Biol Reprod Email Content Delivery
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

BOR - Papers in Press, published online ahead of print February 25, 2004.
Biol Reprod 2004, 10.1095/biolreprod.103.025825
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
71/1/146    most recent
biolreprod.103.025825v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Braun, A. M.
Right arrow Articles by Thomas, P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Braun, A. M.
Right arrow Articles by Thomas, P.
Agricola
Right arrow Articles by Braun, A. M.
Right arrow Articles by Thomas, P.
BIOLOGY OF REPRODUCTION 71, 146–155 (2004)
DOI: 10.1095/biolreprod.103.025825
© 2004 by the Society for the Study of Reproduction, Inc.

Mechanisms of Hormone Action

Biochemical Characterization of a Membrane Androgen Receptor in the Ovary of the Atlantic Croaker (Micropogonias undulatus)1

Alyssa M. Braun2, and Peter Thomas

University of Texas at Austin, Marine Science Institute, Port Aransas, Texas 78373


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Membrane androgen receptors have been biochemically characterized in only a few vertebrate species to date. Therefore, the purpose of the current study was to comprehensively investigate the binding characteristics of a putative membrane androgen receptor in the ovary of the teleost, Atlantic croaker (Micropogonias undulatus). Specific androgen binding to an ovarian plasma membrane fraction was demonstrated using a radioreceptor assay protocol consisting of a short-term incubation with [3H]testosterone (T) and subsequent filtration of bound steroid from free steroid. Saturation and Scatchard analyses of T binding to an ovarian plasma membrane fraction indicated the presence of a single, high-affinity (Kd = 15.32 ± 2.68 nM [mean ± SEM]), low-capacity (Bmax = 2.81 ± 0.31 pmol/mg protein), androgen-binding site. Specific androgen binding to the receptor was readily displaceable, and the association and dissociation kinetics were rapid (half-time = 3.7 ± 1.7 and 4.7 ± 0.2 min, respectively). Competitive binding assays showed that 5{alpha}-dihydrotestosterone, T, and 11-ketotestosterone had relative binding affinities (RBAs) of 193%, 100%, and 13%, respectively, whereas none of the C18 or C21 steroids tested bound with high affinity except for progesterone (RBA = 191%). This androgen-binding moiety with high affinity for progesterone is unlikely to mediate the physiological actions of progestins in croaker, because it has low binding affinity for fish progestin hormones. Androgen-binding sites were also detected in membrane fractions of the brain, liver, kidney, and drumming muscle, whereas little or no binding was detected in the trunk muscle, heart, gills, or intestine. Receptor levels increased 10-fold during ovarian recrudescence, reaching maximum levels in fully mature ovaries, which suggests a likely physiological role for this receptor during the reproductive cycle of female croaker. It is concluded that the androgen-binding moiety identified in the plasma membrane fraction of Atlantic croaker ovarian tissue fulfils all the criteria for its designation as a steroid receptor.

androgen receptor, ovary, steroid hormone receptors, steroid hormones, testosterone


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Steroid hormones are known to regulate cellular processes through two distinct receptor mechanisms. The classically defined, genomic pathway is initiated after the steroid molecule diffuses through the plasma membrane and binds to an intracellular transcription factor belonging to the nuclear steroid receptor superfamily. Subsequently, the steroid/receptor complex translocates to the nucleus and activates or represses the transcription of target genes [1, 2]. Because this mechanism of hormone action involves the genomic processes of transcription and translation, the effects of steroids are not immediate and, typically, become evident only on a time scale of hours. However, many actions of steroids are too rapid to be explained by this mechanism of steroid action. Alterations in the concentrations of intracellular free calcium and other second-messenger molecules can occur within seconds to a few minutes after the application of steroids to the cell surface (for review, see [3]). The subsequent cellular response typically does not involve changes in the rates of transcription. These rapid, cell surface-mediated, nongenomic steroid actions (and, in some cases, the membrane binding sites that mediate them) have been characterized for all classes of steroids in several different tissue types of many vertebrate species (for reviews, see [3–5]).

Overall, rapid actions of androgens and membrane androgen receptors (mARs) have received less attention than those of most other classes of steroids. Nevertheless, several rapid, nongenomic effects of androgens have been described. Intracellular free Ca2+ levels are rapidly regulated by androgen treatment in a variety of tissues and cell-culture systems, including both male and female reproductive tissues [6–9], skeletal muscle [10], osteoblasts [11, 12], and T cells and macrophages [13–15]. Other signaling pathways rapidly activated by androgens include mitogen-activated protein kinases (MAPKs) and protein kinase C [16, 17]. Although these studies have provided indirect evidence for the existence of membrane-associated androgen-binding sites, in many cases the receptors mediating these rapid androgen actions have not been characterized.

Androgen-binding sites have been partially characterized in the membranes of mammalian endothelial cells [18], synaptosomes in the rat [19, 20], olfactory tissues from brown and rainbow trout [21], and testis [22], liver, and prostate from male rats [23]. Through the use of confocal laser-scanning microscopy and flow cytometry, putative testosterone (T) receptors were also identified on the plasma membranes of T cells and macrophages [14, 15]. However, these androgen-binding sites have not been extensively investigated. Therefore, comprehensive information is currently lacking on the binding characteristics of androgen membrane receptors.

Previously, we described a rapid, nongenomic action of androgens on ovarian steroidogenesis in the Atlantic croaker (Micropogonias undulatus) [24]. Evidence was obtained that androgens inhibit in vitro gonadotropin-stimulated production of estradiol by croaker ovaries through a mechanism initiated at the cell surface. Moreover, preliminary binding experiments showed the existence of an androgen-binding site in an ovarian membrane preparation. Previously, a nuclear androgen receptor was identified in croaker ovaries, and its steroid-binding characteristics were described in detail [25–27]. The purpose of the current study was to investigate fully the binding characteristics of the ovarian membrane binding moiety to determine whether it fulfills all the criteria for its designation as a steroid receptor and whether its steroid-binding characteristics are similar to those of the nuclear androgen receptor in this tissue.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals

The radioactive tracer, [1,2,6,7-3H]T (specific activity, 95 Ci/mmol), and the synthetic androgen, 17ß-hydroxy-17{alpha}-methyl-4,9,11-estratiene-3-one (R1881), were purchased from New England Nuclear (Boston, MA). Other unlabeled steroids were purchased from Steraloids (Newport, RI) or Sigma (St. Louis, MO). The synthetic androgen, mibolerone, was a gift from Upjohn Laboratories (Kalamazoo, MI). The antiandrogens, cyproterone acetate and flutamide, were purchased from Sigma. Vinclozolin and its metabolites (M1 and M2) were a gift from L. Earl Gray (U.S. Environmental Protection Agency, Research Triangle Park, NC). All para,para- and ortho,para-congeners of dichlorodiphenyltrichloroethane (DDT), dichlorodiphenyldichloroethylene (DDE), and dichlorodiphenyldichloroethane (DDD) were obtained from Supelco (St. Louis, MO). The steroids, antiandrogens, and xenobiotics were dissolved and stored in 95% ethanol at –20°C. The diagnostic kit for the measurement of 5'-nucleotidase activity, Dulbecco modified Eagle medium (DMEM; nutrient mixture, F-12 HAM), and all other chemicals used to prepare the buffers were of reagent grade and purchased from Sigma. The scintillation cocktail was prepared by dissolving 16 g of 7,5-diphenyl-oxazole (Sigma) and 0.4 g of 1,4-bis[5-phenyl-2-oxazolyl]-benzene (Sigma) in 4 L of reagent-grade toluene.

