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a Department of Marine Science, Marine Science Institute, University of Texas at Austin, Port Aransas, Texas 78373-5015
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Studies in mammalian and nonmammalian vertebrates suggest that estradiol is involved in a variety of testicular functions, including increases in mast cell number in testicular interstitial tissue, Leydig cell development, and the regulation of spermatogenic progression [15–19]. In addition, evidence has been obtained from several laboratories indicating that estrogens can decrease testicular androgen production in vertebrates, possibly by decreasing the activity of one or more of the steroidogenic enzymes that convert progesterone to testosterone ([20, 21], for reviews). However, the precise physiological functions of estrogens and their mechanisms of action in the male reproductive system remain unclear. For example, the involvement of the nuclear estrogen receptor in mediating many actions of estrogens in the testis has not been clearly demonstrated.
The purpose of the present study was to determine whether estrogens are involved in one aspect of testicular function, the regulation of androgen production, in a teleost model, the Atlantic croaker, and to investigate the mechanism of estrogen action. Production of 11-ketotestosterone (11-KT), a predominant androgen in teleosts, from Atlantic croaker testicular fragments was assessed after short-term exposure to estrogens in vitro. Male reproductive function, including testicular steroidogenesis, is susceptible to disruption by xenobiotic compounds. A mixture of polychlorinated biphenyls caused a decrease in androgen production by rat Leydig cells in vitro, and maternal exposure to 4-octylphenol during fetal development of rats caused a reduction in the activity of the steroidogenic enzyme 17
-hydroxylase in the fetuses [22, 23]. Therefore, the effects of several xenoestrogens on testicular 11-KT production were also examined in this study. Finally, the mechanism and cellular site of estrogen action on 11-KT production were explored using a variety of experimental approaches including the characterization of an estrogen-binding moiety on testicular plasma membranes.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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One-year-old mature male Atlantic croaker (35 g) were collected by otter trawl from Redfish Bay near Port Aransas, Texas. Fish were caught in September, during the beginning of gonadal recrudescence, and maintained in 4200-liter circular, recirculating tanks at a temperature of 22–25°C under an 11L:13D photoperiod and fed a diet of commercial pellets and shrimp (3% of body weight per day). Spermiating males with a mean gonadosomatic index (GSI; gonad weight x 100/total weight) of 6.8 ± 2% and 6.4 ± 0.7% were used as testicular tissue donors for the incubation experiments and for analysis of membrane receptor binding, respectively.
Chemicals
[2,4,6,7-3H]Estradiol-17ß ([3H]estradiol, 72 Ci/mmol) was purchased from New England Nuclear (Boston, MA). Nonradioactive steroids were purchased from Sigma Chemical Company (St. Louis, MO) or Steraloids (Wilton, NH). 17ß-Estradiol 17-hemisuccinate-BSA (30 mol steroid/mol BSA), actinomycin D, cycloheximide, hCG, and zearalenone were purchased from Sigma. Tritiated 11-ketotestosterone ([3H]11-KT, 51.9 Ci/mmol) was purchased from Amersham Pharmacia Biotech (Piscataway, NJ). The synthetic estrogen diethylstilbestrol (DES) and the antiestrogens tamoxifen citrate and nafoxidine hydrochloride were purchased from ICN Biomedicals (Aurora, OH). The hydroxylated polychlorinated biphenyl (PCB), 2,2',5'-trichloro-4-biphenylol, was purchased from Ultra Scientific (North Kingston, RI). Derivatives of the pesticide dichlorodiphenyltrichloroethane (DDT) (o,p'-DDT and o,p'-DDE) were purchased from Chem Service (West Chester, PA). The chlorinated pesticide Kepone (chlordecone; formerly manufactured by Allied Chemical, Hopewell, VA) was obtained from the National Institute of Environmental Health Sciences Repository (NIH, Bethesda, MD). The nonionic surfactant 4-nonylphenol was obtained from the Huntsman Corporation (Port Neches, TX). The PCB mixture, Aroclor 1254, was purchased from the Foxboro Co. (North Haven, CT). Mibolerone, a synthetic steroidal androgen, was a gift from Upjohn Laboratories (Kalamazoo, MI). The antiestrogen ICI 182,780 was a gift from Dr. A.E. Wakeling at Zeneca Pharmaceuticals (Cheshire, England). All other chemicals were reagent grade and were purchased from general laboratory suppliers.
