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Endocrinology Vol. 141, No. 7 2683-2690
Copyright © 2000 by The Endocrine Society

ARTICLES

Ontogeny of Corticosterone-Inducible Growth Hormone-Secreting Cells during Chick Embryonic Development1

Ioannis Bossis and Tom E. Porter

Department of Animal and Avian Sciences, University of Maryland, College Park, Maryland 20742

Address all correspondence and requests for reprints to: Dr. Tom E. Porter, Department of Animal and Avian Sciences, University of Maryland, College Park, Maryland 20742. E-mail: tp44{at}umail.umd.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We reported that corticosterone administration into the albumen of fertile chicken eggs on embryonic day (e) 11 induces an increase in the population of GH-secreting cells. The present study evaluated the ontogeny, dose response, localization, and persistence of the glucocorticoid-induced increase in the somatotroph population during chicken embryonic development. Corticosterone (0, 0.02, 0.2, and 2 µg in 300 µl saline) was injected into separate eggs on e9, e10, e11, and e12, and the population of GH-secreting cells was assessed 2 days later using reverse hemolytic plaque assays. Corticosterone treatment on e9 or e10 was unable to increase the population of GH-secreting cells on e11 or e12. In contrast, 0.2 and 2 µg of corticosterone on e11 increased the population of GH-secreting cells on e13 (P < 0.05, n = 3 experiments) to 8.2 ± 0.6 and 6.4 ± 0.5% of all cells, respectively, relative to controls (2.4 ± 0.2%). For e14 embryos treated on e12, only the 2 µg dose increased the proportion of GH-secreting cells (6.4 ± 0.6%) relative to controls (3.6 ± 0.4%). In a second experiment, 0, 0.02, 0.2, 2, and 20 µg of corticosterone were injected on e0, e8, e9, e10, e11, and e12, and the population of GH-secreting cells was assessed on e13 in all groups. No dose of corticosterone was effective when given on e0, e8, e9, or e10. The 0.2 µg and 2 µg doses increased the population of GH-secreting cells (7.6 ± 0.9% and 6.7 ± 0.8%, respectively) relative to controls (2.3 ± 0.4%) when injected on e11 (P < 0.05, n = 4 experiments). The 2-µg dose also increased GH cell abundance when injected on e12 (5.6 ± 0.4%), relative to controls (2.7 ± 0.5%). Treatment with 20 µg on e11 and e12 induced the greatest responses (10.3 ± 1.1% and 8.7 ± 0.9%, respectively). However, in subsequent experiments, administration of 20 µg on e11 resulted in embryonic death by e18. In a third set of experiments, two groups of eggs were injected either with 2 µg of corticosterone in saline or saline alone on e11, and the number of GH-secreting cells was estimated on e13, e16, e19, and the day of hatch (d1). The population of GH-secreting cells in corticosterone treated embryos was significantly higher than in saline treated embryos only on e13 (7.1 ± 0.8% and 2.7 ± 0.3%, respectively). No significant differences were observed on e16 (12.4 ± 1.5% and 13.6 ± 1.2%), e19 (19.0 ± 1.0% and 18.2 ± 1.7%) and d1 (23.8 ± 2.1% and 25.1 ± 1.8%) between corticosterone treated and control embryos, respectively. In a fourth set of experiments, whole mount in situ hybridization indicated that injection of corticosterone on e11 induced GH messenger RNA expression in the caudal part of the pituitary gland on e13, where somatotrophs are located normally later in development. We conclude that corticosterone administration in ovo can increase the population of GH-secreting cells in the caudal anterior pituitary only during a small window of development between e11 and e13 and that this premature increase of GH-secreting cells does not affect the percentage of GH-secreting cells later in development.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
IN RECENT years, considerable progress has been made in identifying pituitary specific transcription factors involved in early pituitary differentiation, such as Lhx-3, Lhx-4, P-OTX, and Prop1, and transcription factors involved in proliferation and terminal differentiation of the five distinct anterior pituitary cell types, such as GATA-2 and Pit-1 (1, 2). The pituitary-specific transactivating factor Pit-1 has been localized in thyrotrophs, somatotrophs, and lactotrophs and has been shown to be necessary for transcription of TSH-ß, PRL, and GH genes. In contrast, limited progress has been made in identifying local or systemic extracellular factors that can influence pituitary cell differentiation and production of hormones on their own or in a synergistic manner with pituitary specific transcription factors. The differentiative effects of glucocorticoids in a number of embryonic tissues has long being recognized (3, 4, 5, 6, 7). In the rat pituitary gland, studies have suggested that induction of terminal somatotroph