Animals and Tissue Collection

Atlantic croaker were captured by gill net or otter trawl from the bays near Port Aransas, TX, during late summer and early fall (1998–2001), which is the period of gonadal recrudescence. Fish were maintained in circular, recirculating tanks under constant photoperiod and temperature (22–25°C) and fed a diet of commercial pellets daily. Before tissue collection, fish were acclimated to laboratory conditions for at least 3 wk. The gonadosomatic index (GSI), calculated by the equation GSI (%) = (gonad weight/somatic weight) x 100, was used to determine the stage of gonadal maturity. The average GSI for females in the present study except for the seasonal experiment, was 14.26% ± 0.74%. The average male GSI was 6.56% ± 1.22%.

Fish were humanely killed by rapid decapitation following procedures approved by the University of Texas Animal Care and Use Committee according to NIH guidelines. Tissues were immediately removed and placed in ice-cold buffer or frozen at –80°C for later analysis. Androgen binding remained constant in frozen tissues for at least 1 yr.

Tissue Preparation

Two-gram aliquots of individual or pooled ovaries were homogenized in 15 ml of HED buffer (25 mM Hepes, 10 mM NaCl, 1 mM dithioerythritol, and 1 mM EDTA, pH 7.6) at 4°C using a Polytron Tekmar Tissumizer (Tekmar, Cincinnati, OH) on a high setting for 5 sec followed by three to five passes with a glass Tenbroek tissue grinder (Wheaton Science Products, Millville, NJ). All the remaining preparation steps were conducted at 4°C. The homogenate was centrifuged at 1000 x g for 7 min to pellet the nuclear material, and the resulting supernatant was centrifuged at 20 000 x g for 20 min to pellet the membrane fraction. The supernatant from this spin was considered to be the crude cytosolic fraction. To further purify the membrane fraction, the membrane pellet was resuspended in 10 ml of HED, layered over an equal volume of 1.2 M sucrose dissolved in HED, and centrifuged at 6900 x g for 45 min based on the method of Peck and Kelner [28]. After this spin, the middle membrane layer was removed with a Pasteur pipette, diluted (1:2) with HED, and centrifuged at 20 000 x g for 20 min. The pellet of this final spin was resuspended in 5 ml of HED and either used immediately in a binding assay or frozen at –80°C for later analysis. Protein concentrations were determined using the Bio-Rad Protein Assay (Bio-Rad Laboratories, Hercules, CA).

Radioreceptor-Binding Assays

All steps of the binding assays were conducted at 4°C. The membrane preparations were diluted to 0.15–0.5 mg protein/ml in HED immediately before all binding assays. Total binding saturation curves were generated by incubating 250 µl of membrane preparation and 250 µl of [3H]T, dissolved in HED for final reaction concentrations ranging from 0.3 to 25 nM, for 40 min. Nonspecific binding was determined from a parallel set of reaction tubes, which also contained 1000-fold excess unlabeled T. The same procedure was used in one-point binding assays except that the membrane preparations were incubated with only one concentration of [3H]T. After the 40-min incubation, the binding reactions were terminated by rapidly filtering 400 µl of the reaction over a presoaked Whatman, glass-fiber filter (pore size, 1 µm) to separate bound steroid from free steroid. The filter was immediately washed twice with 12.5 ml of wash buffer (HED without dithioerythritol) and placed in a scintillation vial. Radioactivity was counted in a liquid scintillation counter (LSC 6000SC; Beckman Instruments, Fullerton, CA). Specific [3H]T binding was calculated as the difference between total and nonspecific binding, and in saturation experiments, specific binding was transformed according to the method of Scatchard [29]. The number of receptors (Bmax) and the affinity constant (Kd) were determined from the nonlinear regression of the specific [3H]T binding curve.

Verification of the Plasma Membrane Preparation

To verify that the androgen binding was in the membrane fraction of the ovarian tissue, 5'-nucleotidase activity was measured as a marker of the plasma membrane according to the protocol provided with the kit by Sigma (procedure no. 265-UV). In addition, each fraction of the ovarian homogenate was diluted to 0.2–0.4 mg protein/ml, and binding was determined subsequently by one-point assays.

Association and Dissociation Rates

The rates of association and dissociation of androgen binding to ovarian membrane preparations were determined at 4°C. For association assays, membrane preparations of individual ovaries or a pool of three ovaries were incubated with 8 or 10 nM [3H]T for various time periods ranging from 1 min to 23 h. A parallel set of reaction tubes contained 1000-fold excess unlabeled T to determine nonspecific binding. Dissociation rates were determined by incubating membrane preparations with 10 nM [3H]T (total binding) or with 10 nM [3H]T plus 10 µM unlabeled T (nonspecific binding) for 40 min before the addition of 1000-fold excess unlabeled T. Reactions were stopped by filtration at various time periods ranging from 1 min to 2 h after addition of the unlabeled T.

Competition Studies

Steroid competitors were dissolved in ethanol, added to the reaction tubes for final reaction concentrations of 0.1 nM to 10 µM, and evaporated under nitrogen. Next, 250 µl of membrane preparation diluted to 0.15–0.5 mg protein/ml in HED and 250 µl of [3H]T (final reaction concentration, 10 nM) were added to the reaction tubes and incubated at 4°C for 40 min. Maximum specific [3H]T binding was defined as the difference between total binding at 10 nM [3H]T (no competitor present) and nonspecific binding (in the presence of 1000-fold excess unlabeled T). Antiandrogens and xenobiotics were tested in competition assays using the same method; however, the range of competitor concentrations was 100 nM to 1 mM. The DDT congeners were tested in the presence of two different concentrations of [3H]T (5 and 10 nM). The concentration of competitor needed to displace 50% of maximum specific [3H]T binding (EC50) was calculated from the one-site competitive binding curve. In this analysis, the top and the bottom of the curve were defined as 100% and 0%, respectively, and held constant. The relative binding affinity (RBA) was calculated from the ratio of the EC50 of T to the EC50 of the competitor and expressed as a percentage (T = 100%).

