Department of Pediatrics, McGill University, Montreal, Quebec, Canada H3Z 2Z3
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ABSTRACT |
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Human tissues express growth hormone receptors (hGHR) by the 3rd mo of gestation. We assessed developmental changes in hGHR function in fibroblasts and liver, testing binding and hormonal response. Fetal cells showed low but reproducible hGH binding. No age-related changes occurred in fibroblasts (9 wk-34 yr). In contrast, there was a fourfold increase in hGH binding in postnatal liver, with a sixfold increase in hGHR mRNA. Both full-length and truncated hGHR mRNAs were detected in all livers. Cross-linking revealed a larger hGH/receptor complex in fetal liver. Fetal hepatocytes produced 10 times more insulin-like growth factor (IGF)-II than IGF-I, and responded to hGH (150 ng/ml) with a significant increase in IGF-II. Fetal hepatocytes secreted three IGF-binding proteins (IGFBPs), including IGFBP1, but not IGFBP3. hGH did not alter fetal hepatocyte IGFBPs but stimulated glucose uptake. Exposure of fibroblasts to hGH decreased hGH binding only in >1-yr postnatal fibroblasts, whereas treatment with dexamethasone (100-400 nM) increased binding only in postnatal cells. Thus, although fetal hepatocytes and fibroblasts possess functional hGHR, these receptors (and/or their signaling pathways) are immature or have adapted to the in utero environment.
fetal; postnatal; insulin-like growth factors; insulin-like growth factor-binding proteins
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INTRODUCTION |
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GROWTH HORMONE (GH) is required for normal postnatal growth, having a critical role in bone growth as well as important regulatory effects on protein, carbohydrate, and lipid metabolism (49). The actions of human (h)GH are primarily through the stimulation of insulin-like growth factor (IGF-I and IGF-II) production in target tissues. Although synthesis of IGF-I has been demonstrated to be hGH- and nutritionally regulated, IGF-II is thought to be more of a constitutive (and fetal) growth factor (6, 10, 45). The IGFs circulate bound to specific IGF binding proteins (IGFBPs) (32) and work in an autocrine, paracrine, or endocrine fashion by binding to specific receptors (39).
In contrast, the role of hGH during fetal development is not clearly defined. Plasma levels of immunoreactive hGH are markedly elevated in the fetus relative to the normal adult, up to 150 ng/ml at midgestation (34). We (19, 54, 66, 67) and others (26, 62) have identified hGH receptors (hGHR) at the mRNA and peptide level in fetal tissues as early as the first trimester. hGH stimulated proliferation and IGF production in cultured fetal hepatocytes but had a variable effect on insulin and IGF production by islets and no effect on fetal myoblasts, fibroblasts, or cartilage explants (13, 24, 46, 56, 57). Despite inconsistencies in the functional studies, the data indicate that hGH may have important functions during differentiation of specific fetal tissues.
Clinical studies also suggest that hGH begins to have an influence
before birth. The median birth length of a large cohort of idiopathic
hGH-deficient children was found to be 0.83 below that of controls, and
21% were >2 standard deviation scores (SDS) shorter than normal
newborns (16). In addition, certain infants with hGHR
defects are shorter than normal controls at term (1.59 SDS), and
their clinical phenotypes indicate that the hGHR is critical for normal
maturation of specific fetal tissues such as skin, liver, kidney,
gonads, and muscle (3, 50, 52).
To examine this question further, we have characterized several functional aspects of the hGHR in human dermal fibroblasts and liver during development, including hGH binding and biological responsiveness to regulatory hormones (hGH, dexamethasone). These tissues were chosen because of their relative availability at both fetal and postnatal stages in development. Our investigations confirm the presence of functional hGHR in fetal fibroblasts and hepatocytes but suggest that the fetal receptors (and/or their activated pathways) are still in an immature form or that they have adapted to the fetal milieu.
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MATERIALS AND METHODS |
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Tissues.
