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Original Articles: Hormones and Reproductive Health |
Departments of Endocrinology and Metabolism (F.R., N.R.B., R.G.V.), Pediatrics (W.H.S.-B., J.-M.W.), and Clinical Chemistry (M.F.), Leiden University Medical Center, 2333AA Leiden, The Netherlands; Division of Endocrinology and Metabolism, National Science Foundation Center for Biological Timing, General Clinical Research Center (J.D.V.); and Department of Internal Medicine, University of Virginia Health Sciences Center (J.D.V.), Charlottesville, Virginia 22908
Abstract
GH secretion is regulated by the interaction of GHRH and somatostatin and is released in 10–20 pulses in each 24-h cycle. The exact roles in pulse generation played by somatostatin, GHRH, and the recently isolated GH-releasing peptide, Ghrelin, are not fully elucidated. To investigate the GHRH-mediated GH secretion in human, we investigated pulsatile, entropic, and 24-h rhythmic GH secretion in two young adults (male, 24 yr; female, 23 yr) from a Moroccan family with a novel inactivating defect of the GHRH receptor gene. Data were compared with values in age- and gender-matched controls. Plasma GH concentration were measured by a sensitive immunofluorometric assay, with a detection limit of 0.01 mU/L. All plasma GH concentrations in the female patient were measurable; in the male patient 30 of 145 samples were at or below the detection limit. GH secretion was pulsatile, with 21 and 23 secretory episodes/24 h in the male and female patients, respectively. The fraction of basal to total GH secretion was raised in both patients by 0.18 and 0.15, respectively. The total 24-h GH production rate was greatly diminished; in the male patient it was 6.9 mU/L (normal values for his age, 26–63 mU/L), and in the female patient it was 4.2 mU/L (normal values for her age, 96–390 mU/L). The nyctohemeral plasma GH rhythm was preserved (P < 0.001), with normal acrophases (0430 and 0218 h in the male and female, respectively). Approximate entropy was greatly elevated in both subjects (0.82 in the male and 1.17 in the female; upper normal values for age and gender, 0.24 and 0.59, respectively). Intravenous injection of 50 µg GHRH failed to increase the plasma GH concentration in both patients, but 100 µg GH-releasing peptide-2 elicited a definite increase (male patient, 0.13 to 1.74 mU/L; female patient, 0.29 to 0.87 mU/L). Both patients had a partial empty sella on magnetic resonance imaging scanning.
In summary, the present studies in two patients with a profound loss of function mutation of the GHRH receptor favor the view that in the human the timing of GH pulses is primarily supervised by intermittent somatostatin withdrawal, and the amplitude of GH pulses is driven by GHRH. In addition, we infer that effectual GHRH input controls the GH cell mass and the orderliness of the secretory process.
GH IS SECRETED in a pulsatile fashion with 10–20 GH secretory events occurring within a 24-h cycle, which arise as small amplitude pulses during the waking hours and larger pulses during sleep. The underlying mechanisms of pulse generation are still unclear, especially in the human. For example, the interactive contributions of the major hypothalamic peptides that regulate pulsatile GH secretion, i.e. GHRH and somatostatin are not well defined. This is illustrated, for instance, by the observation that constant iv infusion of GHRH in healthy subjects amplifies pulsatile GH secretion, suggesting that the timing of GH pulses is driven by intermittent with- drawal of hypothalamic somatostatin and/or the release of cosecretagogues of GHRH (1). On the contrary, a GHRH antagonist suppresses the amplitude of GH release in men, indicating that GHRH is essential to maintain the amplitude of GH pulse generation (2).
Recently, several groups of investigators have reported dwarfism in various countries, caused by an autosomal recessively inherited mutation of the GHRH receptor gene (3, 4, 5). The afflicted members of these families are characterized by severe growth retardation.
The present study examines GH release in two members of a Moroccan family with severe growth retardation attributed to a novel inactivating defect of the GHRH receptor gene. These patients therefore are ideal candidates for investigating spontaneous GH secretion patterns in the absence of an effectual GHRH signal. We hypothesized that GH would still be secreted in very low amplitude bursts, but in a pulsatile fashion due to preserved timing mechanism mediated via intermittent withdrawal of somatostatin and/or release of a non-GHRH cosecretagogue.