Tissue Specificity

Various tissue types were collected from female croaker, and the membrane fraction of each tissue was prepared as described previously. Androgen binding in the membrane fractions of the ovary, liver, brain, drumming muscle, kidney, trunk muscle, intestine, gills, and heart was detected with semiquantitative, one-point assays. Full saturation curves were performed on membrane fractions from brain and testicular tissues.

Biological Relevance: Effects of Stage of Ovarian Development and Hormonal Treatments on Receptor Concentrations

Samples of croaker ovaries were collected throughout the reproductive cycle, and membrane fractions were prepared as described previously. The GSI for each fish was calculated and used as a measure of the reproductive stage of the ovary. Androgen binding in these seasonal samples was detected by one-point assays. In addition, the effects of in vitro steroid treatment on receptor concentrations in mature ovaries were investigated. Aliquots (~1 g) of fresh, mature, ovarian tissue were incubated in 50 ml of DMEM containing 500 nM concentrations of various steroids for 11 h at room temperature. After the incubation, ovarian fragments were removed from the steroid treatment and frozen at –80°C for later analysis. Membrane fractions were prepared as described previously and analyzed for androgen binding by one-point assays.

Statistical Analysis

Values are reported as the mean ± SEM. All linear and nonlinear regression analyses were generated by GraphPad Prism for Windows (Version 3.02; GraphPad Software, San Diego, CA; http://www.graphpad.com). Data were transformed when necessary to conform to assumptions of normality and homoscedasticity, then analyzed by one-way ANOVA and Tukey tests for multiple comparisons (Sigma Stat; SPSS, Chicago, IL).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Determination of Saturation and Verification of Binding in the Membrane Fraction

Nonlinear regression of the specific [3H]T binding curves (Fig. 1) indicates the presence of a high-affinity (Kd = 15.3 ± 2.7 nM), low-capacity (Bmax = 2.8 ± 0.3 pmol/ mg protein or 9.5 ± 1.6 pmol/g tissue), androgen-binding site in the membrane fraction of ovarian tissue (n = 5 preparations, R2 ≥ 0.92). Linear Scatchard transformations indicate a one-site model of androgen binding (Fig. 1, inset).



View larger version (16K):
[in this window]
[in a new window]
  		FIG. 1. Representative saturation curve of [3H]T binding to a membrane preparation of croaker ovarian tissue with Scatchard analysis (inset). The membrane preparation was incubated at 4°C with 0.3–25 nM [3H]T for 40 min before separation by filtration. Specific [3H]T binding (SB) was calculated as the difference between total binding (TB) and nonspecific binding (NSB)

To verify that the androgen binding was located in the membrane preparation, 5'-nucleotidase activity was measured in all subcellular fractions. Enzyme activity was approximately 10-fold higher in the membrane fraction than in the crude nuclear and cytosolic fractions (Fig. 2a). In addition, one-point assays of all fractions indicate that androgen binding was primarily located in the membrane fraction, with values in the cytosolic and nuclear fractions less than 10% of the binding in the membrane fraction (Fig. 2b). Binding was also significantly reduced in boiled membrane fractions (P = 0.01).



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 2. Verification of androgen binding in the membrane fraction of an ovarian homogenate. a) 5'-Nucleotidase activity was measured in all fractions of the ovarian homogenate, and units of activity (U) were normalized to the amount of protein present. b) One-point binding assays at 5 nM [3H]T were conducted on all fractions, including a boiled membrane fraction (Boil Mem). Specific [3H]T binding was calculated as the difference between total binding and nonspecific binding. Values are the mean ± SEM. The number of samples tested is stated above each group. Asterisks denote means significantly different from the membrane fraction (P = 0.05)

Association and Dissociation Rates

The results of the association and dissociation experiments indicated that androgen binding in the membrane fraction was relatively rapid. The half-time of association was 3.7 ± 1.7 min, with maximum binding achieved after 30 min (n = 2) (Fig. 3a). Androgen binding remained constant for at least 23 h. Dissociation was also rapid, with a half-time of 4.7 ± 0.2 min and complete dissociation after 40 min (n = 2) (Fig. 3b). The half-time values were calculated by nonlinear regression analysis of the specific [3H]T binding of association or dissociation (R2 ≥ 0.82).



View larger version (12K):
[in this window]
[in a new window]
  		FIG. 3. Representative association and dissociation curves of androgen binding to a membrane preparation of croaker ovarian tissue at 4°C. a) The rate of association was determined in the presence of 10 nM [3H]T. b) The rate of dissociation was determined after the addition of 1000-fold excess unlabeled T to the binding reaction containing 10 nM [3H]-T. Half-time values were calculated from the nonlinear regression of the specific [3H]T binding curve, which was calculated as the difference between total and nonspecific binding

Competition Studies

Curves used to calculate RBA values (Table 1) accurately reflected the data; coefficients of determination (R2) were greater than or equal to 0.87 except for one assay with estradiol, which had an R2 of 0.52 (n = 2–5). In steroid specificity experiments that tested the binding affinity of steroid hormones found in fish, only T and 11-ketotestosterone substantially displaced 10 nM [3H]T within physiological concentrations (Fig. 4a). The binding affinity of estradiol was two orders of magnitude lower than that of T. Further testing of androgens indicated that in addition to T and 11-ketotestosterone, dihydrotestosterone (DHT) competed for binding, demonstrating approximately twice the affinity of T (Fig. 4b). No other androgens tested effectively competed for binding within this range of concentrations. All the progestogens tested had RBA values ≤ 1.1% except for progesterone (RBA = 191%) (Fig. 4c). The parallel binding curves of T and progesterone suggest that binding is competitive. The antiandrogens flutamide and cyproterone acetate had RBA values of 0.13% and 0.01%, respectively (Fig. 5a). Of the xenobiotics tested, vinclozolin and its metabolites (M1 and M2) displaced [3H]T (Fig. 5a), whereas the highest concentrations of the DDT congeners tested did not compete for binding (Fig. 5b).


View this table:
[in this window]
[in a new window]
  		TABLE 1. Relative binding affinities (RBA) of various competitors for the ovarian membrane androgen receptor



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 4. Representative competition curves of steroid competition for androgen binding in an ovarian membrane fraction. Steroid competition assays of (a) common fish steroids, (b) androgens, and (c) progestogens were conducted in the presence of 10 nM [3H]T (see Table 1 for abbreviations). Competition curves were generated with undefined top and bottom values



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 5. Representative competition curves of antiandrogen and xenobiotic competition for androgen binding in an ovarian membrane preparation. a) The antiandrogenic compounds of flutamide, cyproterone acetate and the xenobiotic fungicide, vinclozolin and its metabolites (M1 and M2) were tested in competition with 5 nM [3H]T. b) The ortho,para- and para,para- congeners of DDT were tested in competition assays with 5 or 10 nM [3H]T. Competition curves were generated with undefined top and bottom values

Tissue Specificity

Substantial androgen binding was detected in membrane fractions of the ovary, brain, drumming muscle, and kidney from female croaker with the semiquantitative, one-point assay, whereas little binding was detected in the trunk muscle, intestine, gills, and heart (Fig. 6). Results of the one-point binding assay with liver tissue indicated very high levels of androgen binding (645 dpm/µg protein). However, the membrane layer from the sucrose pad procedure was not clearly defined in the preparations of the liver; therefore, the accuracy of the binding in this tissue is uncertain.