differentiation (induction of hormone containing phenotype) can be induced by various peptides and glucocorticoids in vitro (8, 9). In addition, treatment of pregnant rats with glucocorticoids induces premature differentiation or detection of GH-containing cells in the pituitaries of their fetuses (10, 11). However fetal-maternal interactions in these experiments could not be ruled out. Our laboratory has been using the chicken embryo as a model to study the mechanisms underlying somatotroph differentiation in the anterior pituitary. Chicken embryonic development is a useful model because the embryos can be easily manipulated in the absence of maternal interactions. In addition, the mechanisms involved in early pituitary differentiation are conserved between mammals and chicks (12), and the pattern of pituitary cell differentiation in chickens is comparable to that in mammals (13). Somatotrophs first appear on or before embryonic day 12 (e12) but become a significant population between e14 and e16 during chicken embryonic development (14). It also has been shown that GH-secreting cells do not differentiate in cultures of embryonic pituitary cells without an extrapituitary signal (15), and GH cell differentiation can be induced in these cultures with corticosterone (16). Furthermore, treatment of chicken embryos in ovo with corticosterone on e11 increases the number of GH-secreting cells on e14 (17). However, nothing is known about the embryonic age at which the somatotroph precursor cells become responsive to glucocorticoid induction or the relative amount of corticosterone necessary to induce somatotroph differentiation at different ages. In the present study, experiments were designed to evaluate the ontogeny and dose response of the corticosterone induced premature increase in GH-secreting cells by administering corticosterone in ovo and estimating the population of somatotrophs at different stages of embryonic development by reverse hemolytic plaque assay (RHPA). In a second set of experiments, we examined whether the corticosterone-induced premature increase in GH-secreting cells can affect the population of somatotrophs during subsequent embryonic development. Finally, we wanted to confirm that corticosterone increases the population of somatotrophs in the area of the pituitary gland where normal somatotroph differentiation occurs.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals, injections, and RHPA
All animals used in the present study were Avian x Avian broiler strain chicken embryos purchased from Allen’s Hatchery (Seaford, DE). Cell culture reagents were obtained from Life Technologies, Inc. (Grand Island, NY), and hormones and other chemicals from Sigma (St. Louis, MO) unless stated otherwise. Eggs were placed in a humidified incubator (G.Q.F. Manufacturing, Savannah, GA) at 37.5 C, and that day is designated as embryonic day 0 (e0). On the day of injections, the eggs were removed from the incubator, corticosterone treatments were injected into the albumen as previously described (17), and eggs were returned to the incubator. In the present studies, corticosterone was used because it is the predominant glucocorticoid in chickens during postembryonic development, and neither corticosterone, cortisol, nor cortisone predominates during embryonic development (18). A corticosterone stock solution (20 mg/ml) was prepared in 100% ethanol and further diluted with normal saline (0.9% NaCl) to appropriate concentrations. At certain stages of embryonic development (as described below), injected eggs were removed from the incubator, and their pituitary glands were isolated under a dissecting microscope. For each experimental group, three to four pituitaries were pooled and monodispersed cells were obtained as previously described (14). Monodispersed cells were subjected to RHPA according to the general method described previously (19) and modifications described later (14), using rabbit antiserum against chicken GH. Briefly, monodispersed anterior pituitary cells were mixed with protein A-coated ovine red blood cells and infused into Cunningham chambers. Cells were allowed to attach for 45 min in a cell culture incubator (37.5 C, 95% air-5% CO2) and rinsed with DMEM to remove unattached cells. Antiserum for chicken GH (1:40) and hGHRH1–40 (10-7 M) in DMEM were then added to the resulting monolayer of cells, and chambers were incubated for 8 h (3 chambers per treatment). Plaque formation was induced by incubation with guinea pig complement (1:40) in DMEM for 45 min. The cells were then fixed with 2% glutaraldehyde in PBS for 10 min and stored in PBS until analyzed. The percentage of pituitary cells that formed plaques was estimated using a light microscope, and at least 200 cells were counted per chamber.

