site and a coenzyme binding sites, but additional experiments will be needed to characterize the substrate specificity. Aromatization of D4 was not observed; however, we cannot rule out the possibility that aromatase may be present in coral tissue at low abundance or may be restricted temporally or spatially within coral polyps. Aromatase activity in mammals outside of the female reproductive organs is generally very low, but localized ‘paracrine’ synthesis of estrogens is biologically important and can result in locally high concentrations of E2 and E1 (Dikkeschei et al., 1996; Simpson and Davis, 2001; Simpson et al., 2002). Use of a cnidarian with larger gonads, such as an anemone, might allow assay of aromatase activity specifically in reproductive tissues. Immunohistochemistry could also be used in the future to determine whether aromatase is present in coral tissue. Polar and non-
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Fig. 3. Mass spectra of estrogens and androgens synthesized during incubations with coral tissue homogenates. Black circles on structures denote 13C atoms. (a) silanized 13C-E1 (b) 13C-D4 (c) 13C-androstanedione.
polar steroid conjugates were not assayed in this study; however, other invertebrates can produce polar and non-polar conjugates of estrogens and androgens (Voogt and Van Rheenen, 1986; Voogt et al., 1987; Fairs et al., 1989; Swevers et al., 1991; Watts et al., 1994). Steroid metabolites produced by coral homogenates in this study are qualitatively similar to those produced by Antarctic soft corals (Slattery et al., 1997). Alcyonium paessleri and Clavularia frankliniana converted D4 into T, epiandrosterone, 5aandrostanedione and an unidentified esterified androgen. T was the major product, with 28% (C. frankliniana) total precursor conversion after 12 h, giving a specific activity of 1700 pmol (g wet weight)y1 hy1. Protein in C. frankliniana is 24%
of dry weight (Slattery and McClintock, 1995); if dry weight is 10–50% of wet weight, specific activity would be 14–71 pmol (mg protein)y1 hy1. In contrast, we observed little reduction of D4, but up to 10% conversion of T to D4 with a specific activity of 9.96 pmol (mg protein)y1 3 hy1 or and average rate of 3.3 pmol (mg protein)y1 hy1. Thus, androgen metabolism in Antarctic alcyonaceans occurred at a higher rate than in Hawaiian scleractinians. Metabolism of estrogens by soft corals has not been described, but in scleractinians we observed specific activities )3000 pmol (mg protein)y1 3 hy1 (Table 3, E2™E1). In the present study, F30% of the substrate was metabolized in any reaction and incubations beyond ;3 h did not result in addi-
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Table 3 Net conversion of tritiated steroid precursors by coral homogenates in June Percent conversion (net)
Specific Activity
Cofactor
NAD(H)
NADP(H)
NAD(H)
NADP(H)
E2 to E1 M. capitata P. damicornis T. coccinea
1.0 mM E2 3.5 8.0 4.7
21 mM E2 2.6 3.7 3.8
1.0 mM E2 31.3 16.4 12.4
21 mM E2 22.6 11.3 11.0
1.0 mM E2 17.7 152 8.0
21 mM E2 306 1420 129
1 mM E2 182 313 21.0
21 mM E2 2620 4540 389
E1 to E2 M. capitata P. damicornis T. coccinea
1.0 mM E1 ND 1.8 ND
21 mM E1 ND 2.9 4.4
1.0 mM E1 0.5 9.7 ND
21 mM E1 4.4 9.6 6.2
1.0 mM E1 ND 34.8 ND
21 mM E1 ND 1160 155
1 mM E1 2.9 186 ND
21 mM E1 539 3850 221
T to D4 M. capitata P. damicornis T. coccinea
1.0 mM T 2.8 2.5 4.9
101 mM T 1.6 ND 2.0
1.0 mM T 4.0 1.7 10.0
101 mM T 2.5 0.6 6.7
1.0 mM T 15.9 48.3 8.4
101 mM T 930 ND 330
1 mM T 23.1 32.7 17.1
101 mM T 1430 1090 1130
D4 to T M. capitata P. damicornis T. coccinea
1.0 mM D4 1.0 1.3 2.3
101 mM D4 0.8 ND 1.4
1.0 mM D4 2.5 2.0 4.7
101 mM D4 1.1 ND 2.8
1.0 mM D4 5.7 24.9 3.9
101 mM D4 420 18.9 235
1 mM D4 14.7 39.0 7.8
101 mM D4 660 ND 474
Coral tissue homogenates (1000=g supernatants from M. capitata, P. damicornis and T. coccinea) were incubated for 3 h with 1.0– 101 mM of a steroid substrate and 0.5 mM of the cofactor indicated. Percent conversion in blank incubations was less than 2% (except E1 to E2, 17%) and has been subtracted. Specific activity given in pmol (milligram protein)y1 3 hy1. ND, activity not detected.
Fig. 4. Conversion of E2 to E1 by M. capitata homogenate (1000=g supernatant) and cytosol (105 000=g supernatant) with NADq or NADPq as cofactor in the absence or presence of T. Assays were run at pH 9.0 with 1.0 mM 3H-E2, 0.5 mM NADq or NADPq and, where noted, 100 mM T. The values are the mean "S.D. of duplicate assays run at room temperature for 330 min.
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Fig. 5. Conversion of E2 to E1 by M. capitata cytosol (solid circles) and M. capitata cytosol dialyzed overnight against a 100-fold excess of 20% GKP buffer (open circles). A 3500mw cut-off membrane was used. Activity was not detected in M. capitata cytosol in the absence of NADPq (triangles). Assays were run at pH 9.0 with 1.0 mM 3H-E2, and 0.5 mM NADPq when added.
tional metabolism. At present, it is not known whether consumption of some unknown cofactor or other component is limiting further conversion. 17b-HSD activity in alcyonacean and scleractinian corals is low relative to activity reported in specific mammalian tissues or cell types we.g. cells derived from a carcinoma of the vulva converted T to D4 with specific activity ;8 nmol (mg protein)y1 30 miny1 (Blomquist et al., 1997)x. It is important to recognize that the coral metabolic rates represent an average rate for an entire animal; rates in some cells or tissues may be considerably higher. Incubations of coral and zooxanthellae with E2 indicate seasonal and interspecies variation in specific activity (Table 3, Fig. 7) and NADPyNAD ratios (Table 4). However, each species during a given month (January or June) was represented by a homogenate from one fragment, so a more complete examination of seasonality is needed. Future experiments should also test the effects of environmental factors (e.g. photoperiod, temperature and exposure to pollutants) on steroid metabolism. Variation in steroid metabolism may cause observed seasonal variation in estrogen concentra-
Fig. 6. Conversion of T to D4 and D4 to T by T. coccinea cytosol. Duplicate assays (mean with the difference between replicates shown) containing 1.0 mM or 101 mM T or D4 and 0.5 mM NADPq or NADPH at pH 9.0 (T to D4) or pH 7.2 (D4 to T) were incubated at room temperature for 5 or 21 h.
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in M. capitata were 20–120 ng (g ash-free dry weight)y1 or ;0.4–3 ng (g wet weight)y1; E2 concentrations were approximately half the E1 concentrations. Thus, to detect endogenous estrogens in coral tissue using GC-MS, it would be necessary to improve the sensitivity by a factor of 10–100 to compare with the sensitivity of radioimmunoassay. This could be accomplished by extracting more tissue or by concentrating the extract on the GC column. These approaches would also increase the concentrations of contaminants, so additional sample preparation would be needed. Additional metabolic assays, protein purification and cloning will help to identify the coral enzymes and their natural substrates. Similarities in substrate specificity are not always related to sequence similarity (Baker, 1994). 17b-HSD sequences have been identified in bacteria and fungi; notably a fungal 17b-HSD catalyzes metabolism of both steroidal and non-steroidal (quinonal) substrates (Lanisnik Rizner et al., 1999). 17b-HSDs are homologous to other oxidoreductases, including bacterial 3a20b- and 3b-HSD and the NodG protein, which is found in the nitrogen-fixing bacterium, Rhizobioum meliloti (Baker, 1994). We now know that estrogens are present in the water column (Atkinson et al., 2003), may be taken up (Tarrant et al., 2001) and metabolized by scleractinian corals (present study) and can affect coral physiology (Tarrant, 2002; Tarrant et al., in review). Environmental pollutants may alter rates of steroid metabolism and consequently, steroid concentrations in tissue and affect physiological
Fig. 7. Time-course of the conversion of E2 to E1 by 1000=g supernatants of M. capitata (solid circles), T. coccinea (open circles), and a sonicate of zooxanthellae from M. capitata (triangles). Sonication was for two 1.0-min intervals in a bathtype sonicator (Heat Systems, Inc.). Assays were run at pH 9.0 with 1.0 mM 3H-E2, and 0.5 mM NADPq.