Human fetal tissues were obtained at the time of therapeutic abortion
[9-20 wk of fetal age (FA)]; FA was determined by foot length
(43). Specimens from premature newborns (23-33 wk FA) and postnatal donors (2 mo-64 yr) were obtained within 6-10 h after death. The three premature newborns died within 1-3 days after birth due to a variety of pathologies (oligohydramnios, diaphragmatic hernia, respiratory distress syndrome). Postnatal liver
samples were obtained from adult transplant donors who had died of
acute head injuries (n = 5). Postnatal skin samples
were obtained from pediatric patients at the time of surgery (inguinal or abdominal hernia) or from our local Genetics Cell Bank collection. Skin samples were placed in explant culture, and liver tissues were
flash frozen in a dry ice-acetone bath and stored at 70°C. Rat
livers were from adult male Sprague-Dawley rats (Charles River, Montreal, QC, Canada). Ethical approval for the study was obtained from local institutional research ethics and animal care committees.
Cultured cells. Dermal fibroblasts from fetal and postnatal donors were grown as monolayer cultures in Ham's F-10-DMEM (1:1; Life Technologies, Gaithersburg, MD) with 10% fetal/newborn calf serum (1:1; Hyclone, Logan, UT), 100 U/ml of penicillin G (Marsam Canada Pharmaceutical, Montreal, QC, Canada), 16 µg/ml of gentamycin sulfate (Schering Canada, Pointe Claire, QC, Canada), and 5 µg/ml of amphotericin B (Squibb Canada, Montreal, QC, Canada). Fibroblasts from the 10th-20th passages were used for experiments. The monolayers were refed every 3 days (until 80% confluence was achieved) as well as 24 h before being tested. IM-9 cells were grown in suspension in RPMI-1640 medium (Life Technologies) with 10% fetal/newborn calf serum (1:1) and antibiotics and tested while in the exponential phase of growth, 24-48 h after refeeding. Cells used for hormonal experiments were preincubated overnight in Ham's-DMEM with 0.1% BSA (Sigma, no. 4378; Missassauga, ON, Canada). Triplicate cultures were treated with recombinant 22-kDa hGH (5-150 ng/ml) (Eli Lilly, Indianapolis, IN) or dexamethasone (100-400 nM; Sabex International, Montreal, QC, Canada) in BSA-medium for 48 h.
Hepatic tissues were minced and dispersed by collagenase, followed by gravity separation, yielding >95% pure hepatocytes (assessed by cytokeratin, albumin, andMicrosomal membrane preparations.
Human and rat livers were homogenized (1:5, wt/vol) on ice in 0.3 M
sucrose containing 0.7 mM 4-(2-aminoethyl)benzenesulfonyl fluoride
(AEBSF; ICN, Mississauga, ON, Canada), filtered through mesh cloth, and
centrifuged at 1,500 g for 10 min at 4°C. The resultant
supernatants were spun first at 15,000 g for 20 min at
4°C, followed by 30 min at 400,000 g at 4°C. The
400,000-g pellets were resuspended in 3 mM
MgCl2, incubated for 15 min on ice to remove endogenously
bound hGH, and then recentrifuged at 400,000 g for 30 min,
resuspended in 25 mM Tris · HCl and 10 mM MgCl2,
and respun at 400,000 g for 30 min at 4°C. Final pellets were resuspended in Tris binding buffer and frozen at 70°C. IM-9 cells were sonicated on ice in 25 mM Tris · HCl, 10 mM
MgCl2, and 0.7 mM AEBSF and spun at 15,000 g for
25 min, followed by centrifugation at 400,000 g for 30 min.
Pellets were treated with MgCl2, rinsed twice, and frozen
at
70°C.
Radioreceptor assays. hGH (National Hormone and Pituitary Program, National Institute of Diabetes and Digestive and Kidney Diseases, Bethesda, MD) and IGF-II (Eli Lilly) were labeled with 125I by the chloramine-T method (specific activity: 42-145 µCi/µg), as previously described (18) and used only if >90% trichloroacetic acid precipitable.