Effects of Estrogens on Testicular Androgen Production
General incubation procedure Spermiating Atlantic croaker were killed, and the testes were rapidly removed and placed in ice-cold incubation buffer. Single tissue fragments, weighing approximately 100 mg, were placed in each well of 24-well polystyrene incubation plates containing 1 ml of Dulbecco's modified Eagle's medium (DMEM) lacking any pH indicator dye, supplemented with sodium bicarbonate (1.2 g/L), penicillin (60 mg/L), and streptomycin (100 mg/L), at a pH of 7.4. Tissues were preincubated for 30–60 min at 25°C under an atmosphere of oxygen. At the end of the preincubation period, the tissue was cut into smaller fragments (quartered, approximately 25 mg); the medium was removed, and 1 ml of fresh medium was added to each incubation well. Tissue fragments were incubated with the test treatments in a Dubnoff shaking incubator (Precision Scientific, Chicago, IL) in a humid environment at 25°C under an atmosphere of oxygen for up to 9 h. This was followed in most experiments by a subsequent 9-h incubation in medium alone (double-incubation procedure). At the end of the incubation period, media were removed and frozen at -20°C until assayed for 11-KT by RIA.
Concentration-dependent effects of estradiol Tissue fragments were incubated for 9 h with 10 IU hCG and concentrations of estradiol ranging from 367 pM to 36.7 µM. Human CG was dissolved directly in the medium, and estradiol was added in 5 µl of ethanol for a final ethanol concentration of 0.5%. Tissues from 3 fish were incubated in duplicate for each estradiol concentration.
Time course of estradiol action Tissue fragments were incubated with hCG (10 IU) and 36.7 µM estradiol dissolved in 5 µl ethanol (final ethanol concentration 0.5%) or hCG (10 IU) and ethanol alone (controls) for time periods ranging from 5 min to 6 h. At the end of the incubation, media were removed, and tissue fragments were washed twice with 1 ml of fresh medium and incubated for an additional 9 h in DMEM without hCG (double-incubation procedure). The final incubation media were assayed for 11-KT content. Tissue fragments from 3 individuals were incubated in duplicate for each time point.
Effects of synthetic estrogens, antiestrogens, and xenobiotics Tissue fragments from 3 fish were incubated in duplicate for 6 h with hCG (10 IU) and each of the test chemicals (xenobiotics, synthetic estrogens, and antiestrogens). Final treatment concentrations ranged from 10 µM to 1 mM and were added in 10 µl of ethanol for a final ethanol concentration of 1% (this concentration of ethanol did not significantly affect androgen production). After the incubation, tissues were washed twice with 1 ml fresh medium and incubated in the absence of test chemicals and hCG for an additional 9 h (double-incubation procedure). The final incubation media were assayed for 11-KT content.
Effects of nonestrogenic steroids Tissue fragments from 4 fish were incubated for 9 h with hCG (10 IU) and each of the steroid treatments. Steroids (17,20ß-dihydroxy-4-pregnen-3-one, 1 µM; 17,20ß,21-trihydroxy-4-pregnen-3-one, 1 µM; progesterone, 10 µM; cortisol, 10 µM; mibolerone, 10 µM; estradiol, 10 µM) were added in 5 µl of ethanol. All incubations had a final ethanol concentration of 0.5%.
Mechanism of Estrogen Action
Effects of various steroid precursors Tissue fragments from 3 fish were incubated in duplicate with estradiol (36.7 µM) or ethanol (1%) for 6 h following a 1-h preincubation in medium alone. Medium was removed, and tissue fragments were washed 5 times with 1 ml fresh medium. Tissue fragments were incubated for an additional 9 h (double-incubation procedure) with 100 ng of steroid precursor (progesterone, 17-hydroxyprogesterone, or androstenedione). The final incubation media were assayed for 11-KT content.
Effects of transcription and translation inhibitors After a 1-h preincubation in medium alone, tissue fragments were incubated with or without 100 µM actinomycin D (transcription inhibitor) or 10 µM cycloheximide (translation inhibitor) for 2 h. Estradiol (36.7 µM) or ethanol (5 µl) was then added, and incubations were continued for 6 h. Medium was then removed, and tissue fragments were washed twice with 1 ml of fresh medium and incubated for an additional 9 h (double-incubation procedure) with progesterone (100 ng/ml) as a substrate. Final incubation media were assayed for 11-KT content. Incubations were performed on tissues from 6 individuals for each treatment.
Effects of BSA-conjugated estradiol The conjugated 17ß-estradiol 17-hemisuccinate-BSA (20 µM) was dissolved directly in incubation medium and incubated with tissue fragments from 4 fish for 9 h in the presence of hCG (10 IU).
