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BOR - Papers in Press, published online ahead of print June 9, 2004.
Biol Reprod 2004, 10.1095/biolreprod.104.030056
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BIOLOGY OF REPRODUCTION 71, 1174–1181 (2004)
DOI: 10.1095/biolreprod.104.030056
© 2004 by the Society for the Study of Reproduction, Inc.

Ovary

Characterization of Growth Hormone Binding Sites in Granulosa and Theca Layers at Different Stages of Follicular Maturation and Ovulatory Cycle in the Domestic Hen

Irina Y. Lebedeva2, Vladimir A. Lebedev2, Roland Grossmann2, Tatiana I. Kuzmina3, and Nahid Parvizi1,2

Department of Functional Genomics and Bioregulation,2 Institute of Animal Science (FAL), Mariensee, 31535 Neustadt, Germany Department of Genetics and Biotechnology,3 Research Institute for Farm Animal Genetics and Breeding, Pushkin, St. Petersburg 196625, Russia


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The currently available evidence points to a possible influence of growth hormone (GH) on avian folliculogenesis, which can be mediated by both hepatic- and ovarian-derived IGF-I. Therefore, the purpose of the present study was to reveal GH-binding sites in granulosa and theca layers of preovulatory follicles and to determine the binding characteristics depending on the degree of follicular maturation and the stage of the ovulatory cycle in the hen. Hens were killed 2 h (stage I), 9 h (stage II), 16 h (stage III), and 23 h (stage IV) after oviposition, and the five largest yellow follicles (from F1 to F5) were removed. GH-binding sites in granulosa and theca layers from F1 to F5 follicles were characterized using a radioreceptor assay. Equilibrium dissociation constants (Kd) and binding capacities (Bmax) were determined by Scatchard analysis of saturation curves, which revealed a single class of high-affinity GH-binding sites in both theca tissue and granulosa cells. In F1, F2, and F5 follicles, Bmax and Kd for GH-binding sites in the granulosa layer changed during the ovulatory cycle, decreasing between stages I and III, to increase again at stage IV, with alterations in Kd being less profound. No significant differences in binding capacities and affinities of GH-binding sites in the theca layer were found between various stages of the cycle. Furthermore, the concentration of GH-binding sites in the granulosa layer rose, whereas that in the theca layer fell with follicular enlargement. These findings indicate the presence of high-affinity GH-binding sites in both granulosa and theca layers of hen preovulatory follicles. Data also demonstrate that GH-binding sites in these tissues are regulated in a tissue-specific manner. Furthermore, the regulation of binding capacity of GH binding in granulosa cells by hormonal factors associated with ovulatory cycle is apparently not dependent on the state of follicular maturation.

chicken, follicle, follicular development, follicular maturation, granulosa cells, growth hormone, growth hormone-binding sites, theca cells, thecal layer


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In mammals, growth hormone (GH) is widely involved in the regulation of ovarian folliculogenesis affecting follicular atresia/apoptosis, steroidogenesis, ovulation, proliferation and differentiation of follicular cells, as well as oocyte maturation (for review, see [1]). Growth hormone actions on follicles can be both direct or mediated by hepatic or local production of insulin-like growth factor-I (IGF-I) [2, 3]. Binding sites for GH or GH receptor mRNA have been detected in follicles of various mammalian species, with the receptor concentration depending on the degree of the follicular maturity [4, 5].

The currently available evidence suggests that GH may be implicated in the control of reproduction in birds as well. In laying hens, the GH and GH-receptor genotypes have been found to be associated with age at first egg and the rate of egg production [6, 7]. Williams et al. reported that the number of small follicles in the domestic hen rises after treatment with ovine GH [8]. In turkeys, higher plasma GH concentrations and pituitary GH mRNA expression were detected in egg-laying than in nonlaying hens [9, 10]. Differences in egg production between ad libitum and restricted fed broiler have been demonstrated to be related to changes in the GH/IGF-I axis [11, 12].