Subjects and Methods
Patients
The patients were referred because of short stature. The boy was aged 16 yr, extremely short (height SD score, -5.7), and puberty had progressed to Tanner stages P3–4, G3, and A2 (testes volume, 16 mL). The girl was 14.9 yr at presentation, had a height SD score of -7.7, and Tanner stages M2, P1, and A1. GH deficiency was confirmed by a deficient rise of GH during arginine infusion; the maximal serum GH concentration (measured by RIA) in the boy was 5 mU/L, and that in the girl was 1.2 mU/L. On computed tomography scanning, both patients had an empty sella configuration. Except for a slightly diminished maximal TSH rise after TRH injection, no other endocrine abnormalities were found, and the diagnosis of familial GH deficiency was made. The patients were treated for, respectively, 5.2 and 5.5 yr and reached their target adult heights.
In the first DNA analysis the GH gene was found to be normal. More recently, after extraction of genomic DNA each exon of the coding sequence of the GHRH receptor gene was amplified by PCR and subjected to single strand conformation polymorphism analysis, followed by complete sequencing. A recessive point mutation G-C in the first position of the donor splicing site of intron 7 was shown, abolishing correct splicing and resulting in an extremely truncated protein (6). Further details will be reported elsewhere.
Endocrine investigations after discontinuation of GH substitution therapy
At chronological ages of 24 yr (male) and 23 yr (female), basal serum concentrations of hormones, including free T4, T3, cortisol, testosterone, estradiol, progesterone, insulin-like growth factor I (IGF-I), and IGF-binding protein-3 were measured. In addition, the following stimulation tests were performed at intervals of 4 or more weeks: TRH test (200 µg, iv), GHRH test (50 µg, iv), and GH-releasing peptide-2 (GHRP-2) test (100 µg, iv). The following hormones were measured; TRH test: TSH, PRL, and GH at -15, 0, 15, 20, 30 45, 60, 90, and 120 min; GHRH test: GH and PRL at 0, 20, 30, 45, 60, and 90 min; and GHRP-2 test: GH, PRL, TSH, and ACTH at -15, 0, 15, 20, 30, 45, 60, 90, 120, 150, and 180 min. To obtain 24-h GH secretion profiles the patients were hospitalized, and an indwelling iv cannula was inserted in a forearm vein for blood withdrawal at 10-min intervals. The patients were free to move around, but not to sleep, during the daytime. Meals were served at 0800, 1230, and 1730 h. Lights were turned off between 2200–2400 h. Five healthy male and seven healthy female volunteers served as controls and underwent an identical sampling study. The mean age of male controls was 27.1 ± 2.5 yr, and that of female controls was 28.4 ± 1.8 yr. The body mass index (BMI) in the male subjects was 24.2 ± 0.7 kg/m2, and that in the female controls was 20.8 ± 0.6 kg/m2.
Assays
Plasma GH was measured with a sensitive time-resolved fluoroimmunoassay (Wallac, Inc., Turku, Finland). The assay is specific for the 22-kDa GH protein. The standard was biosynthetic recombinant human GH (Genotropin, Pharmacia & Upjohn, Inc., Uppsala, Sweden) and was calibrated against the WHO First International Reference Preparation 80/505 (to convert micrograms per L to milliunits per L, multiply by 2.6). The limit of detection of this assay (defined as the value 2 SD above the mean value of the zero standard) was 0.01 mU/L (0.0038 µg/L). The intraassay coefficient of variation varied between 1.6–8.4% in the range of 0.01–18 µg/L GH, and the interassay coefficient of variation was 2.0–9.0% in the same range. PRL was measured with a sensitive time-resolved fluoroimmunoassay (Wallac, Inc., Turku, Finland). The standard was calibrated against the WHO Third International Standard for PRL 84/500 (to convert micrograms per L to milliunits per L, multiply by 36).The limit of detection (defined as 2 SD above the mean value of the zero standard) was 0.04 µg/L. The intraassay coefficient of variation varied from 2.0–3.3% in the assay range from 3.0–80 µg/L, and the corresponding interassay coefficient of variation was 3.4–6.2%.