Experimental protocol
In the first set of experiments, the albumen of eggs containing living embryos was injected with 0, 0.02, 0.2, and 2 µg of corticosterone in 300 µl of saline on e9, e10, e11, and e12 (one embryonic age in each replicate experiment). Two days later (e11, e12, e13, and e14, respectively) the pituitaries from the embryos in each group were isolated, and monodispersed cells were subjected to RHPA to detect GH-secreting cells. This set of experiments was replicated three times for each age. In a second set of experiments, eggs containing embryos on e0, e8, e9, e10, e11, and e12 (one embryonic age in each experiment) were injected with 0, 0.02, 0.2, 2, and 20 µg of corticosterone in 300 µl of saline. Somatotroph differentiation was assessed with RHPA on e13 in all trials. This set of experiments was replicated four times for each age. In a third set of experiments, the highest dose of corticosterone that did not compromise subsequent embryonic survival was determined. Different groups of eggs were injected on e11 with 0, 2, 5, 10, and 20 µg of corticosterone in 300 µl of saline, and embryonic mortality was assessed on d1 (the day of hatching). This experiment was replicated twice. Subsequently, eggs were injected either with 2 µg of corticosterone (the highest dose tested that did not compromise subsequent embryonic survival) in saline or saline alone on e11, and the number of GH-secreting cells was estimated on e13, e16, e19, and d1 by RHPA. This set of experiments was replicated three times. In all of the above experiments, at least three pituitary glands were used per experimental group to obtain monodispersed cells for RHPA. In the final set of experiments, whole mount in situ hybridization (ISH) was performed to localize the site of somatotroph differentiation within the embryonic pituitary glands. Eggs on e11 were injected either with saline or 2 µg of corticosterone in saline. Pituitary glands of both groups were dissected on e13, and whole mount ISH was performed as described below. In addition, whole mount ISH was performed in pituitary glands derived from e16 embryos, to indicate the extent and the site of GH messenger RNA (mRNA) production at this stage.