tions (Tarrant et al., 1999). Variation in the NADPq yNADq ratio is consistent with more than one enzyme catalyzing steroid dehydrogenation (Blomquist, 1995). Changes in steroid metabolism may also result from a variable contribution of algal and animal components to the overall rate. Endogenous estrogens and androgens were not detected by GC-MS in coral homogenates, with a detection limit for steroid metabolites of 40 ng (g wet weight)y1. In contrast, Tarrant et al. (1999) used radioimmunoassay to quantify endogenous E1-immunoreactivity and E2-immunoreactivity. Based on immunoreactivity, the E1 concentrations Table 4 NADPqyNADq ratios for January and two June experiments Experiment
Sample
1.0 mM
21.0 mM
Notes
June Expt. 1
M. capitata P. damicornis T. coccinea M. capitata (homogenate) M. capitata (cytosol) M. capitata T. coccinea Zooxanthellae
10.3 2.1 2.7
8.6 3.2 3
(3 h assay)
June Expt. 2
January Expt.
(5.5 h assay) 5.6 3.8 1.6 3.8 1.1
(2 h assay)
Ratio of specific activity of conversion of E2 to E1 with NADPq as a cofactor to specific activity with NADq as a cofactor. Assays were run at pH 9.0 with 1.0 or 21.0 mM E2 and 0.5 mM NADPq or NADq. Tissue homogenates (1000=g supernatants) were used for M. capitata, P. damicornis, and T. coccinea coral tissue, except in the 2nd June experiment, where both homogenates and cytosol (105 000=g supernatant) were used. Zooxanthellae were sonicated, as described in the text.
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processes such as reproduction. Also, since corals can take up and metabolize steroids, they can alter the concentration or form of steroids available to other reef organisms. Acknowledgments We thank Dr R. Bidigare and B. Benitez-Nelson for assistance with GC-MS analysis. Dr R. Kinzie provided zooxanthellae samples. Support for this project was provided by the EPA STAR fellowship program, the University of Hawaii Sea Grant College Program and HealthPartners Research Foundation. The views expressed herein are solely those of the authors. References Atkinson, S., Atkinson, M., Tarrant, A., 2003. Estrogens from sewage in the coastal marine environment. Environ. Health Perspect. 111, 531–535. Atkinson, S., Atkinson, M.J., 1992. Detection of estradiol-17b during a mass coral spawn. Coral Reefs 11, 33–35. Baker, M., 1994. Sequence analysis of steroid- and prostaglandin-metabolizing enzymes: application to understanding catalysis. Steroids 59, 248–258. Blomquist, C., Leung, B., Beaudoin, C., Tremblay, Y., 1997. Intracellular regulation of 17b-hydroxysteroid dehydrogenase type 2 catalytic activity in A431 cells. J. Endocrinol. 153, 453–464. Blomquist, C., 1995. Kinetic analysis of enzymatic activities: prediction of multiple forms of 17b-hydroxysteroid dehydrogenase. J. Steroid Biochem. Mol. Biol. 55, 515–524. Blomquist, C., Bealka, D., Hensleigh, H., Tagatz, G., 1994. A comparison of 17b-hydroxysteroid oxidoreductase type 1 and type 2 activity of cytosol and microsomes from human term placenta, ovarian stroma and granulosa-luteal cells. J. Steroid Biochem. Mol. Biol. 49, 183–189. Bose, R., Majumdar, C., Bhattacharya, S., 1997. Steroids in Achatina fulica (Bowdich): steroid profile in haemolymph and in vitro release of steroids from endogenous precursors by ovotestis and albumen gland. Comp. Biochem. Physiol. C 116, 179–182. Callard, G., Pudney, J., Kendall, S., Reinboth, R., 1984. In vitro conversion of androgens to estrogen in amphioxus gonadal tissues. Gen. Comp. Endocrinol. 56, 53–58. Dikkeschei, L., Wolthers, B., Bos-Zuur, I., Brutel de la Riviere, G., Nager, G., van der Kolk, D., et al., 1996. Optimization of a classical aromatase activity assay and application in normal, adenomatous and malignant breast parenchyma. J. Steroid Biochem. Mol. Biol. 59, 305–313. Fairs, N., Evershed, R., Quinlan, P., Goad, L., 1989. Detection of unconjugated and conjugated steroids in the ovary, eggs and haemolymph of the decapod crustacean Nephrops norvegicus. Gen. Comp. Endocrinol. 4, 199–208. Gassman N.J., 1992. Aspects of the steroid biochemistry of the scleractinian coral, Favia fragum. Ph.D. Dissertation,
Department of Marine Biology and Fisheries, University of Miami, 158 pp. Ghosh, D., Ray, A.K., 1992. Evidence for physiological responses to estrogen in freshwater prawn, Macrobrachium rosenbergii. J. Inland Fish. Soc. India 24, 15–21. Ghosh, D., Ray, A.K., 1993. Subcellular action of estradiol17b in a freshwater prawn, Macrobrachium rosenbergii. Gen. Comp. Endocrinol. 90, 273–281. Gottfried, H., Dorfman, R., Wall, P., 1967. Production of oestrogens by an accessory reproductive tissue of the slug Arion ater rufus (Linn.). Nature 215, 409–410. Hines, G.A., Bryan, P.J., Wasson, K.M., McClintock, J.B., Watts, S.A., 1996. Sex steroid metabolism in the antarctic pteropod Clione antarctica (Mollusca: Gastropoda). Invert. Biol. 115, 113–119. Hines, G.A., Watts, S.A., McClintock, J.B., 1994. Biosynthesis of estrogen derivatives in the echinoid Lytechinus variegatus Lamarck. In: David, B., Guille, A., Feral, J.P., Roux, M. (Eds.), Echinoderms Through Time. Balkema, Rotterdam, pp. 711–716. Hines, G.A., Watts, S.A., Walker, C.W., Voogt, P.A., 1992. Androgen metabolism in somatic and germinal tissues of the sea star Asterias vulgaris. Comp. Biochem. Physiol. B 102, 521–526. Lanisnik Rizner, T., Moeller, G., Thole, H., Zakelj-Mavrik, M., Adamski, J., 1999. A novel 17b-hydroxysteroid dehydrogenase in the fungus Cochiobolus lunatus: new insights into the evolution of steroid hormone signaling. Biochem. J. 337, 425–431. Le Curieux-Belfond, O., Moslemi, S., Mathieu, M., Seralini, G., 2001. Androgen metabolism in oyster Crassostrea gigas: evidence for 17b-HSD activities and characterization of an aromatase-like activity inhibited by pharmacological compounds and a marine pollutant. J. Steroid Biochem. Mol. Biol. 78, 359–366. Lupo di Prisco, C., Dessi’Fulgheri, F., 1975. Alternative pathways of steroid biosynthesis in gonads and hepatopancreas of Aplysia depilans. Comp. Biochem. Physiol. B 50, 191–195. Maragos, J., 1977. Order scleractinia: stony corals. In: Devaney, D.M., Eldredge, L.G. (Eds.), Reef and Shore Fauna of Hawaii, Section 1: Protozoa Through Ctenophora. Bishop Museum Press, Honolulu, Hawaii, pp. 158–241. Maragos J., 1995. Revised checklist of extant shallow-water stony coral species from Hawaii (Cnidaria: Anthozoa: Scleractinia). Bishop Museum Occasional Papers, 42, pp. 54– 55. Matsumoto, T., Osada, M., Osawa, Y., Mori, K., 1997. Gonadal estrogen profile and immunohistochemical localization of steroidogenic enzymes in the oyster and scallop during sexual maturation. Comp. Biochem. Physiol. B 118, 811–817. Miller, W., 1988. Molecular biology of steroid hormone synthesis. Endocr. Rev. 9, 295–318. Oberdorster, E., Cheek, A., 2001. Gender benders at the beach: endocrine disruption in marine and estuarine organisms. Environ. Toxicol. Chem. 20, 23–36. Schoenmakers, H., Voogt, P., 1980. In vitro biosynthesis of steroids from progesterone by the ovaries and pyloric caeca
