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Growth Hormone Receptor Messenger Ribonucleic Acid in Normal and Pathologic Human Adrenocortical Tissues—An Analysis by Quantitative Polymerase Chain Reaction Technique¹

Research Laboratory of Department of Internal Medicine (C.J.L., B.B.M., I.C.M.G, W.N., S.M.F.V.), Division of Endocrinology, and Division of Urology (A.M.L.), Hospital das Clínicas, School of Medicine, University of São Paulo, Caixa Postal 8091,CEP 01065–970, São Paulo, Brazil

Address all correspondence and requests for reprints to: Chin Jia Lin, Research Laboratory of Department of Internal Medicine, Division of Endocrinology, Hospital das Clínicas, School of Medicine, University of São Paulo, Caixa Postal 8091,CEP 01065–970, São Paulo, Brazil.

	Abstract

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GH receptor (GHR) has been reported to express in both normal rat and human adrenals. However, no study examined GHR expression in diseased human adrenal cortex. We quantitated, with RT-PCR, GHR messenger RNA (mRNA) in both normal and diseased human adrenal cortex with the following results: GHR mRNA levels in four histologically normal, not steroid-stimulated, control adrenal cortices was 1.5–11 x 10⁴ molecules/µg total RNA; in three diffusely hyperplastic adrenals (DH): 6.7–17.7 x 10⁴; in two nonfunctioning tumors (NF): 0.84–1.9 x 10⁴; in five androgen-producing neoplasms (AP): 4.6–34 x 10⁴; and in five glucocorticoid-producing neoplasms (GP): 6.7–87 x 10⁴. GHR transcript levels among adrenal cortices, DH, NF, AP, and GP reached statistically significant difference (P < 0.03). The GP group exhibited higher GHR mRNA levels than controls (P < 0.006). NF, as well as GP and AP, tumors had less GHR mRNA than their histologically normal adjacent cortex (P < 0.05). A positive correlation between urinary cortisol and GHR messenger RNA (mRNA) levels from GP and DH was observed (r = 0.93, P < 0.003).

Our data suggest that GHR is expressed in both normal and diseased adrenal cortex and that GHR mRNA accumulation is less efficient in adrenocortical neoplasm than their adjacent nonneoplastic cortex. GHR expression in adrenal cortex provides an evidence of direct GH action in this tissue.

	Introduction

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THE GH HAS somatic growth-promoting effect, through its action on the insulin-like growth factor-1 (IGF-1) synthesis. Direct GH action also has been reported. GH has been demonstrated to be a potent growth and mitogenic factor for the pancreatic ß-cell (1). GH also has been reported to induce expression of c-fos and c-jun mRNA in 3T3-F442A preadipocytes (2).

The GH receptor (GHR) is a member of the cytokine receptor superfamily receptors. Its gene consists of 10 exons and encodes a 620-amino acid protein with a single centrally located transmembrane domain (3). Cloning of GHR complementary DNA (cDNA) by Leung et al. (4) has provided researchers with a powerful and sensitive tool for the study of direct GH action on the cells and detection of GHR gene expression. Since then, GHR mRNA has been detected in several cell lines. Expression of GHR gene in nonprimary GH-action target human and animal tissues has been reported. The GHR mRNA has been demonstrated in rat muscle, heart, kidney, ovary, digestive tract (5), and pituitary gland (6). In humans, it has been described in ovary (7), pancreas, different segments of the digestive tract (8), heart, spleen, lung, uterus, aorta, bladder (9), skin, thymus, testis (10), and thyroid (11). GHR mRNA also has been detected both in rat (5) and human normal adrenal gland (9).

The present study was undertaken to evaluate GHR gene expression in normal and diseased human adrenocortical tissues.

	Materials and Methods

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Patients

Informed consent was obtained from all patients, and this study was approved by the Ethics Committee of our institution.

Adrenocortical tissue samples were obtained from 17 patients. Their clinical, hormonal, and histological data are summarized in Tables 1 and 2. Their ages ranged from 1.7–56 yr. Patients 1–5 were evaluated because of signs and symptoms of virilization. Patients 6 and 7 had an incidentally discovered adrenal mass when referred to endocrinological evaluation; neither patient had specific endocrinological complaint. Patients 8–15 presented with hypercortisolism. Patients 8–12 had a glucocorticoid-producing adrenocortical neoplasm. Patients 13 and 14 had been treated unsuccessfully for Cushing’s disease. No hypercortisolemic patient has been treated with adrenal steroidogenesis blockade medication. Patients 16–18, who had a urological problem and no endocrine abnormalities, had their adrenal glands removed because of the technical difficulties during nephrectomy. Patient 10 died, after a 5-month follow-up, because of metastatic dissemination of his adrenocortical neoplasm. The others are under current ambulatory follow-up with no evidence of disease recurrence or metastasis.