The hen ovulatory cycle is characterized by alterations in plasma levels of ovarian steroid hormones and LH [13]. In mammals, ovarian steroid hormones have been shown to be regulators of GH-receptor synthesis in various tissues [14, 15], whereas LH can modulate GH-receptor gene expression in the ovary [16]. In vitro experiments have shown that GH exerts a stimulatory influence on both synthesis and secretion of IGF-I by hen granulosa cells, implying that GH can regulate the local production of this growth factor in the chicken ovary [17] and pointing to the presence of functional GH receptors on these cells. Furthermore, the ability of follicular theca to produce IGF-I observed by different authors [18, 19] indicates that the thecal tissue may also be a target for GH action. Therefore, the purpose of the present study was to reveal GH-binding sites in granulosa and theca layers of hen preovulatory follicles and to determine their binding characteristics depending on the degree of the follicular maturation and the stage of the ovulatory cycle.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Experimental Design

Single-comb white Leghorn hens were kept in individual cages under a lighting regimen of 12L:12D with free access to food and water. Egg laying was recorded daily from the first oviposition for each hen and a total of 22 birds (41–47 wk old) laying more than seven eggs per sequence were selected for the experiment. Time of oviposition in the hens was monitored throughout the egg sequence preceding the sequence under study and during the latter to the time of slaughtering in 15-min intervals. Time of ovulation was estimated on the basis that it occurs 30 min after oviposition [20]. Under the lighting regimen used, the time lapse between successive ovipositions from the second to the last but one egg in the sequences was 24.4 ± 0.1 h. This time was taken as the duration of the ovulatory cycle, which was divided into four successive stages. Stage I covers processes occurring in preovulatory follicles immediately after ovulation, when the second largest follicle becomes the first follicle, the third largest follicle becomes the second, etc. Stage II spans the time lapse between 8 and 12 h after ovulation, in the course of which the persistent increase in the FSH-binding ability of the theca of all preovulatory follicles and the short rise in the plasma level of FSH proceed [21]. Stage III directly precedes the preovulatory LH surge, which occurs 4–6 h before the ovulation [21]. Stage IV encompasses the period of time between the LH peak and ovulation. Therefore, the birds were killed at 2 h (stage I; n = 5), 9 h (stage II; n = 6), 16 h (stage III; n = 5), and 23 h (stage IV; n = 6) after oviposition, corresponding to 1.5, 8.5, 15.5, and 22.5 h after assumed ovulation. To obtain tissues for characterization of the GH-binding assay, birds were killed at 1000 h regardless of time of ovulation. Hens were killed by cervical dislocation in the middle of the egg sequence. Birds were used in accordance with procedures approved by the Institute of Animal Science and Hannover Government Animal Care and Use Committee (approval 509C 42502 00/394).

Blood and Tissue Collection

Immediately after slaughtering, blood samples were withdrawn from the heart into heparin-coated tubes. Plasma samples were frozen and stored at –20°C for estradiol-17ß and progesterone assays.

Ovaries were removed and rinsed in saline. The first (F1), second (F2), third (F3), fourth (F4), and fifth (F5) largest yellow follicles were detached from the ovaries. Theca and granulosa layers were separated as described by Gilbert et al. [22]. The thecal tissues were washed several times in saline and weighed.

Livers, removed in some hens, were immediately frozen and stored at –70°C until used as control tissue.

Tissue and Cell Preparation

The thecal tissue was minced with scissors and mechanically homogenized in ice-cold Dulbecco PBS (DPBS; 1:5, w/v) containing 1 mM PMSF. The homogenate was filtered through a screen cup (50 mesh), further homogenized with an ultrasonic cell disruptor (cell disruptor B15; Branson Sonic Power, Schwäbisch Gemünd, Germany), filtered through a screen cup (200 mesh), and centrifuged at 2300 x g for 30 min. The pellet was washed in DPBS and centrifuged again. The pellet volume was brought up to 3 ml by DPBS and an aliquot of the suspension was taken for protein determination by the method of Bradford [23]. The suspension was centrifuged and the final pellet, subsequently referred to as a crude particulate membrane preparation (CPMP), was then resuspended in the assay buffer of the following composition: 50 mM Tris-HCl (Sigma Chemical Co, St. Louis, MO), 150 mM NaCl (AppliChem, Darmstadt, Germany), 0.1% BSA (fraction V; Serva, Heidelberg, Germany), 0.02% thimerosal (Serva, Heidelberg), pH 5.0.

The granulosa layer was washed in DPBS to remove adhering yolk, minced with scissors and pipetted with DPBS. Granulosa cells were filtered through a screen cup (200 mesh) and centrifuged at 500 x g for 15 min. The cell pellet was washed in DPBS and centrifuged again. The final pellet was resuspended in the above-mentioned assay buffer and the cell concentration calculated with a hemocytometer.