A.M. Tarrant et al. / Comparative Biochemistry and Physiology Part B 136 (2003) 473–485 of the starfish Asterias rubens. Gen. Comp. Endocrinol. 41, 408–416. Simpson, E., Clyne, C., Rubin, G., Boon, W., Robertson, K., Britt, K., et al., 2002. Aromatase—a brief overview. Ann. Rev. Physiol. 64, 461–470. Simpson, E., Davis, S., 2001. Minireview: aromatase and the regulation of estrogen biosynthesis—some new perspectives. Endocrinology 142, 4589–4594. Slattery, M., Hines, G.A., Watts, S.A., 1997. Steroid metabolism in Antarctic soft corals. Polar Biol. 18, 76–82. Slattery, M., McClintock, J., 1995. Population structure and feeding deterrence in three shallow-water antarctic soft corals. Mar. Biol. 122, 461–470. Swevers, L., Lambert, J.G.D., De Loof, A., 1991. Metabolism of vertebrate-type steroids by tissues of three crustacean species. Comp. Biochem. Physiol. B 99, 35–41. Tarrant A., 2002. Estrogen Action in Scleractinian Corals: Sources, Metabolism and Physiological Effects. Ph.D. Dissertation, Department of Oceanography, University of Hawaii at Manoa, 186 pp. Tarrant A., Atkinson M., Atkinson S., Effects of steroidal estrogens on coral growth and reproduction. Mar. Ecol. Prog. Ser., In Review. Tarrant, A.M., Atkinson, M.J., Atkinson, S., 2001. Uptake of estrone by a coral reef community. Mar. Biol. 139, 321–325. Tarrant, A.M., Atkinson, S., Atkinson, M.J., 1999. Estrone and estradiol-17b concentration in tissue of the scleractinian
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coral, Montipora verrucosa. Comp. Biochem. Physiol. A 122, 85–92. Voogt, P., Lambert, J., Granneman, J., Jansen, M., 1992. Confirmation of the presence of oestradiol-17b in the sea star Asterias rubens by GC-MS. Comp. Biochem. Physiol. B 101, 13–16. Voogt, P.A., Den Besten, P.J., Kusters, G.C.M., Messing, M.W.J., 1987. Effects of cadmium and zinc on steroid metabolism and steroid level in the sea star Asterias rubens L. Comp. Biochem. Physiol. C 86, 83–89. Voogt, P.A., Van Rheenen, W.A., 1986. Androstenedione metabolism in the sea star Asterias rubens L. studies in homogenates and intact tissue: biosynthesis of the novel steroid fatty-acyl testosterone. Comp. Biochem. Physiol. B 85, 497–501. Voogt, P.A., Van Rheenen, W.A., Lambert, J.G.D., De Groot, B.F., Mollema, C., 1986. Effects of different experimental conditions on progesterone metabolism in the sea star Asterias rubens L. Comp. Biochem. Physiol. B 84, 397–402. Watts, S., Hines, G., Byrum, C., McClintock, J., Marion, K., 1994. Tissue- and species-specific variations in androgen metabolism. In: David, B., Guille, A., Feral, J.P., Roux, M. (Eds.), Echinoderms Through Time. Balkema, Rotterdam, pp. 155–161. Young, N., Quinlan, P., Goad, L., 1992. Progesterone metabolism in vitro in the decapod crustacean, Penaeus monodon. Gen. Comp. Endocrinol. 87, 300–311.
Metabolism of estrogens and androgens by scleractinian corals Ann M. Tarranta,*, C.H. Blomquistb,c, P.H. Limab, M.J. Atkinsond, S. Atkinsone a Department of Oceanography, University of Hawaii at Manoa, Honolulu, USA Department of Obstetrics and Gynecology, HealthPartners Regions Hospital, St Paul, USA c Department of Obstetrics and Gynecology, University of Minnesota Medical School, Minneapolis, USA d Hawaii Institute of Marine Biology, University of Hawaii at Manoa, Kaneohe, USA e Alaska SeaLife Center, University of Alaska Fairbanks, Seward, USA b
Received 9 April 2003; received in revised form 2 August 2003; accepted 3 August 2003
Abstract Estrogens and androgens are steroids that act as reproductive hormones in vertebrates. These compounds have also been detected in reef-building corals and other invertebrates, where they are hypothesized to act as bioregulatory molecules. Experiments were conducted using labeled steroid substrates to evaluate metabolism of estrogens and androgens by coral homogenates. GC-MS analysis of 13C-labeled steroids showed that Montipora capitata coral homogenates or fragments could convert estradiol to estrone and testosterone to androstenedione and androstanedione, evidence that M. capitata contains 17b-hydroxysteroid dehydrogenase and 5a-reductase. When homogenates from three coral species and symbiotic dinoflagellates (zooxanthellae) were incubated with tritiated steroid substrates, metabolites separated by thin-layer chromatography confirmed that 17b-hydroxysteroid dehydrogenase activity occurred in all species tested. NADPq was the preferred cofactor in dehydrogenation reactions with coral homogenates. Reduction of estrone and androstenedione occurred at lower rates and aromatization of androgens was not observed. It is unclear whether estrogens detected previously in coral tissues are produced endogenously or sequestered in coral tissue from dietary or environmental sources. Previous studies have demonstrated that corals can take up estrogens from the water column overlying coral reefs. Considered in total, these observations suggest corals could alter the concentration or form of steroids available to reef organisms. 䊚 2003 Elsevier Inc. All rights reserved. Keywords: Androgen; Coral; Dehydrogenase; Estrogen; Invertebrate; Metabolism; Scleractinia; Steroid
1. Introduction Estrogens and androgens are steroids with broad hormonal activity, studied particularly with respect to regulation of vertebrate reproduction. These steroids have been identified in diverse invertebrates, but the physiological roles of estrogens and *Corresponding author. Present address: Biology Department, Woods Hole Oceanographic Institution MS-32, Woods Hole, MA 02543, USA. Tel.: q1-508-289-3398; fax: q1508-457-2134. E-mail address: [email protected] (A.M. Tarrant).
androgens in invertebrates are poorly understood. Invertebrates, including echinoderms (Schoenmakers and Voogt, 1980; Voogt et al., 1986), mollusks (Lupo di Prisco and Dessi’Fulgheri, 1975; but see also Young et al., 1992; Hines et al., 1996), and cnidarians (Gassman, 1992; but see also Slattery et al., 1997), can synthesize androgens, such as testosterone (T) and androstenedione (D4), from progesterone. Evidence for biosynthesis of estrogens by invertebrates is less widespread. Protochordate (Callard et al., 1984), and mollusk (Gottfried et al., 1967; Bose et al., 1997; Matsu-
1096-4959/03/$ - see front matter 䊚 2003 Elsevier Inc. All rights reserved. doi:10.1016/S1096-4959(03)00253-7
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Fig. 1. Metabolism of androgen and estrogen substrates by coral. Solid arrows indicate metabolic transformations that were observed during experimental incubations. Dotted arrows indicate transformations that were not observed during these experiments but occur in other animals (reviewed in Miller, 1988). Numbers represent enzymes or families of enzymes. (1-2) 17b-hydroxysteroid dehydrogenase, showing dehydrogenation in (1) and reduction in (2), (3) 5a-reductase, (4) aromatase.
