Somatostatins (SSs), a diverse family of peptide hormones, have been shown to inhibit the release of growth hormone (GH) from the pituitary. In this study, we used rainbow trout to determine whether or not SSs affect growth in an extrapituitary manner, in particular, by decreasing GH sensitivity in liver. SS-14 significantly decreased hepatic GH binding in fish implanted (5.8 × 10−11 mol/h) for 15 days and in isolated hepatocytes. The processing of 125I-labeled trout GH (tGH) by isolated hepatocytes was investigated to determine whether or not the decrease in GH binding capacity resulted from receptor internalization. The internalization of 125I-labeled tGH was time dependent. By 6 h, 100 ng/ml SS-14 increased internalization of 125I-labeled tGH 58% over that observed in controls. Steady-state levels of mRNAs encoding the two hepatic growth hormone receptors (GHRs) of trout, GHR 1 and GHR 2, were measured to determine whether or not decreased GH binding capacity also resulted from decreased GHR synthesis. SS-14 directly inhibited steady-state levels of GHR 1 and GHR 2 mRNA in isolated hepatocytes in a concentration-dependent manner. The inhibitory effects of SS-14 on steady-state levels of GHR mRNAs resulted from reduced GHR mRNA transcription and not from altered mRNA stability. These results indicate that SSs regulate hepatic GH sensitivity by increasing GHR internalization and by altering GHR expression and suggest that SSs coordinate growth at the level of the pituitary, as well as at extrapituitary levels.
- growth hormone receptor
- rainbow trout
a primary growth-promoting factor in vertebrates is growth hormone (GH), which has been isolated from the pituitary gland from representatives in every extant class of vertebrate (18). Considerable evidence indicates that many of the growth-promoting actions of GH in fishes and other animals are mediated by insulin-like growth factor-1 (IGF-1) (4, 7, 11, 23). The complexity of the GH-IGF-1 axis has been expanded by several observations. First, two different forms of GH and GH receptor (GHR) occur in several teleost fish, including rainbow trout (8, 22, 37). Second, the extrapituitary production of GH has been noted in mammals and fish (22). Third, IGF binding proteins (IGFBPs) bind to IGFs and can modify their action (10). Fourth, GH binding proteins, the roles of which have not been fully established, occur in mammals and fish (1, 32). Lastly, GH as well as IGFBPs have direct, non-IGF-1-dependent effects on growth (6, 10).
Much of the research on the regulation of animal growth has focused on the production and release of GH (17). In most species studied, including mammals and fish, pituitary GH release is under dual antagonistic control from the hypothalamus as well as from systemically borne hormones. Major inhibitors of GH release are somatostatins (SSs), a structurally and functionally diverse family of peptides derived from multiple genes with widespread anatomical distribution (25, 29). Interestingly, not all SSs inhibit GH release. For example, in fish, SS-14 and the N-terminally extended SS-28 peptide inhibit GH secretion from isolated pituitary fragments, but salmonid SS-25, which contains [Tyr7,Gly10]-SS-14 at its C-terminus does not (36). GH releasing hormone, dopamine, gonadotropin releasing hormone, thyrotropin releasing hormone, cholecystokinin, and ghrelin, on the other hand, stimulate GH release (6, 17). Given the possibility of both a direct and indirect mode of action of GH and the complexity of the GH-IGF-1 axis, it is becoming increasingly clear that the regulation of growth may occur at many levels outside the pituitary.
The GHR, which mediates the effects of GH in tissues, has received increasing attention as a target of growth regulation (28). GHRs have been characterized from several species of mammals, birds, and fish, as well as from turtle and frog; salmonid fish appear to have two distinct forms of GHR (37). As members of the class I cytokine receptor superfamily, GHRs contain a single transmembrane domain and conserved intracellular regions involved in signal transduction through the JAK-STAT and other pathways (21). In both mammals and fish, high affinity GH binding and GHRs are widespread in numerous tissues (e.g., brain, gill, gut, muscle, pancreas, spleen), but among peripheral tissues, they are most highly concentrated in the membranes of the liver (14, 37). Given the importance of GHR to the growth-promoting and other actions of GH, a heightened understanding of the factors regulating GHR expression is warranted.
