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Department of Poultry Science, Texas A&M University (C.E.D.), College Station, Texas 77843; and the Department of Animal and Avian Sciences, University of Maryland (T.E.P.), 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
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Abstract |
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Introduction |
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The purpose of the present study was to investigate the role of extracellular factors in regulation of somatotroph differentiation and GH secretion during embryonic development, using the chick embryo as a model system. The chicken embryo is a useful model for studying the influence of extracellular factors on pituitary cell differentiation due to its isolation from maternal influences and the relative ease with which the endocrine environment of individual embryos can be manipulated. Somatotrophs first appear between embryonic day (e-) 12 and e-14 and become a significant population by e-16 of chick development (15, 16, 17). This ontogenic profile of somatotroph differentiation correlates with the capacity of embryonic serum to induce GH cell differentiation in vitro (18). Full differentiation of functional chicken somatotrophs continues gradually from e-16 through at least e-20 of the 21-day incubation period (19). The mean proportion of GH cells in pituitaries from a mixed sex population of e-20 chicks was determined to be 19.5% (19). Typical somatotroph percentages in male and female chickens between 4 and 5 weeks of age are about 20% and 13% of the total pituitary cells, respectively (20). Thus, practically all somatotrophs present in the posthatch chicken are differentiated during the embryonic period. Chicken GH cells do not differentiate in culture without an extrapituitary signal (18); however, somatotroph differentiation in the chicken embryonic pituitary can be induced in vitro by glucocorticoids, and corticosterone is responsible for the GH cell-differentiating activity of e-16 chicken serum (21). A role for induction of somatotroph differentiation by corticosterone in vivo is supported by reports that an increase in circulating corticosterone levels occurs before GH cell differentiation on e-16 (22). Pituitary GH release in chickens, as in other vertebrates, is controlled by stimulatory and inhibitory factors from the hypothalamus. GHRH and TRH are thought to be the primary GH secretagogues, whereas somatostatin decreases GH secretion, possibly by antagonizing the effects of GHRH and TRH. Synthetic human GHRH (hGHRH) has been shown to release GH in vivo in both young (23, 24) and adult chickens (24). hGHRH also stimulated GH release from chicken pituitaries in vitro (24, 25). Although TRH is an effective GH secretagogue in immature and anesthetized birds, it has little effect on GH secretion in conscious adult chickens (26). The effects of hGHRH and TRH on in vitro GH release during the embryonic period depend on developmental stage. Between 50–70% of initial somatotrophs present on e-16 released GH in response to hGHRH-(1–40) (15, 19), whereas only 30% of GH cells responded to TRH (19). By e-20, the proportions of somatotrophs that responded to hGHRH1–40 and TRH were approximately equal at about 40% (19). hGHRH and TRH were shown to increase plasma GH concentrations approximately equally on e-18 (27).
To date, the effect of corticosterone on GH cell differentiation in the chicken has been evaluated using only e-12 pituitary cells, an age when somatotrophs are rare. Moreover, potential interactions between glucocorticoids and other extracellular factors that could affect somatotroph differentiation and GH secretion have not been investigated. Therefore, the objective of the present study was to evaluate the effects of corticosterone and GHRH on GH secretion and somatotroph differentiation at different stages of embryonic development, using pituitary cells from e-12, -14, and -17 chicks.
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Materials and Methods |
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Extended cell cultures
Anterior pituitary cells were cultured according to the
procedure described previously (18). Cells were plated (2.0 x
105 cells/well) in poly-L-lysine-coated 12-well
tissue culture plates and allowed to attach for 45 min. Wells were then
filled (2 ml) with serum-free medium alone or medium containing
corticosterone (10-9 M) or synthetic
hGHRH-(1–40) (10-10-10-7 M;
Sigma Chemical Co., G8770) alone or in combination. The
medium consisted of a 1:1 mixture of phenol red-free medium 199 and
Ham’s F-12 nutrient mixture supplemented with 0.1% BSA, 5 µg/ml
human transferrin, 5 µg/ml bovine insulin, 100 U/ml penicillin G, and
100 µg/ml streptomycin sulfate. Culture plates were prepared in
triplicate, and after culture intervals of 2, 3, and 6 days in a
humidified incubator (37.5 C; 95% air-5% CO2), cells were
harvested for detection of GH-secreting cells by reverse hemolytic
plaque assay (RHPA). Culture medium was aspirated from the wells and
replaced with fresh medium and treatments after 3 days of incubation.