moto et al., 1997; Le Curieux-Belfond et al., 2001) tissues can synthesize estrogens from androgen precursors. Biosynthesis of estrogens by echinoderms has not been documented despite repeated efforts (Hines et al., 1992; Voogt et al., 1992; Watts et al., 1994). In the few published studies that address metabolism of androgen precursors by cnidarians (Slattery et al., 1997) and crustaceans (Swevers et al., 1991), biosynthesis of estrogens has not been observed, although biological effects of exogenous estrogens have been found in both groups (Ghosh and Ray, 1992, 1993; Tarrant et al., in review). In several studies of estrogen and androgen metabolism by invertebrates, 17b-hydroxysteroid dehydrogenase (17b-HSD), which interconverts estradiol (E2) and estrone (E1), and T and D4 (Fig. 1), was the enzyme with the highest activity (Swevers et al., 1991; Hines et al., 1994; Watts et al., 1994; Matsumoto et al., 1997). Activity of 17b-HSD and other steroid-metabolizing enzymes helps to determine the concentrations of individual steroid hormones in animal tissues. Exposure to environmental chemicals can disrupt steroid
metabolism and cause reproductive abnormalities in invertebrates, although disruption of steroid metabolism has not been shown to cause impaired reproduction (Oberdorster and Cheek, 2001). Scleractinian corals contain a variety of steroids, including estrogens; however, few studies have directly investigated steroid metabolism in corals or other cnidarians. Scleractinian corals can convert progesterone to a variety of steroid products, including T (Gassman, 1992). Alcyonacean octocorals can also produce metabolites of labeled progesterone and D4, indicating the presence of several enzymes, including 5a-reductase, 3bhydroxysteroid dehydrogenase, 17b-HSD and acyl transferase (Slattery et al., 1997). Annual profiles of E1 and E2 concentrations in the scleractinian coral, Montipora capitata, are consistent with metabolism of E2 to E1 by coral (Tarrant et al., 1999). Therefore, we hypothesized that corals would contain 17b-HSD and would be able to metabolize estrogens and androgens. To our knowledge, no previous studies have described metabolism of both androgens and estrogens by scleractinian corals.
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While the endogenous roles of estrogens and androgens in coral tissues remain to be clarified, it has been shown that exogenous estrogens can impair coral growth and reproduction (Tarrant et al., in review). Estrogens are present in the nearshore marine environment (Atkinson and Atkinson, 1992; Atkinson et al., 2003), and corals can take up dilute estrogens at rates approaching a maximum uptake rate, predicted by mass transfer theory (Tarrant et al., 2001). Rates of steroid metabolism in coral tissue may determine the effects of these ‘environmental’ steroids on corals or other reef biota. In this study, we describe the metabolism of androgens and estrogens by scleractinian corals. 2. Materials and methods Two types of experiments were conducted: (1) incubations of coral homogenates or fragments with stable-isotope-labeled steroids and analysis with gas chromatography coupled with mass spectroscopy (GC-MS), and (2) incubations of coral homogenates with radioisotope-labeled steroids and analysis with thin-layer chromatography (TLC). The stable-isotope experiments were conducted with M. capitata (previously M. verrucosa), (Maragos, 1977, 1995), a reef-building coral we have used in previous studies of coral steroids (Tarrant et al., 1999, 2001). These stable-isotope experiments were conducted to provide conclusive structural characterization of products formed during incubations. The radioisotope experiments were conducted using a variety of coral species and were aimed at providing a better understanding of reaction kinetics and the variability in enzyme activity. Details of these experiments follow in subsequent sections. 2.1. Preparation of homogenates Coral fragments (M. capitata, Pocillopora damicornis, Tubastrea coccinea) were collected by a snorkeller from the reef flat adjoining Coconut Island (Kaneohe Bay, Oahu, Hawaii). M. capitata and P. damicornis are common reef-building corals which contain photosynthetic dinoflagellate endosymbionts (zooxanthellae). T. coccinea is a fully heterotrophic coral, which lacks zooxanthellae. Coral fragments were stored in an outdoor tank with continuously flowing unfiltered seawater for up to one week. Zooxanthellae (‘flat plate’ strain,
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cultured from M. capitata) were provided by Dr R. Kinzie, University of Hawaii at Manoa. Coral fragments were weighed and ground in a chilled mortar and pestle. The resulting slurry was scraped into a 200-ml beaker. One of two buffers (100 ml per 8 g coral wet weight) was added to the ground tissue. Artificial seawater buffered with 20 mM HEPES was used in most stable-isotope experiments (HEPES buffer). A potassium phosphate buffer with 20% glycerol was used to prepare homogenates for radioisotope experiments (GKP buffer). In all experiments, buffer pH was adjusted to 7.2 for aromatization and ketosteroid or 5areduction and to 9.0 for hydroxysteroid dehydrogenation. Tissue was homogenized for three 10-s bursts, with 1 min cooling on ice between bursts (to reduce protein degradation and maintain enzyme activity), using the lowest setting of a stand-mounted Ultra-Turrax T25 homogenizer. The mixture was centrifuged for 5 min at 1380=g. For stable-isotope experiments, the supernatant was used immediately. For radioisotope experiments, the supernatant was packed on ice and shipped to the HealthPartners Regions Hospital (St. Paul, MN; typical shipping time was 48 h). 2.2. Stable-isotope experiments with GC-MS detection All solvents were GC-MS grade and obtained from Fisher Scientific (Houston, TX). Glassware was combusted for 5 h at 450 8C or extracted serially with several rinses of methanol, dichloromethane and hexane. Incubations of M. capitata coral homogenate with steroid precursors were conducted during August–September 2001. For each incubation, 70 ml of homogenate was measured into a 500-ml round-bottomed flask. Because radioisotope experiments indicated higher metabolism of steroid precursors with NADP(H) than with NAD(H) as a cofactor, NADP(H) was used as a cofactor in all GC-MS experiments. To each incubation, 0.5 mM of the appropriate cofactor was added (27 mg NADPq for dehydrogenation reactions or 29 mg NADPH for aromatizationy reduction reactions). The stable-isotope label was added as 0.25–0.5 ml of a 0.5 mg mly1 solution of 13C-labeled steroid (E2-3,4-13C2 or T-3,4-13C2, Cambridge Isotope Laboratories Inc., Andover MA) in methanol. In initial experiments 0.25 ml of the stock solution was added (assay concentrations: 5.8 mM E2, 6.2 mM T); the amount of
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substrate added was doubled in subsequent experiments (assay concentrations 11.6 mM E2, 12.4 mM T) to improve assay precision. The sample was incubated with continuous stirring for 4 h and then extracted for 30 min with 90 ml of chloroform: methanol (2:1). The mixture was transferred quantitatively into a 500-ml separatory funnel and shaken vigorously. Aqueous sodium chloride (100 ml of a 5% solution) was added to increase the ionic strength of the polar phase. The lower organic phase was removed quantitatively into a clean flask and centrifuged for 5 min at 1380=g to further separate the chloroform extract from remaining methanol and a fatty layer present at the solvent interphase. After centrifuging, the lower chloroform fraction was dried for )8 h over sodium sulfate, concentrated to -5 ml using a rotary evaporator and dried further through a 5-ml sodium sulfate column. The crude extract was evaporated to dryness under a gentle stream of prepurified nitrogen, reconstituted in 0.5 ml hexane and applied to a silica gel column that was pre-equilibrated with hexane. The sample was fractionated by increasing the polarity of the mobile phase by adding an increasing proportion of ethyl acetate (steroid metabolites eluted in 50–66% ethyl acetate in hexane) to obtain a purified steroid fraction. An n-alkane (n-C23) was added as an internal standard to allow quantitation of steroids. Trimethylsilyl (TMS) derivatives were synthesized according to the manufacturer’s instructions (BSTFA solution, Alltech, Deerfield, IL). The GC-MS system consisted of a Hewlett-Packard 6890 GC system coupled to Hewlett Packard 5973 mass selective detector. A HP 7683 autoinjector was used to apply 0.5 ml of sample directly onto a J and W DB-5 column (60 m, 0.32-mm internal diameter, 0.25-mm film thickness) through a cooled oncolumn inlet. The initial GC oven temperature was 50 8C (1 min hold) followed by a ramp of 20 8C miny1 to 250 8C (0 min hold), and a 4 8C miny1 ramp to 320 8C (20 min hold). Steroids were identified through the use of authentic standards (Steraloids, Newport, RI) andyor by their mass spectra. During two additional experiments, intact M. capitata coral fragments (8 g) were incubated with 13 C-E2 or 13C-T in 100 ml HEPES buffer for 4 h. At the end of the incubation, fragments were removed from the incubation buffer and homoge-