All patients with hypercortisolism were given 100 mg hydrocortisone at the anesthetic induction. Their adrenals were removed 40–60 min later. The exogenous glucocorticoid was tapered gradually on the subsequent days. No glucocorticoid has been given preoperatively.

We included, in the control group (CT), adrenal cortices that were both histologically normal on the light microscopy and free of influences of hypersecreted steroid hormones or ACTH. By fulfilling both the criteria, adrenal cortex of patients 16 and 17, as well as neoplasm-free cortex adjacent to NF of patients 6 and 7, were used as control tissues. Adrenal cortices surrounding the steroid-producing tumors (patients 5, 11, and 12) were categorized as adjacent adrenal because of their steroid-rich microenvironment.

Tissue processing and RNA extraction

Human adrenal tissues were obtained in the operating room during the adrenalectomies. The removed adrenals were dissected using a scalpel and pincers. Fat, fibrous tissue, and tumor capsule were removed before tissue collection. Adrenal cortex was collected in regions where we could leave a clearly visible margin of cortical tissue separating the medulla from our sample. Tumoral samples were collected in areas with no necrosis or hemorrhage. Nonneoplastic adrenal cortex was collected in regions that were not contiguous to the tumor. The tissue samples were promptly frozen and stored in liquid nitrogen until used. The nature of our samples was confirmed by microscopical examination before undergoing RNA extraction.

Total cellular RNA was extracted using the single-step acid guanidinium thiocyanate-phenol-chloroform method described by Chomczynski and Sacchi (13) and stored at -80 C. The integrity of each sample was checked by the presence of bands corresponding to 28s and 18s of ribosomal RNA after electrophoresis using a 1% agarose gel stained with ethidium bromide. The purity of RNA samples was assessed by the 260/280 ratio (between 1.6 and 1.9) and by the absence of bands corresponding to contaminating DNA in the agarose electrophoresis. The RNA concentration was assessed by spectrophotometric absorbance at 260 nm (Ultrospect III, Pharmacia LKB, Uppsala, Sweden).

Quantitative PCR

Quantitative PCR consists of the coamplification of the target molecule (whose concentration is to be determined) with a known amount of a molecule with the same flanking sequences and denominated as internal control (IC) molecule. The same pair of PCR primers (one of which is labeled with ³²P) are used to amplify both target and IC molecules (14). The concentration of the target molecule can then be calculated by extrapolating the relationship between the initial concentration of the IC and the radioactivity incorporated into the corresponding PCR products.

Quantitative PCR has been used to quantitate GHR mRNA in human liver and muscle (15). The use of IC molecule minimizes the effect of external interference and assures the coherence of assay procedure. This mRNA quantitation method presented high sensitivity, specificity, and good reproducibility (14, 15). For GHR mRNA, the reported detection limit of quantitative PCR was about 500 molecules/µg total RNA, and the interassay variability was 3.0% (15).

The IC molecule we used is a synthetic RNA, which consists of a mutant human GHR transcript created by the insertion of a fragment of a foreign DNA. The plasmid used in the synthesis of the IC RNA is the same one used by Martini et al. (15) and was a kind donation of Dr. Postel-Vinay. One of us (S. M. F. Villares) participated in the preparation of this plasmid at INSERM U-344, Faculté de Médicine Necker Enfants Malades. Its preparation procedure has been described previously (15). Briefly, this plasmid was prepared by inserting a 50-bp fragment of rat PRL receptor cDNA into the NcoI site located in the region of human GHR cDNA that encodes the transmembrane domain of this receptor (exon 8). After the insertion of rat PRLR fragment, the mutated GHR cDNA was then digested with EcoRI and EcoR V to generate a 545-bp fragment containing about two thirds of the exon 6, the entire exons 7, 8, and 9, and the initial portion of exon 10 (48 bp). This 545-bp fragment was subcloned into the SmaI-EcoRI sites of a Bluescript vector containing an oligo d(A) tail inserted at the HindIII site.