Whole tissue mount in situ hybridization
The ISH procedure was performed according to the method for the whole mount ISH of mouse embryos (20) with minor modifications. The whole procedure was carried out in 1.5 ml microfuge tubes. The dissected pituitary glands were immediately fixed with 3.7% formaldehyde in 0.1 M phosphate buffer (pH 7.0) at 4 C for 1 h. The pituitaries were subsequently washed three times with PBT (phosphate buffer saline with 0.1% Triton X-100) for 5 min each, dehydrated in a series of increasing methanol concentrations (50, 75, 95, 100%) in PBT (1 min in each solution), and stored overnight at -20 C in 100% methanol. The following day, the pituitary glands were rehydrated and permeabilized with proteinase K digestion (10 µg/ml in PBT) for 5 min. Subsequently, they were washed once with PBT, once with PBT containing 2 mg/ml glycine, and again with PBT, and then they were postfixed with 2% formaldehyde in PBS for 10 min. After brief washes in PBT, the glands were hybridized in the presence of digoxigenin-labeled GH cRNA at a concentration of 500 ng/ml hybridization buffer [50% formamide, 300 mM NaCl, 1x Denhardt’s solution (0.02% BSA, 0.02% Ficoll 400, 0.02% polyvinylpyrrolidone), 0.1% Triton X-100, 500 µg/ml transfer RNA, 50 µg/ml heparin, 2 mM EDTA (pH = 8), 25 mM Tris-HCl (pH 7.4)] for 16 h at 55 C. At the end of the incubation the tissues were washed three times for 30 min each with 2 xSSC, 0.1% CHAPS (20 x SSC is 3 M NaCl, 0.3 M sodium citrate, pH 7) and three times for 30 min each in 0.2 x SSC, 0.1% CHAPS at 58 C. The tissues were then washed once with buffer A (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 10 mM KCl, 1% Triton X-100), blocked with 10% heat inactivated sheep serum in buffer A for 2 h at room temperature, and incubated with a sheep antidigoxigenin antibody (1:2000) conjugated to alkaline phosphatase (Roche Molecular Biochemicals, Mannheim, Germany) for 16 h at 4 C in buffer A. At the end of the incubation, tissues were washed six times with buffer A for 10 min each at room temperature and four times for 1 h each at 4 C. The tissues were then washed twice for 5 min each in buffer B (100 mM Tris-HCl, pH 9.5, 100 mM NaCl, 50 mM MgCl2, 0.1% Triton X-100). To visualize GH mRNA, the tissues were incubated in the dark with buffer B containing 335 µg/ml of 4-nitro blue tetrazolium chloride (Roche Molecular Biochemicals) and 160 µg/ml 5-bromo-4-chloro-3- indolylphosphate (Roche Molecular Biochemicals). The pituitaries from the different groups were exposed for the same amount of time in the alkaline phosphatase substrate (15 min). The plasmid construct used for synthesis of digoxigenin labeled chicken GH (cGH) riboprobes was produced in this laboratory. A cGH complementary DNA (cDNA) (21) was provided by Dr. D. Foster (University of Minnesota, St. Paul, MN). This plasmid was digested with EcoRI and XbaI, and the resulting cGH cDNA fragment was ligated into the EcoRI and XbaI sites of the pGEM-4Z plasmid vector (Promega Corp., Madison, WI). Plasmid linearization with EcoRI and in vitro transcription using T7 polymerase results in a 320 bp antisense probe (296 bp GH sequence), whereas digestion with HindIII and in vitro transcription with SP6 results in a 344 bp sense probe (296 bp GH sequence). In vitro transcriptions were carried out using digoxigenin-UTP (Roche Molecular Biochemicals), according to instructions by the manufacturer.

Statistical analysis
Data are reported as the mean ± SEM from three or four replicate trials as described above. In each replicate, GH plaque-forming cells (% of all pituitary cells) were determined for each experimental group using three replicate chambers (at least 600 pituitary cells were counted for each treatment group per replicate). Data were analyzed using ANOVA. Where necessary, log-transformed data were used to remedy for nonhomogeneity of variance. Tukey’s test was used to compare differences between treatments. Differences were considered significant at P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Exp 1
This experiment was designed to define the ages during embryonic development that injection of corticosterone in ovo can induce premature appearance of GH-secreting cells. Three doses of corticosterone were injected (0, 0.02, 0.2, and 2 µg) on e9, e10, e11, and e12, and the percentage of GH-secreting cells was assessed in each group 2 days later, on e11, e12, e13, and e14, respectively. None of the doses injected on e9 or e10 was able to increase the population of GH-secreting cells in pituitaries derived from e11 and e12 embryos (Fig. 1Go, a and b), relative to control. On e11, the population of GH-secreting cells is below 1% (Fig. 1aGo), indicating that only occasional GH-secreting cells exist at this stage of embryonic development, as assessed by RHPA. The percentage of GH-secreting cells is slightly increased on e12 (~2%), but no statistically significant effect of any corticosterone dose injected on e10 could be observed (Fig. 1bGo). On e13, both the 0.2 and 2 µg doses of corticosterone injected on e11 (Fig. 1cGo) increased significantly the proportion of GH-secreting cells to 8.2 ± 0.6% and 6.4 ± 0.5%, respectively, relative to controls (2.4 ± 0.2%). Interestingly, injection of 0.2 µg elicited a greater response than injection of 2 µg of corticosterone. For e14 embryos treated on e12 (Fig. 1dGo), only the 2-µg dose increased the proportion of GH-secreting cells (6.4 ± 0.6%), relative to controls (3.6 ± 0.4%). It is worth noting that the 0.2 µg dose was effective on e11 but not on e12.