nized in a fresh 70-ml aliquot of HEPES buffer, extracted and chromatographed as described above. The HEPES buffer used during the incubation was extracted separately. 2.3. Radioisotope experiments using TLC Radiolabeled steroid precursors were as follows: w6,7-3HxE2 (1.5 TBqymmol, 40 Ciymmol), w2,4,6,7-3HxE1 (4.0 TBqymmol, 108 Ciymmol), w1,2,6,7-3HxD4 (4.1 TBqymmol, 110 Ciymmol), Amersham Corp (Arlington Heights, IL); w1,23 x H T (1.9 TBqymmol, 52.5 Ciymmol), w1b3 x H D4 (0.9 TBqymmol, 24.1 Ciymmol) DuPont NEN Products (Boston, MA). Two sets of experiments were conducted: one in June 2001 (summer) and a second in January 2002 (winter). All experiments conducted during a given month used the same tissue extracts. Homogenates were stored at 4 8C and all experiments were completed within three weeks of original collection of tissue fragments. Homogenates were used in incubations, except where cytosol is specified. Cytosol was prepared by centrifuging homogenates at 105 000=g for 60 min and collecting the supernatant. To measure 17b-HSD-like activity, homogenates or cytosol were incubated with cofactor (0.5 mM NAD(H) or NADP(H), as specified) and a tritiated steroid substrate (1.0, 21.0 or 101 mM, as specified). All assays were incubated at room temperature. Reaction mixtures were fractionated into substrate and product by thin-layer chromatography using benzene-acetone (4:1 v:v) as the mobile phase (Blomquist et al., 1994). To test the substrate specificity of the conversion of E2 to E1, the rate of production of radiolabeled E1 from E2 was measured in the presence and absence of a 100fold excess of unlabeled T. E1 was confirmed as product of E2 metabolism through recrystallization. Aromatase was assayed based on the NADPHdependent release of 3H2O from 1b-w3HxD4. Microsomes (105 000=g pellet) were incubated at pH 7.2 in 0.5-ml reaction mixtures prepared by combining 290 ml of 100 mM HEPES, pH 7.2, containing 0.5 mM glucose-6-phosphate, 0.5 unity ml glucose-6-phosphate dehydrogenase and 1.0 mM NADPH with 200 ml of microsomes. The reaction was started by the addition of 10 ml of 10 mM w3HxD4. Reaction was stopped by the addition of 0.5 ml of 20% trichloroacetic acid and 1.0 ml of activated charcoal (5% wyv). After
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centrifugation at 1000=g for 10 min, 1.0 ml of the supernatant was taken for liquid scintillation counting of 3H2O. Microsomes prepared from M. capitata, P. damicornis and T. coccinea were incubated in triplicate for 22.5 and 43.5 h. Human placental microsomes were used as a positive control. 3. Results 3.1. Stable-isotope experiments with GC-MS detection When GKP and HEPES buffers with coral homogenate were extracted as blanks, GKP buffer resulted in a higher baseline. Omitting glycerol from the GKP buffer recipe and solvent–rinsing the potassium phosphate salts reduced the concentration of contaminants. HEPES buffer was used in most incubations to minimize the contaminants co-eluting with steroids and to facilitate comparisons with other studies (i.e. Slattery et al., 1997). Androgens and estrogens were only detected in buffers or coral homogenates when added experimentally; unlabelled (endogenous) androgens or estrogens were not detected. In all buffers, an unknown, non-steroidal (fatty acid) contaminant co-eluted with E1. The concentration of the fatty acid was lower in HEPES buffer than in GKP buffer; this concentration was measured in blanks and was subtracted from peak areas from incubations to determine E1 concentration. Although elution times for E1 and the contaminant overlapped, spectra of E1 could be clearly identified. Steroid substrates and products were identified based on peak elution time and mass spectra (Table 1). Coral homogenate spiked with 13C-T and 13CE2 and immediately extracted showed no evidence of labeled steroid metabolites (Table 2). Similarly, when coral rubble was incubated with E2, labeled steroid metabolites were not detected. However, after a 4-h incubation of 13C-E2 with coral homogenate, approximately 2% of the precursor was converted to E1. Incubations with 13C-T resulted in up to 4% conversion of precursor into D4 and androstanedione (Fig. 2, Table 2). Mass spectra of 13 C-labeled products are shown in Fig. 3. 3.2. Radioisotope experiments using TLC As a first approach to assessing 17b-HSD-like activity, assays were run with E2, E1, T and D4 at
477
Table 1 Steroid standards (E1, E2), substrates (13C-E2, 13C-T), and products (13C-E1, 13C-D4, 13C-Androstanedione) detected by GC-MS Compound
Time
MW
TMS
E1 13 C-E1 E2 (mono-silane) 13 C-E2 (mono-silane) 13 C-D4 E2 13 C-E2 13 C-T 13 C-Androstanedione
22.10 22.10 22.32 22.34 22.50 22.61–22.66 22.64 22.98–23 22.76–22.79
342 344 344 346 288 416 418 362 290
1 1 1 1 0 2 2 1 0
Time is peak elution time, MW is molecular mass (measured as the mass to charge ratio), TMS is the number of trimethylsilyl groups attached during derivatization. Note that both mono- and bis-TMS forms of E2 were detected. Unlabeled and 13 C-steroids coeluted; however, endogenous unlabeled estrogens and androgens were not detectable in coral homogenates.
a low concentration (1.0 mM) and a high concentration (21.0 mM E2 or E1, the solubility limit and 101 mM T or D4) of steroid substrate. These conditions have been shown to be optimal for detecting various isoforms of 17b-HSD in human tissues and subcellular fractions (Blomquist, 1995). During the summer experiments, coral homogenates converted E2 to E1, indicating the presence of a 17b-HSD-like enzyme (Table 3). Approximately 10-fold greater conversion occurred when NADPq was used as a cofactor than when NADq was used. When homogenates were incubated for 3 h with 1.0 mM E2 and 0.5 mM NADPq, conversion to E1 ranged from 12% for T. coccinea to 31% for M. capitata. Protein concentrations varied among samples, so the specific activity of conversion (pmol (mg protein)y1 3 hy1) of E2 to E1 was lowest for T. coccinea and highest for P. damicornis. Reduction of E1 to E2 by coral homogenates ranged from below detection to 9.7% conversion. Coral homogenates also converted T to D4 (Table 3). As with the estrogens, NADPq was preferred as a cofactor over NADq. When homogenates were incubated for 3 h with 1.0 mM testosterone and 0.5 mM NADPq, conversion to D4 ranged from 1.7% for P. damicornis to 10% for T. coccinea (Table 3). With 101 mM T, the specific activity increased, but only 0.6–6.7% of the precursor was converted. Reduction of D4 to T by coral homogenates ranged from below detection to 4.7%. Coral microsomes incubated with
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478 Table 2 Conversion of Time (h)
pH
13
C-labeled steroid substrates by M. capitata
Sample
Wet weight (g)
Substrate (mg)
6.5 5.5 5.8 5.4 5.6
110 220 110 220 220
9
220
Recovery E2
E1
84 117 55 197 136
ND ND 0.99 4.8 2.5 0.03 (0.5)
Conversion (%)
Specific activity pmol gy1
pmolymg proteiny1
– – 1.8 2.4 1.8
– – 500 2600 1300
– – 290 1500 760
1.2
870
510
– 10.5 3.1 1.6 3.8
– 7800 200 3200 1200
– 4600 120 1900 740
1.7
1600
970
13
C-Estradiol substrate 0 9 Homogenate 4 9 Rubble 4 9 Homogenate 4 9 Homogenate 4 9 Homogenate* Fragment 4 8.2 (buffer)
13
C-Testosterone substrate 0 9 Homogenate 4 9 Homogenate 4 9 Homogenate 4 7.2 Homogenate 4 7.5 Homogenate Fragment 4 8.2 (buffer)
6.5 5.4 5.7 5.5 5.6
125 252 252 252 252
8.4
252
12 (32) T
D4
A
20 125 13 29 76 0.44 (4.3)
ND ND 0.4 6.1 2.4 ND (0.07)
ND 14.6 ND 0.2 0.7 ND
Sample types: homogenatesM. capitata homogenized in buffer, rubblesM. capitata skeletal fragment homogenized in HEPES buffer, fragmentsM. capitata live coral fragment incubated in artificial seawater with 20 mM HEPES (fragment and buffer incubated together for 4 h with a labeled steroid substrate and extracted separately). One sample homogenized in GKP buffer is marked with an asterisk. Substrate added and steroids recovered are given in milligram; AsAndrostanedione. Percent conversion is given as a percent of labeled steroids recovered. ND indicates no detectable conversion. Specific activity (per 0- or 4-h incubation) is given in pmol (gram wet weight)y1 and pmol (milligram protein)y1. Protein concentration was estimated as 1.7 mg protein per gram wet weight (Tarrant et al., 1999). Activity is normalized to estimated protein concentration to allow comparison with radioisotope results.