TSH was measured with a time-resolved fluoroimmunoassay (Wallac, Inc.). The detection limit of this assay was 0.05 mU/L. The interassay coefficient of variation was less than 5%, and the intraassay coefficient of variation was 4.4% in the 0.15–3.2 mU/L range. ACTH was measured by an immunoradiometric assay, using reagents obtained from Nichols Institute Diagnostics (San Juan Capistrano, CA). The detection limit of this assay is 3.0 ng/L, and the intra- and interassay coefficients of variation ranged from 2.8–7.5%. The plasma cortisol concentration was measured by RIA (Sorin Biomedica, Milan, Italy). The detection limit of this assay is 25 nmol/L. The intra- and interassay coefficients of variation ranged from 2.0–4.0%.
Total IGF-I was determined by RIA (INCSTAR Corp., Stillwater, MN) after extraction and purification on ODS-silica minicolumns. The intra- and interassay coefficients of variation were less than 11%. The detection limit was 1.5 nmol/L. Age-related normal data were determined in the same laboratory. The measurement of IGF-binding protein-3 was performed by RIA (Nichols Institute Diagnostics). The limit of detection of this assay was 0.08 mg/L, and the interassay coefficient variation was below 6.8%.
Analytical techniques
For the GH time series, multiple parameter deconvolution analysis was used to estimate various specific measures of pulsatile secretion and half-life based on all plasma hormone concentrations and their dose-dependent intrasample variances considered simultaneously (7, 8). Results were expressed as milliunit (mU) per L distribution volume. The total 24-h production rate was calculated using a nominal GH distribution volume of 7.9% of body weight (9).
The minute to minute regularity or serial orderliness of GH secretion was quantitated by the approximate entropy (ApEn) statistic, a scale- and model-independent metric (10). Normalized ApEn parameters of m = 1 and r = 20% of the intraseries SD were applied, as previously validated for GH series of this length (11). This member of the ApEn set is designated ApEn (1, 20%). ApEn estimates the regularity of subordinate (nonpulsatile) patterns in the data, and as such yields information complementary to deconvolution and cosine-dependent techniques (12, 13).
The diurnal rhythmicity of plasma GH concentrations was appraised by cosinor analysis. The latter entails trigonometric regression of a 1440-min periodic cosine function on the full 24-h serum GH concentration profile vs. time.
Results
Baseline studies
At baseline (
2 yr after GH replacement therapy was completed),
the patients had no complaints related to GH deficiency. The male
patient had reached a height of 167.4 cm, a body weight of 55 kg, and a
BMI of 19.7 kg/m2. The height of the female
patient was 156.8 cm, her weight was 48.5 kg, and her BMI was 19.6
kg/m2. The female patient reported regular
menstrual cycles. The plasma testosterone concentration in the male was
18.4 nmol/L (normal range, 13–32 nmol/L). Plasma free
T4 and T3 concentrations
were normal in both subjects. The plasma IGF-I concentration in the
female subject was 3 nmol/L (normal value for age, 20–30 nmol/L), and
that in the male subject was 10 nmol/L (normal values for age, 18–32
nmol/L).
Specific studies: effects of GHRH,GHRP-2, and TRH stimulation
After GHRH injection, serum GH concentrations failed to increase
(see Fig. 1
), in contrast to an abrupt
increase after GHRP-2 infusion. Nevertheless, the latter stimulated
values remained far below the normal maximum for this age group, which
approximate 80- to 100-fold. Thus, in the male subject plasma GH
increased from 0.13 mU/L to a maximal of 1.74 mU/L; respective values
for the female patient were 0.29 and 0.87 mU/L after GHRP-2
administration. GHRH did not stimulate PRL release, but GHRP-2 induced
a definite, although small, increase (see Fig. 2). In the male patient, PRL increased
from 8.7 to 12 µg/L, and in the female patient PRL increased from 16
to 19 µg/L. TRH caused a normal increase in plasma concentrations of
TSH and PRL and no change in the circulating GH concentration, as
illustrated in Fig. 3.