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  		Figure 1. Ontogeny and dose response of the corticosterone-induced premature increase of GH-secreting cells 2 days after treatment. Corticosterone (0, 0.02, 0.2, and 2 µg in 300 µl of saline) was injected into separate eggs on e9, e10, e11, and e12. The population of GH- secreting cells (% of total pituitary cells) was assessed 2 days later using reverse hemolytic plaque assays. Data represent the mean ± SEM of three replicate experiments. Means with different letters (a, b, c) are significantly different (P < 0.05).

 
Exp 2
Results obtained in the first experiment indicated that the greatest responses to corticosterone injections are observed on e13 following treatment on e11. In the second experiment, the effect of the timing of corticosterone injection was evaluated further. Four different doses of corticosterone (0, 0.02, 0.2, 2, and 20 µg) were injected on e0, e8, e9, e10, e11, and e12, and the population of GH-secreting cells was assessed on e13 in all groups. Administration of 2 and 20 µg on e0 resulted in 100% mortality. Based on the morphology of the dead embryos on e13 (the day that the eggs were opened), death occurred between e4 and e6. In addition, injection of 20 µg of corticosterone on e8, e9, and e10 resulted in some degree of mortality by e13, with the incidence being more frequent following treatment of the younger embryos. Hence, results presented from this dose of corticosterone should be interpreted cautiously. No other dose of corticosterone resulted in mortality or in any compromise of development as assessed macroscopically. None of the corticosterone doses was effective when given on e0, e8, e9, and e10, relative to controls (Fig. 2Go, a–d). Injection of 0.2, 2, and 20 µg of corticosterone on e11 (Fig. 2eGo) increased the population of GH-secreting cells (7.6 ± 0.9%, 6.7 ± 0.8% and 10.3 ± 1.1%, respectively), relative to controls (2.3 ± 0.4%). The 2 and 20 µg doses also increased GH-secreting cells when injected on e12 (5.6 ± 0.4% and 8.7 ± 0.9%, respectively), relative to controls (2.7 ± 0.5%). Apparently, treatment with 20 µg on both e11 and e12 induced the greatest responses. However, in subsequent experiments (see below), administration of 20 µg on e11 resulted in embryonic death by e18. As in Exp 1, the 0.2 µg dose was only effective when administered on e11, not on e12.



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  		Figure 2. Effects of corticosterone injections at earlier ages on somatotroph abundance on e13. Corticosterone (0, 0.02, 0.2, 2, and 20 µg in 300 µl of saline) was injected into separate eggs on e0, e8, e9, e10, e11, and e12. The population of GH-secreting cells (% of total pituitary cells) was assessed on e13 in all groups using reverse hemolytic plaque assays. Data represent the mean ± SEM of four replicate experiments. Means with different letters (a, b, c) are significantly different (P < 0.05).

 
Exp 3
This experiment was designed to examine whether corticosterone-induced premature increases in GH-secreting cells persist during subsequent embryonic development. From the results in the second experiment, it became apparent that the highest dose (20 µg) elicited the greatest responses. However, preliminary trials indicated that administration of 20, 10, and 5 µg of corticosterone on e11 resulted in 100, 80, and 40% mortality by e18, respectively, while administration of 2 µg neither resulted in any significant mortality nor appeared to compromise normal development. Thus, two groups of eggs were injected either with 2 µg of corticosterone in saline or saline alone on e11, and the number of GH-secreting cells was estimated on e13, e16, e19, and d1. The population of GH-secreting cells in corticosterone treated embryos was significantly greater than in saline treated embryos only on e13 (7.1 ± 0.8% and 2.7 ± 0.3%, respectively). No significant difference was observed on e16 (12.4 ± 1.5% and 13.6 ± 1.2%), e19 (19.0 ± 1.0% and 18.2 ± 1.7%) and d1 (23.8 ± 2.1% and 25.1 ± 1.8%) between corticosterone-treated and control embryos, respectively (Fig. 3Go).



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  		Figure 3. Eggs were injected either with 2 µg of corticosterone in saline or saline alone on e11, and the number of GH secreting-cells was estimated on e13, e16, e19, and d1 (day of hatch) using reverse hemolytic plaque assays. Data represent the mean ± SEM of three replicate experiments. Means with different letters (a, b, c, d, e) are significantly different (P < 0.05).