All procedures were performed at 4°C. The samples of CPMP and granulosa cells were frozen and stored at –20°C for no more than 3 mo. Preliminary experiments revealed that the freezing-thawing procedure does not affect GH-binding ability of theca CPMP and granulosa cells.

Hepatic membrane preparations were obtained by the method of Krishnan et al. [24], with some modifications. Briefly, the frozen liver was homogenized in 50 mM Tris-HCl containing 0.3 M sucrose, 1 mM EDTA, and 2 mM PMSF, filtered, and centrifuged at 2500 x g for 20 min. The supernatant was recovered and centrifuged at 11 000 x g for 30 min. The final supernatant was then recentrifuged at 47 000 x g for 150 min and the resulting pellet was resuspended.

Binding Assay

Ovine GH (oGH; NIDDK-oGH-15) used for iodination was a gift of the National Institute of Diabetes and Digestive and Kidney Diseases and used as a tracer. Ovine GH had been shown to be an effective competitor for chicken GH [24]. Iodination of oGH was performed to a specific activity of 28–44 µCi/µg, using lactoperoxidase.

The general procedure for GH-binding assay was as follows. Aliquots of theca CPMP or granulosa cells were incubated with 125I-labeled oGH (150 000–200 000 cpm, equivalent to 4.5–5.3 ng) in a total volume of 250 µl in the above-mentioned assay buffer. Nonspecific binding of the labeled hormone was determined in the presence of an excess of unlabeled recombinant bovine GH (40 µg; Monsanto, St. Louis, MO). Incubation was conducted in polystyrene tubes in triplicate (total binding) or duplicate (nonspecific binding) in a shaking water bath at 37°C for 24 h. The reaction was terminated by the addition of 1 ml of the ice-cold assay buffer followed by centrifugation at 2300 x g for 30 min at 4°C. The supernatant was discarded, the pellet was washed twice with 1 ml of the cold buffer, and the radioactivity in the pellet was determined after the third centrifugation. The specific binding was determined by subtracting nonspecific binding from total binding.

Saturation analyses were performed by incubating aliquots of CPMP (7–10 µg of protein) or granulosa cells (0.08–0.12 x 106 cells) with increasing quantities of 125I-labeled oGH (0.03–0.6 pM) with or without excess of bovine GH. Equilibrium dissociation constants (Kd) and binding capacities (Bmax) were determined by Scatchard analysis of saturation curves [25] using the MultiCalc program (Perkin Elmer Wallac, Freiburg, Germany).

Characterization of GH-Binding Assay

The influence of pH on GH-specific binding was assessed by incubating 25 µg of CPMP protein or 0.8 x 106 cells with 125I-labeled oGH at pH ranging from 4.0 to 7.0 at 37°C for 24 h. To determine a possible occupancy of GH-binding sites by endogenous hormone, an aliquot of CPMP (150 µg of protein) was incubated in 0.3 ml 4 M MgCl2 at 23°C for 15 min, and then 3 ml ice-cold assay buffer, with pH 7.0, was added. After centrifugation, the resulting pellet was washed with 3 ml of the same assay buffer, centrifuged, and then binding assay was performed at pH 7.0. To investigate the effect of incubation time and temperature on the binding, the mixture of granulosa cells (0.15 x 106 and 0.5 x 106 cells) or theca CPMP (10 and 25 µg of protein) and 125I-labeled oGH was incubated (pH 5.0) at 37°C for different time scales (2–24 h). The mixture containing 25 µg of CPMP protein and radioligand was also incubated at 4 and 23°C for 2–69 h. The effect of increasing concentrations of CPMP and granulosa cells was examined by incubating theca CPMP (from 3.1 to 100.0 µg of protein) or granulosa cells (from 0.05 x 106 to 2.00 x 106 cells) with 125I-labeled oGH at 37°C and pH 5.0 for 24 h. Specificity of binding sites for GH was confirmed by testing the ability of bovine FSH and LH (Biogenesis Ltd., Poole, UK) and ovine prolactin (oPRL: NIDDK-oPRL-21) to compete with 125I-labeled oGH for binding sites.

Hormone Determination

Plasma concentrations of estradiol-17ß were determined by RIA as previously described [26]. The intra-assay coefficient of variation was 10%, and sensitivity of the assay was 1.5 pg/ml. Plasma progesterone concentrations were measured by an enzyme immunoassay [27]. The intra-assay coefficient of variation was 11%. The lowest level of detection was 0.5 ng/ml plasma.