1b-w3HxD4 did not produce significant quantities of 3H2O; thus, aromatase activity was not detected in these experiments (data not shown). When extracts were fractionated into cytosol and microsomes by centrifugation at 105 000 g for 60 min, most of the activity with E2 as a substrate was recovered in the cytosol, NADPq was the preferred cofactor, and T in 100-fold excess had only a slight inhibitory effect (Fig. 4). To confirm the requirement for a pyridine nucleotide cofactor, samples of M. capitata cytosol were dialyzed overnight against a 100-fold excess of 20% GKP buffer and then assayed with and without NADPq. No activity was observed in the absence of NADPq. Dialysis with a 3500-mw cutoff membrane had little effect on activity in the presence of NADPq; thus, the removal of small molecules had little effect on the activity, which is most likely mediated through an enzyme (Fig. 5). Because T. coccinea homogenates showed the greatest capacity for androgen metabolism in preliminary experiments (Table 3), T. coccinea homogenate and cytosol were incubated with low (1.0
mM) and high (101 mM) concentrations of T or D4 for up to 21 h. Activity with T and D4 was also recovered largely in the cytosolic fraction. With both a low (1.0 mM) and a high (101 mM) concentration of T or D4, the reaction was essentially complete by 5 h with no further product formation during an overnight incubation (Fig. 6). Winter experiments quantified conversion of E2 to E1 in the presence of NADPq by extracts of M. capitata, T. coccinea and the zooxanthellae culture (Fig. 7). In marked contrast with the results of the summer experiments, activity in T. coccinea cytosol greatly exceeded that in the M. capitata sample. Activity in both coral species, particularly M. capitata, was lower in winter than in summer. The highest specific activity was found in the zooxanthellae. The ratio of specific activity measured with NADPq as cofactor to the specific activity with NADq as cofactor (NADPq yNADq ratio) can be used to identify different enzymes that act on the same steroid substrate (Blomquist, 1995). In converting E2 to E1, the T. coccinea tissue homogenate
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Fig. 2. Chromatographs of extracted M. capitata homogenates. Labels represent elution times for 13C-labeled substrates and products. (a) E2 and T added and immediately extracted, 22.64s13C E2, 23s13C T (b) 4-h incubation of M. capitata rubble homogenate with E2, 22.33s13C E2 (mono-silane), 22.65s13C E2 (c) 4-h incubation of M. capitata homogenate with E2, showing E1 as a product, 22.1s13C E1, 22.3s13C E2 (mono-silane), 22.67s13C E2 (d) 4-h incubation of M. capitata homogenate with T, showing androstanedione as a product, 22.77s13C androstanedione, 23.02s13C T.
had similar NADPq yNADq ratios in winter and summer (Table 4). The corresponding NADPq y NADq ratios for M. capitata ranged from approximately 10 in the first summer experiment to -2 in the winter. The ratio for the zooxanthellae sonicate was 1.1. 4. Discussion These experiments clearly demonstrate that corals (zooxanthellate and azooxanthellate) and their dinoflagellate symbionts contain a functional 17bHSD and a 5a-reductase. Reduction (E1™E2 and D4™T) occurred at a lower rate than dehydrogenation (E2™E1 and T™D4). NADPq was the preferred cofactor in dehydrogenation reactions and dehydrogenation of E2 was slightly inhibited by an excess of T (Fig. 4). These results are consistent with metabolism by an enzyme with
both a steroid recognition site and a coenzyme binding sites, but additional experiments will be needed to characterize the substrate specificity. Aromatization of D4 was not observed; however, we cannot rule out the possibility that aromatase may be present in coral tissue at low abundance or may be restricted temporally or spatially within coral polyps. Aromatase activity in mammals outside of the female reproductive organs is generally very low, but localized ‘paracrine’ synthesis of estrogens is biologically important and can result in locally high concentrations of E2 and E1 (Dikkeschei et al., 1996; Simpson and Davis, 2001; Simpson et al., 2002). Use of a cnidarian with larger gonads, such as an anemone, might allow assay of aromatase activity specifically in reproductive tissues. Immunohistochemistry could also be used in the future to determine whether aromatase is present in coral tissue. Polar and non-
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Fig. 3. Mass spectra of estrogens and androgens synthesized during incubations with coral tissue homogenates. Black circles on structures denote 13C atoms. (a) silanized 13C-E1 (b) 13C-D4 (c) 13C-androstanedione.
polar steroid conjugates were not assayed in this study; however, other invertebrates can produce polar and non-polar conjugates of estrogens and androgens (Voogt and Van Rheenen, 1986; Voogt et al., 1987; Fairs et al., 1989; Swevers et al., 1991; Watts et al., 1994). Steroid metabolites produced by coral homogenates in this study are qualitatively similar to those produced by Antarctic soft corals (Slattery et al., 1997). Alcyonium paessleri and Clavularia frankliniana converted D4 into T, epiandrosterone, 5aandrostanedione and an unidentified esterified androgen. T was the major product, with 28% (C. frankliniana) total precursor conversion after 12 h, giving a specific activity of 1700 pmol (g wet weight)y1 hy1. Protein in C. frankliniana is 24%
of dry weight (Slattery and McClintock, 1995); if dry weight is 10–50% of wet weight, specific activity would be 14–71 pmol (mg protein)y1 hy1. In contrast, we observed little reduction of D4, but up to 10% conversion of T to D4 with a specific activity of 9.96 pmol (mg protein)y1 3 hy1 or and average rate of 3.3 pmol (mg protein)y1 hy1. Thus, androgen metabolism in Antarctic alcyonaceans occurred at a higher rate than in Hawaiian scleractinians. Metabolism of estrogens by soft corals has not been described, but in scleractinians we observed specific activities )3000 pmol (mg protein)y1 3 hy1 (Table 3, E2™E1). In the present study, F30% of the substrate was metabolized in any reaction and incubations beyond ;3 h did not result in addi-
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481
Table 3 Net conversion of tritiated steroid precursors by coral homogenates in June Percent conversion (net)
Specific Activity
Cofactor
NAD(H)
NADP(H)
NAD(H)
NADP(H)
E2 to E1 M. capitata P. damicornis T. coccinea
1.0 mM E2 3.5 8.0 4.7
21 mM E2 2.6 3.7 3.8
1.0 mM E2 31.3 16.4 12.4
21 mM E2 22.6 11.3 11.0
1.0 mM E2 17.7 152 8.0
21 mM E2 306 1420 129
1 mM E2 182 313 21.0
21 mM E2 2620 4540 389
E1 to E2 M. capitata P. damicornis T. coccinea
1.0 mM E1 ND 1.8 ND
21 mM E1 ND 2.9 4.4
1.0 mM E1 0.5 9.7 ND
21 mM E1 4.4 9.6 6.2
1.0 mM E1 ND 34.8 ND
21 mM E1 ND 1160 155
1 mM E1 2.9 186 ND
21 mM E1 539 3850 221
T to D4 M. capitata P. damicornis T. coccinea
1.0 mM T 2.8 2.5 4.9
101 mM T 1.6 ND 2.0
1.0 mM T 4.0 1.7 10.0
101 mM T 2.5 0.6 6.7
1.0 mM T 15.9 48.3 8.4
101 mM T 930 ND 330
1 mM T 23.1 32.7 17.1
101 mM T 1430 1090 1130
D4 to T M. capitata P. damicornis T. coccinea
1.0 mM D4 1.0 1.3 2.3
101 mM D4 0.8 ND 1.4
1.0 mM D4 2.5 2.0 4.7
101 mM D4 1.1 ND 2.8
1.0 mM D4 5.7 24.9 3.9
101 mM D4 420 18.9 235
1 mM D4 14.7 39.0 7.8
101 mM D4 660 ND 474
Coral tissue homogenates (1000=g supernatants from M. capitata, P. damicornis and T. coccinea) were incubated for 3 h with 1.0– 101 mM of a steroid substrate and 0.5 mM of the cofactor indicated. Percent conversion in blank incubations was less than 2% (except E1 to E2, 17%) and has been subtracted. Specific activity given in pmol (milligram protein)y1 3 hy1. ND, activity not detected.