The GH content of medium samples collected at the end of each
incubation interval was assessed by RIA (28). All samples were assayed
in a single RIA, and the intraassay coefficient of variation was 5.6%.
The sensitivity of this assay was 2.5 ng/ml.
RHPA
The RHPA procedure allows for detection of hormone secretion
from individual cells (29). The assays were performed according to the
protocol described in detail previously (30), using rabbit antiserum
against chicken GH and modifications described previously (15).
Briefly, recovered anterior pituitary cells were mixed with an equal
volume of an 18% suspension of protein A-coated ovine erythrocytes and
infused by capillary action into previously constructed Cunningham
chambers. After cells were allowed to attach for 45 min (37.5 C; 95%
air-5% CO2), chambers were rinsed with DMEM to remove
unattached cells. DMEM containing GH antiserum (1:40) and hGHRH-(1–40)
(10-7 M) was then added to the resulting
monolayer of cells, and replicate chambers were incubated for 8 or
20 h (three chambers per treatment per time point). Plaque
formation was subsequently induced by a 45-min incubation with guinea
pig complement (1:40, in DMEM). The cells were then fixed with 2%
glutaraldehyde in 0.9% saline and stained with methyl green. Chambers
were analyzed in two ways using a light microscope. First, the
percentage of all pituitary cells that formed plaques was determined,
with at least 200 pituitary cells counted/assay chamber. Second, the
area of each plaque formed was measured with the aid of an ocular
reticle.
Statistical analysis
The data reported are the mean ± SEM from at
least 3 completely separate experiments, with the number of replicate
experiments provided in the legend to each figure. For each replicate
experiment, percentages of GH plaque-forming cells were determined for
each combination of treatment and time point using the three replicate
chambers for that combination (for a total of at least 600 pituitary
cells analyzed for each combination). All data were analyzed using the
general linear models procedure of SAS (31). Differences between
treatments were tested using Tukey’s Studentized range test and were
considered significant at P < 0.05.
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Results |
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and 2 for e-12 and e-17 pituitary cells,
respectively. As fresh medium and treatments were added after 3 days of
incubation, the 6-day data represent accumulation of GH during the
final 3 days of culture only. Neither corticosterone nor GHRH alone
significantly changed the amount of GH released by e-12 pituitary cells
after 2 and 6 days of incubation relative to basal control levels (Fig. 1). In contrast, the combination of corticosterone and GHRH increased
GH release by e-12 cells in a concentration-dependent manner to levels
significantly higher than control values or either hormone alone after
2 and 6 days. The results were similar for e-12 cells exposed to
corticosterone and GHRH alone and in combination for 3 days (data not
shown). Data for e-14 cultures (not shown) were essentially the same as
those for e-12 cells. Corticosterone alone did not affect GH release
from e-17 cells (Fig. 2); however, GHRH alone significantly increased
medium GH content relative to basal controls after 2 and 3 days.
Corticosterone plus GHRH also increased GH release from e-17 cells in a
concentration-dependent manner after 2 days, but this treatment was no
more effective than GHRH alone.
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Treatment with corticosterone alone for 2, 3, and 6 days increased
(P < 0.05) the percentage of GH-secreting cells to
8.8 ± 0.8%, 8.9 ± 1.2%, and 9.6 ± 0.8% of all
pituitary cells, respectively, compared with basal levels of 0.6
± 0.1%, 0.8 ± 0.1%, and 0.6 ± 0.1% of cells in cultures
derived from e-12 pituitaries. The effect of corticosterone alone was
not significantly augmented by incubating the cells longer than 2 days.