View larger version (10K):
[in this window]
[in a new window]
  		FIG. 6. Tissue distribution of androgen binding in membrane preparations. Androgen binding in various tissue types was determined in one-point binding assays with 9 nM [3H]T. Specific [3H]T binding, the difference between total and nonspecific binding, was normalized to the amount of protein in the preparations. Binding values are the results of one assay of a pool of six fish each for brain, kidneys, intestine, gills, and drumming muscle. Binding values for ovary and trunk muscle are the means of two different pools of three fish each

Saturation analysis of pooled brain tissue of both males and females indicates that androgen binding in the membrane fraction of the brain is of significantly higher affinity (Kd = 1.55 ± 0.51 nM) and lower capacity (Bmax = 0.60 ± 0.003 pmol/mg protein or 3.12 ± 0.63 pmol/g tissue) compared to the binding in the ovary (n = 3, R2 ≥ 0.80 for the nonlinear regression of the specific [3H]T binding curves, P < 0.01) (Fig. 7a). Scatchard analysis indicates a one-site model of binding (Fig. 7a, inset). In contrast, the affinity of binding in the testes was lower than that in the ovary (Kd = 27.94 ± 2.01 nM, R2 ≥ 0.98 for the nonlinear regression of the specific [3H]T binding curves, P < 0.05, n = 2). Androgen-binding capacity in the membrane fraction of testicular tissue was significantly lower (Bmax = 0.36 ± 0.03 pmol/mg protein or 1.56 pmol/g tissue) as compared to the ovary (P < 0.01) (Fig. 7b).



View larger version (19K):
[in this window]
[in a new window]
  		FIG. 7. Representative saturation curves of androgen binding in the membrane fraction of brain and testicular tissues. a) The membrane preparation of brain tissue was incubated with 0.01–4 nM [3H]T. b) The testicular membrane fraction was incubated over the range of 0.25–27 nM [3H]T. Specific [3H]T binding (SB) was calculated as the difference between total binding (TB) and nonspecific binding (NSB). Scatchard transformations are inset on both graphs

Biological Relevance: Effects of Stage of Ovarian Development and Hormonal Treatments on Receptor Concentrations

The amount of androgen binding in croaker ovarian membranes was dependent on the reproductive stage of the fish (Fig. 8a). One-point assays indicate that binding was 10-fold greater in fully mature ovaries than in regressed tissue. Androgen-binding concentrations in ovarian fragments were not altered after an 11-h treatment with steroid hormones in vitro (Fig. 8b). One-way ANOVA detected no significant differences among the different treatment groups (P > 0.05).



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 8. Seasonal changes and in vitro regulation of membrane androgen receptor concentrations in croaker ovaries. a) Specific androgen binding in seasonal samples of croaker ovaries was determined by one-point assays with 15 nM [3H]T. The GSI = 0.94% ± 0.05% for regressed ovaries, 3.66% ± 0.70% for early recrudescing ovaries (Recrud.), and 18.61% ± 1.76% for fully mature ovaries (Mature). Asterisk denotes significant difference from recrudescing and mature ovaries (n = 4 fish/reproductive stage). b) Ovarian aliquots were incubated with 500 nM T, DHT, estradiol, or control treatments for 11 h (n = 4 fish/treatment). One-point assays of the isolated membrane fractions with 15 nM [3H]T were used to determine androgen binding. Specific [3H]T binding was calculated as the difference between total binding and nonspecific binding. Values are the mean ± SEM


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The results of the present study indicate the presence of a low-capacity, high-affinity mAR in the ovary of the Atlantic croaker. The androgen binding corresponds to the presence of 5'-nucleotidase, a marker of the plasma membrane, and is not detected in the nuclear or cytosolic fractions of the tissue homogenate using the filtration-binding assay. The association and dissociation of binding are rapid, with half-times under 5 min, which are values typical of membrane steroid receptors. The results of the competition experiments suggest that this binding moiety differs from most androgen receptors characterized to date in that it also binds progesterone but not synthetic androgens that have high affinities for nuclear androgen receptors. The mAR shows tissue specificity with high levels of binding in reproductive tissues such as the ovary and also in the brain. Binding was also detected in the kidney and drumming muscle, whereas very little binding was detected in the heart, trunk muscle, gills, or intestine. Ovarian mAR levels are maximal in reproductively mature gonads, which suggests this receptor has a physiological role during the female reproductive cycle. The demonstration of high affinity, saturable and reversible binding, steroid specificity, tissue specificity, and biological relevance fulfills the criteria for this membrane androgen-binding site to be designated as a steroid receptor.

To our knowledge, only a few reports describe partial biochemical characterizations of androgen-binding sites associated with cell membranes. The brown trout (Salmo trutta) olfactory mAR [21] demonstrates higher binding affinity (Kd = 0.5–1.9 nM) and lower capacity (Bmax = 30–60 fmol/mg protein) than the croaker ovarian mAR (Kd = 15.3 nM, Bmax = 2.8 pmol/mg protein). In contrast, the androgen-binding site identified in the membrane fraction of rat brain tissue [20] demonstrated an affinity of the same order of magnitude (Kd = 31.8 nM) as the croaker ovarian mAR, although it had a considerably higher binding capacity (202 pmol/mg protein). Thus, there appear to be marked species and/or tissue differences in the binding affinities and capacities of the few membrane androgen-binding moieties investigated so far, although the physiological significance of these differences remains unclear.

The rapid association and dissociation of membrane androgen binding in the croaker ovary is similar to that of other membrane androgen-binding sites, which show complete association after 16 min in brain tissue [20] and after 45 min in fish olfactory tissue [21]. Additionally, both the membrane maturation-inducing steroid (MIS) receptor in spotted seatrout (Cynoscion nebulosus) and the membrane estrogen receptor in Atlantic croaker associate rapidly, reaching equilibrium within 5 and 30 min, respectively [30, 31]. In contrast, complete androgen association to the nuclear androgen receptor in the croaker ovary is only observed after an 8-h incubation [25]. The rapid rate of association of ligands to steroid membrane receptors would theoretically enable steroids acting at the cell surface to initiate cellular responses within a few minutes and, therefore, is consistent with the rapid changes in the intracellular levels of second-messenger molecules frequently reported within minutes of steroid application.

