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Endocrinology Vol. 142, No. 6 2649-2659
Copyright © 2001 by The Endocrine Society

ARTICLES

Genomic Structure and Transcriptional Regulation of the Human Growth Hormone Secretagogue Receptor1

Stephan Petersenn, Anja C. Rasch, Martina Penshorn, Frank U. Beil and Heinrich M. Schulte

IHF Institute for Hormone and Fertility Research, University of Hamburg (S.P., A.C.R., M.P.), 22529 Hamburg, Germany; Department of Medicine, University of Hamburg (S.P., F.U.B.), 20251 Hamburg, Germany; and Endokrinologikum Hamburg (H.M.S.), 22767 Hamburg, Germany

Address all correspondence and requests for reprints to: Dr. S. Petersenn, University Clinic Essen, Department of Endocrinology, Hufelandstrasse. 55, 45122 Essen, Germany.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Synthetic GH secretagogues stimulate GH release through binding to a recently cloned specific GH secretagogue receptor (GHS-R). The endogenous ligand of this receptor may be part of a new endocrine pathway controlling GH secretion. Two different receptor variants, type 1a and 1b, have been described that differ in their 3'-terminal amino acids. We investigated the genomic structure and transcriptional regulation of the human GHS-R. An 18-kb genomic clone including sequences encoding for the two GHS-R variants was isolated. Sequencing revealed that the two variants originate from specific RNA processing of a single gene that spans approximately 4.1 kb. The transcription start site was defined by 5'-inverse PCR analysis at position -227. RT-PCR analysis points to differential transcriptional initiation and processing. Type 1a is encoded by two exons; 2152 bp of intronic sequence are removed by splicing at position 796/797 relative to the translation start site. Type 1b is encoded by a single exon. A putative polyadenylation signal consensus motif was identified at position +4118; 2.7 kb of the 5'-flanking region were sequenced, and putative transcription factor binding sites were identified. Transcriptional regulation was investigated by transient transfections using promoter fragments ranging in size from 168-1745 bp; 1745 bp of the GHS-R promoter directed significant levels of luciferase expression in GH4 rat pituitary cells, whereas no activity was detected in monkey kidney COS-7 cells, human endometrium Skut-1B cells, mouse hypothalamic LHRH neuronal GT1–7 cells, or mouse corticotroph pituitary AtT20 cells. A minimal 309-bp promoter allowed pituitary-specific expression. Its activity in COS-7 cells was enhanced by cotransfection of the pituitary-specific transcription factor Pit-1. We did not find any regulation of the GHS-R promoter by forskolin, somatostatin, insulin-like growth factor I, or 12-O-tetraphorbol 12-myristate 13-acetate. Thyroid hormone and estrogen lead to a significant stimulation; glucocorticoids lead to a significant inhibition. Further mapping suggests a thyroid hormone-responsive element, an estrogen-responsive element, and a glucocorticoid-responsive element located between -309 and the translation start codon. These studies demonstrate the nature of the human GHS-R gene and identify its 5'-flanking region. Furthermore, pituitary-specific activity of the promoter and regulation by various hormones are demonstrated.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE SECRETION of GH by the anterior pituitary is under complex control. Small synthetic molecules termed GH secretagogues (GHS) act on the pituitary and the hypothalamus to stimulate and amplify pulsatile GH release. These compounds appear to mimic a putative endogenous ligand that activates a receptor distinct from that of GH-releasing hormone (GHRH) and somatostatin and whose function is probably critical in the regulation of normal GH secretion. Analogs studied to date include GHRP-6, hexarelin, and MK-0677 (1). An endogenous specific ligand of 28 amino acids has recently been purified from rat stomach; it has been termed ghrelin (2). Expression in stomach was demonstrated by Northern blot analysis, and more sensitive RT-PCR revealed the presence of transcripts in the brain. Immunohistochemical analysis demonstrated that ghrelin-immunoreactive neurons were localized in the hypothalamic arcuate nucleus. Interestingly, considerable plasma concentrations were found in healthy human blood.

The GHS receptor (GHS-R) belongs to the family of G protein-coupled receptors (3). It is expressed by a single highly conserved gene in the human, chimpanzee, pig, cow, rat, and mouse. The gene is found at the chromosomal location 3q26.2 (4). Two types of GHS-R complementary DNAs (cDNAs) 1a and 1b, have been identified from human and pig (3) and rat (4, 5). Their sequences do not show significant homology with other known receptors; the closest relatives are the neurotensin receptor and the TRH receptor, with 59% and 56% similarity, respectively. The human GHS-R type 1a consists of 366 amino acids with 7 transmembrane regions; its molecular mass is approximately 41 kDa. Type 1b consists of 289 amino acids with only 5 predicted transmembrane regions, the nucleotide sequence of codons 1–265 is identical to that in type 1a. Beyond that, the type 1b cDNA diverges in its nucleotide sequence and is fused to a short conserved reading frame of 24 amino acids. Type 1a was demonstrated to confer high affinity, specific binding of GHS and lead to Ca2+ release through stimulation of the G protein subunit G{alpha}11. In contrast, type 1b failed to bind GHS and to respond to GHS (3). The binding affinities of various GHS are correlated with the GH stimulatory effect (6). The intracellular signaling is thought to involve activation of phospholipase C (7). Subsequent generation of inositol triphosphate (8) causes an increase in free intracellular Ca2+ by redistribution from intracellular stores. Activated phospholipase C may also lead to tyrosine phosphorylation of a potassium channel, resulting in inhibition of that channel. Depolarization of the membrane and activation of L-type voltage-gated calcium channels elicit GH secretion (7). Expression of the cloned GHS-R was shown in hypothalamus and pituitary, consistent with its role in regulating GH release (3). Expression was also demonstrated in various other regions of the central nervous system and in the pancreas, possibly indicating its involvement in as yet undefined physiological functions (9). Studies using a photoactivatable ligand suggest a second distinct GHS-R subtype in pituitary cells with a molecular mass of 57 kDa (10) and a third subtype in heart with a molecular mass of 84 kDa (11). The cDNAs of these additional subtypes have not yet been isolated.

Characterization of the endogenous GHS and its receptors may allow new insight into the regulation of GH secretion and additional physiological roles of these agents. GHS may also offer practical diagnostic and therapeutic value in humans. To understand the expression of GHS-R we isolated a genomic clone of GHS-R and investigated the structure and regulation of the GHS-R gene.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Forskolin, 12-O-tetraphorbol 12-myristate 13-acetate (TPA), somatostatin, hydrocortisone, T3, and ß-estradiol were purchased from Sigma (St. Louis, MO). Insulin-like growth factor I (IGF-I) was purchased from BIOZOL-Diagnostics (Eching, Germany).