In this study, rainbow trout were used as a model system to investigate the role(s) of SSs in regulating hepatic GH sensitivity. Specifically, we tested the hypothesis that SSs regulate the internalization and synthesis of GHRs in liver. The rationale for this work stemmed from studies in mammals and fish that revealed periods of reduced growth, resulting from food deprivation (mammals and fish) and premature transfer to seawater (salmonid fish), can occur in the face of normal or elevated GH and in association with elevated plasma SS and reduced hepatic GH binding (4, 15, 33, 36).
Experimental animals and conditions.
Juvenile rainbow trout of both sexes were obtained from Dakota Trout Ranch near Carrington, ND, and transported to North Dakota State University, where they were maintained in 800 liter circular tanks supplied with recirculated (10% replacement volume per day) dechlorinated municipal water at 14°C under a 12:12-h light-dark photoperiod. Fish were fed to satiation twice daily with AquaMax Grower (PMI Nutrition International, Brentwood, MO), except 24–36 h before experimental manipulations. Animals were acclimated to laboratory conditions for at least 4 wk prior to experiments. All procedures were performed in accordance with the Guide for the Care and Use of Laboratory Animals (National Research Council, Washington, DC) and were approved by the North Dakota State University Institutional Animal Care and Use Committee.
For in vivo experiments, fish were anesthetized in 0.5% (vol/vol) 2-phenoxyethanol, weighed (initial weight was 58.3 ± 1.3 g), individually tagged, and implanted intraperitoneally with Alzet miniosmotic pumps (Alza, Palo Alto, CA) as previously described (35). The minipumps contained either 0.75% (wt/vol) NaCl (control) or 4 × 10−4 M SS-14-I (treated) and had a flow rate established to be 0.135 μl/h; this would deliver ∼5.8 × 10−11 mol SS-14-I/h. SS-14-I, obtained from Sigma, St. Louis, MO (cat. no. S-9129), has the sequence first described by Brazeau et al. (5), which is highly conserved among vertebrates and which is derived from the precursor designated preprosomatostatin I (PPSS I)(28). After 15 days, the fish were anesthetized as before, and the plasma and livers were removed and frozen at −90°C for later analyses. SS-14-I was determined by a specific radioimmunoassay previously validated for use with trout plasma (11).
In vitro experiments were conducted on isolated hepatocytes. Fish were anesthetized with 0.5% (vol/vol) 2-phenoxyethanol, and the hepatocytes were isolated by the in situ perfusion method described by Mommsen et al. (24). The viability of the cells was assessed by trypan blue dye exclusion and ranged between 93 and 97% for all experiments. The isolated cells (∼1 × 106 cells/ml) were allowed to recover in incubation medium [in mM: 136.9 NaCl, 5.4 KCl, 0.81 MgSO4, 0.44 KH2PO4, 0.33 Na2HPO4, 10 HEPES, and 5 NaHCO3, 1.5 CaCl2, pH 7.6, with 2% (wt/vol) defatted BSA, 3 mM glucose, 2 ml GIBCO MEM amino acid mix (50×)/100 ml, and 1 ml GIBCO nonessential amino acid mix (100×)/100 ml] for 3 h at 14°C with gyratory shaking (100 rpm) under 100% O2. After the recovery period, hepatocytes were collected by centrifugation (100 g for 3 min at 14°C) and resuspended at a final concentration of 1 × 106 cells/ml in incubation medium. Cells were incubated in medium alone (control) or in medium containing various concentrations of SSs [PPSS I-derived peptides: SS-14-I and SS-28-I (cat. no. S 6135; Sigma); PPSS II-derived peptides, containing [Tyr7,Gly10]-SS-14 at their COOH terminus, custom synthesized by Dr. D. Smith: SS-14-II, SS-25-II, and SS-28-II] for various times under the same conditions as those used for recovery. Incubations were stopped by centrifugation (100 g for 3 min at 14°C), and the cells were rinsed with incubation medium, and surface-bound hormone was removed by a 3-min treatment with acidified (pH 4.5) incubation medium. After treatment, the cells were washed twice with incubation medium and either incubated further for analysis (e.g., whole-cell GH binding/processing) or subjected to RNA extraction.
GH binding analyses.