The steroid specificity of the croaker ovarian mAR is unusual in several respects. To our knowledge, the high affinity of progesterone for this receptor has not been observed in other studies of androgen binding to membrane preparations, in which progesterone failed to compete for androgen binding [19, 20, 23]. Furthermore, the common synthetic androgen, R1881, does not bind to the croaker ovarian mAR, which is similar to the lack of R1881 competition for androgen binding in membranes prepared from of rat liver and prostate cells [23]. Interestingly, a nuclear androgen receptor has been described in the shark testis that binds progesterone, but not R1881, with high affinity [32]. Moreover, it was suggested that this shark androgen receptor is related to a common ancestral nuclear-receptor molecule. The possible phylogenetic significance of the high binding affinity of progesterone to the croaker ovarian mAR cannot be determined in the absence of information regarding the structures of steroid membrane receptors. However, from a physiological standpoint, this binding is unlikely to be significant, because progesterone is not present at physiologically relevant concentrations in fish. The mAR appears to mediate only the actions of androgens, because the progestins 17,20ß,21-trihydroxy-4-pregnen-3-one and 17,20ß-dihydroxy-4-pregnen-3-one, the maturation-inducing steroids in fish, do not bind to this receptor. The lack of binding of two other progestins, 17-hydroxy-4-pregnene-3,20-dione and 20ß-hydroxy-4-pregnen-3-one, to the receptor further indicates it is not a progestin receptor and has a distinctly different steroid specificity from that of progestin membrane receptors.

Possible binding of antiandrogens to other mARs can only be inferred indirectly from studies of their effects on rapid androgen actions. For example, cyproterone acetate and flutamide do not reverse the rapid effects of androgens on intracellular calcium in macrophages [11, 15], suggesting that these compounds do not act as androgen-receptor antagonists in this system. However, both flutamide and cyproterone acetate displace T from the croaker ovarian mAR at high concentrations, indicating that these compounds, acting as either agonists or antagonists, could potentially alter androgen signaling via this receptor. Interestingly, flutamide has been shown to mimic the nongenomic actions of androgens on MAPKs in human breast cancer cells [17].

Androgen binding to plasma membrane fractions was tissue specific in the Atlantic croaker. In addition to the ovary, significant binding was detected in other tissues, including the brain, where rapid, nongenomic effects of androgens have been demonstrated previously in other vertebrates [33–36]. Studies concerning nongenomic steroid actions in the brain have shown that steroids are capable of modulating the actions of gamma-aminobutyric acid (GABA) by noncompetitively binding to its receptor (for review, see [37]). Evidence suggests that the androgen, 5{alpha}-androstane-3{alpha},17ß-diol, inhibits sexual behavior via this mechanism in rats [38]. In croaker, both GABA and androgens decrease LH secretion in fully mature females [39, 40]. However, to our knowledge, competition studies to determine whether androgens and GABA bind to the same receptor in croaker brain have not been conducted. Therefore, it is not known whether androgens could potentially influence LH secretion through binding to the GABA-receptor complex in this species. Interestingly, T binding in the croaker brain was of a higher affinity and lower capacity compared to that of the ovarian mAR, which suggests these two binding sites may represent two distinct receptors or the same receptor in two different conformations. Additionally, preliminary research indicates that in contrast to the ovarian mAR, T binds to the membrane site in the brain with higher affinity than DHT (unpublished results), which is similar to the higher binding affinity of T to the nuclear receptor identified in the croaker brain [25]. However, a complete characterization of the androgen-binding site in the brain will be necessary to confirm the existence of two distinct croaker mARs.

The presence of specific androgen-binding sites in croaker testicular membranes suggests the testis is also a site of nongenomic androgen actions initiated at the cell surface, although the concentrations of androgen-binding sites in the testis were significantly lower than those in the ovaries and brain. The binding capacity of the membrane binding site in croaker testis was 360 fmol/mg protein, which is similar to the levels of membrane androgen binding observed in the rat testes (Bmax = 328 fmol/mg protein) [22]. However, the rat testicular binding site had a higher binding affinity (Kd = 1.6 nM) than that of the croaker testicular binding site (Kd = 27.94 nM). Although the cellular localization and physiological role of the croaker testicular membrane androgen-binding site is yet unknown, androgens have been shown to rapidly alter Ca2+ signaling in rat Sertoli cells [6, 7].

One-point assays revealed that androgen binding was also present in membrane fractions of croaker kidneys and drumming (sonic) muscle, although the saturation points and binding affinities of these binding sites were not determined. It is interesting to note that a role of androgens has been established in both these tissues in certain species of fish. Androgen treatment increases the sonic muscle mass in male weakfish (C. regalis), a species closely related to Atlantic croaker, mimicking the changes observed in this species during the reproductive season [41]. Androgens also cause hypertrophy of the stickleback kidney, which is associated with production of the nest-building glue protein, spiggin [42, 43]. To our knowledge, the type of receptor mediating these androgen actions has not yet been determined. Therefore, the involvement of nuclear androgen receptors and/or mARs remains a possibility in these tissues, although an mAR has yet to be identified in the stickleback kidney.

Because steroid membrane receptors have not been as thoroughly characterized as their nuclear counterparts, relatively little information is available regarding their binding affinities for xenobiotic chemicals. Certain metabolites of the pesticide DDT were found to bind to a membrane estrogen receptor in the testis of the Atlantic croaker [31], and o,p'-DDE was found to bind to the membrane progestogen receptor on croaker sperm [44]. In addition, o,p'-DDD was found to bind to the membrane MIS receptor in croaker ovaries and to disrupt the process of final oocyte maturation (for review, see [45]). However, the DDT metabolites, including p,p-DDE, a xenobiotic antiandrogen [46, 47] that binds to the nuclear androgen receptor in the croaker ovary [26], fail to displace T binding from the ovarian mAR at high concentrations. Other xenobiotics that compete for binding to nuclear androgen receptors in mammals and Atlantic croaker ovaries include the metabolites of the fungicide vinclozolin, M1 and M2 [26, 48]. In contrast to the results with the DDT metabolites, M1 and M2 were capable of binding to the croaker ovarian mAR, which suggests that these xenobiotics have the potential to disrupt androgen signaling through this receptor.