Fibroblast monolayers were washed four times (15 min, 37°C, 5% CO2) with 5 ml of binding buffer (DMEM, 25 mM HEPES, 25 mM Tris · HCl, pH 7.4, and 0.1% BSA). A total of 15,000 cpm/ml (0.1 ng/ml) of iodinated peptide with or without excess unlabeled hGH (22 kDa, Eli Lilly) was added to each dish and incubated for 2 h at 30°C. The reaction was terminated by washing the monolayers three times with 4 ml of ice-cold PBS-0.1% BSA. The fibroblasts were solubilized with 0.6 N NaOH for 6-12 h, and aliquots were analyzed for radioactivity and protein. All assays were performed in quadruplicate. Binding assays with IM-9 cells were run in parallel with the use of triplicate tubes of 750 µg (protein) of cells in suspension per 0.5 ml of final volume. The reaction was terminated by adding ice-cold PBS-0.1% crude BSA and centrifuging at 3,000 rpm for 30 min. Viability of the fibroblasts and IM-9 cells was monitored using light microscopy as well as the trypan blue exclusion test. Radioreceptor assays with liver or IM-9 microsomal membranes were carried out in triplicate for 18-22 h at 4°C. A total of 100 µg/ml of IM-9 membranes or 200 µg/ml of liver membranes was incubated in 150 µl of binding buffer [25 mM Tris · HCl, pH 7.6, 10 mM MgCl2, 0.1% BSA, 1 µM aprotonin, 0.7 mM AEBSF, 21 µM leupeptin, 1 µM pepstatin, and 100,000 cpm of 125I-labeled hGH (0.7 ng)]. Nonspecific binding was determined by addition of excess unlabeled hGH (10 µg/ml). The assay was terminated by adding ice-cold Tris · HCl and MgCl2 buffer with 0.1% BSA and centrifuging at 15,000 g for 30 min at 4°C. Radioreceptor assays with primary hepatocytes were not carried out due to the limited amounts of fetal and adult liver samples available.Affinity cross-linking.
Microsomal membranes (200-1,000 µg) were incubated with
0.2-1.0 × 106 cpm of 125I-hGH in 0.5 ml of binding buffer for 18-22 h at 4°C with or without excess
(10 µg/ml) unlabeled hGH. Binding was terminated by addition of
ice-cold Tris · HCl buffer and centrifugation at 400,000 g for 30 min. The pellets were resuspended in PBS containing
0.5 mM disuccinimidyl suberate (Pierce, Rockford, IL) and
incubated at room temperature for 15 min. Cross-linking was stopped by
addition of 1 mM Tris-0.2 M EDTA and recentrifugation at 400,000 g for 30 min. Pellets were dissolved in sample buffer (0.05 M Tris · HCl, pH 6.8, 10% glycerol, 0.05% bromphenol blue,
10% SDS, and 5% -mercaptoethanol) and run on 7.5% polyacrylamide
gels. The gels were dried, and the receptor complexes were revealed by
autoradiography. Liver, but not fibroblast, microsomal membranes gave
sufficiently high specific binding for cross-linking results.
-Fetoprotein and IGF RIAs.
-Fetoprotein levels were measured using the Amerlex-M RIA kit
(Johnson and Johnson Clinical Diagnostics, Rochester, NY). IGF-I and
IGF-II concentrations were determined by DSL RIA kits (Webster, TX)
after formic acid-acetone extraction (7).
IGFBP Western ligand blots. IGFBP profiles were analyzed according to the method of Hossenlopp et al. (29), with modifications. Conditioned media were concentrated 10-fold and electrophoresed through a 12.5% SDS-polyacrylamide gel, and the proteins were transferred to a nitrocellulose membrane. Blots were probed overnight with 125I-IGF-II, and the IGFBPs were visualized by autoradiography.
[14C]glucose uptake studies. Assays were carried out in triplicate on day 5, following 72 h with or without 22-kDa hGH treatment. Cultures were incubated in Krebs-Ringer phosphate with 0.1% BSA with or without test factors (150 ng/ml hGH, 300 nM insulin) for 30 min, and then 0.375 µCi of D-[U-14C]glucose (25 mCi/mmol; New England Nuclear) was added to each well for 30, 60, or 90 min. Glucose uptake was stopped by rinsing 3 times with ice-cold PBS. One milliliter of 0.6 N NaOH was added to each well, the cells were solubilized overnight at 37°C, and aliquots were analyzed for radioactivity and protein (Bio-Rad, Hercules, CA).