 
Exp 4
Despite the fact that the RHPA is a very sensitive technique and the only one available that allows evaluation of secreted hormones by individual cells, it does not provide any information on the location and distribution of these cells in the pituitary gland. Whole mount ISH can provide information on the location and the relative intensity of a signal (e.g. GH mRNA) in response to a differentiating or trophic factor (e.g. corticosterone). This experiment verified that corticosterone administration increased abundance of GH mRNA in the caudal part of the pituitary gland (where somatotrophs are located normally later in development). From a total of seven pituitaries derived from control embryos on e13, two developed weak staining (4 c) and two developed moderate staining (4 d), whereas three displayed no staining at all. In contrast, from a total of eight pituitaries derived from embryos on e13 injected with 2 µg of corticosterone on e11, six displayed strong staining similar to that in Fig. 4Go, a and b, one displayed a moderate intensity, and one a weak intensity. The intensity of staining in a pituitary gland derived from an embryo on e16 is presented in Fig. 4eGo to indicate the location of GH mRNA during normal development. For all pituitaries, staining for GH mRNA was restricted to the caudal portion of the anterior pituitary. No specific signal was observed when pituitaries were hybridized with a sense probe (4 f).



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  		Figure 4. Whole anterior pituitary mount in situ hybridization for GH mRNA. A and B are pituitary glands from embryos on e13, injected with corticosterone on e11. C and D are pituitary glands from embryos on e13 injected with saline on e11. E and F are pituitary glands from untreated embryos on e16, hybridized with antisense and sense probes, respectively.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In the present study, we defined the ability of corticosterone to prematurely induce GH-secreting cells during embryonic development of the chicken. Perhaps the most intriguing result of our experiments was the fact that injection of corticosterone before e11 did not induce any significant increase in the number of GH-secreting cells. While injection of as little as 0.2 µg of corticosterone on e11induced a substantial increase in GH-secreting cells by e13, injection of higher doses on e10 (2 and 20 µg) did not induce any increase in the number of somatotrophs by e12 or e13. This finding suggests that corticosterone cannot induce somatotroph differentiation before e13 and that corticosterone administered before e11 is rapidly cleared before it can elicit this response. As has been reported previously, injection of cortisone in the same mode and around the same period of embryonic development results in a significant increase of blood glucose within three hours (22) and a significant decrease in retina DNA synthesis within 8 h (23). These observations indicate that upon injection, glucocorticoids enter the embryonic circulation relatively fast. The half-life of corticosterone in posthach chickens is 22 min (24) and 25 and 15 min for total and free corticosterone, respectively, in 3-week-old rats (25). The half-life of corticosterone during early stages of embryonic development is probably even shorter. Studies indicate that the increase in corticosteroids in the late stages of embryonic development and early postnatal life in sheep, horse, and rat is at least partially due to reduced metabolic clearance (26, 27) as a consequence of increasing levels of corticosteroid-binding globulin (CBG). Levels of CBG increase significantly around day 15 during chicken embryonic development (28). These studies, together with our results, indicate that a single injection of corticosterone will increase circulatory levels for less than a day and that this period of elevated glucocorticoids on e11 but not on e10 is sufficient to induce an increase in GH-secreting cells. This conclusion is supported by the report that incubation of e18 rat pituitaries with glucocorticoids for 5 h is sufficient to induce an increase in GH mRNA levels 24 h later (9). In addition, incubation of e12 chicken pituitary cells with corticosterone for 48 h was sufficient to induce GH-secreting cells that were present even after 5 days in culture with corticosterone-free medium (unpublished observations).

The reason for the apparent difference in responsiveness to glucocorticoids between e10 and e11 during chick embryonic development can only be hypothesized at this point. The Pit-1 transcription factor is an absolute requirement for GH expression. However, it is unlikely that Pit-1 expression can account for the absence of responsiveness to glucocorticoids before e11. We have identified Pit-1 expression in chicken embryonic pituitaries as early as e8 by fluoresence immunocytochemistry and in greater than 30% of all pituitary cells by e10 (unpublished results). In addition, treatment of e18 rat pituitaries with glucocorticoids does not affect Pit-1 or Pit-1 mRNA levels (9). Nevertheless, the presence of other factors that act synergistically with Pit-1 might be required for activation of the GH promoter.