Fig. 4. Conversion of E2 to E1 by M. capitata homogenate (1000=g supernatant) and cytosol (105 000=g supernatant) with NADq or NADPq as cofactor in the absence or presence of T. Assays were run at pH 9.0 with 1.0 mM 3H-E2, 0.5 mM NADq or NADPq and, where noted, 100 mM T. The values are the mean "S.D. of duplicate assays run at room temperature for 330 min.
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Fig. 5. Conversion of E2 to E1 by M. capitata cytosol (solid circles) and M. capitata cytosol dialyzed overnight against a 100-fold excess of 20% GKP buffer (open circles). A 3500mw cut-off membrane was used. Activity was not detected in M. capitata cytosol in the absence of NADPq (triangles). Assays were run at pH 9.0 with 1.0 mM 3H-E2, and 0.5 mM NADPq when added.
tional metabolism. At present, it is not known whether consumption of some unknown cofactor or other component is limiting further conversion. 17b-HSD activity in alcyonacean and scleractinian corals is low relative to activity reported in specific mammalian tissues or cell types we.g. cells derived from a carcinoma of the vulva converted T to D4 with specific activity ;8 nmol (mg protein)y1 30 miny1 (Blomquist et al., 1997)x. It is important to recognize that the coral metabolic rates represent an average rate for an entire animal; rates in some cells or tissues may be considerably higher. Incubations of coral and zooxanthellae with E2 indicate seasonal and interspecies variation in specific activity (Table 3, Fig. 7) and NADPyNAD ratios (Table 4). However, each species during a given month (January or June) was represented by a homogenate from one fragment, so a more complete examination of seasonality is needed. Future experiments should also test the effects of environmental factors (e.g. photoperiod, temperature and exposure to pollutants) on steroid metabolism. Variation in steroid metabolism may cause observed seasonal variation in estrogen concentra-
Fig. 6. Conversion of T to D4 and D4 to T by T. coccinea cytosol. Duplicate assays (mean with the difference between replicates shown) containing 1.0 mM or 101 mM T or D4 and 0.5 mM NADPq or NADPH at pH 9.0 (T to D4) or pH 7.2 (D4 to T) were incubated at room temperature for 5 or 21 h.
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483
in M. capitata were 20–120 ng (g ash-free dry weight)y1 or ;0.4–3 ng (g wet weight)y1; E2 concentrations were approximately half the E1 concentrations. Thus, to detect endogenous estrogens in coral tissue using GC-MS, it would be necessary to improve the sensitivity by a factor of 10–100 to compare with the sensitivity of radioimmunoassay. This could be accomplished by extracting more tissue or by concentrating the extract on the GC column. These approaches would also increase the concentrations of contaminants, so additional sample preparation would be needed. Additional metabolic assays, protein purification and cloning will help to identify the coral enzymes and their natural substrates. Similarities in substrate specificity are not always related to sequence similarity (Baker, 1994). 17b-HSD sequences have been identified in bacteria and fungi; notably a fungal 17b-HSD catalyzes metabolism of both steroidal and non-steroidal (quinonal) substrates (Lanisnik Rizner et al., 1999). 17b-HSDs are homologous to other oxidoreductases, including bacterial 3a20b- and 3b-HSD and the NodG protein, which is found in the nitrogen-fixing bacterium, Rhizobioum meliloti (Baker, 1994). We now know that estrogens are present in the water column (Atkinson et al., 2003), may be taken up (Tarrant et al., 2001) and metabolized by scleractinian corals (present study) and can affect coral physiology (Tarrant, 2002; Tarrant et al., in review). Environmental pollutants may alter rates of steroid metabolism and consequently, steroid concentrations in tissue and affect physiological
Fig. 7. Time-course of the conversion of E2 to E1 by 1000=g supernatants of M. capitata (solid circles), T. coccinea (open circles), and a sonicate of zooxanthellae from M. capitata (triangles). Sonication was for two 1.0-min intervals in a bathtype sonicator (Heat Systems, Inc.). Assays were run at pH 9.0 with 1.0 mM 3H-E2, and 0.5 mM NADPq.
tions (Tarrant et al., 1999). Variation in the NADPq yNADq ratio is consistent with more than one enzyme catalyzing steroid dehydrogenation (Blomquist, 1995). Changes in steroid metabolism may also result from a variable contribution of algal and animal components to the overall rate. Endogenous estrogens and androgens were not detected by GC-MS in coral homogenates, with a detection limit for steroid metabolites of 40 ng (g wet weight)y1. In contrast, Tarrant et al. (1999) used radioimmunoassay to quantify endogenous E1-immunoreactivity and E2-immunoreactivity. Based on immunoreactivity, the E1 concentrations Table 4 NADPqyNADq ratios for January and two June experiments Experiment
Sample
1.0 mM
21.0 mM
Notes
June Expt. 1
M. capitata P. damicornis T. coccinea M. capitata (homogenate) M. capitata (cytosol) M. capitata T. coccinea Zooxanthellae
10.3 2.1 2.7
8.6 3.2 3
(3 h assay)
June Expt. 2
January Expt.
(5.5 h assay) 5.6 3.8 1.6 3.8 1.1
(2 h assay)
Ratio of specific activity of conversion of E2 to E1 with NADPq as a cofactor to specific activity with NADq as a cofactor. Assays were run at pH 9.0 with 1.0 or 21.0 mM E2 and 0.5 mM NADPq or NADq. Tissue homogenates (1000=g supernatants) were used for M. capitata, P. damicornis, and T. coccinea coral tissue, except in the 2nd June experiment, where both homogenates and cytosol (105 000=g supernatant) were used. Zooxanthellae were sonicated, as described in the text.