Currently, much research is focused on the molecular nature of the membrane steroid receptors. Unlike the nuclear receptors, current evidence suggests that the proteins mediating the rapid, nongenomic effects of steroid hormones cannot be classified into a single superfamily of shared molecular characteristics. In some tissues, the membrane-binding sites appear to be structurally related to the nuclear receptors, as found in some studies of membrane estrogen binding [49, 50]. However, the binding results from the current study provide indirect evidence that the structures of the binding pockets of the nuclear androgen receptors and mARs in the croaker ovary differ. Competition studies show that the synthetic androgens, mibolerone and R1881, do not bind to the mAR, whereas these androgens compete for binding to the ovarian nuclear androgen receptor, AR2, with RBA values of 42% and 62%, respectively [25, 27]. The mAR demonstrates little affinity for 4-androstene-3,11,17-trione, whereas this steroid displays a biphasic binding curve in AR2 competition assays and binds with high affinity (RBA = 2000%; [27]). Furthermore, the high affinity of the mAR for progesterone is not a characteristic of the nuclear androgen receptor [25]. In addition, the mAR has a lower affinity and a higher capacity than the nuclear androgen receptor (Kd = 0.62 nM, Bmax = 0.38 pmol/g tissue [25]). Although these differences in binding characteristics suggest that the croaker ovarian mAR is a novel receptor, as in the case of the T receptors in the plasma membranes of T cells [14], without specific antibodies directed against these proteins or the protein or nucleic acid sequences the possibility remains that the androgen receptor on the membrane is a modification of the nuclear receptor. The finding that the croaker ovarian mAR is colocalized with the nuclear receptor described by Sperry and Thomas [25] in croaker ovarian, testicular, and brain tissues supports this possibility.

In addition to modified nuclear receptors, membrane steroid-binding sites can be novel proteins and are often G protein-coupled receptors (GPCRs) [4]. A membrane progestin receptor with the characteristics of a GPCR has been discovered in the ovary of spotted seatrout, a species closely related to the croaker [51, 52]. However, preliminary studies indicate that the croaker ovarian mAR is not coupled to a G protein (unpublished results). Alternatively, other proteins could potentially mediate the rapid, nongenomic actions of steroids initiated at the cell surface. Androgens can increase the growth of prostate cancer cells through a nongenomic mechanism mediated by the steroid hormone-binding globulin (SHBG) [53]. Whereas an SHBG has been identified in the plasma of the spotted seatrout [54], this fish SHBG demonstrated a higher affinity for androgens (Kd = 4.89 nM), a substantially higher affinity for estradiol (Kd = 3.13 nM), and a lower affinity for progesterone [54] compared to the mAR currently identified in the ovary of the Atlantic croaker. Because evidence suggests that the steroid-binding characteristics of the SHBG do not change once it has bound to its receptor on the plasma membrane [55], the mAR described in the ovary of the Atlantic croaker is unlikely to be the SHBG/receptor complex.

A rapid, nongenomic action of androgens initiated at the cell surface to down-regulate ovarian estradiol production in croaker has been described previously [24]. It is possible that the mAR currently described is responsible for mediating this nongenomic effect of androgens. The current study supports a role for the mAR in the reproductive cycle of female Atlantic croaker. Moreover, a positive relationship appears to exist between plasma sex-steroid levels and mAR concentrations during the reproductive cycle. The highest concentrations of mAR were detected in fully mature ovaries, which produce large amounts of androgens, whereas only a small amount of binding was present in regressing ovaries, which synthesize lower amounts of steroids. These results suggest that the nongenomic actions of androgens mediated by the mAR are of greatest importance at the end of ovarian and oocyte growth. Therefore, the mAR may have an important role in the changes of ovarian estradiol production observed at the end of this phase of the reproductive cycle. However, mibolerone, which is an inhibitor of estradiol production in the in vitro system [24], does not bind to the mAR, suggesting that this synthetic steroid is not acting via the mAR. In this case, mibolerone may be acting through the nuclear receptor, because only DHT was clearly shown to act at the cell surface through a rapid, nongenomic mechanism to inhibit in vitro estradiol production [24]. Therefore, it is possible that androgens decrease estradiol synthesis through binding to both membrane and nuclear receptors in the croaker ovary. The finding that the membrane receptor has a lower affinity and higher capacity than the nuclear receptor [25] raises the possibility that these two receptors are activated under different androgen concentrations and that the rapid effects of androgens in the croaker ovary are not always turned on but, rather, are initiated only at times of higher androgen concentrations.

In conclusion, an androgen-binding site has been characterized in a membrane fraction of Atlantic croaker ovaries. This moiety demonstrates high-affinity binding that is both reversible and of finite capacity. Androgen binding in the membrane is both steroid and tissue specific and likely of biological relevance in the reproductive cycle of female croaker. Thus, it fulfills all the binding requirements and should be designated as a steroid receptor. This is one of the first reports providing conclusive evidence for the existence of mARs in vertebrates.


   ACKNOWLEDGMENTS
 
The authors thank L. Earl Gray for the gift of vinclozolin and its metabolites as well as Upjohn Laboratories for the donation of mibolerone. We also thank Susan Lawson for animal care.


   FOOTNOTES
 
1 Supported by an E.J. Lund Fellowship to A.B. from the University of Texas Marine Science Institute and by Environmental Protection Agency STAR grants R826126 and R82902401 to P.T. Back

2 Correspondence and current address: Alyssa M. Braun, University of Nevada, Las Vegas, Department of Biological Sciences, 4505 Maryland Pkwy, Las Vegas, NV 89154-4004. FAX: 702 895 2396; alyssa.braun{at}ccmail.nevada.edu Back

Received: 21 November 2003.

First decision: 13 December 2003.