After linealization by digestion with Sal I, the plasmid containing chimeric hGHR/rat PRL receptor cDNA was used as template for in vitro transcription. T7 RNA polymerase (Promega Corporation, Madison, WI) was used in the synthesis of RNA for IC molecule. The procedures of in vitro transcription followed the manufacturer’s protocol. IC RNA was extracted with phenol/chloroform and further purified with chromatography in oligo d(T) column. After purification, IC RNA was suspended in ribonuclease-free water (treated with diethylpyrocarbonate). The integrity of IC RNA was assessed with electrophoresis in 1.5% agarose gel. A single band with expected size (639 bp) was seen on the gel. Through the determination of the optical density we estimated the concentration (µg/µL) of IC RNA solution (1 OD = 40 µg/mL). The absolute number of IC RNA molecules was then estimated using the formula below:

The value for molecular weight of IC RNA is 216,600 g/mol and for Avogadro’s number is 6.02 x 10²³ molecules/mol. The IC RNA solution was diluted with diethylpyrocarbonate-treated water to give a final concentration of 1 x 10⁶ molecules/µL. This solution was aliquoted and stored at -80 C.

Oligonucleotide primers, designed for the amplification of the human GHR transcript, were purchased from Gibco BRL (Gaithersburg, MD). The forward primer, GHR-23, spanned exon 7 [5'-CCCTATATTGACAACATCAGTTCC-3': nucleotides (nt) 624–647] (4) and the reverse primer, GHR-15, spanned exon 9 (5'-CTTGAGGAGATCTGG-3': nt 931–954 (4)). The resulting amplification product is a fragment comprising a sequence that encodes the entire transmembranic region and a short segment of both extracellular and cytoplasmic domains (exons 7, 8, and 9) of the GHR (3). Primer GHR-23 was 5'-end labeled with [-³²P]ATP (>5000 Ci/mmol, New England Nuclear, Boston, MA) using T4 polynucleotide kinase (Gibco BRL). Unicorporated nucleotides were removed by a Sephadex G-25 medium column (Pharmacia).

Four to 6 micrograms total RNA and 1.0 x 10⁶ molecules IC RNA were reversely transcribed with 200 U Moloney Murine Leukemia Virus RT (M-MLV-Reverse Transcriptase, Gibco BRL). The RT was primed with 20 pmol of a primer that recognized the boundary between exons 9 and 10 (5'-TTCACCTCCTCTAAT-3', primer 14 (4) nt 955–969) and was carried out in 20 µL (total vol) of cDNA buffer (50 mmol/L Tris-HCl, pH = 8.3, 75 mmol/L KCl, 3 mmol/L MgCl₂), 0.5 mmol/L of each deoxynucleotide (dNTP) (dNTP mix, Pharmacia), 1 mmol/L DTT (Gibco BRL), and 10 U ribonuclease inhibitor (rRNasin Ribonuclease Inhibitor, Promega). The reaction mix was incubated at 37 C for 1.5 h. The products were frozen at -20 C until used.

Three-fold serial dilution of the RT products (1–81 x) were amplified with PCR. The reaction was carried out using 10 µL of each diluted RT mixture in PCR buffer (50 mmol/L KCl, 1.5 mmol/L MgCl₂, 10 mmol/L Tris-HCl, pH = 9.0), 200 µmol/L dNTPs, 25 pmol of forward and reverse primers, 1.0 x 10⁶ cpm ³²P-labeled primer 12, and 2.5 U Taq polymerase (Taq DNA Polymerase, Pharmacia) in a total vol of 50 µL. PCR was performed for 30 cycles in a sample-sensing thermocycler (GeneAmp PCR System 9600, Perkin Elmer Cetus, Norwalk, CT). The amplification protocol consisted of 30 sec of denaturation at 94 C, 1 min 15 sec of annealing at 55 C, and 1.5 min of extension at 72 C, after an initial denaturation at 94 C for 3 min. Amplification was completed with an additional extension step at 72 C for 10 min.

RT-PCR was performed with total RNA and all reagents, but no RT, to assure that there was no contaminating DNA. To assure that no contamination occurred during the course of RT-PCR procedure, two kinds of negative control were prepared. The first negative control was made by omitting the total RNA in the reverse transcription. The second one was prepared by replacing the cDNA mix with water in PCR reaction. The PCR was considered useful only if no band was observed in the negative control lanes on a 5% polyacrylamide gel.