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processes such as reproduction. Also, since corals can take up and metabolize steroids, they can alter the concentration or form of steroids available to other reef organisms. Acknowledgments We thank Dr R. Bidigare and B. Benitez-Nelson for assistance with GC-MS analysis. Dr R. Kinzie provided zooxanthellae samples. Support for this project was provided by the EPA STAR fellowship program, the University of Hawaii Sea Grant College Program and HealthPartners Research Foundation. The views expressed herein are solely those of the authors. References Atkinson, S., Atkinson, M., Tarrant, A., 2003. Estrogens from sewage in the coastal marine environment. Environ. Health Perspect. 111, 531–535. Atkinson, S., Atkinson, M.J., 1992. Detection of estradiol-17b during a mass coral spawn. Coral Reefs 11, 33–35. Baker, M., 1994. Sequence analysis of steroid- and prostaglandin-metabolizing enzymes: application to understanding catalysis. Steroids 59, 248–258. Blomquist, C., Leung, B., Beaudoin, C., Tremblay, Y., 1997. Intracellular regulation of 17b-hydroxysteroid dehydrogenase type 2 catalytic activity in A431 cells. J. Endocrinol. 153, 453–464. Blomquist, C., 1995. Kinetic analysis of enzymatic activities: prediction of multiple forms of 17b-hydroxysteroid dehydrogenase. J. Steroid Biochem. Mol. Biol. 55, 515–524. Blomquist, C., Bealka, D., Hensleigh, H., Tagatz, G., 1994. A comparison of 17b-hydroxysteroid oxidoreductase type 1 and type 2 activity of cytosol and microsomes from human term placenta, ovarian stroma and granulosa-luteal cells. J. Steroid Biochem. Mol. Biol. 49, 183–189. Bose, R., Majumdar, C., Bhattacharya, S., 1997. Steroids in Achatina fulica (Bowdich): steroid profile in haemolymph and in vitro release of steroids from endogenous precursors by ovotestis and albumen gland. Comp. Biochem. Physiol. C 116, 179–182. Callard, G., Pudney, J., Kendall, S., Reinboth, R., 1984. In vitro conversion of androgens to estrogen in amphioxus gonadal tissues. Gen. Comp. Endocrinol. 56, 53–58. Dikkeschei, L., Wolthers, B., Bos-Zuur, I., Brutel de la Riviere, G., Nager, G., van der Kolk, D., et al., 1996. Optimization of a classical aromatase activity assay and application in normal, adenomatous and malignant breast parenchyma. J. Steroid Biochem. Mol. Biol. 59, 305–313. Fairs, N., Evershed, R., Quinlan, P., Goad, L., 1989. Detection of unconjugated and conjugated steroids in the ovary, eggs and haemolymph of the decapod crustacean Nephrops norvegicus. Gen. Comp. Endocrinol. 4, 199–208. Gassman N.J., 1992. Aspects of the steroid biochemistry of the scleractinian coral, Favia fragum. Ph.D. Dissertation,
Department of Marine Biology and Fisheries, University of Miami, 158 pp. Ghosh, D., Ray, A.K., 1992. Evidence for physiological responses to estrogen in freshwater prawn, Macrobrachium rosenbergii. J. Inland Fish. Soc. India 24, 15–21. Ghosh, D., Ray, A.K., 1993. Subcellular action of estradiol17b in a freshwater prawn, Macrobrachium rosenbergii. Gen. Comp. Endocrinol. 90, 273–281. Gottfried, H., Dorfman, R., Wall, P., 1967. Production of oestrogens by an accessory reproductive tissue of the slug Arion ater rufus (Linn.). Nature 215, 409–410. Hines, G.A., Bryan, P.J., Wasson, K.M., McClintock, J.B., Watts, S.A., 1996. Sex steroid metabolism in the antarctic pteropod Clione antarctica (Mollusca: Gastropoda). Invert. Biol. 115, 113–119. Hines, G.A., Watts, S.A., McClintock, J.B., 1994. Biosynthesis of estrogen derivatives in the echinoid Lytechinus variegatus Lamarck. In: David, B., Guille, A., Feral, J.P., Roux, M. (Eds.), Echinoderms Through Time. Balkema, Rotterdam, pp. 711–716. Hines, G.A., Watts, S.A., Walker, C.W., Voogt, P.A., 1992. Androgen metabolism in somatic and germinal tissues of the sea star Asterias vulgaris. Comp. Biochem. Physiol. B 102, 521–526. Lanisnik Rizner, T., Moeller, G., Thole, H., Zakelj-Mavrik, M., Adamski, J., 1999. A novel 17b-hydroxysteroid dehydrogenase in the fungus Cochiobolus lunatus: new insights into the evolution of steroid hormone signaling. Biochem. J. 337, 425–431. Le Curieux-Belfond, O., Moslemi, S., Mathieu, M., Seralini, G., 2001. Androgen metabolism in oyster Crassostrea gigas: evidence for 17b-HSD activities and characterization of an aromatase-like activity inhibited by pharmacological compounds and a marine pollutant. J. Steroid Biochem. Mol. Biol. 78, 359–366. Lupo di Prisco, C., Dessi’Fulgheri, F., 1975. Alternative pathways of steroid biosynthesis in gonads and hepatopancreas of Aplysia depilans. Comp. Biochem. Physiol. B 50, 191–195. Maragos, J., 1977. Order scleractinia: stony corals. In: Devaney, D.M., Eldredge, L.G. (Eds.), Reef and Shore Fauna of Hawaii, Section 1: Protozoa Through Ctenophora. Bishop Museum Press, Honolulu, Hawaii, pp. 158–241. Maragos J., 1995. Revised checklist of extant shallow-water stony coral species from Hawaii (Cnidaria: Anthozoa: Scleractinia). Bishop Museum Occasional Papers, 42, pp. 54– 55. Matsumoto, T., Osada, M., Osawa, Y., Mori, K., 1997. Gonadal estrogen profile and immunohistochemical localization of steroidogenic enzymes in the oyster and scallop during sexual maturation. Comp. Biochem. Physiol. B 118, 811–817. Miller, W., 1988. Molecular biology of steroid hormone synthesis. Endocr. Rev. 9, 295–318. Oberdorster, E., Cheek, A., 2001. Gender benders at the beach: endocrine disruption in marine and estuarine organisms. Environ. Toxicol. Chem. 20, 23–36. Schoenmakers, H., Voogt, P., 1980. In vitro biosynthesis of steroids from progesterone by the ovaries and pyloric caeca
A.M. Tarrant et al. / Comparative Biochemistry and Physiology Part B 136 (2003) 473–485 of the starfish Asterias rubens. Gen. Comp. Endocrinol. 41, 408–416. Simpson, E., Clyne, C., Rubin, G., Boon, W., Robertson, K., Britt, K., et al., 2002. Aromatase—a brief overview. Ann. Rev. Physiol. 64, 461–470. Simpson, E., Davis, S., 2001. Minireview: aromatase and the regulation of estrogen biosynthesis—some new perspectives. Endocrinology 142, 4589–4594. Slattery, M., Hines, G.A., Watts, S.A., 1997. Steroid metabolism in Antarctic soft corals. Polar Biol. 18, 76–82. Slattery, M., McClintock, J., 1995. Population structure and feeding deterrence in three shallow-water antarctic soft corals. Mar. Biol. 122, 461–470. Swevers, L., Lambert, J.G.D., De Loof, A., 1991. Metabolism of vertebrate-type steroids by tissues of three crustacean species. Comp. Biochem. Physiol. B 99, 35–41. Tarrant A., 2002. Estrogen Action in Scleractinian Corals: Sources, Metabolism and Physiological Effects. Ph.D. Dissertation, Department of Oceanography, University of Hawaii at Manoa, 186 pp. Tarrant A., Atkinson M., Atkinson S., Effects of steroidal estrogens on coral growth and reproduction. Mar. Ecol. Prog. Ser., In Review. Tarrant, A.M., Atkinson, M.J., Atkinson, S., 2001. Uptake of estrone by a coral reef community. Mar. Biol. 139, 321–325. Tarrant, A.M., Atkinson, S., Atkinson, M.J., 1999. Estrone and estradiol-17b concentration in tissue of the scleractinian
485
coral, Montipora verrucosa. Comp. Biochem. Physiol. A 122, 85–92. Voogt, P., Lambert, J., Granneman, J., Jansen, M., 1992. Confirmation of the presence of oestradiol-17b in the sea star Asterias rubens by GC-MS. Comp. Biochem. Physiol. B 101, 13–16. Voogt, P.A., Den Besten, P.J., Kusters, G.C.M., Messing, M.W.J., 1987. Effects of cadmium and zinc on steroid metabolism and steroid level in the sea star Asterias rubens L. Comp. Biochem. Physiol. C 86, 83–89. Voogt, P.A., Van Rheenen, W.A., 1986. Androstenedione metabolism in the sea star Asterias rubens L. studies in homogenates and intact tissue: biosynthesis of the novel steroid fatty-acyl testosterone. Comp. Biochem. Physiol. B 85, 497–501. Voogt, P.A., Van Rheenen, W.A., Lambert, J.G.D., De Groot, B.F., Mollema, C., 1986. Effects of different experimental conditions on progesterone metabolism in the sea star Asterias rubens L. Comp. Biochem. Physiol. B 84, 397–402. Watts, S., Hines, G., Byrum, C., McClintock, J., Marion, K., 1994. Tissue- and species-specific variations in androgen metabolism. In: David, B., Guille, A., Feral, J.P., Roux, M. (Eds.), Echinoderms Through Time. Balkema, Rotterdam, pp. 155–161. Young, N., Quinlan, P., Goad, L., 1992. Progesterone metabolism in vitro in the decapod crustacean, Penaeus monodon. Gen. Comp. Endocrinol. 87, 300–311.