11-KT RIA Media samples were assayed for 11-KT content by RIA according to Singh et al. [24]. Briefly, 150 µl of each sample plus 50 µl of recovery tracer ([3H]11-KT, 1500 dpm) was extracted with 2 ml hexane/ethyl acetate (70/30). Extracts were resuspended in 300 µl of phosphate buffer, and duplicate 100-µl aliquots were analyzed for 11-KT content. Aliquots (50 µl) were counted in a liquid scintillation counter (LS 6000SC; Beckman Instruments, Fullerton, CA) to determine extraction efficiencies (mean extraction efficiency was 82%). The mean detection limit of the RIA was 6.4 ± 0.7 pg 11-KT. Cross-reactivity of the 11-KT antibody (Helix Biotech Ltd., Warminster, PA) was 0.002% for estradiol, 14% for testosterone, and 17% with dihydrotestosterone. The traces of estradiol carried over from the treatments into the final incubation media did not interfere with the measurement of 11-KT in the RIA. No cross-reactivity occurred with progesterone. The intraassay and interassay coefficients of variation were 6% and 15%, respectively.
Characterization of Estrogen Membrane Binding
Tissue preparation Tissue was prepared for the estrogen membrane-binding assay according to the general procedure described for the 20ß-S ovarian membrane receptor [25]. Briefly, 3–4.5 g of fresh testicular tissue was homogenized in 15 ml HA buffer (25 nM Hepes, 10 mM NaCl, 1 mM dithioerythritol, pH 7.8) with a Polytron tissuemizer (Tekmar, Cincinnati, OH) and centrifuged at 20 000 x g for 20 min. The supernatant was discarded; the pellet was resuspended in 15 ml HA buffer and centrifuged at 500 x g for 7–10 min to remove the nuclear fraction. The supernatant was centrifuged at 20 000 x g for 20 min. The washing and centrifugation step was repeated. The final pellet that contained the plasma membrane fraction was resuspended in 2.5 ml HA buffer and assayed for binding activity. The final pellet was further fractionated by centrifugation through a sucrose pad (1.2 M) for more complete separation of the plasma membrane from other cellular components prior to measuring binding activity in several assays.
Measurement of Kd and Bmax Radioactive estradiol, with final concentrations ranging from 5 nM to 0.125 nM, was dissolved in 250 µl HA buffer with or without 100- or 1000-fold excess nonradioactive estradiol. Testicular membrane preparation (250 µl) was added, and the mixture was vortexed and incubated for 30 min at 4°C. The reaction was stopped, and bound steroid was separated from free steroid by filtration through presoaked glass microfiber filters (Whatman G/F B, pore size 1 µm; Clifton, NJ). The filters were immediately washed with 25 ml washing buffer (HA buffer without dithioerythritol). Radioactivity was counted for 5 min in a liquid scintillation counter. Dissociation constant (Kd) and binding capacity (Bmax) were determined from the Scatchard plot of specific binding [26].
Association and dissociation kinetics To determine the association kinetics of total binding, membrane extracts (250 µl) were incubated in triplicate with 2 nM [3H]estradiol for periods of time ranging from 15 sec to 90 min. Nonspecific binding was determined simultaneously by incubating membrane extracts with 200 nM nonradioactive estradiol. Specific binding was calculated by subtracting nonspecific binding from total binding at each time point.
Dissociation kinetics were determined after incubation of membrane extracts for 30 min with 2 nM [3H]estradiol in triplicate. Dissociation began with the addition of 200 nM nonradioactive estradiol and was measured at time points ranging from 15 sec to 60 min. Dissociation was calculated as a percentage of the specific binding present in a 30-min control incubation.
Steroid specificity and xenobiotic binding Plasma membrane extracts (250 µl) were incubated in triplicate with [3H]estradiol (final concentration 2 nM) in HA buffer with or without competitor. Competitor concentration ranged from 1 nM to 1 mM, and competitors were added in 5 µl ethanol. Free steroid was removed by filtration. Maximum specific binding was defined as the fraction of total binding suppressed by 200 nM nonradioactive estradiol for estrogens, antiestrogens, and xenobiotics and 1 µM nonradioactive estradiol for other steroids. Competitor binding was expressed as a percentage of maximum specific binding.