Plasmids
The plasmid construct pCMV-hpit1 contains the full-length human Pit-1 cDNA under control of the cytomegalovirus-1 (CMV-1) promoter. Plasmid pGL3-Basic is a luciferase vector lacking eukaryotic promoter and enhancer sequences (Promega Corp., Madison, WI). pGL3-Control contains a simian virus 40 (SV40) promoter and an SV40 enhancer inserted into the structure of pGL3-Basic (Promega Corp.). phGH344/luc contains 344 bp of the human GH promoter and includes two nucleotides of the transcribed sequence. pSV-ß-GAL contains an SV40 promoter and an SV40 enhancer. Both promoter and enhancer drive transcription of the lacZ gene, which encodes the ß-galactosidase enzyme (Promega Corp.).

Isolation of GHS-R cDNA probe and screening of genomic DNA library
Total RNA was extracted from a human somatotropic pituitary tumor. RNA (0.5 µg total) was reverse transcribed and amplified by the PCR using a GeneAmp RNA PCR Kit (Perkin-Elmer Corp., Norwalk, CT). The DNA fragment containing residues 67–215 of the human GHS-R cDNA (numbering of residues as in Ref. 3 , relative to the translation start codon) was amplified (95 C for 30 sec, 62 C for 60 sed, 72 C for 60 sec, 35 cycles) using G1 (5'-GCT-TCC-CCC-GGC-AAC-GAC-TC-3') and G2 (5'-CGG-AAG-CGC-GAC-ACC-ACC-AG-3') as primers. The PCR product was fractionated on an 0.8% agarose gel and subsequently cloned into pCRII (GHSR/pCRII) using the TA-Cloning Kit (Invitrogen, San Diego, CA). Sequencing analysis confirmed the identity of the amplified DNA. GHSR/pCRII was used to synthesize a digoxigenin-labeled probe by PCR using a GeneAmp PCR Kit (Perkin-Elmer Corp.), primers G1 and G2 (95 C for 30 sec, 62 C for 60 sec, 72 C for 60 sec, 30 cycles), and digoxigenin-11-deoxy-UTP (Roche Molecular Biochemicals, Mannheim, Germany). An amplified human placenta {lambda} FIX II genomic DNA library (Stratagene, La Jolla, CA) was screened for a GHS-R fragment by adapting a PCR-based method (12). Aliquots containing approximately 10,000 plaque-forming units were distributed into each of the 96 wells of a microplate. Samples from each well in a column and from each well in a row were combined. Pools were screened by PCR using primers G1 and G2 (95 C for 30 sec, 62 C for 60 sec, 72 C for 60 sec, 35 cycles) and were analyzed by gel electrophoresis. Screening of pools allowed identification of single wells, which were further investigated using the same PCR conditions. Aliquots of positive wells were plated and screened with the digoxigenin-labeled probe. Hybridization and detection were performed using a DIG Luminescent Detection Kit (Roche Molecular Biochemicals) following the manufacturer’s protocol. Briefly, prehybridization was performed for 1 h at 42 C in 50% formamide, 5 x SSC (standard saline citrate), 2% blocking reagent, 0.02% SDS, and 0.1% N-lauroylsarcosine. The prehybridization solution was exchanged against fresh solution for hybridization. The probe was denatured at 100 C for 4 min before being added to the hybridization solution at 42 C overnight. The filters were washed twice for 5 min each time with 2 x SSC/0.1% SDS at room temperature and twice for 15 min each time in 0.1 x SSC/0.1% SDS at 68 C. The hybridized probe was immunodetected with antidigoxigenin and was then visualized with the chemiluminescence substrate CPD1. The light emission was recorded on Kodak XAR-5 film (Eastman Kodak Co., Rochester, NY). Positive recombinant plaques were purified by replating twice and were grown in liquid culture. Phage DNA was prepared with a QIAGEN Lambda Midi Kit (QIAGEN, Hilden, Germany).

Subcloning of phage DNA
The phage DNA was digested with various restriction enzymes and separated on a 0.7% agarose gel. Subsequently, genomic fragments were purified by a QIAEX Gel Extraction Kit (QIAGEN) and subcloned into Bluescript SKII+ vector (Stratagene, La Jolla, CA). Plasmids were prepared by QIAGEN Plasmid Maxi Kits (QIAGEN).

Nucleotide sequence determination
Double stranded plasmid DNA was sequenced by fluorescent sequencing using dye-labeled terminators (ABI PRISM Dye Terminator Cycle Sequencing Ready Reaction Kit, PE Applied Biosystems, Warrington, UK) and Applied Biosystems instrumentation. Sequences were assembled using Lasergene computer software (DNASTAR, Madison, WI). To avoid errors, all sequences were determined by sequencing both strands of the DNA. The nucleotide sequence data reported in this paper has been submitted to GenBank and assigned Accession No. AF 369786.

Determination of the genomic structure
Exon/intron boundaries were determined by a loss of identity between the genomic and cDNA nucleotide sequences and also by the presence of consensus donor and acceptor signals at the point of divergence. Transcription factor-binding sites in the 5'-region were identified using TFSEARCH on the internet, which searches for sequence fragments vs. TFMATRIX, the transcription factor-binding site profile database by E. Wingender, R. Knueppel, P. Dietze, and H. Karas (GBF-Braunschweig). Polyadenylation signals in the 3'-region were identified by a search for the consensus sequences AATAAA.