In contrast, GHRH alone did not alter the percentage of GH-secreting
cells compared with basal levels after 2, 3, or 6 days
(P > 0.05). Treatment with corticosterone plus GHRH
increased the percentage of somatotrophs relative to basal levels after
2, 3, and 6 days, but was not more effective than corticosterone alone
at 2 days. By 3 days, the combination of corticosterone and GHRH was
significantly more effective than corticosterone alone for inducing
somatotroph differentiation, but only at the highest concentration
(10-7 M) of GHRH tested. However, percentages
of GH-secreting cells detected after 6 days of combined treatment with
corticosterone plus all concentrations of GHRH tested were
significantly increased (to as much as 15.9% of all pituitary cells),
relative to those after treatment with corticosterone alone, indicating
at least an additive induction of somatotroph differentiation by these
hormones on e-12 pituitary cells. The effects of corticosterone and
GHRH on proportions of e-14 GH cells (data not shown) were similar to
the effects on e-12 cells.
The effects of corticosterone and GHRH on proportions of e-17 GH cells
were markedly different from those on e-12 and e-14 cells (Fig. 2).
First, treatment with corticosterone alone did not change percentages
of e-17 somatotrophs relative to basal levels of 12.3 ± 2.4%,
11.2 ± 1.6%, and 8.5 ± 2.3% after 2, 3, and 6 days,
respectively. In contrast, GHRH decreased the percentages of GH cells
detected in a concentration-dependent manner after all three incubation
intervals. Treatment with corticosterone plus GHRH tended to counteract
the reduction of somatotroph proportions induced by GHRH alone. The
percentages of e-17 GH cells detected after combined treatment with
corticosterone and GHRH for 2 and 6 days did not differ from those
after treatment with corticosterone alone.
Effects of corticosterone and GHRH on GH secretion in RHPA
To determine the influence of exposure to treatments during
extended culture on subsequent GH secretion, we measured plaque area as
an index of the relative amount of GH secreted from individual
somatotrophs. These results are shown for e-12 and e-17 cells in Figs. 1 and 2, respectively. Cells from e-12 that were exposed to GHRH alone
in culture for 2, 3, and 6 days subsequently released more GH in the
RHPA than cells that were cultured under basal conditions. These
increases were apparent for cells cultured with each concentration of
GHRH for 2 days and with at least one concentration of GHRH tested for
3 and 6 days. Cells cultured with corticosterone alone and
corticosterone in combination with GHRH for 2, 3, and 6 days
subsequently released more GH in RHPA than cells cultured under basal
conditions or with GHRH alone. Finally, cells exposed to corticosterone
in combination with GHRH for 2 and 3 days, but not 6 days, released
more GH in RHPA than cells cultured with corticosterone alone (Fig. 1).
Similarly, e-14 pituitary cells cultured with corticosterone alone and
in combination with GHRH for 2, 3, and 6 days subsequently released
more GH in RHPA than cells cultured under basal conditions, and
exposure to both corticosterone and GHRH was significantly more
effective than that to either hormone alone for increasing subsequent
GH secretion (data not shown).