Statistics
Statistical significance was determined using ANOVA and the Tukey honestly significant difference (HSD) multiple range test for the experiments with synthetic estrogens and antiestrogens, steroids, xenobiotics, and transcription and translation inhibitors. Statistical significance was determined using the Student's t-test for paired means in all other experiments.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Concentration-dependent effects of estradiol Human CG-simulated testicular production of 11-KT in vitro was decreased in response to treatment with estradiol in a concentration-dependent manner (Fig. 1). However, 11-KT production by testicular tissues from the three donor fish (GSIs 5.4%, 5.9%, and 6.9%) varied considerably in this experiment. Although an apparent decline in 11-KT synthesis was observed with estradiol concentrations as low as 36.7 nM, this decrease was not significant until estradiol concentrations reached 367 nM (control: 23.6 ± 3.1 pg/mg tissue; 367 nM estradiol: 17.8 ± 1.8 pg/mg tissue, P < 0.05).
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There was no evidence of toxic effects of estradiol on 11-KT production. Tissue fragments incubated with the highest concentration of estradiol used in these experiments (37 µM) retained their responsiveness to hCG stimulation (untreated controls: 1.59 ± 0.04 pg; hCG plus estradiol: 3.03 ± 0.13 pg 11-KT/mg tissue; N = 3). Tissue fragments also retained their steroidogenic capacity to convert a steroid precursor, 17-hydroxyprogesterone (10 ng/ml), to 11-KT after estradiol (37 µM) treatment in the absence of gonadotropin stimulation (1.76 pg vs. 0.39 pg 11-KT/mg tissue without precursor).
Time course of estradiol action Five-minute exposure to estradiol was sufficient to significantly (P < 0.05) decrease 11-KT production from 11.47 ± 1.55 pg to 8.02 ± 0.83 pg/mg tissue (Fig. 2). Significant decreases in steroid production were also seen after 30 min, 3 h, and 6 h of estradiol treatment. Although a similar trend was evident at the 15-min and 1-h treatment times, these values were not significantly different from the respective control values.
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Effects of synthetic estrogens and antiestrogens Synthetic estrogens and antiestrogens had variable effects on testicular 11-KT production (Fig. 3). As observed in the previous experiments, estradiol treatment (37 µM) caused a dramatic and significant decline in 11-KT production to approximately 35% of control levels. The same concentration of the synthetic estrogen DES also caused a significant (P < 0.05) decrease in 11-KT production. Interestingly, the antiestrogen ICI 182,780 at a lower concentration (10 µM) also caused a decrease in 11-KT production, from 5.40 ± 0.85 pg/mg tissue in control incubations to 1.96 ± 0.28 pg/mg tissue. The antiestrogens tamoxifen and nafoxidine did not significantly affect 11-KT production at concentrations of 10 µM.
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Effects of nonestrogenic steroids Testicular tissue fragments were incubated with several other steroids in two separate experiments to determine whether the observed decrease in 11-KT production was specific to estrogens (Fig. 4, A and B). Progesterone, cortisol, and the synthetic androgen mibolerone did not cause significant decreases in testicular 11-KT production at the same treatment concentration as estradiol (10 µM, Fig. 4B). The C21 maturation-inducing steroid, 20ß-S (17,20ß,21-trihydroxy-4-pregnen-3-one), significantly decreased 11-KT production by an amount similar to that with estradiol at a 10-fold lower concentration, from control levels of 11.49 ± 2.45 pg/mg tissue to 3.16 ± 1.03 pg/mg tissue. The other teleostean maturation-inducing steroid, 17,20ß-P (17,20ß-dihydroxy-4-pregnen-3-one), also tended to decrease 11-KT production (6.53 ± 1.5 pg/mg tissue), but the decline was not significant at a concentration of 1 µM (Fig. 4).
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Effects of xenobiotics A variety of xenobiotics that bind to the nuclear estrogen receptor in this species [3] were tested for their ability to alter testicular 11-KT production (Fig. 5). The chlorinated pesticide, Kepone, and the hydroxylated PCB, 2,2',5'-trichloro-4-biphenylol, caused significant decreases in 11-KT production at concentrations of 1 mM and 100 µM, respectively. 4-Nonylphenol (500 µM) also significantly decreased 11-KT production from 5.40 ± 0.85 pg to 2.60 ± 0.51 pg/mg tissue. There was a trend of lower 11-KT production after treatment with o,p'-DDE at a concentration of 500 µM, but the decrease was not significant. Ortho, para-DDT and the PCB mixture Aroclor 1254 did not alter 11-KT production at the concentrations tested (1 mM and 300 µM, respectively).