Determination of the transcription start site by 5'-inverse PCR
We used an adapted inverse PCR method to clone 5'-cDNA regions (13). Briefly, 5 µg total RNA obtained from a human pituitary tumor were reverse transcribed (51 C for 30 min, 95 C for 5 min) by use of 200 U Superscript reverse transcriptase (Life Technologies, Inc., Grand Island, NY) and 20 pmol antisense primer G3 (5'-TAC-TGC-CAG-AGG-CGA-ACG-A-3', position +318). The second strand was synthesized with Escherichia coli DNA polymerase I (Life Technologies, Inc.) and T4 DNA ligase (Life Technologies, Inc.); simultaneously RNA was degraded by ribonuclease H (16 C for 8 h, 75 C for 5 min). cDNA was purified using a PCR purification kit (Roche Molecular Biochemicals). Blunt ends were generated with T4 DNA polymerase (11 C for 15 min; Life Technologies, Inc.). After phenol extraction, cDNA self-ligation was performed with T4 DNA ligase (16 C, overnight). The ligation reaction was used for 40 cycles of PCR (95 C for 1 min, 59 C for 1 min, 72 C for 2 min) with sense primer G4 (5'-AGC-ATG-GCC-TTC-TCC-GAT-CT-3', position +250) and antisense primer G5 (5'-GCG-ATA-CCC-ACC-ACG-AAG-AGT-3', position +176). PCR products were fractionated on an 0.8% agarose gel and subsequently cloned into pCRII using the TA-Cloning Kit (Invitrogen). Transcription start sites were determined by sequencing analysis and comparison with genomic and G3 sequences. Transcripts of the GHS-R were studied by RT-PCR analysis. Total RNA (800 ng) obtained from a pituitary tumor or normal pituitary messenger RNA (mRNA; 100 ng; CLONTECH Laboratories, Inc., Palo Alto, CA) was reverse transcribed and amplified (95 C for 30 sec, 60 C for 60 sec, 72 C for 60 sec, 40 cycles). Primer pairs were GP8 (5'-CTT-CTG-CCT-CTC-ACC-TCC-CTC-TC-3', position -168) and G6 (5'-CCA-GGG-GCA-TGC-AGA-GGA-AG-3', position +295), GP9 (5'-GGG-AAG-TGC-GAG-ATG-GAA-3', position -386) and G5, or G7 (5'-TGG-CCG-ACC-TGG-ACT-GGG-ATG-CT-3', position +47) and G8 (5'-GGG-TCG-GTG-CCG-TTC-TCG-TGC-TC-3', position +575). The PCR fragments obtained were sequenced.

Construction of luciferase expression vectors containing upstream sequence
Upstream sequences were obtained by amplification of the genomic clone using GP1 (5'-CTC-AGC-TGA-ACA-GGC-TCT-GGG-AC-3', starting at position -9) as antisense primer and GP2 (5'-GGT-AGT-AGA-GGT-GGT-ACA-AAC-T-3', starting at position -1745), GP3 (5'-GGT-TGA-AGA-CCT-TGC-CCA-AGG-3', starting at position -1137), GP4 (5'-ACC-CCG-CCA-CCA-GGT-TTG-CAT-C-3', starting at position -951), GP5 (5'-CCA-AGT-CGA-GTT-GGT-CAA-GCT-CG-3', starting at position -643), GP6 (5'-GCT-TCG-GAG-AGA-GCG-CCT-CAC-3', starting at position -460), GP7 (5'-CCG-GAG-GAT-GTG-TTG-GGA-GAA-AC-3', starting at position -309), and GP8 as sense primers. The PCR products were fractionated on a 1.0% agarose gel. Fragments of the correct size were subsequently cloned into pCRII using the Zero Blunt PCR Cloning Kit (Invitrogen). Sequencing analysis confirmed the identity and orientation of the amplified DNA. DNA fragments were isolated by restriction digestion with KpnI and NotI, purified using a QIAEX Gel Extraction Kit (QIAGEN), and inserted upstream of the luciferase reporter gene into pGL3-Basic mammalian expression vector (Promega Corp.). Plasmids were prepared using QIAGEN Plasmid Maxi Kits.

Cell culture, transient transfection, luciferase assay, and ß-galactosidase assay
Rat mammosomatotroph pituitary GH4 cells, mouse corticotroph pituitary AtT20 cells, monkey kidney COS-7 cells, and mouse hypothalamic LHRH neuronal GT1–7 cells were grown in DMEM (Life Technologies, Inc.) containing 10% FCS (Serva, Heidelberg, Germany). Human endometrium Skut-1B cells were grown in DMEM/Ham’s F-12 medium containing 10% FCS. Cells were maintained at 37 C in 5% CO2. Cells (5 x 105/well) were seeded in six-well plates for transfection. The medium was changed 3 h before transfection. Experimental and control plasmids were mixed and transfected in triplicate by CaPO4-DNA coprecipitation. Transfections included 3 µg reporter gene construct and 2 µg pSV-ß-GAL as an internal control of transfection efficiency. For cotransfection studies 1.5 µg pCMV-hPit1 were added. The total amount of DNA was maintained constant with nonspecific DNA. After 16 h in the presence of DNA, cells were shocked for 2 min at room temperature with 15% glycerol in PBS, which was then replaced by serum-free DMEM containing 3% of BSA. Cells were harvested 64 h after transfection in lysis buffer (Promega Corp.). The luciferase assay was performed in a final volume of 120 µl, containing 20 µl cell extract, following the protocol for the luciferase assay system (Promega Corp.). Luciferin was added just before measuring light units, which were measured during the first second of the reaction at 25 C in a Luminometer. The ß-galactosidase assay was performed following the protocol for the ß-galactosidase assay system (Promega Corp.). Cell extract (50 µl) was incubated with 50 µl assay buffer until color developed (30–120 min), and the reaction was stopped by adding 150 µl 1 M Na2CO3. Absorbance was then read at 405 nM. Luciferase light units were normalized to the activity of ß-galactosidase. Data are expressed as the mean ± SEM. All experiments were repeated at least three times.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Isolation of a genomic clone of the human GHS-R
Screening of approximately 1,000,000 phage clones of a human genomic library with a 5'-GHS-R cDNA probe resulted in the isolation of several positive clones. Phage DNA from 1 positive clone was prepared and digested with NotI restriction enzyme to release the insert. Southern analysis confirmed hybridization of the 5'-GHS-R cDNA probe to an approximately 18-kb insert. Restriction digestion of the genomic clone with NotI and SstI released five fragments of approximately 1.6, 2.8, 3, 4.8, and 5.5 kb. The genomic fragments were individually subcloned into a pBluescript SKII+ vector and designated p1.6/SKII, p2.8/SKII, p3/SKII, p4.8/SKII, and p5.5/SKII, respectively. The nucleotide sequences of the fragments were determined on both DNA strands.

Genomic structure of the GHS-R
The genomic structure of GHS-R type 1a and type 1b was determined by DNA sequence comparison. The 5.5-kb fragment was shown to contain nucleotides 1–796 of the open reading frame of GHS-R type 1a, followed by 1958 nucleotides of intronic sequence (Fig. 1Go). The 3-kb fragment contained 194 nucleotides of intronic sequence, followed by nucleotides 797-1101 of GHS-R type 1a and approximately 2 kb of the 3'-flanking region. Additional intronic sequence between these fragments was excluded by amplification of genomic DNA obtained from lymphocytes and sequencing of an overlapping PCR fragment. Nucleotides 1–796 of the open reading frame of GHS-R type 1b are identical to type 1a and are therefore contained in the 5.5-kb fragment. Nucleotides 797–870 of GHS-R type 1b were shown to align to the following 74 nucleotides of intronic sequence of type 1a.