RNA extractions. The guanidium-thiocyanate-phenol method (9) was used to extract total RNA from human fetal and postnatal tissues.
Semiquantitative RT-PCR.
Five micrograms of total RNA were reverse transcribed for 1 h at
48°C in the presence of 2.5 U of AMV-RT (Life Technologies), 80 U of
RNAsin (Promega, Madison, WI), 71.4 ng/µl random primers (Life
Technologies), 0.48 mM deoxyribonucleotides (dNTPs) (Pharmacia Biotech,
Baie D'Urfe, QC, Canada), 10 mM MgCl2, 10 mM
dithiothreitol, 100 mM Tris · HCl, pH 8.3, and 50 mM KCl. Two
microliters of RT product were amplified for 23 cycles with varying
amounts of internal standard (0.9-48.0 × 104
fmol), 2.5 U of Taq DNA polymerase (Life Technologies), 0.5 mM dNTPs, 0.5 µM hGH receptor sense (exon 7: 5'-CCA GTG TAC TCA TTG AAA GTG GAT-3') and antisense (exon 10: 5'-GTC TGA TTC CTC AGT CTT TTC
ATC-3') primers, 3 mM MgCl2, 20 mM Tris · HCl, pH
8.4, and 50 mM KCl. The first cycle consisted of 3 min at 92°C, 1 min at 61°C, and 3 min at 72°C; subsequent cycles were 30 s at
92°C, 1 min at 61°C, and 1.5 min at 72°C, terminating with a
final elongation of 5 min at 72°C. As a control, RNA samples were
incubated in the absence of AMV-RT and then amplified in the presence
of 1.4 × 10
3 fmol of internal standard. Initial
experiments (data not shown) showed that the PCR reaction was in an
exponential phase at 23 cycles. The internal PCR standard was
constructed according Jin et al. (33), as previously
described (66). PCR fragments were electrophoresed through
1.5-2% agarose gels and transferred to 0.45-µm positively
charged nylon membranes (Schleicher & Schuell, Keene, NH), as
previously described (66, 67). Blots were hybridized overnight using the nested oligonucleotide (exon 10: 5'-GCT AAG ATT GTG
TTC ACC TCC TC-3') end labeled with [
-32P]ATP (NEN),
and the bands were quantified using a Fuji phosphorimager (Stamford,
CT) to obtain the molecules of hGHR mRNA per microgram of total RNA, as
described (66).
Full-length vs. truncated hGHR mRNA. Five micrograms of hepatic total RNA were reverse transcribed using Superscript II (Life Technologies) and a specific exon 9 reverse primer (9AS) designed to recognize both the full-length and 1-279 hGHR mRNAs (5'-TAATCTTTGGAACTGGAACT-3'). RT products were amplified using an exon 7 sense (7S) primer (5'-ATAAGGAATATGAAGTGCGTGTGAG-3') and the exon 9AS primer (see Fig. 2A), under the following conditions for 35 cycles: 94°C for 30 s, 52°C for 30 s, and 72°C for 1 min, ending with 72°C for 10 min. PCR products were separated on 10% polyacrylamide gels (Fig. 2B) and stained with Sybr gold, and densitometric analyses were carried out using the Bio-Rad GelDoc system (Mississauga, ON, Canada).
Statistical analyses. Differences between groups (by age or treatment) were analyzed by Student's t-test if there were two groups or by ANOVA followed by Duncan's multiple range test if multiple groups.
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RESULTS |
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Developmental changes in tissue hGHR
mRNA and microsomal membrane binding.
Using semiquantitative RT-PCR assays, we determined total hGHR mRNA
levels in four pairs of human fetal and postnatal tissues. Although
there was a general decrease in postnatal lung (P < 0.05), kidney, and small intestine compared with their fetal
counterparts, the hGHR mRNA in postnatal liver increased sixfold
(P < 0.01; Fig.