Accepted: 24 February 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Beato M. Gene regulation by steroid hormones. Cell 1989 56:335-344[CrossRef][Medline]
  2. Mangelsdorf DJ, Thummel C, Beato M, Herrlich P, Schutz G, Umesono K, Blumberg B, Kastner P, Mark M, Chambon P, Evans RM. The nuclear receptor superfamily: the second decade. Cell 1995 83:835-839[CrossRef][Medline]
  3. Falkenstein E, Tillmann H-C, Christ M, Feuring M, Wehling M. Multiple actions of steroid hormones—a focus on rapid, nongenomic effects. Pharmacol Rev 2000 52:513-555[Abstract/Free Full Text]
  4. Watson CS, Gametchu B. Membrane-initiated steroid actions and the proteins that mediate them. Proc Soc Exp Biol Med 1999 220:9-19[Abstract]
  5. Schmidt BMW, Gerdes D, Feuring M, Falkenstein E, Christ M, Wehling M. Rapid, nongenomic steroid actions: a new age?. Front Neuroendocrinol 2000 21:57-94[CrossRef][Medline]
  6. Lyng FM, Jones GR, Rommerts FFG. Rapid androgen actions on calcium signaling in rat Sertoli cells and two human prostatic cell lines: similar biphasic responses between 1 picomolar and 100 nanomolar concentrations. Biol Reprod 2000 63:736-747[Abstract/Free Full Text]
  7. Gorczynska E, Handelsman DJ. Androgens rapidly increase the cytosolic calcium concentration in Sertoli cells. Endocrinology 1995 136:2052-2059[Abstract]
  8. Steinsapir J, Socci R, Reinach P. Effects of androgen on intracellular calcium of LNCaP cells. Biochem Biophys Res Commun 1991 179:90-96[CrossRef][Medline]
  9. Machelon V, Nomé F, Tesarik J. Nongenomic effects of androstenedione on human granulosa luteinizing cells. J Clin Endocrinol Metab 1998 83:263-269[Abstract/Free Full Text]
  10. Estrada M, Liberona JL, Miranda M, Jaimovich E. Aldosterone- and testosterone-mediated intracellular calcium response in skeletal muscle cell cultures. Am J Physiol Endocrinol Metab 2000 279:E132-E139[Abstract/Free Full Text]
  11. Lieberherr M, Grosse B. Androgens increase intracellular calcium concentration and inositol 1,4,5-triphosphate and diacylglycerol formation via a pertussis toxin-sensitive G-protein. J Biol Chem 1994 269:7217-7223[Abstract/Free Full Text]
  12. Armen TA, Gay CV. Simultaneous detection and functional response of testosterone and estradiol receptors in osteoblast plasma membranes. J Cell Biochem 2000 79:620-627[CrossRef][Medline]
  13. Benten WPM, Lieberherr M, Sekeris CE, Wunderlich F. Testosterone induces Ca2+ influx via nongenomic surface receptors in activated T cells. FEBS Lett 1997 407:211-214[CrossRef][Medline]
  14. Benten WPM, Lieberherr M, Giese G, Wrehlke C, Stamm O, Sekeris CE, Mossmann H, Wunderlich F. Functional testosterone receptors in plasma membranes of T cells. FASEB J 1999 13:123-133[Abstract/Free Full Text]
  15. Benten WPM, Lieberherr M, Stamm O, Wrehlke C, Guo Z, Wunderlich F. Testosterone signaling through internalizable surface receptors in androgen receptor-free macrophages. Mol Biol Cell 1999 10:3113-3123[Abstract/Free Full Text]
  16. Peterziel H, Mink S, Schonert A, Becker M, Klocker H, Cato A.C. Rapid signaling by androgen receptor in prostate cancer cells. Oncogene 1999 18:6322-6329[CrossRef][Medline]
  17. Zhu X, Li H, Liu J-P, Funder JW. Androgen stimulates mitogen-activated protein kinase in human breast cancer cells. Mol Cell Endocrinol 1999 152:199-206[CrossRef][Medline]
  18. Liu D, Dillon JS. Dehydroepiandrosterone activates endothelial cell nitric oxide synthetase by a specific plasma membrane receptor coupled to G{alpha}i2,3. J Biol Chem 2002 277:21379-21388[Abstract/Free Full Text]
  19. Towle AC, Sze PY. Steroid binding to synaptic plasma membrane: differential binding of glucocorticoids and gonadal steroids. J Steroid Biochem 1983 18:135-143[CrossRef][Medline]
  20. Ramirez VD, Zheng J, Siddique KM. Membrane receptors for estrogen, progesterone and testosterone in the rat brain: fantasy or reality. Cell Mol Neurobiol 1996 16:175-198[CrossRef][Medline]
  21. Pottinger TG, Moore A. Characterization of putative steroid receptors in the membrane, cytosol, and nuclear fractions from the olfactory tissue of brown trout. Fish Physiol Biochem 1997 16:45-63[CrossRef]
  22. Campo S, Pellizzari E, Cigorraga S, Monteagudo C, Nicolau G. Androgen binding to subcellular particles of rat testis. J Steroid Biochem 1982 17:165-173[CrossRef][Medline]
  23. Konoplya EF, Popoff EH. Identification of the classical androgen receptor in male rat liver and prostate cell plasma membranes. Int J Biochem 1992 24:1979-1983[CrossRef][Medline]
  24. Braun AM, Thomas P. Androgens inhibit estradiol-17ß synthesis in Atlantic croaker (Micropogonias undulatus) ovaries by a nongenomic mechanism initiated at the cell surface. Biol Reprod 2003 69:1642-1650[Abstract/Free Full Text]
  25. Sperry TS, Thomas P. Characterization of two nuclear androgen receptors in Atlantic croaker: comparison of their biochemical properties and binding specificities. Endocrinology 1999 140:1602-1611[Abstract/Free Full Text]
  26. Sperry TS, Thomas P. Identification of two nuclear androgen receptors in kelp bass (Paralabrax clathratus) and their binding affinities for xenobiotics: comparison with Atlantic croaker (Micropogonias undulatus) androgen receptors. Biol Reprod 1999 61:1152-1161[Abstract/Free Full Text]
  27. Sperry T, Thomas P. Androgen binding profiles of two distinct nuclear androgen receptors in Atlantic croaker (Micropogonias undulatus). J Steroid Biochem Mol Biol 2000 73:93-103[CrossRef][Medline]
  28. Peck EJ, Kelner KL. Receptor measurement. In: Lajtha A (ed.), Handbook of Neurochemistry, vol. 2 Experimental Neurochemistry, 2nd ed. New York: Plenum Press; 1982:53–75
  29. Scatchard G. The attractions of proteins for small molecules and ions. Ann N Y Acad Sci 1949 51:660-672[CrossRef]
  30. Patino R, Thomas P. Characterization of membrane receptor activity for 17{alpha},20ß,21-trihydroxy-4-pregnen-3-one in ovaries of spotted seatrout (Cynoscion nebulosus). Gen Comp Endocrinol 1990 78:204-217[CrossRef][Medline]
  31. Loomis AK, Thomas P. Effects of estrogens and xenoestrogens on androgen production by Atlantic croaker testes in vitro: evidence for a nongenomic action mediated by an estrogen membrane receptor. Biol Reprod 2000 62:995-1004[Abstract/Free Full Text]
  32. Cuevas ME, Callard G. Androgen and progesterone receptors in shark (Squalus) testis: characteristics and stage-related distribution. Endocrinology 1992 130:2173-2182[Abstract]
  33. Pike CJ. Testosterone attenuates ß-amyloid toxicity in cultured hippocampal neurons. Brain Res 2001 919:160-165[CrossRef][Medline]
  34. Orsini JC, Barone FC, Armstrong DL, Wayner MJ. Direct effects of androgens on lateral hypothalamic neuronal activity in the male rat: I. A microiontophoretic study. Brain Res Bull 1985 15:293-297[CrossRef][Medline]
  35. Kubli-Garfias C, Canchola E, Arauz-Contreras J, Feria-Velasco A. Depressant effect of androgens on the cat brain electrical activity and its antagonism by ruthenium red. Neuroscience 1982 7:2777-2782[CrossRef][Medline]
  36. Yamada Y. Effects of testosterone on unit activity in the rat hypothalamus and septum. Brain Res 1979 172:165-168[CrossRef][Medline]
  37. Zinder O, Dar DE. Neuroactive steroids: their mechanism of action and their function in the stress response. Acta Physiol Scandinavia 1999 167:181-188
  38. Frye CA, Duncan JE, Basham M, Erskine MS. Behavioral effects of 3{alpha}-androstanediol II: hypothalamic and preoptic area actions via a GABAergic mechanism. Behav Brain Res 1996 79:119-130[CrossRef][Medline]
  39. Khan IA, Thomas P. GABA exerts stimulatory and inhibitory influences on gonadotropin II secretion in the Atlantic croaker. Neuroendocrinology 1999 69:261-268[CrossRef][Medline]
  40. Khan IA, Hawkins MB, Thomas P. Gonadal stage-dependent effects of gonadal steroids on gonadotropin II secretion in the Atlantic croaker (Micropogonias undulatus). Biol Reprod 1999 61:834-841[Abstract/Free Full Text]
  41. Connaughton MA, Taylor MH. Effects of exogenous testosterone on sonic muscle mass in the weakfish, Cynoscion regalis. Gen Comp Endocrinol 1995 100:238-245[CrossRef][Medline]
  42. Jakobsson S, Borg B, Haux C, Hyllner SJ. An 11-ketotestosterone induced kidney-secreted protein: the nest building glue from male three-spined stickleback, Gasterosteus aculeatus. Fish Physiol Biochem 1999 20:79-85
  43. Jones I, Lindberg C, Jakobsson S, Hellqvist A, Hellman U, Borg B, Olsson P-E. Molecular cloning and characterization of spiggin: an androgen-regulated extraorganismal adhesive with structural similarities to Von Willebrand factor related proteins. J Biol Chem 2001 276:17857-17863[Abstract/Free Full Text]
  44. Thomas P, Breckenridge-Miller D, Detweiler C. The teleost sperm membrane progestogen receptor: interactions with xenoestrogens. Mar Environ Res 1998 46:163-166[CrossRef]
  45. Thomas P. Nontraditional sites of endocrine disruption by chemicals on the hypothalamus-pituitary-gonadal axis: Interactions with steroid membrane receptors, monoaminergic pathways, and signal transduction systems. In: Naz RK (ed.), Endocrine Disruptors: Effects on Male and Female Reproductive Systems. Boca Raton: CRC Press; 1999:3– 38
  46. Kelce WR, Stone CR, Laws SC, Gray LE, Kemppainen JA, Wilson EM. Persistent DDT metabolite p,p'-DDE is a potent androgen receptor antagonist. Nature 1995 375:581-585[CrossRef][Medline]
  47. Maness SC, McDonnell DP, Gaido KW. Inhibition of androgen receptor-dependent transcriptional activity by DDT isomers and methoxychlor in HepG2 human hepatoma cells. Toxicol Appl Pharmacol 1998 151:135-142[CrossRef][Medline]
  48. Kelce WR, Monosson E, Gamcsik MP, Laws SC, Gray LEJ. Environmental hormone disruptors: evidence that vinclozolin developmental toxicity in mediated by antiandrogenic metabolites. Toxicol Appl Pharmacol 1994 126:276-285[CrossRef][Medline]
  49. Norfleet AM, Thomas ML, Gametchu B, Watson C. Estrogen receptor-{alpha} detected on the plasma membrane of aldehyde-fixed GH/B6/F10 rat pituitary tumor cells by enzyme-linked immunocytochemistry. Endocrinology 1999 140:3805-3814[Abstract/Free Full Text]
  50. Razandi M, Pedram A, Greene G, Levin ER. Cell membrane and nuclear estrogen receptors (ERs) originate from a single transcript: studies of ER{alpha} and ERß expressed in Chinese hamster ovary cells. Mol Endocrinol 1999 13:307-319[Abstract/Free Full Text]
  51. Thomas P, Zhu Y, Pace M. Progestin membrane receptors involved in the meiotic maturation of teleost oocytes: a review with some new findings. Steroids 2002 67:511-517[CrossRef][Medline]
  52. Zhu Y, Rice CD, Pace M, Thomas P. Cloning, expression, and characterization of a membrane progestin receptor and evidence it is an intermediary in meiotic maturation of fish oocytes. Proc Natl Acad Sci U S A 2003 100:2231-2236[Abstract/Free Full Text]
  53. Nakhla AM, Rosner W. Stimulation of prostate cancer growth by androgens and estrogens through the intermediacy of sex-hormone-binding globulin. Endocrinology 1996 137:4126-4129[Abstract]
  54. Laidley C, Thomas P. Partial characterization of a sex-steroid binding protein in the spotted seatrout (Cynoscion nebulosus). Biol Reprod 1994 51:982-992[Abstract]
  55. Rosner W, Hryb DJ, Khan MS, Nakhla AM, Romas NA. Androgen and estrogen signaling at the cell membrane via G-proteins and cyclic adenosine monophosphate. Steroids 1999 64:100-106[CrossRef][Medline]