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  		Figure 1. Partial genomic sequence of GHS-R type 1a and type 1b. The translation start site is defined as +1. Nucleotide positions are shown on the left. The coding sequence of GHS-R type 1a is shown in capital letters; the intronic sequence and 3'-untranslated sequence are shown in small letters. The amino acid sequence is depicted below the nucleotide sequence. Splicing positions are marked by vertical lines. For GHS-R type 1a, 2145 nucleotides of intronic sequence are removed by splicing. The coding sequence and amino acid sequence of GHS-R type 1b differing from type 1a are shown in bold letters. A putative polyadenylation signal in the 3'-region of the GHS-R is indicated by a horizontal line.

 
RNA protection analysis and primer extension analysis did not allow for determination of the transcription start site, possibly because of low expression levels of the GHS-R. Therefore, the GHS-R transcription start site was determined by 5'-inverse PCR using total RNA extracted from a human somatotroph pituitary tumor (Fig. 2AGo). We analyzed 15 PCR clones, of which 10 contained 227 bp of 5'-flanking region, determining an adenosine residue as a major transcription start site. The sequence obtained by 5'-inverse PCR was identical to the genomic sequence, excluding introns in the 5'-untranslated region. The remaining clones contained various length of 5'-flanking region, possibly indicating further transcription start sites or artificial early termination of RT. Analysis of the 5'-inverse PCR sequence revealed that a few nucleotides of the primer used for RT next to the putative transcription start site were replaced. Because of the variability of the nucleotide changes at a constant position and the presence of the correct sequence 5' of the primer, these changes were considered artifacts, possibly occurring during ligation. A putative polyadenylation signal containing the homology sequence AATAAA was identified at +4118 in the 3'-flanking region.



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  		Figure 2. Determination of the transcription start site of the human GHS-R gene by 5'-inverse PCR (A) and analysis of transcripts by RT-PCR (B). A, An adapted 5'-inverse PCR method was used to clone 5' cDNA regions (13 ). RNA obtained from a human pituitary tumor was reverse transcribed. After second strand synthesis and blunt end generation, the DNA strand was self-ligated and subjected to PCR cloning. A major transcription start sites (10 of 15 clones) was determined at bp -227 by sequencing analysis and comparison with genomic sequence. The position of primer G3 is indicated. B, GHS-R transcripts were amplified from normal (P) and adenomatous (T) pituitary tissue using primers GP9 (-386) and G5 (+176) or GP8 (-168) and G6 (+295). Primers G7 (+47) and G8 (+575) were used to confirm expression of the GHS-R. To exclude contamination of the tumor RNA with genomic DNA, amplification was performed in parallel with omission of the RT. N, Negative controls; M, 100-bp ladder.

 
To study transcripts in more detail, we performed RT-PCR analysis of both pituitary tumor RNA as well as commercially available pituitary mRNA from normal tissue (Fig. 2BGo). Using GP8 (-168) and G6 (+295) as primers, we obtained PCR products from both normal and adenomatous pituitary tissue. Contamination with genomic DNA was excluded by PCR of RNA. Sequencing of the RT-PCR fragment confirmed that genomic and cDNA sequence were identical in the 5'-untranslated region, therefore excluding an intron in that transcript. By RT-PCR analysis using GP9 (-386) and G5 (+176), we obtained a PCR product of approximately 350 nucleotides, again from both normal and adenomatous pituitary tissue. The sequence was partially identical to the 5'-sequence of the GHS-R but did not include the sequence between -328 and -133. A PCR product containing both the sequence between -386 and -227 and the putative intronic sequence between -328 and -133 was not observed. Therefore, we found evidence for transcripts starting at -227, without any splicing occurring in the 5'-untranslated region. We also found evidence for transcripts starting upstream of -386. In that case, intronic sequence between -328 and -133 is spliced out.

These data reveal that GHS-R type 1a is encoded by at least two exons; 2152 nucleotides of intronic sequence are removed from the pre-mRNA by splicing at position 796/797 relative to the translation start codon. Exon 1 is composed of the 5'-untranslated region as well as the first 265 amino acids encoding for the transmembrane regions I–V, exon 2 encodes for 101 amino acids (transmembrane regions VI and VII) and includes the 3'-untranslated region. The nucleotide sequence of the intron revealed no deviations from the consensus sequences for splice site in the 5'-donor and 3'-acceptor splice sites; the intron-exon splice junctions followed the GT/AG rule (14). The coding sequence of GHS-R type 1b is encoded by a single exon, which encodes for 289 amino acids, allowing for 5 transmembrane regions. The 265 amino-terminal amino acids are identical to GHS-R type 1a. The genomic sequence obtained matched the previously described cDNA sequences for the open reading frame of GHS-R type 1a and type 1b (3), except for a nucleotide substitution (C instead of T) at position 171 relative to the translation start codon, which does not result in an amino acid change.

Sequence analysis of the 5'-untranscribed sequence
The 5.5-kb fragment contained approximately 2.7 kb of 5'-flanking region of the GHS-R gene (Fig. 3Go). No potential TATA box was identified in the appropriate location. A comparison of the 5'-flanking sequence of the GHS-R gene with the initiator consensus sequence PyPyANA/TPyPy raises the possibility of a putative initiator element located at position -215 or -237 relative to the translation start codon. A number of other putative response elements were identified by homology comparison. These include putative binding sites for the enhancer factor AP-1 at bp -2363, -1620, -1526, -1211, -967, -767, -504, and -336; for AP-4 at bp -2131, -1885, -1506, -1270, -910, -779, and -596; the consensus sequences for the nuclear factor NF-1 at bp -2213, -1394, -1356, and -1297; and for SP1 at -2250 and -2043, and a binding site for the upstream regulatory factor (USF) at bp -2239. Furthermore, several binding sites for tissue-specific transcription factors, such as the POU domain factors Oct-1 at bp -2377, -2326, -2277, -1832, -1153, and -924; Pit-1 at bp -2618, -2476, -2399, -2162, -1756, and -1083; and Brn-2 at bp -1867 were identified. In addition, the promoter region contains consensus motifs corresponding to inducible promoter elements that are known to bind transcription factors induced by exogenous stimuli. These include binding sites for the estrogen receptor at bp -2420, -1319, -634, and -242, and for the glucocorticoid receptor at bp -1464.