Statistical Analysis

Results are expressed as means ± SEM. Data concerning the GH-binding characteristics of follicles with various degrees of maturation at different stages of the ovulatory cycle were analyzed by two-way repeated-measures ANOVA using SAS software. One-way repeated-measures ANOVA was employed for statistical verification of GH-binding characteristics during the ovulatory cycle. The independent variables were stage of ovulatory cycle and/or follicular category (repeated factor). Plasma levels of estradiol-17ß and progesterone in different hens were analyzed by one-way ANOVA. The residual error term was used to test the effect of stage of ovulatory cycle. Significant differences were determined using a Tukey test, which protects the significance tests of all combinations of pairs. A probability of P < 0.05 was considered to be statistically significant. Correlations between different reproductive parameters were estimated by Pearson correlation coefficient (r), employing the SigmaStat software package (SPSS Inc.).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Characterization of GH-Binding Sites

To characterize GH-binding sites, different experiments were performed using pooled samples of hen theca CPMP or granulosa cells from F1–F5 follicles. The procedures used for preparing theca CPMP and granulosa cells avoided the employment of enzyme digestion, which damages binding sites, and enabled us to process low quantities of thecal and granulosa tissues isolated from F4 and F5 follicles. Furthermore, under centrifugation conditions used in these procedures and binding assays, cytosolic proteins and most organelles are not precipitated [28].

Specific binding of 125I-labeled oGH to the hen theca CPMP and granulosa cells was strongly pH dependent. At pH 7.0, the binding to CPMP was only significant (4.6 ± 0.9% of total counts) when no less than 150 µg of protein per tube was incubated with radioligand at 37°C for 24 h. When incubated at 23°C for 48 h, no specific binding of 125I-labeled oGH to theca CPMP (200 µg of protein) was detected, whereas the binding to hen liver membranes (200 µg of protein) was about 4%. There was a striking 200-fold rise in 125I-labeled oGH binding to CPMP (25 µg of protein) and granulosa cells (0.8 x 106 cells) at 37°C as pH decreased from 7.0 to 5.0 (Fig. 1). The treatment of CPMP with 4 M MgCl2 at pH 7.0 did not cause a considerable increase in oGH binding (data not shown). Thus, the rise in specific binding did not result from the removal of the endogenously bound hormone at low pH.



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  		FIG. 1. Effect of pH on specific binding of 125I-labeled oGH to hen theca CPMP (25 µg of protein) and granulosa cells (0.8 x 106 cells). Each point represents mean ± SEM of three experiments performed in triplicate

Specific binding of 125I-labeled oGH to CPMP (25 µg of protein) was time and temperature dependent (Fig. 2). The specific binding increased with time at 37°C, reaching a maximum by 13 h, and then was stable for at least 11 h. In contrast, at 4°C and 23°C, the binding increased very slowly and did not attain an equilibrium even after 60 h of incubation. Specific binding of 125I-labeled oGH to granulosa cells (0.5 x 106 cells) also reached a steady state by 13 h of incubation at 37°C. The decrease in the concentration of CPMP protein and granulosa cells up to 10 µg and 0.12 x 106 cells, respectively, resulted in deceleration of the binding, with the equilibrium being established by 22– 24 h (data not shown).



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  		FIG. 2. Time course of 125I-labeled oGH specific binding to hen theca CPMP (25 µg of protein) and granulosa cells (0.5 x 106 cells). Each point represents mean ± SEM of three experiments performed in triplicate at 37°C or means of triplicate determinations in one experiment performed at 4°C and 23°C

Specific binding of 125I-labeled oGH to theca CPMP rose in a linear fashion, with increasing concentration of protein from 3.1 to 25 µg/tube, reaching saturation at the protein concentration of 50–100 µg/tube (Fig. 3A). The binding to granulosa cells also increased almost linearly at the cell concentration of 0.05 x 106 to 0.5 x 106 cells/tube (Fig. 3B). Thus, 7–10 µg of CPMP protein or (0.08–0.12) x 106 granulosa cells were used in the following saturation experiments.



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  		FIG. 3. Relationship of 125I-labeled oGH-specific binding to concentrations of hen theca CPMP (A) and granulosa cells (B). Each point represents mean ± SEM of three experiments performed in triplicate

Bovine LH and FSH at concentrations up to 4 µg/ml exhibited virtually no competition with 125I-labeled oGH for binding to theca CPMP (data not shown), bPRL and oPRL at a concentration of 4 µg/ml reduced slightly (by 2%), confirming the specificity of binding sites for GH and the low cross-reactivity of chicken GH with ovine and bovine PRL observed in the chicken liver [24].