1A). We also detected mRNAs
for both the full-length and the major truncated (GHR279) GHR in fetal
(n = 8; 14-18 wk) and postnatal (n = 3; 21-54 yr) liver tissues. The full-length transcript was
always predominant (Fig. 2B).
Although there was a trend for fetal liver to have a twofold higher
ratio of truncated to full-length mRNAs, there was two- to threefold
variability within each age group, and the difference did not reach
statistical significance (Fig. 2C).
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Affinity cross-linking.
Cross-linking experiments showed that 125I-hGH bound to
different molecular mass species of hGHR in human fetal, human adult, or rat adult liver (Fig. 3). In human
fetal liver, the 125I-hGH/hGHR band was ~112 kDa; excess
unlabeled 22-kDa hGH decreased, but did not eliminate,
125I-hGH binding (Fig. 3A). In human adult
liver, the 125I-hGH/receptor complex was a broad band at
~80-100 kDa, which disappeared with excess unlabeled hGH
(Fig. 3B). In rat liver, the hGH tracer was part of an
~65-kDa receptor complex; excess unlabeled hGH completely eliminated
125I-hGH binding to this band (Fig.
3C).
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Fetal hepatocyte -fetoprotein, IGF, and
IGFBP production and
[14C]glucose uptake.
Fetal hepatocyte cultures were monitored for
-fetoprotein production
on day 5 in vitro. Levels in the conditioned serum-free media were quite consistent [n = 9; 149.4 ± 23 ng/ml (means ± SE); blank medium = <6 ng/ml]. IGF-I and
IGF-II concentrations were also assessed on day 5. There was
a striking 10-fold greater production of IGF-II than IGF-I
(n = 9; 15.3-20 wk; 4.2 ± 0.3 vs. 0.43 ± 0.05 ng/ml). Pretreatment with 150 ng/ml of hGH for 72 h
resulted in a small, significant increase in IGF-II (5.2 ± 0.4;
P < 0.04) but not in IGF-I (0.42 ± 0.05).
Western ligand blot analyses of the same day 5-conditioned
media revealed the presence of three IGFBPs (32, 28, and 24 kDa; data
not shown). The middle band was identified as IGFBP1 by Dr. J. W. van Neck (personal communication) (53), whereas the two
others correspond to the molecular masses of IGFBP2 and IGFBP4,
respectively. IGFBP3 was readily detectable in the control normal serum
pool but was not present in the fetal hepatocyte-conditioned media
(data not shown). hGH (50-150 ng/ml) treatment for 72 h had
no effect on IGFBP profiles. In contrast, hepatocytes pretreated for
72 h with 150 ng/ml hGH showed a small, significant,
time-dependent (P < 0.01) increase in
[14C]glucose uptake (Fig.
4A). The level of uptake
was similar to that observed after acute (30-min) treatment with 300 nM
insulin (P < 0.05; Fig. 4B).
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Fibroblast radioreceptor assays.
Previous studies demonstrated hGHR mRNA in human fetal (10-38 wk)
and postnatal (4 mo-35 yr) skin as well as cultured fibroblasts through
12 generations in vitro (67). In the present study, we
determined that human dermal fibroblasts specifically bound 125I-hGH from as early as the 9th wk of fetal life. When a
large fetal (n = 23; 9-33 wk FA) and postnatal
(n = 11; 2 mo-34 yr) series was compared, we
observed a small increase in both total (2.2 ± 0.1 vs. 3.2 ± 0.3%) and nonspecific (0.9 ± 0.09 vs. 1.6 ± 0.2%)
binding of 125I-hGH (P 0.01) but no
significant change in specific binding (1.3 ±0.1 vs. 1.6 ±0.3%) as a
function of donor age. Control IM-9 cells exhibited much higher total
(8.8 ±0.3%; n = 34) and specific (4.9 ± 0.3%)
binding of 125I-hGH, although the percentage of nonspecific
(3.7 ± 0.2%) binding was similar. The low level of fibroblast
binding precluded Scatchard analyses.