Culture of e-17 pituitary cells with GHRH alone tended to decrease
subsequent GH secretion, with exposure to the highest concentration for
2 days resulting in a significant decrease relative to control values
(Fig. 2). Both short and long term exposures to corticosterone alone
failed to increased subsequent GH secretion by e-17 cells relative to
controls. Finally, e-17 cells exposed to corticosterone plus GHRH for 6
days, but not for 2 and 3 days, subsequently released significantly
more GH than cells exposed to corticosterone alone.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Pituitary cells were subjected to GH RHPA after hormone exposure for 2–6 days to obtain two types of information. First, percentages of GH-secreting cells were determined as a measure of how corticosterone and GHRH affected the extent of somatotroph differentiation. Second, the mean plaque area was measured as an index of the relative amount of GH secreted by the somatotrophs after acute (8-h) GHRH stimulation. The results indicate that both corticosterone and GHRH affected e-12 and e-17 pituitary cells differently. Furthermore, the responses of cells from the two developmental stages were affected by the duration of exposure to the hormones. Short and long term exposures to corticosterone dramatically increased somatotroph percentages among pituitary cells from e-12, an age when somatotrophs are normally rare. Glucocorticoids have also been shown to induce premature differentiation of fetal rat somatotrophs in vitro (4, 5, 6). Corticosterone also increased GH cell proportions on e-14, but not e-17, in the present study, suggesting that the induction of somatotroph differentiation by glucocorticoids is not possible at this later stage of development in the chicken. Treatment of e-12 pituitary cells with corticosterone alone raised somatotroph proportions to as high as 9.6%, which was less than the maximum percentage of e-17 GH cells (12.3%) detected under basal conditions; however, corticosterone increased the proportion of e-14 somatotrophs to as much as 15.7%. Thus, corticosterone was capable of increasing somatotroph proportions on e-14 to levels greater than those observed under basal conditions on e-17 and to levels comparable with those observed in posthatch chickens. Although treatment of e-12 pituitary cells with GHRH alone did not affect somatotroph proportions, combined treatment with corticosterone and GHRH for 6 days increased e-12 GH cell percentages to levels exceeding those observed after treatment with corticosterone alone. The effects of corticosterone and GHRH were truly synergistic, because GHRH alone was ineffective, and the combination increased somatotroph proportions to as much as 15.9% compared to 9.6% with corticosterone alone. One possible explanation for this effect is that GHRH stimulated mitosis of somatotrophs that had been induced to differentiate by corticosterone. Indeed, GHRH has been shown to stimulate the proliferation of differentiated rat somatotrophs in vitro (12). Interestingly, the synergistic effect of corticosterone and GHRH required exposure to the hormones for more than 3 days. Perhaps cells were first induced to express GHRH receptors by corticosterone and then stimulated to release GH by GHRH. The mechanism of this effect cannot be determined from this study, because no attempt was made to evaluate somatotroph proliferation. The synergistic effect of corticosterone and GHRH on GH cell proportions was essentially lost by e-14, about the time when somatotroph differentiation normally begins in vivo.
The effects of corticosterone and GHRH on e-17 pituitary cells were markedly different from those on e-12 and e-14 cells. Although corticosterone did not increase the percentages of e-17 GH cells, treatment with GHRH decreased the percentages of e-17 somatotrophs detected in a concentration-dependent manner; however, this apparent reduction in somatotroph proportions probably reflects decreased sensitivity of the cells to GHRH rechallenge in the RHPA. This phenomenon, referred to as homologous desensitization, was observed when rat anterior pituitary cells were pretreated for 24 h with GHRH (33). When those cells were rechallenged with GHRH, maximal GH secretion was significantly lower than the control value, and releasable GH pools were reduced by 5-fold. Recently, GHRH was shown to inhibit the production of its own receptor in as few as 4 h by reduction of GHRH receptor messenger RNA accumulation in cultures of rat anterior pituitary cells (34). In the present study, treatments were removed from cells for approximately 3 h during the transition from cell culture to RHPA. During this time, cells were recovered from the culture wells, rinsed, and allowed to attach to the RHPA chambers. As all RHPAs were performed under stimulatory conditions in this study (i.e. in the presence of GHRH), cells that were initially cultured with GHRH were, in effect, rechallenged with GHRH during the RHPA. Because detection of somatotrophs by RHPA depends on GH secretion, desensitization to GHRH rechallenge might explain the apparent dose-related reduction in GH cell percentages determined by RHPA. In other words, exposure to GHRH in culture may have decreased the percentage of somatotrophs that released GH when rechallenged with GHRH during RHPA. Treatment of e-17 pituitary cells with corticosterone and GHRH in combination appeared to moderate or eliminate the loss of responsiveness caused by prolonged exposure to GHRH. This finding could reflect corticosterone-induced sensitization of somatotrophs to GHRH. Glucocorticoids have been shown to enhance GHRH-induced GH secretion in primary cultures of rat anterior pituitary cells (35, 36) and in freely moving rats (37). Furthermore, glucocorticoids have been shown to increase GHRH receptor gene expression (38) and the number of GHRH-binding sites on rat anterior pituitary cells (39). Another possibility is that corticosterone increased GH protein synthesis in e-17 cells, an effect that has also been documented in rat anterior pituitary cells (4, 5, 6, 40). Thus, corticosterone could be important for sustaining releasable GH pools and regulating sensitivity to secretagogues during late embryonic development when chicken somatotrophs become responsive to GHRH and TRH (19).