No loss in the binding ability of 125I-labeled oGH was observed after its preincubation in the assay buffer alone or with theca CPMP at 37°C for 20 h, indicating the absence of the radioligand degradation during the established incubation period. Moreover, there was not significant degradation of GH-binding sites after preincubation of CPMP and granulosa cells under the same conditions (data not shown).

Saturation curves and Scatchard plots of specific binding of 125I-labeled oGH to theca CPMP and granulosa cells from F1–F5 follicles are presented in Figure 4 (A and B). The Scatchard analysis revealed a single class of high-affinity oGH-binding sites in both theca CPMP and granulosa cells. Dissociation constants (Kd) for specific binding of 125I-labeled oGH to CPMP and granulosa cells were 723 ± 46 pM (n = 6) and 737 ± 81 pM (n = 6), respectively. Maximum binding capacity (Bmax) was 15.2 ± 1.2 pM per milligram protein for theca CPMP and 610 ± 44 fM per 106 cells for granulosa cells.



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  		FIG. 4. Saturation curves (A) and Scatchard plots (B) of specific binding of 125I-labeled oGH to theca CPMP (10 µg of protein) and granulosa cells (0.1 x 106 cells) from hen preovulatory follicles F1–F5. Each point represents mean of triplicate determinations of one pooled sample

Changes in Binding Characteristics of GH-Binding Sites During Follicular Development

Two-way repeated-measures ANOVA revealed that there is a significant (P ≤ 0.05) relation between ovulatory cycle, follicular category, and concentration of GH-binding sites. The GH-binding capacity of granulosa cells from preovulatory follicles with various degrees of maturity changed during the hen ovulatory cycle (Fig. 5A). In F1 and F2 follicles, concentration of GH-binding sites was high at stage I, decreased at stages II and III to increase again at stage IV. F5 follicles also possessed a significantly higher (P ≤ 0.01) Bmax for GH at stages IV and I when compared with stage III (Fig. 5A). Furthermore, the GH-binding capacity of granulosa cells rose (P ≤ 0.001) as the follicles moved from the fifth to the first position in the hierarchy, with a considerable difference in Bmax for GH-binding sites between F5 and F1 as well as between F4 and F1 follicles being observed at all stages of the ovulatory cycle (Fig. 5B). In addition, a negative correlation (r = –0.44; P < 0.001) was found between Bmax for GH in granulosa cells of different follicular category and the time to expected ovulation (Table 1).



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  		FIG. 5. Binding capacities (Bmax) of GH-binding sites in granulosa cells from hen preovulatory follicles (means ± SEM). A) Comparison of follicles with various degrees of maturation at different stages of the ovulatory cycle. Means for follicular categories within a stage and for stages within a follicular category with different letters differ significantly (at least P ≤ 0.05). Numbers of follicles F1–F5: n = 5 (stages I and III) and n = 6 (stages II and IV). B) The mean binding capacities of GH-binding sites in granulosa cells during ovulatory cycle. Means with different letters differ significantly (at least P ≤ 0.01). Number of follicles F1–F5: n = 22. Data were analyzed by repeated-measures two-way (A) and one-way (B) ANOVA


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  		TABLE 1. Correlations between binding characteristics and hen repro ductive parameters

As shown in Figure 6A, the pattern of alterations in Kd for GH in granulosa cells of F5, F2, and F1 follicles was, as a whole, similar to that of Bmax, but less profound. The mean affinity of GH-binding sites during the cycle was highest in granulosa cells from F4 and F2 follicles (Fig. 6B).



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  		FIG. 6. Dissociation constants (Kd) for GH-binding sites in granulosa cells from hen preovulatory follicles (means ± SEM). A) Comparison of follicles with various degrees of maturation at different stages of the ovulatory cycle. Means for follicular categories within a stage and for stages within a follicular category with different letters differ significantly (at least P ≤ 0.05). Numbers of follicles F1–F5: n = 5 (stages I and III) and n = 6 (stages II and IV). B) The mean dissociation constants for GH-binding sites in granulosa cells during ovulatory cycle. Means with different letters differ significantly (at least P ≤ 0.05). Number of follicles F1–F5: n = 22. Data were analyzed by repeated-measures two-way (A) and one-way (B) ANOVA