This article has been cited by other articles:


Home page
 EndocrinologyHome page
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]

Home page
 EndocrinologyHome page
Y. Katsu, R. Ichikawa, T. Ikeuchi, S. Kohno, L. J. Guillette Jr., and T. Iguchi
Molecular Cloning and Characterization of Estrogen, Androgen, and Progesterone Nuclear Receptors from a Freshwater Turtle (Pseudemys nelsoni)
Endocrinology, January 1, 2008; 149(1): 161 - 173.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
E. Filardo, J. Quinn, Y. Pang, C. Graeber, S. Shaw, J. Dong, and P. Thomas
Activation of the Novel Estrogen Receptor G Protein-Coupled Receptor 30 (GPR30) at the Plasma Membrane
Endocrinology, July 1, 2007; 148(7): 3236 - 3245.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
J. W. Gatson and M. Singh
Activation of a Membrane-Associated Androgen Receptor Promotes Cell Death in Primary Cortical Astrocytes
Endocrinology, May 1, 2007; 148(5): 2458 - 2464.
[Abstract] [Full Text] [PDF]

Home page
 Hum Mol GenetHome page
Y. Takeshita, R. Fujinaga, C. Zhao, A. Yanai, and K. Shinoda
Huntingtin-associated protein 1 (HAP1) interacts with androgen receptor (AR) and suppresses SBMA-mutant-AR-induced apoptosis
Hum. Mol. Genet., August 1, 2006; 15(15): 2298 - 2312.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
J. W. Gatson, P. Kaur, and M. Singh
Dihydrotestosterone Differentially Modulates the Mitogen-Activated Protein Kinase and the Phosphoinositide 3-Kinase/Akt Pathways through the Nuclear and Novel Membrane Androgen Receptor in C6 Cells
Endocrinology, April 1, 2006; 147(4): 2028 - 2034.
[Abstract] [Full Text] [PDF]

Home page
 J Mol EndocrinolHome page