Fibroblast responses to hormonal treatments.
When fibroblast cultures were exposed to hGH (5-150 ng/ml) for
48 h, there were significant age-related differences in their ability to subsequently bind 125I-hGH (Fig.
5A). The fetal and early (<1
yr) postnatal cells showed no change in percentage of specific
125I-hGH binding, even when the highest hGH concentration,
150 ng/ml, was tested. In contrast, the older (>1 yr) postnatal
fibroblast series showed a significant (P < 0.05)
dose-related decrease in binding. Control IM-9 cells showed the
expected highly significant (P < 0.01) decrease in
125I-hGH binding (40).
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DISCUSSION |
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hGH acts by binding to its high-affinity receptor on target cells (47). The hGHR, a single-chain (620 amino acid) polypeptide member of the hGH/human prolactin/cytokine receptor superfamily, dimerizes in the presence of one molecule of hGH, activating specific intracellular signal transduction pathways and leading to gene regulation (14, 21, 35). The hGHR is encoded by a single gene spanning >150 kb on chromosome 5p13.1-12 (5, 17, 19, 41). The coding region is defined by exons 2-10, with exon 2 contributing the translation start site and signal sequence. Three hGHR isoforms have been reported. Exon 3-deficient mRNA can be detected in fetal tissues from as early as the 8th wk of fetal life (11, 59, 66, 67). This transcript, expressed in an individual- rather than a tissue-specific manner, translates into a high-affinity hGHR that binds hGH and related hormones with similar affinities as the full-length receptor (55, 60, 63, 67). Two truncated forms of hGHR, 1-277 and 1-279, have also been demonstrated, with 1-279 being the major (10- to 20-fold more prevalent) form (2, 51). mRNAs for these truncated forms are produced at low levels in all human tissues examined to date due to variant splicing within exon 9 (4). The truncated receptors bind hGH with high affinity but do not dimerize or activate intracellular signal transduction pathways and thus may act as dominant negative receptors (51).
Although pivotal roles for hGH and its receptor have been well established in postnatal tissues, their function during human gestation remains unclear (22). Fetal serum GH levels are high in both primates and subprimates (15). However, although numerous studies have shown that onset of significant GHR mRNA expression occurs only around the time of birth in subprimates (1, 8, 36, 42, 58, 61, 65), ubiquitous transcription of the hGHR gene has been documented from the first trimester in the human fetus (19, 66, 67). In addition, immunohistochemical studies have identified hGHR protein in human tissues from 8 wk FA and shown that, by midgestation, the pattern of immunostaining is often identical to that found in the adult (26, 54). Thus both hGH and its receptor are present at very early stages in human development.
Whether the fetal hGHR are functional has been a controversial issue due to the variability of results from different research groups and the lack of correlation between hGHR mRNA/immunopeptide data and hGH responsiveness in certain tissues. There are reports of low hGH binding in fetal liver, chondrocytes, lung, and fibroblast cell lines, but no binding in muscle (25, 37, 44, 62). In one study, hGH treatment of fetal hepatocytes increased cell proliferation and IGF production (56). However, there have been inconsistent effects reported on pancreatic islet cells and negative effects observed with fetal myoblasts, cartilage explants, and dermal fibroblasts (13, 23, 46, 57). In the present study, we have addressed this controversy by focusing on two cell types, hepatocytes and dermal fibroblasts, and wherever possible, comparing hGHR functional characteristics in fetal vs. postnatal cells.
Our initial RT-PCR analysis demonstrated changes in total hGHR mRNA in four different sets of tissues: liver, kidney, lung, and small intestine. All but liver showed a decrease in hGHR mRNA in postnatal compared with fetal samples. In lung, this change was statistically significant and paralleled the previous observation of low hGH binding in fetal lung membranes and no binding in postnatal samples (37). In contrast, postnatal liver showed a striking increase in hGHR mRNA and a parallel increase in hGH specific binding. Thus there are marked tissue-specific differences in regulation of hGHR expression during development.