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  		Figure 3. Nucleotide sequence of the 2.7-kb 5'-flanking region of the GHS-R. The translation start site is defined as +1. Nucleotide positions are shown on the left. Potential transcriptional regulatory sequences identified by a computer-assisted analysis are underlined by arrows. Consensus sequences for initiator elements near the transcription start site are boxed. The transcription start site identified is indicated by an arrow.

 
Transient expression analysis of the 5'-flanking region
To determine whether the GHS-R 5'-flanking region can direct cell-specific expression, the proximal 168 and 1745 bp were inserted into a transient expression vector. pGL3-Basic, which contains bacterial luciferase as the reporter gene, and the resulting plasmids (-168GHSR/luc and -1745GHSR/luc) were transiently transfected into various cultured cell lines. Gene transfer studies were performed by calcium-phosphate transfection, and luciferase enzyme activity was measured in light units as an indication of promoter activity. Cells were cotransfected with pSV-ß-GAL as an internal control for transfection efficiency. As shown in Fig. 4Go, 1745 bp of the GHS-R promoter directed significant levels of luciferase expression in GH4 rat pituitary cells compared with the promoter-less pGL3-Basic luciferase vector. In contrast, we observed no significant activity of -1745GHSR/luc in COS-7 monkey kidney cells, human endometrium Skut-1B cells, mouse hypothalamic LHRH neuronal GT1–7 cells, or mouse corticotroph pituitary AtT20 cells. One hundred and sixty-eight base pairs of the GHS-R promoter directed no significant activity in any of the cell lines tested. The relative activities of pGL3-Control containing an SV40 viral promoter in COS-7, Skut-1B, GT1–7, AtT20, and GH4 cells were 7000-, 120-, 13-, 10-, and 12-fold, respectively.



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  		Figure 4. Promoter activity of the GHS-R 5'-flanking region in various cell lines. The -168GHSR/luc ({square}) and -1745GHSR/luc ({blacksquare}) constructs were transfected in parallel with pGL3-Basic, which lacks promoter activity, into monkey kidney cells COS-7, human endometrium Skut-1B cells, mouse hypothalamic LHRH neuronal GT1–7 cells, mouse corticotroph pituitary AtT20 cells, or GH4 rat pituitary cells. Cotransfection with CMV-ß-galactosidase was used to control for transfection efficiency. The luciferase activity of each construct was normalized with the ß-galactosidase activity, and values were expressed as fold induction relative to the activity of the promoter-less construct pGL3-Basic. Values represent the mean ± SEM of at least three determinations.

 
Elements required for GHS-R promoter activity are located within 309 bp 5' of the translation start codon
To further analyze the 5'-flanking region of the GHS-R gene for constitutive promoter activity in GH4 rat pituitary cells, varying lengths of 5'-flanking regions created by PCR were placed upstream of the luciferase reporter gene (left panel of Fig. 5Go). Only background activity was obtained with the construct containing 168 nucleotides 5' to the translation start codon (right panel of Fig. 5Go), whereas a significant increase in activity was observed with the construct containing 309 bp. The construct containing 460 bp demonstrated significantly lower activity. Another increase in activity was observed with the constructs containing 643 and 951 bp of 5'-flanking region, which is comparable to the activity obtained with the SV40 viral promoter. With the construct containing 1137 bp the level of luciferase activity was slightly decreased, further decreased activity was found for the construct containing 1745 bp. These results suggest that the regions between -951 and -643, between -643 and -460, and 3' of bp -309 contain positive regulators of GHS-R expression in GH4 rat pituitary cells. In contrast, the regions between -1745 and -951 and between -460 and -309 may contain negative regulators.



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  		Figure 5. Promoter activities of various deletion constructs of the GHS-R 5'-flanking region. The schematic diagram on the left represents a series of GHS-R promoter-luciferase gene chimeric plasmids with variable 5'-ends (from -1745 to -168) and the same 3'-end (-9). Each construct was transiently transfected into GH4 rat pituitary cells. Promoter activity is normalized for transfection efficiency by the ß-galactosidase activity and is expressed relative to the activity of the pGL3-Basic control (right panel). Data are the mean ± SEM of at least three independent experiments performed in triplicate.

 
The GHS-R promoter is under control of the pituitary-specific transcription factor Pit-1
Selective expression of the GHS-R 5'-flanking region in GH4 pituitary cells suggests that an interaction between pituitary-specific factors and the GHS-R promoter may facilitate expression of the GHS-R. The POU domain transcription factor Pit-1 is strictly pituitary specific in expression and is required for determination of the somatotroph cell line (15). COS-7 monkey kidney cells do not produce any significant amount of Pit-1. GHS-R 5'-promoter deletion constructs transiently transfected into COS-7 cells were used to determine the effect of Pit-1 cotransfection and the approximate location of the Pit-1-responsive element(s) in the GHS-R promoter. The transcriptional activity of the GHS-R promoter measured in COS-7 cells is approximately 100-fold lower than that in GH4 cells. As shown in Fig. 6Go, cotransfection of Pit-1 did not enhance the activity of the construct containing -168 bp of 5'-flanking region compared with the mock-cotransfected cells. In contrast, cotransfection of Pit-1 to any of the larger constructs ranging from 309 to 1745 bp of 5'-flanking region resulted in significantly increased promoter activity. These results suggest that a Pit-1-responsive region is localized downstream of position -309. The effect of Pit-1 cotransfection on the GHS-R promoter was lower than that on 344 nucleotides of the human GH promoter. The -344 GH/luc construct demonstrated a 37-fold higher activity than the promoterless construct in the presence of Pit-1 compared with only 3.6-fold increased activity in the absence of Pit-1.



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  		Figure 6. Effect of Pit-1 cotransfection on the activities of various deletion constructs of the GHS-R 5'-flanking region. The indicated series of GHS-R promoter deletions was cotransfected with ({blacksquare}) or without ({square}) pCMV-hpit1 transiently into COS-7 monkey kidney cells, as described in Materials and Methods. Promoter activity is normalized for transfection efficiency by the ß-galactosidase activity and is expressed relative to the activity of the pGL3-Basic control, respectively. Results are the mean ± SEM of three transfections.

 
Activity of the GHS-R promoter is stimulated by thyroid hormone and estrogen and inhibited by glucocorticoids in GH4 cells
To investigate the hormonal regulation of the GHS-R 5'-flanking region, we analyzed the effects of various agents on 1745 bp of GHS-R promoter in GH4 rat pituitary cells. As shown in Fig. 7Go, treatment with 10-6 M forskolin, 10-8 M somatostatin, 10-7 M TPA, and 13 nM IGF-I did not significantly influence activity of the GHS-R promoter region analyzed. Treatment with 10-9 M T3 or 10-9 M ß-estradiol significantly enhanced the activity of 1745 bp of GHS-R 5'-flanking region. In contrast, treatment with 10-7 M hydrocortisone significantly inhibited 1745 bp of GHS-R promoter in GH4 rat pituitary cells.