The only significant difference in binding capacities and affinities of GH-binding sites in theca CPMP were found between various stages of the hen ovulatory cycle was a reduction in Kd of F2 follicles in stages III and IV when compared with stages I and II (Figs. 7A and 8A). In contrast with the granulosa cells, the mean concentration of GH-binding sites during the cycle decreased in theca CPMP with the follicular enlargement from F5 to F1 (Fig. 7B), whereas the affinity of the sites did not differ between follicles with various degrees of maturity (Fig. 8B). Further analysis of the data revealed the existence of a positive correlation between Bmax of GH-binding sites in theca CPMP of different follicular categories and the time to expected ovulation (Table 1; r = 0.33; P ≤ 0.01).



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  		FIG. 7. Binding capacities (Bmax) of GH-binding sites in theca CPMP from hen preovulatory follicles (mean ± SEM). A) Comparison of follicles with various degrees of maturation at different stages of the ovulatory cycle. Numbers of follicles F1–F5: n = 4 (stages I–IV). B) The mean binding capacities of GH-binding sites in theca CPMP during ovulatory cycle. Means with different letters differ significantly (P ≤ 0.05). Number of follicles F1–F5: n = 16. Data were analyzed by repeated-measures two-way (A) and one-way (B) ANOVA



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  		FIG. 8. Dissociation constants (Kd) for GH-binding sites in theca CPMP from hen preovulatory follicles (means ± SEM). A) Comparison of follicles with various degrees of maturation at different stages of the ovulatory cycle. Numbers of follicles F1–F5: n = 4 (stages I–IV). B) The mean dissociation constants for GH-binding sites in granulosa cells during ovulatory cycle. Number of follicles F1–F5: n = 16. Data were analyzed by repeated-measures two-way (A) and one-way (B) ANOVA

The highest binding capacity and the lowest mean affinity of GH-binding sites in granulosa cells were found in stage I, while stage III was characterized by the lowest binding capacity and the highest affinity of the binding sites during the hen ovulatory cycle (Table 2).


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  		TABLE 2. Alterations in binding characteristics of GH-binding sites in granulosa and theca layers of preovulatory follicles during the hen ovulatory cycle.*

It should be mentioned that the content of CPMP protein in the thecal tissue of preovulatory follicles varied during the hen ovulatory cycle and was lowest at stage III (Table 3). In addition, the mean CPMP protein content during the cycle increased in the theca with follicular enlargement from F5 to F1. Furthermore, the mean binding capacity of the thecal tissue from F4 and F5 follicles, producing substantial quantities of estradiol [29], positively correlated with the plasma estradiol-17ß concentration (Table 3) in individual hens (Table 1; r = 0.83; P ≤ 0.001).


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  		TABLE 3. Alterations in plasma ovarian steroid levels and protein content of theca CPMP during the hen ovulatory cycle.*


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study has demonstrated the presence of a single class of high-affinity GH-binding sites in both granulosa and theca layers of preovulatory follicles in the domestic hen. The method developed has allowed determination of binding characteristics of GH to follicular tissues not only from large F1–F3 follicles but also from small individual F4 and F5 follicles. The affinity of GH-binding sites in granulosa and thecal layers was similar to the affinity of GH-binding sites in the rabbit ovary [30] but slightly lower than those measured in the chicken liver [present study and 24]. This seems to be rather a difference between tissues than a result of different studies. Binding capacity of GH in granulosa cells and theca CPMP was high, with the latter being in the same region as Bmax of unmasked related binding sites for prolactin in detergent-treated membrane preparations of luteinized rat ovaries [31]. Indeed, our preliminary experiments have shown that an increase in specific binding of 125I-labeled oGH to theca CPMP with decreasing pH is due to a rise in Bmax rather than a decline in Kd. Apparently, most of the GH-binding sites in follicular tissues are masked, at least for in vitro binding, and can be revealed at low pH.

There is no data in the literature elucidating characteristics of follicular GH-binding sites during the maturation of avian ovarian follicles. Our findings for Bmax in granulosa cells from follicles with different degrees of maturity as well as the negative correlation revealed between this and the time to expected ovulation point to an increase in GH responsiveness of the cells with the follicular development. This increase is consistent with a higher ability of GH to stimulate IGF-I production by granulosa cells in F1 than in F3 follicles [17]. The data obtained are in contrast with those for pigs [32] and cows [33], in which the concentration of GH biding sites in granulosa cells has been shown to decrease during the last stages of follicular maturation.