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Mechanism of Estrogen Action
Effects of various steroid precursors
Tissue fragments were incubated with steroid precursors of 11-KT including progesterone, 17-hydroxyprogesterone, and androstenedione after estradiol treatment to determine whether estradiol was disrupting the steroidogenic pathway by inhibiting one or more of the steroidogenic enzymes that convert progesterone to 11-KT, including 17-hydroxylase, 17–20 desmolase, or 17ß-dehydrogenase. Addition of the steroid precursors did not reverse the effects of estradiol on 11-KT production. 11-KT values for estradiol-treated tissues were 54%, 48%, and 57% of control values for incubations with progesterone, 17-hydroxyprogesterone, and androstenedione, respectively.
Effects of transcription and translation inhibitors To determine whether the estradiol-induced decrease in 11-KT production occurs via a genomic mechanism, tissue fragments were incubated in the presence of transcription and translation inhibitors in combination with estradiol treatment (Fig. 6, A and B). Actinomycin D, an inhibitor of transcription, did not block the estradiol-induced decrease in 11-KT production using a progesterone substrate as a precursor and did not influence androgen production in the absence of estradiol (Fig. 6A).
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Cycloheximide, a translation inhibitor, did not block the decrease in 11-KT production caused by estradiol treatment (Fig. 6B). This compound did, however, cause a significant increase in 11-KT production when incubated without estradiol (Fig. 6B).
Effects of BSA-conjugated estradiol To determine whether the site of estradiol action on 11-KT production was at the cell surface, tissue fragments were incubated with estradiol conjugated to BSA, which cannot pass through the cell membrane. Tissue fragments incubated with 20 µM 17ß-estradiol 17 hemisuccinate-BSA produced significantly less 11-KT compared to control incubations: 7.68 ± 1.42 pg/mg tissue and 11.74 ± 2.14 pg/mg tissue, respectively (P < 0.05, N = 4).
Characterization of Estrogen Membrane Binding
Saturation analysis Saturation analysis of [3H]estradiol binding to testicular plasma membrane preparations showed a single class of high-affinity saturable estradiol-binding sites with a dissociation constant (Kd) of 1.6 nM and a maximum binding capacity (Bmax) of 0.031 nM or 26 fmol/g testis (N = 3, representative plot shown in Fig. 7). The binding component saturated between 1 and 2 nM [3H]estradiol.
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Association and dissociation kinetics The time course of specific binding of [3H]estradiol to the membrane preparation is shown in Figure 8. The binding component had a t1/2 of 5 min, and association was complete after 30 min. Dissociation of [3H]estradiol from the membrane fraction had a t1/2 of 30 min, and approximately 60% of the binding had dissociated after 60 min (results not shown).
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Steroid-binding specificity Competitive binding assays showed that the synthetic estrogen DES bound with affinity similar to that of estradiol, whereas the two natural estrogens estriol and estrone bound with an order of magnitude less affinity (Fig. 9A, Table 1). The antiestrogen ICI 182,780 bound with approximately 3% of the affinity for estradiol (EC50 9 x 10-8 M, compared to 2.8 x 10-9 M for estradiol, Table 1). The affinity of tamoxifen for the receptor was less than 0.1% that of estradiol, and nafoxidine did not displace 50% of specific binding at concentrations up to 10 µM (Fig. 9A, Table 1). None of the other steroids tested caused competitive displacement of [3H]estradiol binding at concentrations up to 10 µM (Fig. 9B).
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Xenobiotic binding A variety of xenoestrogens bound to the estradiol membrane-binding component (Fig. 9C). Zearalenone and nonylphenol bound to the membrane-binding component with 10 and 1000 times less affinity than estradiol, respectively (Fig. 9C, Table 1). The hydroxylated PCB 2,2',5'-trihydroxy-4-biphenylol, o,p'-DDT, and o,p'-DDE displaced 50% of specific binding at concentrations between 50 and 150 µM. Aroclor 1254 displaced less than 30% of specific binding at concentrations up to 500 µM. Kepone was also tested for binding, but high concentrations (1 mM) interfered with the separation of bound from free [3H]estradiol.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Although a decline in 11-KT production by Atlantic croaker testis was distinguishable with the 36.7 nM estradiol treatment, significant decreases were observed only at higher concentrations (367 nM and higher). Similarly, it has been found that relatively high concentrations of estradiol are required to decrease androgen production in vitro by testes of other vertebrate species [20, 27, 28]. In contrast, similar effects of estradiol on androgen production in vivo were observed with more modest increases in circulating estradiol concentrations. Implantation of rats with estradiol capsules, which caused only a 2- to 5-fold increase in plasma and testicular estradiol concentrations, was sufficient to cause a decrease in testicular androgen production when the treated testes were incubated in vitro [33, 34]. The finding that several other types of steroids at similar concentrations did not cause the same inhibition of 11-KT production as estradiol in the present study suggests that this effect is specific to estrogens. We have previously demonstrated that the testis is a site of estrogen production in male Atlantic croaker [3]. Interestingly, a variety of xenobiotic estrogens, which previously have been shown to impair male reproductive function, also significantly impaired Atlantic croaker androgen production in vitro. Taken together, these results suggest that the inhibition of androgen production from Atlantic croaker testis by estrogenic compounds is of both physiological and toxicological significance.