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The 24-h serum GH concentration profiles of the patients are
depicted in Fig. 4, together with that of
a representative gender- and age-matched control. The figure
illustrates the extremely low plasma GH values in the patients compared
with normal individuals. In the female patient all plasma samples were
measurable, but in the male patient 30 of 145 samples were at or below
the limit of detection, especially in the late afternoon and beginning
of the evening. Importantly, GH secretion was clearly pulsatile, as
assessed objectively by deconvolution analysis. The results of the
deconvolution analysis are detailed in Table 1 and
show a slightly increased number of secretory events and relatively
accentuated basal GH secretion, as illustrated by the high normal value
of the ratio of basal to total secretion. Cosinor analysis also
established a significant 24-h rhythmicity (P <
0.001), with acrophases of 0430 h (male) and 0218 h
(female).
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In the male patient, ApEn for GH was 0.82 (normal median for age- and gender- matched controls, 0.19; range, 0.10–0.24). Likewise, GH ApEn in the female patient, which was 1.17, was also greatly increased. Normal values for her age and gender are: median, 0.44; and range, 0.13–0.59.
Discussion
The genetic defect in our patients was caused by a point mutation in the donor splicing site of intron 7, resulting in a severely truncated GHRH receptor protein. The defect caused severe growth retardation in both patients. The growth retardation was successfully treated with rhGH replacement therapy in combination with a GnRH agonist for 2.5 yr, because of their advanced pubertal stage at presentation. Complete loss of function of the mutated GHRH receptor was inferred by the absence of membrane anchoring and signal transduction regions in the resultant mutant protein and was established further by the absence of any significant increase in plasma GH (and PRL) concentrations after GHRH administration.
The pulsatile secretion of GH is regulated predominantly by two hypothalamic peptides, GHRH and somatostatin (14). Several studies in animals, including the rat, sheep, and pig, have demonstrated a pulsatile mode of release of these peptides into the hypophyseal portal blood, but failed to document a universal association of peripheral GH pulses with portal blood GHRH pulses or somatostatin withdrawal (15, 16, 17, 18, 19).
GHRH receptor-blocking agents in the human greatly diminish the amplitude of pulsatile GH secretion, thereby demonstrating that GHRH is a key substance for amplifying GH pulses, but not for the timing of discrete pulse generation (2). The amplifying role of GHRH is consistent with observations in the human that 24-h GHRH infusions in normal volunteers increases the amplitude, but not the number, of GH pulses (1). Accordingly, a plausible inference is that somatostatin is involved in the timing of pulse generation. Indeed, in both rats and humans, acute somatostatin withdrawal evokes a rebound-like pulse of GH secretion, although the absolute amount produced is generally small compared with maximal exogenous GHRH-induced GH release (20, 21). Interestingly, in one study this effect was not blocked by pretreatment with a GHRH antagonist, suggesting that the increase in GH is GHRH independent (22). The observation of low amplitude spontaneous pulsatile GH release in our patients would support the foregoing idea of GHRH-dependent GH pulse amplitude and somatostatin-generated pulses, as the patients had no functional GHRH receptor. In brief, the present data support a bipartite mechanism of amplitude control by GHRH and pulse timing control by somatostatin or a non-GHRH cosecretagogue (below).
During an 8-h somatostatin infusion in healthy subjects GH pulse frequency decreased, but GH pulses were still present in several subjects (23). This observation seems to contradict the hypothesis that in the human GH pulse timing is regulated by somatostatin withdrawal in hypophyseal portal blood. However, it is feasible that the relative contribution of hypothalamic somatostatin is more important than that of somatostatin reaching the somatotrope via the systemic circulation. In effect, the physiological role of somatostatin in GH pulse timing can only be addressed with certainty in subjects with nonfunctional pituitary somatostatin receptors.
Two decades ago, Bowers developed GH-releasing peptides by modification of the met-enkephalin molecule (24). A recently developed peptide is GHRP-2, one of the most potent GHRPs (25), which acts via the specific cloned GHRP receptor (26). Maximal GH-releasing effects, however, are observed in vivo rather than in vitro, suggesting that hypothalamic factors play a role in potentiating the effect of GHRH and/or suppressing somatostatin release (27, 28, 29). Indeed, GHRH and GHRPs typically synergize in vivo, and maximal GH responsiveness to GHRPs require an intact hypothalamic-pituitary unit (14). GHRP-2 administration caused a significant increase in circulating GH concentration in our patients (3-fold in the female patient and 12-fold in the male patient), but the released amount was clearly very low (expected values, 80- to 100 fold in age- and gender-matched controls). We have no a priori reason to suppose that the GHRP receptor is also altered in this condition, as the PRL response to GHRP-2 was preserved. Comparable results were obtained in humans with functionally down-regulated GHRH receptors, as achieved by sustained GHRH infusions. In this experimental setting there was partial loss of responsiveness to a bolus injection of GHRH, but not GHRP (30). In the present studies more severe loss of GHRH responsiveness probably accounts in part for blunted GHRP-2 actions (below).