Of particular interest are the findings concerning variations in the concentration of GH-binding sites in granulosa cells during the hen ovulatory cycle. The data disclosed that the GH-binding capacity of the cells in F1, F2, and F5 follicles decreases between stages I and III to increase again at stage IV following the preovulatory LH surge. This provides reason to assume that the rise in the concentration of GH-binding sites is caused by the preceding elevation in the blood LH level. Indeed, the treatment of hypophysectomized ewes with LH has been shown to raise expression of GH receptor mRNA in the corpus luteum [16]. Furthermore, the synergistic action of LH and GH on IGF-I production by granulosa cells observed in vitro [17] might be due to an upregulating influence of LH on GH-binding sites. Alternatively, the increase in the concentration of GH-binding sites in granulosa cells may be triggered by the preovulatory enhancement of follicular production of estradiol-17ß and/or progesterone [34], regulating GH receptor expression in different mammalian tissues [15]. In the present study, no correlation was found between GH-binding capacity of granulosa cells and plasma concentrations of estradiol-17ß and progesterone in hens. Nevertheless, a possibility of the joint upregulating action of LH, estradiol, and progesterone on GH-binding sites in granulosa cells cannot be ruled out. This assumption is corroborated by the data that alterations in the GH-binding capacity of granulosa cells occur not only in F1 and F2 follicles, which show the highest expression of LH-receptor mRNA in the granulosa layer, but in F5 follicles as well, in which this expression is low [35], but estradiol production is higher than in F1–F4 follicles [29]. It should be mentioned that GH downregulates GH receptor expression and binding activity in the chicken liver [36, 37]. The available data on plasma concentrations of GH in the hen are rather contradictory. One report states that the GH concentration does not vary considerably during the ovulatory cycle [38], whereas others have found a significant increase in the GH level between 2 h before and 2 h after ovulation, which remained elevated for about 4 h thereafter [39].

There was a strong correlation between changes in Kd with changes in Bmax (r = 0.637, P < 0.01) during the ovulatory cycle. In contrast, changes in Kd and Bmax in the course of follicular enlargement were not connected.

The findings of the present study in conjunction with the evidence of other authors for GH-driven production of IGF-I by granulosa cells [17] suggest a potential implication of GH in the regulation of the follicular development in the domestic hen. It seems likely that the increase in the concentration of GH-binding sites in granulosa cells from F2 follicles after the preovulatory LH surge can be important for the enhancement of IGF-I production stimulating progesterone secretion by the cells [17]. In turn, the respective rise in IGF-I production by F5 follicles in response to GH may be required to accelerate the growth of granulosa and thecal tissues at this stage of maturation [40]. The increase in GH-binding capacity of granulosa cells in F1, follicles especially at the last stage of their development, provides reason to assume that GH might play a facilitatory role in ovulation of hen follicles, similar to its action in mammals [1]. In addition, a positive correlation revealed between the mean binding capacity of the thecal tissue from F4–F5 follicles and the plasma estradiol-17ß concentration in individual hens reflects a direct stimulatory influence of GH on estradiol-17ß production by the theca because IGF-I has been shown to inhibit this production [40].

In conclusion, the findings of the present study indicate the presence of high-affinity GH-binding sites in both granulosa and theca layers of hen preovulatory follicles. Data also demonstrate that GH-binding sites in these tissues are regulated in a tissue-specific manner. Furthermore, the regulation of binding capacity as well as affinity of GH-binding sites in granulosa cells by hormonal factors associated with ovulatory cycle occurs independently of maturational factors.


   ACKNOWLEDGMENTS
 
We are grateful to the NIDDK (National Institute of Diabetes and Digestive and Kidney Disease) and Monsanto (St. Louis, MO) for the gifts of ovine GH, ovine and bovine prolactin, and recombinant bovine GH. We thank R. Wittig for expert technical assistance with labeling of GH and RIA and P. Aldag for help with enzyme immunoassay.


   FOOTNOTES
 
1 Correspondence: N. Parvizi, Department of Functional Genomics and Bioregulation, Institute of Animal Science FAL, Mariensee, Höltystrasse 10, 31535 Neustadt, Germany. FAX: 49 0 5034 871247; parvizi{at}tzv.fal.de Back

Received: 25 March 2004.

First decision: 11 April 2004.

Accepted: 27 May 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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