PCR products (20 µL) were separated on 5% nondenaturing polyacrylamide gel and stained with ethidium bromide. The bands corresponding to each specific PCR product were excised from gels, and the amount of incorporated radioactivity was determined in a ß scintillation counter (LS100C, Beckman, Irvine, CA). Radioactivity (cpm) was plotted against the amount of template (IC RNA or target molecule). Gel pieces of the negative control were excised at the position of the same size as that of each positive band. The radioactivity of these negative control gel values served as background. Both the amount of template (total and IC RNA) and the radioactivity counting of their respective RT-PCR products underwent logarithm transformation. These log-transformed values were used to calculate the linear regression for total and IC RNA dilutional curve. The extrapolated radioactivity for 1 µg total RNA was calculated using the linear regression equation for total RNA. This extrapolated value was then used in the linear regression equation for IC RNA to yield the estimated absolute number of target molecules contained in each µg total RNA. Results were expressed as number of GHR mRNA molecules/µg of total RNA.

Statistical analysis

The differences of GHR transcript levels among adrenal cortices groups [CT, diffusely hyperplastic cortices (DH), NF, androgen-producing neoplasms (AP), and glucocorticoid-producing neoplasms (GP)] were evaluated with the Kruskal-Wallis test. This test was followed by Dunn’s test to assess the difference of GHR mRNA levels between two different adrenocortical tissue groups. Tumoral cortices of patients 5, 6, 7, 11, and 12 were compared with their neoplasm-free adjacent tissue using the Wilcoxon matched-pairs test. The Spearman rank order correlation was used to test the correlation between total urinary cortisol and tissue GHR mRNA abundance. A P value less than 0.05 was considered significant.

	Results

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All RNA samples were submitted to conventional RT-PCR. After PAGE, a 330-bp band, corresponding to the amplification product of GHR mRNA, was visible in the lanes of each RT-PCR product (data not shown).

The results of quantitative RT-PCR are presented in Fig. 1 and Table 1. A great individual variability of the GHR mRNA levels was evident. In the not-hormone-stimulated control adrenal group (CT), the observed GHR mRNA levels were 1.5–11 x 10⁴ molecules/µg total RNA. The nonfunctioning neoplasms (NF) and the hyperplastic adrenal cortices (DH) presented, respectively, 0.84–1.9 x 10⁴ and 6.7–17.7 x 10⁴ GHR mRNA molecules/µg total RNA. The AP group was comprised only of patients under the age of 3 yr. This group exhibited 4.6–34 x 10⁴ GHR mRNA molecules/µg total RNA. The GP group exhibited 6.7–87 x 10⁴ molecules/µg total RNA. The GHR mRNA levels, in the adrenal cortices adjacent to functioning tumors, were 41–188 x 10⁴ GHR mRNA molecules/µg total RNA.

View larger version (11K): [in this window] [in a new window]   Figure 1. Distribution of GHR mRNA abundance according to steroid-secretory activity and histological diagnosis.

Tumoral cortices of patients 5, 6, 7, 11, and 12 were compared with their adjacent adrenal cortices, to evaluate the quantitative difference in GHR mRNA abundance between neoplastic and nonneoplastic adrenal cortex. The results of this analysis are presented in Fig. 2. The neoplastic tissue, in contrast with its neoplasm-free counterpart, showed less GHR mRNA abundance (P < 0.05, Table 1).

View larger version (30K): [in this window] [in a new window]   Figure 2. Comparative study between GHR mRNA abundance observed in neoplastic adrenal cortex and its neoplasm-free counterpart. The results were normalized using neoplasm-free cortex value as 100%.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
There is clinical evidence suggesting that the adrenal cortex is a target of GH action. In children with isolated GH deficiency, the 11-deoxycortisol responsiveness to acute ACTH 1–17 test could be modified by both short- (increasing responsiveness) and long-term recombinant GH (r-GH) treatment (decreasing responsiveness), whereas long-term r-GH administration also caused an increase in cortisol responsiveness (16). High levels of urinary 17-ketosteroids (17-KS) with an increased adrenal responsiveness to ACTH have been reported in acromegalic patients (17, 18). It was not possible to differentiate, in either report, the direct vs. IGF-1-mediated GH action, because the serum IGF-1 levels increased, along with the GH levels, both in r-GH-treated and acromegalic patients. The demonstration of GHR or its mRNA in the adrenal cortex is essential to prove a direct GH action on the adrenal cortex.