That treatment of e-12 and e-14 pituitary cells with GHRH alone failed to significantly increase the medium GH content was not surprising, as only small percentages of somatotrophs were detected at these ages. Others have failed to observe GHRH-stimulated GH release from chicken pituitaries until after e-14 (41), when the adenohypophyseal vascular sinusoids attain their adult structure (42). That e-17 cells did release GH in response to GHRH was also not surprising, as 50–70% of somatotrophs become responsive to GHRH by e-16 (15, 19). Unexpectedly, the medium GH content was not increased by treatment of e-12 and e-14 cells with corticosterone alone, even though corticosterone substantially increased somatotroph percentages at these ages. This result was surprising in view of previous findings that somatotrophs induced to differentiate prematurely by corticosterone synthesized and released GH in the absence of GHRH (18, 21). However, others have reported that dexamethasone had little effect on basal GH release (38, 43). Although corticosterone alone failed to increase GH secretion in the present study, combined treatment with corticosterone and GHRH increased GH secretion from pituitary cells of all ages in a concentration-dependent manner. This finding confirms that somatotrophs, whether differentiated normally in vivo or induced to differentiate prematurely with corticosterone in vitro, are responsive to GHRH.
The effects of treatments in culture on subsequent GH release in RHPA were determined by measuring plaque area. Interestingly, the responses of e-12 and e-17 cells were different under most conditions. Treatment with GHRH may have stimulated the few somatotrophs present on e-12 to synthesize more GH, thus increasing the pool available for release during RHPA. Indeed, GHRH increased GH gene transcription (44) and GH synthesis (33) in rat anterior pituitary cells in vitro. The increased mean plaque area of e-12 cells cultured with corticosterone alone might also reflect increased GH synthesis as well as greater responsiveness to GHRH. Glucocorticoids have been shown to induce GH synthesis (4, 5, 6, 40), to increase GHRH binding capacity (39), and to enhance GHRH-induced GH secretion in cultures of rat anterior pituitary cells (35, 36). In contrast, the reduced plaque area of e-17 cells after treatment with GHRH could reflect decreased sensitivity due to down-regulation of GHRH receptors. The increased capacity of e-12 cells to secrete GH after exposure to corticosterone and GHRH in combination for 2 days suggests an additive effect of the treatments; however, 6 days of combined treatment failed to increase GH secretion more than corticosterone alone, suggesting that the latter treatment exerted a maximal effect by 6 days.
In conclusion, corticosterone induced GH cell differentiation in vitro on e-12 and e-14, and the somatotroph-differentiating activity of corticosterone was magnified by concomitant exposure to GHRH. Corticosterone did not induce GH cell differentiation in vitro on e-17, but sustained the capacity of e-17 somatotrophs to release GH in response to GHRH. Somatotrophs normally appear during chicken embryonic development after a rise in circulating glucocorticoid levels (22), and corticosterone is responsible for the GH cell-differentiating activity of e-16 chicken serum (21). Taken together, these findings provide strong evidence that glucocorticoids are involved in GH cell differentiation during embryonic development, and that GHRH may act in concert with corticosterone to stimulate differentiation and expansion of the somatotroph population.
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Footnotes |
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Received July 20, 1998.
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