Although a variety of other nonestrogenic steroids were incapable of altering 11-KT production—including a synthetic androgen, progesterone, and cortisol—the teleost maturation-inducing steroid 20ß-S caused an equivalent decrease in 11-KT production at a concentration 10-fold lower than that of estradiol. Recently, 20ß-S membrane receptors have been characterized in the testes of Atlantic croaker and spotted seatrout (Cynoscion nebulosus), a closely related species [35, 36]; and the possibility exists that activation of the 20ß-S membrane receptor in croaker testes by the teleost maturation-inducing steroids also results in a decrease in androgen production. The relative binding affinity of estradiol for the 20ß-S membrane receptor is low, <0.43% [36], so it is unlikely that estradiol is also acting via this receptor.
It is generally accepted that estradiol exerts its inhibitory effect on gonadotropin-stimulated androgen production in mammals by altering the activity of steroidogenic enzymes including 17-hydroxylase, C17,20-lyase (desmolase), and 17ß-dehydrogenase [21, 33, 34, 37]. However, the role of the nuclear estrogen receptor in the estrogen regulation of these enzymes in mammals remains equivocal. Melner and Abney [38] did not find a causal relationship between depletion of the testicular cytoplasmic estrogen receptor and the direct action of estradiol on LH-stimulated testosterone production in rat testes, whereas the experiments of Nozu et al. [39] suggest the estrogen nuclear receptor is involved in the estrogen-induced decrease in 17-hydroxylase/17,20-desmolase activity in rat Leydig cells.
In contrast, the results of the current study suggest that estradiol is acting on steroidogenic enzyme(s) further along the pathway for 11-KT synthesis after production of androstenedione. In addition, estradiol does not appear to be acting through the classical nuclear estrogen receptor to influence androgen production in Atlantic croaker, but instead it is acting at the cell surface probably via a membrane estrogen receptor. The time-course study shows that 5-min treatment with estradiol is sufficient to significantly decrease 11-KT production in subsequent testicular incubations. A rapid action is characteristic of a membrane, or cell-surface, steroid receptor-mediated mechanism [29, 40]. Moreover, the demonstration that the action of estradiol was not blocked by the transcription and translation inhibitors actinomycin D and cycloheximide suggests that it is acting via a nongenomic mechanism. The finding that BSA-conjugated estradiol caused a significant decrease in 11-KT production similar to that observed with nonconjugated estradiol provides further evidence that the steroid is acting at the cell surface and not via a classic nuclear estrogen receptor or by directly inhibiting the steroidogenic enzymes. Finally, the identification of a membrane receptor in the testis of Atlantic croaker provides a mechanistic explanation for these observed effects of estrogens on 11-KT production. Overall, the activities of estrogens, antiestrogens, xenoestrogens, and other steroids in inhibiting 11-KT production by croaker testes in vitro correlated with their binding affinities for the estrogen membrane receptor. Among all the compounds tested, only the maturation-inducing steroids 20ß-S and 17,20ß-P, the synthetic antiestrogen ICI 182,780, and o,p'-DDT produced unexpected results. The inhibitory actions of 20ß-S and 17,20ß-P could be mediated by binding to another membrane receptor, specific for 20ß-S and other progestogens, which has recently been characterized in the testes and sperm of Atlantic croaker and spotted seatrout [35, 36]. Our results suggest that ICI 182,780, which is a pure nuclear estrogen receptor antagonist [41], acts as an estrogen agonist when it binds to the testicular estrogen membrane receptor to inhibit 11-KT production. Interestingly, ICI 182,780 has also been shown to have agonist activities in bioassays of rapid, membrane-mediated nongenomic estrogen actions on mammalian neuroblastoma and smooth muscle cells [42, 43]. Finally, the low solubility of o,p'-DDT in aqueous solutions may be a partial explanation for its lack of activity in the testicular incubation system.