The presence of specific receptor for GH in the adrenal cortex was first detected on the bovine zona fasciculata adrenal cell (19). More recently, through Northern blot assay and RT-PCR, GHR mRNA was demonstrated in rat (5) and human normal adrenals (9). In this study, we demonstrated the presence of GHR mRNA and successfully quantitated it in both normal and pathologic human adrenal cortex. These data provided a strong argument favoring the hypothesis of a direct GH action in the adrenal cortex.

As reported for human liver and muscle (15), GHR mRNA measured in adrenal cortices also showed wide individual variation. A great individual variance also was reported for serum GHBP (20, 21). Because serum GHBP levels have been considered to reflect the status of tissue GHR expression (20), such individual variance may be a characteristic feature of tissular GHR expression. Adrenal cortex presented lower GHR mRNA levels than reported for other human tissue. The GHR transcript levels of our control adrenals (1.5–11 x 10⁴) was about 2–4 times lower than those reported for normal muscle (6.0–22 x 10⁴) (15) and was almost 100 x less abundant than those reported for normal liver (50–140 x 10⁴) (15). The quantities of GHR mRNA in all but one GP neoplasm were lower than normal human liver, although this group exhibited the highest GHR transcript levels among the adrenal cortices (43–87 x 10⁴).

The expression of GHR in human neoplastic tissue has been evaluated in liver and mammary gland (22, 23). Both works studied the expression of exon 3-intact and exon 3-deleted GHR mRNA isoforms in normal and tumoral tissues; no relationship between the qualitative pattern of GHR mRNA expression and tissue differentiation state was reported in neither paper. This paper showed that adrenocortical functioning neoplasms (specially the GP tumors) presented higher GHR transcript levels than not-hormone-stimulated control CT. We hypothesize that GHR mRNA accumulation in adrenal cortex can be stimulated by steroid hormones. However, GHR transcript levels in tumoral tissues were lower than their adjacent neoplasm-free cortex, when compared in a matched-pair manner. This decrease in the quantitative GHR expression in tumoral tissue seems to be a consequence of a constitutionally lower GHR mRNA accumulation efficiency in tumors, relative to their adjacent nonneoplastic cortex. The mechanism underlying this differential GHR mRNA accumulation and its pathophysiological meaning need further investigation.

Our data suggest an influence of glucocorticoid on GHR expression in human tissue. This was demonstrated by high GHR mRNA abundance observed among GP and the positive correlation between TUC and GHR mRNA levels in CT from hypercortisolemic patients. The role of glucocorticoid on GHR expression has been investigated. An increase of GHR mRNA levels in the liver and growth plate of dexamethasone-treated rabbits has been reported (24). Recently, prepartum cortisol surge was shown to play an important role in the ontogeny of sheep hepatic GHR gene expression (25). A positive correlation between log plasma cortisol concentration in utero and the abundance of sheep fetal hepatic GHR mRNA also was reported (25). DH and CT exhibited similar range of GHR mRNA levels. This is probably caused by the small size of the DH group.

In conclusion, the data presented here suggest that GHR is expressed in both normal and diseased adrenal cortex, that glucocorticoid may stimulate the accumulation of mRNA for GHR in adrenal cortex, and that GHR mRNA accumulation is less efficient in adrenocortical neoplasm than its adjacent nonneoplastic cortex. The exact meaning of this differential expression and physiologic role of GHR in the adrenal cortex remain to be elucidated.

	Acknowledgments

 
The authors thank Dr. Marie C. Postel-Vinay for donation of the plasmid containing chimeric hGHR/rat RPRL cDNA, Dr. Ana Claudia Latronico for donating samples of normal adrenal cortex, Dr. Ana Elisa Correia Billerbeck for helpful discussion, and Dr. José Antonio Miguel Marcondes and Dr. Maria Adelaide Albergaria Pereira for referring patients to this study. We also thank Miriam Yumie Nishi and Maria Aparecida Medeiros for technical assistance, the house staff of Division of Endocrinology for help in tissue collection, and Sonia Strong for assistance in English usage.

	Footnotes

 
¹ This work was partially supported by FAPESP (process N°: 2828/3–94).

Received November 13, 1996.

Revised December 18, 1996.

Revised April 29, 1997.

Accepted May 6, 1997.

	References

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This Article Abstract Full Text (PDF) Submit a related Letter to the Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Copyright Permission Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Lin, C. J. Articles by Villares, S. M. F. Search for Related Content PubMed PubMed Citation Articles by Lin, C. J. Articles by Villares, S. M. F.