A prominent reason for the low GH response to GHRP-2 is the severe diminution of somatotrope mass as indirectly inferred from the empty sella configuration on magnetic resonance imaging. The estimated loss of volume was about 70%, suggesting that the GH cell population was sparingly developed. The latter is not surprising, as GHRH also controls GH cell mass in healthy subjects and in patients with GHRH-producing tumors (31). The animal counterpart of this gene defect is the lit-lit mouse, described in 1976, the pituitary gland of which exhibits a 70% reduction of the GH cell population (32, 33). Additional studies have revealed that the GHRH receptor gene in this model contains a missense mutation, resulting in the loss of binding properties of the receptor for its natural ligand (34, 35).
Another contributing factor to the diminutive release of GH after GHRP-2 injection in GHRH receptor-deficient patients, could be a small releasable pool per GH cell, as GH stores are also GHRH controlled (36). Nevertheless, plasma GH concentrations increased severalfold after GHRP-2 infusion, which contrasts with responses reported in four dwarfs of Sindh, whose serum GH concentrations failed to increase in response to a high dose of hexarelin, although PRL and cortisol increased normally (37) The two studies are not strictly comparable, as an ultrasensitive GH assay to quantitate GH responsiveness was not used in the earlier analysis.
Noteworthy is the increased disorderliness of GH secretion, as assessed by elevated ApEn in the patients. This statistic measures the minute to minute regularity of GH secretion, and monitors the strength of feedback and feedforward signals of a hormone system. Blockade of input signals may be associated with increased irregularity of release, as observed for autonomous GH-secreting pituitary tumors (38, 39). In our patients, therefore, the observed increase in ApEn is probably attributable to an absent GHRH signal. Lowered feedback, due to reduced circulating IGF-I concentrations, might also play a role. Increases in ApEn, although generally less marked, have also been reported in other forms of GH deficiency (40, 41), which may share a mechanistic basis. However, lower mean GH levels per se do not account for disorderly GH secretion, as the entropy measure is scale-invariant (10).
Both patients maintained a significant diurnal rhythm for GH (and also for TSH and PRL; data not shown), with maximal GH secretion during the early night hours. Akin to data in other GH-deficient patients of various etiologies, the acrophase occurred at a normal time. This finding suggests that the sleep-wake cycle and circadian system are not greatly altered in this syndrome. This clinical observation is relevant, because the lit-lit mouse has underdeveloped suprachiasmatic nuclei (42).
Recently, a natural ligand for the GHRP receptor, Ghrelin, was isolated from the rat stomach and cloned in this species and the human. This Ser3-octanoylated 28-amino acid peptide circulates in human blood, selectively stimulates GH release from the pituitary in vitro, and is as potent as GHRH (43). At present, it is not possible to delineate the physiological role of Ghrelin. The present results do not exclude a role for ligands of this family in GH regulation. Indeed, bioassay of sheep portal blood revealed episodic release of an unidentified GHRP receptor agonist (44). However, short- and long-term administration of GHRPs and nonpeptidyl mimetics also amplify pulsatile GH secretion, without altering GH pulse frequency, suggesting that GH pulse timing is not governed by GHRPs (28, 45, 46).
In summary, the present studies in two patients with a profound loss of function mutation of the GHRH receptor favor the view that in the human the timing of GH pulses is primarily supervised by intermittent somatostatin withdrawal and the amplitude of GH pulses is driven by GHRH. In addition, we infer that effectual GHRH input controls the GH cell mass and the orderliness of the secretory process.
Footnotes
Address all correspondence and requests for reprints to: Dr. F. Roelfsema, Department of Endocrinology and Metabolism, Leiden University Medical Center, Albinusdreef 2, 2333AA, Leiden, The Netherlands.
Received June 20, 2000.
Revised October 4, 2000.
Revised December 5, 2000.
Accepted December 20, 2000.
References
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