The biological response of cells to glucocorticoids is affected by the concentration of glucocorticoid receptors (29, 30, 31). No report exists on the ontogeny of glucocorticoid receptors during chicken pituitary development. The receptor can be detected in the pituitary gland as early as e13 (32) in mice and by e15 in rats (33). Thus, in the present experiment acquisition of responsiveness to corticosterone after e11 might result from increased expression of glucocorticoid receptor. However, expression of the receptor does not always control the developmental acquisition of responsiveness to glucocorticoids. Studies in different tissues have indicated that expression of glucocorticoid receptors precedes substantially the developmental acquisition of responsiveness to glucocorticoids (3, 34, 35, 36). The different mechanisms that control this process have not been clarified to date. However, expression of proteins that interact with the glucocorticoid receptor are likely involved.

It is unlikely that glucocorticoids directly activate GH gene transcription. The chicken GH gene has been cloned (37), and the proximal promoter region (488 bp upstream from the transcription initiation site) is lacking a distinct GRE (glucocorticoid response element). In addition, premature induction of GH mRNA by dexamethasone in rat embryonic pituitary glands is completely blocked by a protein synthesis inhibitor (9), indicating that the effects of glucocorticoids on GH gene expression are mediated by a protein synthesized in response to glucocorticoids. Preliminary studies conducted in our lab with chicken embryonic pituitary cells are in agreement with these findings. This protein might well be the GHRH receptor, as many studies indicate that glucocorticoids stimulate GHRH receptor gene expression (38, 39, 40, 41). The glucocorticoid-induced increase in GHRH receptor gene expression is inhibited by actinomycin D but not cyclohexamide (41, 42), suggesting a direct action. In addition, the promoter region of the human GHRH receptor can be regulated by glucocorticoids (42), indicating that glucocorticoids induce GHRH receptor mRNA by directly stimulating transcription. However, we have reported previously that GHRH alone is incapable of inducing somatotroph differentiation, whereas corticosterone can induce somatotrophs in the absence of GHRH (43). Therefore, if GHRH receptor expression is involved in the response to corticosterone, it would appear to be independent of GHRH activation.

In addition to increased expression of GHRH receptors, glucocorticoids may stimulate GH expression through other mechanisms. Glucocorticoid-induced transcription of many genes requires induction of the transcription factor CCAAT/enhancer binding protein {alpha} (C/EBP{alpha}). C/EBP{alpha} mediates glucocorticoid activation of the angiotensinogen gene (44), the rat tyrosine aminotransferase gene (45), the rat {alpha}-1 acid glycoprotein gene (46), the rat p21 gene (47), and the chicken glutamine synthetase gene (36). It has also been shown that C/EBP{alpha} can strongly regulate rat GH promoter activity synergistically with Pit-1 in cell lines that do not express the endogenous or transfected GH genes, and transient expression of C/EBP{alpha} in GHFT1–5 cells (pituitary cells transformed at a stage of development just before GH expression) induces an increase in GH promoter activity (48). It is of interest that the chicken GH promoter contains several consensus C/EBP{alpha} response elements and that a putative Pit-1 response element (37) is adjacent to one of them. Thus, glucocorticoids may increase or induce GH gene expression indirectly through induction of C/EBP{alpha} and/or GHRH receptor.

A close correlation between pituitary cell differentiation and vasculogenesis during chicken embryonic development has been previously indicated (49). Vessels begin to enter the gland at e6 but from e10 they markedly increase in number and size. Thus, supply of the pituitary gland with blood-borne signals is likely increased during this period, and the differentiating effect of corticosterone may be more pronounced as a result. We also cannot rule out the possibility that corticosterone injection before e11 induces some increase in GH expression that cannot be detected by RHPA. It must be kept in mind that the RHPA detects only GH-secreting cells (fully differentiated somatotrophs) and not GH- containing cells. It is our belief that only when a cell acquires the ability to secrete GH should it be considered as a terminally differentiated somatotroph. In favor of this argument, dexamethasone-induced GH cells in e17 rat fetuses displayed more immature features than those induced on e18, and cytological changes (observed by electron microscopy) indicating acquisition of secretory ability, such as enlargement of the Golgi system, development of the rough endoplasmic reticulum and accumulation of secretory granules, were evident only in e19 fetuses (50). Thus, the RHPA is the most appropriate method to detect terminally differentiated hormone secreting cells.