Cross-linking experiments indicate structural differences between the fetal and postnatal hepatic hGHR: the fetal receptor runs as a higher molecular mass band than either the human or rat adult liver GHR (27, 28, 30). Because fetal hepatic hGHR mRNAs are homologous to those found in postnatal liver (4, 67, present study), it is unlikely that this size difference is due to increased length of the basic receptor protein. Two other explanations seem more appropriate: increased glycosylation of the extracellular domain of the fetal hGHR and/or complexing with additional molecules.
The fetal hepatocytes produced IGFs and IGFBPs in an appropriate manner, with significantly higher IGF-II than IGF-I and no IGFBP3 (32, 39, 48). When pretreated with hGH, fetal hepatocytes showed an unusual response: no change in IGF-I but an increased production of IGF-II. Strain et al. (56) previously reported increased IGF-I secretion by human fetal hepatocytes after exposure to similar concentrations of hGH. However, the antibody they used to assay IGF-I has since been shown to also recognize IGF-II (6). Because IGF-II is the predominant IGF found in fetal tissues and circulation throughout gestation (6, 38, 45), it is likely that it was, in fact, IGF-II that was measured. There was no effect of hGH on IGFBP profiles. IGFBP3, the one IGFBP known to be responsive to hGH, is not produced until the final third of gestation (39, 48). Our data suggest that either the early hepatocytes cannot be induced to synthesise IGFBP3 after only 72 h of exposure to hGH or that hGH is not the appropriate inducing factor. The hepatocytes did respond to hGH with enhanced [14C]glucose uptake. This increase was similar to the effect observed with acute insulin exposure, but much lower than has been observed in postnatal liver (12).
Dermal fibroblasts showed no age-related changes in hGH binding. However, there were marked changes in their responsiveness to both hGH and dexamethasone. Chronic exposure to hGH is well known to cause desensitization of hGH target cells due to loss of hGHR from the cell surface (40, 64). This response was exhibited by postnatal fibroblasts, but only those from infants >1 yr of age, suggesting that maturation of the downregulatory effect of hGH is not linked to birth and independence from the in utero environment. In contrast, the ability of dexamethasone to increase hGH binding was observed in postnatal fibroblasts as early as 4 mo after birth.
In conclusion, our studies demonstrate that fetal hepatocytes and fibroblasts do have functional hGHR, in that these cells specifically bind hGH and have biological responses after chronic exposure to hGH. Certain of the responses appear to be an adaptation to the fetal environment (increased IGF-II rather than IGF-I), whereas others (low glucose uptake, no change in hGHR in response to hGH or dexamethasone) may be simply due to immature cell systems (internalization/receptor cycling, signal transduction pathways, gene regulatory mechanisms), none of which has been investigated to date. The fact that the fetal hepatic hGHR is a different molecular size is also intriguing and needs to be explored to determine what role this difference may play in restricting hGH responsiveness during fetal development.
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ACKNOWLEDGEMENTS |
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We thank the operating room staff at the Montreal Children's Hospital and Hôpital Maisonneuve-Rosemont for their help in providing tissues. We also acknowledge the expert assistance of Jean Parodo, Sharon Lerner, and Andrew Khalil with the IGF assays, the IGFBP1 immunoblot analysis by Dr. J. W. van Neck, and the expert advice on microsomal membrane preparations from Dr. John Bergeron and on fibroblast cultures from Gail Dunbar.
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FOOTNOTES |
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R. M. O. Figueiredo was supported by a research fellowship from Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (Brazil) and the McGill University-Montreal Children's Hospital Research Institute. G. Zogopoulos was the recipient of an Fonds pour la Formation de Chercheurs et d'Aide à la Recherche studentship. This research was funded by the Medical Research Council of Canada (C. G. Goodyer).
Address for reprint requests and other correspondence: C. G. Goodyer, Endocrine Research Laboratory, 4th Floor, Place Toulon, McGill University-Montreal Children's Hospital Research Institute, 4060 Ste. Catherine St. West, Westmount, Quebec, Canada H3Z 2Z3 (E-mail: cindy.goodyer{at}muhc.mcgill.ca).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 2 February 2001; accepted in final form 11 July 2001.
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