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  		Figure 7. Hormonal regulation of the human GHS-R promoter. A GHS-R deletion constructs -1745GHSR/luc ({square}) and the promoter-less vector pGL3-Basic ({blacksquare}) were transiently transfected into GH4 rat pituitary cells. Regulation by various agents was tested by treatment with 10-6 M forskolin (For), 10-8 M somatostatin (SS), 10-7 M TPA, 13 nM IGF-I, 10-9 M T3, 10-9 M ß-estradiol (E2), and 10-7 M hydrocortisone (HC). Activity is expressed as fold induction relative to that driven by each construct transfected alone in the absence of treatment and represents the mean ± SEM of three independent experiments.

 
We next performed 5'-deletion analysis to identify specific sequences that regulate hormone-induced promoter activity. Plasmids containing progressively decreasing amounts of GHS-R 5'-flanking region upstream of the luciferase gene were transiently transfected into GH4 rat pituitary cells, with or without hormonal treatment, and promoter activity was measured by luciferase assay. Stimulation of the GHS-R promoter by thyroid hormone or estrogen was maintained for all deletion constructs investigated, with no significant changes among the various constructs (Fig. 8Go, A and B). Therefore, positive thyroid- and estrogen-responsive elements may be located in the region between -309 and the transcription start site. Similarly, inhibition of the GHS-R promoter by glucocorticoids was maintained for all deletion constructs investigated, with no significant changes between the various constructs (Fig. 8CGo). A negative glucocorticoid-responsive element may be located downstream of bp -309.



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  		Figure 8. Mapping of the cis elements required for activation of the GHS-R promoter by thyroid hormone (A) or estrogen (B) or inhibition of the GHS-R promoter by glucocorticoids (C). Regulation was mapped by transfecting 3 µg of the indicated GHS-R promoter deletion constructs or the pGL3-Basic vector into GH4 pituitary cells, followed by treatment with the indicated agents. Activity is expressed as fold induction relative to that driven by each construct transfected alone in the absence of treatment and represents the mean ± SEM of three independent experiments.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
To gain insight into the developmental and differential regulation of the GHS-R, characterization of its promoter and gene are essential. We present data regarding the structure and regulation of the human GHS-R. In a recent report the 5'-flanking region of the GHS-R gene has been isolated (16), but a systematic study of regulation of the GHS-R gene has not yet been described.

The sequence obtained from an isolated genomic clone of the GHS-R confirms the published sequences for the open reading frames of GHS-R subtypes 1a and 1b (3) and extends them at the 5'- and 3'-ends. By 5'-inverse PCR using total RNA obtained from a human somatotropic pituitary tumor a major transcription start site was located 227 nucleotides upstream from the ATG translation initiation codon. Ten of 15 clones confirmed that transcription start site; occasional shorter sequences may represent additional transcription start sites or artifacts. Comparison of genomic sequence and sequences obtained by 5'-inverse PCR in our study did not reveal any differences in the 5'-untranslated region, virtually excluding any introns in that region for these transcripts. Kaji et al. suggested a major transcription start site at position -453 by rapid amplification of 5'-cDNA ends (5'-RACE) analysis using commercially prepared pituitary gland cDNA (16). As they obtained at least 13 RACE products with different 5'-termini, the researchers discussed the possibility of multiple additional transcription start sites. They also described an intron in the 5'-untranslated region, as PCR products obtained by 5'-RACE in their studies lacked the sequence from -328 to -133. To understand transcriptional initiation and processing in more detail, we performed RT-PCR analysis of normal and adenomatous pituitary tissue. The results suggest that there are both transcripts with a short unspliced 5'-untranslated region, supporting our observation of a transcription start site at -227, as well as transcripts starting upstream of -386. In that case, intronic sequence between -328 and -133 is spliced out, as suggested by Kaji et al. The functional significance of differential transcriptional initiation and processing remains to be determined. We did not observe any differences between normal and adenomatous pituitary tissue.

The gene encoding for the human GHS-R type 1a spans approximately 4.3 kb, as demonstrated by the location of the transcription start site and the polyadenylation signal. The mRNA results from a splicing event that removes 2152 nucleotides of intronic sequence. In the GHS-R type 1b mRNA, the intron is not removed. As a result, type 1b contains 24 differing amino acids at the carboxyl-terminal region and uses an alternate stop codon. Splicing of type 1a mRNA is very similar for the rat, where 2034 nucleotides are removed at position +790/791. Because the intronic sequence in the rat differs from the human sequence, a putative type 1b receptor contains an 80-amino acid extension (4).

The isolation of a human genomic GHS-R clone allowed characterization of approximately 2.7 kb of the proximal promoter sequence. It matches the 1234 bp of 5'-flanking region recently reported by Kaji et al. (16), except for a few deviations, and extends it at the 5'-end. No TATA box was evident in the 5'-flanking region analyzed, but consensus sequences for initiator elements were found at locations -215 and -237 surrounding the transcription start site determined. In TATA-less genes, GC-rich domains or initiator elements have been proposed to direct gene transcription.