We have demonstrated previously that a significant increase in the population of GH-secreting cells occurs between days 14 and 16 of chicken embryonic development, although occasional somatotrophs can be detected as early as day 10 by RHPA (14) and day 6.5 by immunocytochemistry (51). Low levels of corticosterone (the primary glucocorticoid in avian species) can be detected in the circulation by day 10 of embryonic development (52, 18), and adrenals from day 8 embryos but not from day 6 have secretory capabilities after culture in vitro for 48 h (53). Around day 13–14, a substantial increase in plasma corticosterone concentrations is observed, which coincides with increased mitotic activity in the adrenals and acquisition of responsiveness to ACTH (18, 54, 55, 56). We believe that this increase in circulatory corticosterone drives the substantial increase in the population of GH- secreting cells between days 14 and 16 of embryonic development. Occasional somatotrophs can be detected as early as day 15 during rat embryonic development (57). However a substantial increase in the population of somatotrophs occurs around day 19–20 (10, 58). Fetal plasma corticosterone concentrations reach a peak on day 19 of rat embryonic development (59), supporting a role for endogenous glucocorticoids in augmenting somatotroph abundance. In favor of this theory, culture of pituitary cells from e12 chicken embryos with corticosterone or serum from e16 embryos, but not with serum from e12 embryos, increased the proportion of GH-secreting cells (16). In addition, administration of corticosterone in ovo to embryos on e11 resulted in a substantial increase of GH-secreting cells by embryonic day 14 (17). In the present study, administration of various doses of corticosterone on e11 or e12 resulted in increased proportions of GH-secreting cells and increased abundance of GH mRNA by e13. Similarly, treatment of pregnant rats with dexamethasone on day 16 or 17 of pregnancy, induced premature expression of GH in the pituitaries of their fetuses by days 17 and 18, respectively (10, 11). Taken together with our current findings, these observations indicate that somatotroph recruitment occurs in response to increased adrenal glucocorticoid production and that administration of glucocorticoids before this endogenous increase results in premature somatotroph differentiation.

A reduction in response to corticosterone was observed between e11 and e12. The 0.2 µg dose was effective when administered on e11 but not on e12. However, higher doses of corticosterone (2 and 20 µg) injected on e12 were still able to increase the population of GH-secreting cells. As was mentioned earlier, an increase in CBG occurs around this time of embryonic development. This increase can reduce the amount of bioavailable corticosterone and explain the shift in sensitivity to corticosterone between e11 and e12.

With the use of whole pituitary mount ISH for GH mRNA, we confirmed that the corticosterone-induced increase in GH cells occurs in the caudal lobe of the pituitary gland, where normal somatotroph differentiation takes place. Furthermore, the corticosterone-induced premature increase in GH cells did not affect the population of somatotrophs later in development. Injection of corticosterone on e11 increased the number of GH-secreting cells in pituitaries from e13 but not from e16 and e19 embryos and d1 chicks. These observations indicate that glucocorticoids increase GH gene expression and GH secretion in a pool of cells that are predetermined to become somatotrophs.

In conclusion, corticosterone can increase the population of GH-secreting cells only during a small window of development between e11 and e13. This window is defined by the absence of responsiveness to corticosterone before e11 and by the endogenous increase of corticosterone and CBG on e13-e14 and normal somatotroph differentiation by e14-e16. The premature increase in the abundance of GH cells induced by in ovo corticosterone administration occurs in the caudal part of the pituitary gland and does not affect the somatotroph population later in development.


   Footnotes
 
1 This work was supported by USDA Grant 97–35206-5086(to T.E.P). Back

Received December 23, 1999.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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