The binding moiety identified in testicular plasma membranes fulfills all the criteria for its designation as a hormone receptor. The testicular membrane preparation has a single class of high-affinity binding sites with a Kd of 1.6 nM, which is typical for steroid receptors. The binding was displaceable, saturated with low concentrations of estradiol (1–2 nM), and exhibited a low finite number of binding sites with a Bmax of 0.03 nM, equivalent to 26 fmol/g testis. This testicular membrane-binding component also exhibits rapid association with a t1/2 of 5 min, which is similar to that observed with other steroid membrane receptors [25, 35]. In addition, competition studies showed that the binding was highly specific for estrogens, antiestrogens, and several xenoestrogens, whereas no binding was observed with a variety of other steroids. Finally, the incubation studies clearly implicate this binding moiety in the estrogen regulation of androgen production, thereby indicating its possible biological significance and fulfilling the most difficult criteria for its designation as a hormone receptor.
A nuclear estrogen receptor has also recently been characterized in the Atlantic croaker testis [3]. A comparison of the rank order of the binding affinities of estrogens and antiestrogens for the two receptors and their EC50s indicates that they are broadly similar. DES showed 5 times higher binding affinity than estradiol for the nuclear receptor, whereas its affinity for the membrane receptor appeared to be similar to that of estradiol based on a limited number of competition assays. The effective concentrations of DES and estradiol for 50% displacement of radiolabeled estradiol from the membrane receptor were 20-fold and 3-fold, respectively, of those required to displace the radioligand from the testicular nuclear receptor, but similar to the values obtained for the hepatic nuclear receptor in this species [3]. Estriol and estrone displayed 10-fold lower affinities than estradiol for the membrane receptor with EC50 values similar to those for the nuclear estrogen receptor. In contrast, the antiestrogen ICI 182,780 had a much lower affinity for the membrane receptor with an EC50 approximately 35 times that for the nuclear receptor. The antiestrogen had a higher affinity than estriol and estrone for the nuclear receptor, but a lower affinity for membrane receptor. Tamoxifen showed low affinity and nafoxidine even lower affinity for both receptors. Finally, testosterone showed some affinity for the nuclear estrogen receptor, but none for the membrane estrogen receptor at a concentration of 10-5 M. Other binding characteristics such as the rate of association differ markedly between the two testicular estrogen receptors. However, the competition studies suggest that the steroid specificities of the testicular nuclear and membrane estrogen receptors in this species are similar. It will be interesting to see whether this similarity is shared at the structural level when the deduced amino acid sequence of the steroid-binding domain of a membrane estrogen receptor finally becomes available.
The results provide further evidence that xenobiotics can disrupt endocrine function by binding to steroid membrane receptors and interfering with the nongenomic actions of steroids. Recent studies in our laboratory have shown that a variety of xenoestrogens bind competitively to 20ß-S membrane receptors on the ovary, testes, and sperm of Atlantic croaker and spotted seatrout and that they can antagonize 20ß-S induction of final oocyte maturation and stimulation of sperm motility in these species [44–46]. The results of the present study show that xenobiotics can also bind to an estrogen membrane receptor and appear to act as agonists on a membrane receptor-mediated steroid action, thereby suggesting that this novel type of endocrine disruption is not limited to progestogen membrane receptors. The binding affinities of xenoestrogens for the three steroid membrane receptors identified in these fish species appear to be similar, based on the limited number of xenoestrogens investigated to date. Xenoestrogen concentrations in the 10-5–10-4 M range were effective in displacing approximately 50% of the radioligand from each of the receptors [44–46]. Nonylphenol was an effective competitor at lower concentrations (10-6 M) for the estrogen membrane receptor but has not been tested in the 20ß-S membrane receptor assays. These values for xenobiotic binding to membrane receptors are comparable to those reported for nuclear steroid receptors, including the nuclear estrogen receptor characterized in Atlantic croaker testicular and hepatic tissues [3]. Both the rank order of xenoestrogen binding for the nuclear and membrane estrogen receptors in Atlantic croaker testicular tissues and their EC50s are similar. Nonylphenol shows the highest affinity with EC50s of 5 x 10-6 M and 1.3 x 10-6 M for the membrane and nuclear estrogen receptors, respectively. Ortho, para-DDE and the hydroxylated PCB have intermediate affinities for both receptors with EC50s in the range of 1.0–5.0 x 10-5 M. A significant difference in binding affinity was observed only with o,p'-DDT, which appears to have a 10-fold lower affinity for the membrane estrogen receptor—identical to its affinity for the hepatic nuclear receptor, which, in general, demonstrates 2–5 times lower affinity for estrogens and xenoestrogens than the testicular nuclear receptor [3]. Taken together, these results suggest that steroid hormone actions mediated by binding to steroid membrane receptors are just as susceptible as nuclear steroid receptors to disruption by several major environmental estrogens and warrant further study.