Specific expression of the GHS-R is largely confined to the central nervous system and pituitary. By ribonuclease protection assay analysis Guan et al. demonstrated GHS-R expression in the pituitary gland, hypothalamus, and hippocampus. A weak signal was observed in pancreas (9). Studies of human pituitary adenomas revealed functional expression in nearly all somatotroph, lactotroph, and corticotroph adenomas, but absence in most gonadotroph, thyrotroph, and silent adenomas (17, 18, 19, 20, 21, 22). Furthermore, expression of GHS-R was found in endocrine bronchial tumors, especially carcinoids, in gastrinomas, and in insulinomas, but not in nonendocrine bronchial tumors (19, 20). Putative binding sites for several transcription factors that may be involved in tissue specificity were identified on the GHS-R promoter. The USF has been implicated in the regulation of tissue-specific and developmentally regulated genes (23). The transcription factor Pit-1 is specifically expressed in the pituitary and is capable of activating both PRL and GH promoter in nonpituitary cells (15, 24). The octamer-binding protein Oct-1 differs in its ubiquitous tissue distribution and its ability to activate certain eukaryotic promoters that lack a TATA box (25), but is also coexpressed in cells of the anterior pituitary. The neuron-specific transcription factor Brn-2 is expressed in a distinct spatial and temporal pattern in the brain (26). In our transfection studies we found significant activity of 1.7 kb of 5'-flanking region of the GHS-R gene in a rat somatotroph pituitary cell line, but not in a monkey kidney cell line and a human endometrium cell line. These findings suggest a tissue-specific regulation of the GHS-R gene, but we cannot exclude that elements in the intron identified or 5' of the investigated promoter region may allow for expression in other tissues. Similar to our studies, Kaji et al. described significant activity of the -734 5'-flanking region in rat pituitary GH3 cells, whereas they could not detect any significant activity in HeLa human epithelioid cervix carcinoma cells or EP1 neuroblastoma cells (16). Expression of the GHS-R in somatotroph pituitary cells may partly account for the observed direct effects of GHS on GH release by these cells (27). In a mouse corticotroph pituitary cell line we did not observe any significant promoter activity. In agreement, the GHS stimulation of ACTH is probably mediated through central hypothalamic control rather than through a direct effect on the pituitary, possibly involving arginine vasopressin (28). Our studies of the mouse hypothalamic LHRH neuronal cell line also did not demonstrate significant promoter activity of the GHS-R gene. Supporting this observation, clinical studies did not find any modulation of LH and FSH secretion by GHS (29).

Our studies further show that 309 bp of the 5'-flanking region contain an element(s) that supports gene expression preferentially in a GH4 pituitary cell line, whereas a minimal 168-bp promoter did not allow for considerable transcription. The regions between positions -309 and -460 and between positions -951 and -1745 may contain repressor elements, whereas enhancer elements may be located between positions -460 and -643 and between positions -643 and -951. In the transfection studies by Kaji et al., the minimal promoter construct studied in GH3 pituitary cells contained 475 nucleotides of 5'-flanking region. These researchers found only low activity of constructs containing 475, 531, or 608 nucleotides of 5'-flanking region, whereas significant activity was observed for 669 nucleotides. By analysis of larger fragments, binding of repressor elements between positions -734 and -1224 was suggested (16). We cannot exclude that transcriptional activation of the GHS receptor promoter may differ in GH3 and GH4 cells.

Cotransfection of a Pit-1 expression vector did not change the activity of the -168GHSR/luc construct in COS-7 cells, but the activity of constructs containing at least -309 bp of 5'-flanking region was enhanced by Pit-1. No Pit-1 consensus sequence could be identified between -309 and the transcription start site, the precise mechanisms of Pit-1 activation of the GHS-R promoter are currently investigated. Pit-1 determines specific expression of GH in somatotropic pituitary cells (15). The enhancing effect of Pit-1 was much lower for the GHS-R promoter than for 344 nucleotides of the human GH promoter, in agreement with the significantly lower expression levels of the GHS-R in somatotroph pituitary cells.

Results from several systems suggest that the GHS-R gene may be under regulatory control. Therefore, hormonal regulation of the GHS-R promoter was studied. Regulation of the GHS-R by GHS itself has been demonstrated. A significant suppression of pituitary GHS-R mRNA was observed by Kineman et al. in rats infused with GHS, suggesting that GHS-R synthesis can be rapidly down-regulated by its own ligand (30). In contrast, Nass et al. described stimulation of pituitary GHS-R mRNA expression by hexarelin in both neonate and adult GHRH-deficient rats (31). The contrasting data reported by Kineman et al. and Nass et al. may be due to differences in the time course and delivery of GHS. At the hypothalamic level, continuous infusion of GHS did not alter GHS-R expression in arcuate and ventromedial nuclei of the rat (32). The underlying mechanisms are still unclear. In our studies we observed no significant effect of GHS on GH secretion by GH4 rat pituitary cells (data not shown), possibly indicating absent or low expression of the GHS-R or alteration of GHS-R coupling to the intracellular signal cascade. Similar, Bercu et al. did not find any stimulation of GH secretion by GHS in GH3 cells (33). Therefore, we used TPA as an activator of protein kinase C, which is an essential element of the signaling cascade regulated by the GHS-R (34). In our cell transfection studies we did not observe any significant changes in GHS-R promoter activity by TPA using the -1745GHSR/luc construct in GH4 pituitary cells. We cannot exclude that physiologically important transcription factor-binding sites may be located 5' of the analyzed promoter region. It also remains to be determined whether the suppression of GHS-R occurs by a direct effect of GHS at the pituitary level, and whether it is due to a transcriptional or a posttranscriptional mechanism.

A significant increase in pituitary GHS-R mRNA has been demonstrated in rats infused with GHRH, which was absent in in vitro studies (30). The promoter of the GHS-R does not contain any putative cAMP response element for binding of the transcription factor cAMP response element-binding protein, which could transduce the signaling cascade induced by GHRH to the GHS-R promoter. As GH4 cells do not possess any endogenous GHRH receptor, we used forskolin as an activator of protein kinase A in our transfection studies. We did not find any significant regulation of 1.7 kb of 5'-flanking region of the GHS-R gene by forskolin. Additional factors may be required for GHRH to alter pituitary GHS-R mRNA levels. For somatostatin, no changes in GHS-R mRNA levels were found after treatment with somatostatin antiserum (30). In agreement, incubation with somatostatin did not alter the activity of the -1745GHSR/luc construct in GH4 pituitary cells in our study. Insulin-like growth factor I (IGF-I) is known to exert a negative feedback on GH secretion at the level of the pituitary and the hypothalamus. In humans, IGF-I was shown to inhibit the GH response to GHS (35). It is presently unclear whether the effect of IGF-I involves any alteration of pituitary GHS-R expression. In our study we did not observe any effect of IGF-I on 1.7 kb of the 5'-flanking region of the GHS-R gene.

Glucocorticoids have long been known to regulate GH secretion by influencing both hypothalamic and pituitary functions. Thomas et al. found markedly decreased expression of the GHS-R gene in the arcuate and ventromedial nuclei of the rat hypothalamus after adrenalectomy (36), suggesting that adrenal steroids are necessary for normal GHS-R expression. In contrast, Ono et al. did not observe any effect of adrenalectomy on either pituitary or hypothalamic GHS-R gene expression in rats (37). Studying primary pituitary cultures, Tamura et al. described increased GHS-R mRNA levels after treatment with dexamethasone. The effect was blocked by the transcriptional inhibitor actinomycin D, indicating that dexamethasone increases GHS-R transcription at the pituitary level (38). These researchers also observed that adrenalectomy decreased pituitary GHS-R expression, whereas dexamethasone replacement was able to restore GHS-R mRNA levels. The reason for the conflicting observations is presently unclear. Our studies of a transient expression system indicate a negative glucocorticoid-responsive element located downstream of bp -309. A search of 2.7 kb of 5'-flanking region of the GHS-R vs. TFMATRIX did not identify any putative glucocorticoid receptor response elements between position -309 and the transcription start site, but located a response element at position -1464. Further analysis will characterize the glucocorticoid-responsive element within the GHS-R gene more precisely. In a recent abstract, Kaji et al. demonstrated inhibition of the GHS-R promoter by glucocorticoids in GH3 cells (39). Activity of a -961 promoter construct was significantly inhibited, whereas no effect was seen on a -734 promoter construct. The underlying mechanisms are not yet characterized. The reason for the different locations of the negative glucocorticoid element in our study and in the analysis presented by Kaji et al. is unknown and might involve differences in cell systems, promoter constructs, and transfection conditions. The contrasting data of expression analysis and promoter studies may be due to species differences or lack of the appropriate promoter elements in the transient transfection experiments.

Thyroid hormones are essential for growth in mammals. Thyroidectomy in rats resulted in a marked decrease in pituitary GHS-R mRNA levels without affecting those in the hypothalamus. Thyroid hormone replacement restored the pituitary GHS-R mRNA levels to control values (37). We identified a positive thyroid hormone-responsive element 3' of bp -309 by transient transfection studies. A search of 2.7 kb of 5'-flanking region of the GHS-R vs. TFMATRIX did not identify any putative thyroid hormone receptor response elements. The nature of the thyroid-responsive element identified is currently investigated. A reason for the discrepancy between promoter studies and the lack of corresponding regulatory sequences may be that the promoter scanning program did not identify all possible sequence motifs. Other scanning programs may reveal additional regulatory sequences depending on the mode of recognition.

GH secretion exhibits marked sexual dimorphism in many species. Kamegai et al. observed that pituitary GHS-R mRNA levels were greater in adult female rats compared with males (40). Levels of hypothalamic GHS-R mRNA were gender independent in this study. By in situ hybridization studies Bennett et al. observed higher GHS-R mRNA levels within the female ventromedial nucleus of the hypothalamus, whereas expression of the GHS-R in the arcuate nucleus did not differ between male and female rats (32). In a transient expression system we observed significant stimulation of the GHS-R promoter by ß-estradiol. Our studies suggest a positive estrogen-responsive element located downstream of bp -309. Interestingly, a search of the 5'-flanking region of the GHS-R gene vs. TFMATRIX identified a putative estrogen receptor response elements at position -242 that is currently being studied in more detail. Up-regulation of GHS-R expression in females by estrogen could explain the observed gender differences in GHS-R mRNA levels.

In summary, our initial characterization of the promoter region of the human GHS-R gene demonstrates that this gene contains a TATA-less promoter region with specific activity in a pituitary cell line. A number of transcription factor-binding sites were identified by sequence homology, the Pit-1 dependence of promoter activity was demonstrated. Regulation of the GHS-R promoter by thyroid hormone, estrogen, and glucocorticoids was shown. Given that many genes are regulated by elements in the intronic sequence, analysis of the intronic sequence reported here and use of these sequences in promoter-reporter constructs may reveal additional essential elements. Further study is necessary and will provide insight into the mechanisms that regulate the expression of the GHS-R gene. Analysis of the structure and regulation of the GHS-R gene may provide tools to investigate this new system of GH regulation in more detail.


   Acknowledgments
 
We thank A. Pauer for excellent technical assistance. The pCMV-hpit1 construct was provided by Garcia DiMattia, the AtT20 cells by Hartwig Schmale, and the Gt1–7 cells by James Olcese.


   Footnotes
 
1 Presented in part at the 81st Annual Meeting of The Endocrine Society, San Diego, California, 1999. It is based in part on the doctoral study by A.R. performed at the Faculty of Biology, University of Hamburg, and on the doctoral study by M.P., performed at the Faculty of Medicine, University of Hamburg. This work was supported by grants from the Deutsche Forschungsgemeinschaft (Schu 669/5–1 and GRK336). Back

Received October 24, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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M. C. Machado, S. Valeria de Sa, M. L. Correa-Giannella, R. R. Giorgi, M. A. A. Pereira, V. A. S Cescato, D. Giannella-Neto, and L. R. Salgado
Association between tumoral GH-releasing peptide receptor type 1a mRNA expression and in vivo response to GH-releasing peptide-6 in ACTH-dependent Cushing's syndrome patients
Eur. J. Endocrinol., May 1, 2008; 158(5): 605 - 613.
[Abstract] [Full Text] [PDF]

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 EndocrinologyHome page
R. M. Luque and R. D. Kineman
Gender-Dependent Role of Endogenous Somatostatin in Regulating Growth Hormone-Axis Function in Mice
Endocrinology, December 1, 2007; 148(12): 5998 - 6006.
[Abstract] [Full Text] [PDF]

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 J EndocrinolHome page
M. Sato, K. Nakahara, M. Miyazato, K. Kangawa, and N. Murakami
Regulation of GH secretagogue receptor gene expression in the rat nodose ganglion
J. Endocrinol., July 1, 2007; 194(1): 41 - 46.
[Abstract] [Full Text] [PDF]

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 J EndocrinolHome page
S. E Mitchell, R. Nogueiras, K. Rance, D V. Rayner, S. Wood, C. Dieguez, and L. M Williams
Circulating hormones and hypothalamic energy balance: regulatory gene expression in the Lou/C and Wistar rats.
J. Endocrinol., September 1, 2006; 190(3): 571 - 579.
[Abstract] [Full Text] [PDF]

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 J. Clin. Endocrinol. Metab.Home page
J. D. Veldhuis, D. M. Keenan, A. Iranmanesh, K. Mielke, J. M. Miles, and C. Y. Bowers
Estradiol Potentiates Ghrelin-Stimulated Pulsatile Growth Hormone Secretion in Postmenopausal Women
J. Clin. Endocrinol. Metab., September 1, 2006; 91(9): 3559 - 3565.
[Abstract] [Full Text] [PDF]

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 J Mol EndocrinolHome page
R. M Luque, M. D Gahete, R. J Valentine, and R. D Kineman
Examination of the direct effects of metabolic factors on somatotrope function in a non-human primate model, Papio anubis.
J. Mol. Endocrinol., August 1, 2006; 37(1): 25 - 38.
[Abstract] [Full Text]