In conclusion, these data show that estradiol, as well as several xenobiotics, can influence testicular function in Atlantic croaker by altering androgen production. The proposed mechanism for this action is through a cell-surface estrogen receptor. The identification of a testicular estradiol membrane receptor not only provides a mechanistic explanation for the observed effect of estradiol, but also indicates a potential additional site of xenoestrogen disruption of male reproductive function.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 This study was funded by NIEHS Grant No. ESO 4214 and EPA STAR Grant No. R826125. 
2 Correspondence: Peter Thomas, UTMSI, 750 Channel View Drive, Port Aransas, TX 78373-5015. FAX: 361 749 6777; thomas{at}utmsi.utexas.edu
Accepted: November 19, 1999.
Received: June 25, 1999.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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-hydroxylase/C17–20 lyase in the fetal rat testis is reduced by maternal exposure to exogenous estrogens. Endocrinology 1996; 137:1063–1070.[Abstract],20ß,21-trihydroxy-4-pregnen-3-one in ovaries of spotted seatrout Cynoscion nebulosus. Gen Comp Endocrinol 1990; 78:204–217.[CrossRef][Medline]-hydroxylase/C-17,20-lyase by estrogens. Endocrinology 1981; 109:763–767.[Abstract]This article has been cited by other articles:
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  J. Teng, Z.-Y. Wang, E. R. Prossnitz, and D. E. Bjorling The G Protein-Coupled Receptor GPR30 Inhibits Human Urothelial Cell Proliferation Endocrinology, August 1, 2008; 149(8): 4024 - 4034. [Abstract] [Full Text] [PDF]
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Y. Pang, J. Dong, and P. Thomas Estrogen Signaling Characteristics of Atlantic Croaker G Protein-Coupled Receptor 30 (GPR30) and Evidence It Is Involved in Maintenance of Oocyte Meiotic Arrest Endocrinology, July 1, 2008; 149(7): 3410 - 3426. [Abstract] [Full Text] [PDF]
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 L. Remage-Healey and A. H. Bass Plasticity in Brain Sexuality Is Revealed by the Rapid Actions of Steroid Hormones J. Neurosci., January 31, 2007; 27(5): 1114 - 1122. [Abstract] [Full Text] [PDF]
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 Y. Wu, W. G Foster, and E. V Younglai Rapid effects of pesticides on human granulosa-lutein cells Reproduction, February 1, 2006; 131(2): 299 - 310. [Abstract] [Full Text] [PDF]
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 A.L. Filby and C.R. Tyler Molecular Characterization of Estrogen Receptors 1, 2a, and 2b and Their Tissue and Ontogenic Expression Profiles in Fathead Minnow (Pimephales promelas) Biol Reprod, October 1, 2005; 73(4): 648 - 662. [Abstract] [Full Text] [PDF]
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P. Thomas, Y. Pang, E. J. Filardo, and J. Dong Identity of an Estrogen Membrane Receptor Coupled to a G Protein in Human Breast Cancer Cells Endocrinology, February 1, 2005; 146(2): 624 - 632. [Abstract] [Full Text] [PDF]
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A. M. Braun and P. Thomas Biochemical Characterization of a Membrane Androgen Receptor in the Ovary of the Atlantic Croaker (Micropogonias undulatus) Biol Reprod, July 1, 2004; 71(1): 146 - 155. [Abstract] [Full Text] [PDF]
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L. Remage-Healey and A. H. Bass Rapid, Hierarchical Modulation of Vocal Patterning by Steroid Hormones J. Neurosci., June 30, 2004; 24(26): 5892 - 5900. [Abstract] [Full Text] [PDF]
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 C. S. Watson and B. Gametchu Proteins of Multiple Classes May Participate in Nongenomic Steroid Actions Experimental Biology and Medicine, December 1, 2003; 228(11): 1272 - 1281. [Abstract] [Full Text] [PDF]
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