Binding of 125I-labeled trout GH (tGH) to hepatic membranes prepared from frozen liver was performed as described previously (14). Whole cell binding assays were conducted for 3 h at 14°C by incubating 400 μl of hepatocyte suspension (∼4 × 105 cells) and 50 μl of 125I-labeled tGH [∼40,000 cpm diluted with 0.25 mM Tris buffer] with either 50 μl of incubation medium (for total binding) or 50 μl of unlabeled tGH (diluted in incubation medium), in concentrations from 10 to 1,000 nM. The incubation was stopped by centrifugation at 1,000 g for 5 min at 14°C. Unbound hormone was removed by aspiration, and the cells were washed three times with cold incubation medium. The tips of the tubes containing the pellets were cut off, placed individually in 12 × 75-mm tubes, and counted in a Beckmann 5500 gamma counter. The percentage of specific binding was calculated as B/T × 100, where B is the difference between total binding and nonspecific binding (binding observed in the presence of 1,000 nM tGH), and T is the amount of radioactivity originally added to each tube.
Processing of bound tGH and internalization was evaluated according to Simón et al. (31), except 125I-labeled tGH was added to hepatocytes suspensions (∼35,000 cpm per 106 cells) and incubated at 14°C under 100% O2 with gyratory shaking (100 rpm).
tGH [equivalent to GH 1, the most predominant form of GH in trout and other salmonids (22) (GroPep; Adelaide, Australia)] was iodinated for use in binding assays by a procedure modified from Gray et al. (14). Five micrograms of tGH were labeled using lactoperoxidase and 200 μCi of 125I-labeled NaI (MP Biomedicals, Irvine, CA); 125I-labeled tGH was purified by gel filtration chromatography on Sephadex G-75 (0.6 × 12 cm).
GHR mRNA expression.
Total RNA was extracted using TRI-Reagent as specified by the manufacturer (Molecular Research Center, Cincinnati, OH). Isolated RNA was dissolved in 75 μl RNase-free deionized water. Total RNA was routinely quantified by ultraviolet (A260) spectrophotometry and diluted to 100 ng/μl in RNase-free deionized water. RNA quality was examined with the Agilent 2100 Bioanalyzer (Santa Clara, CA). From 200 ng total RNA, endogenous poly(A)+ RNA was reverse transcribed according to the manufacturer's instructions in a 10-μl reaction using a TaqMan reverse transcription reagent kit containing MultiScribe reverse transcriptase and oligo(dT)17 as a primer (Applied Biosystems, Foster City, CA). mRNA levels of the two tGHR forms were determined by real-time RT-PCR using TaqMan chemistry and an ABI PRISM 7000 sequence detection system (Applied Biosystems) as described previously (37). Briefly, real-time reactions were carried out for samples, standards, and no-template controls in a 10-μl volume, containing 1 μl cDNA from the reverse transcription reactions, 5 μl TaqMan Universal PCR Master Mix, and 1 μl of each forward primer, reverse primer, and probe at concentrations optimized for each RNA species. Cycling parameters were as follows: 95°C for 10 min and 45 cycles of 95°C for 15 s, and 60°C for 1 min. Sample copy number was calculated from the threshold cycle number (CT) and relating CT to a gene-specific standard curve, followed by normalization to β-actin. No difference (P > 0.05) was observed in β-actin expression among the various treatment groups.
Analysis of GHR mRNA transcription and stability.
The transcription of GHR mRNAs was assessed by quantification of nascent transcripts in nuclei isolated from rainbow trout hepatocytes (∼2 × 107 cells/sample). Following isolation (16), the nuclei were stored at −90°C in glycerol storage buffer [in mM: 10 Tris·HCl, 5 MgCl2, 0.1 EDTA, 40% (vol/vol) glycerol, pH 8.3]. The run-on assay was conducted as described previously (27). Briefly, nuclei were thawed and mixed with run-on buffer reagents [in mM: 25 Tris·HCl, pH 8.0, 12.5 MgCl2, 750 KCl, 1.25 each of ATP, GTP, and CTP, and 0.6% (wt/vol) Sarkosyl, 100 μCi [32α-P]UTP (Amersham Life Science; Arlington Heights, IL)] in 2-ml microcentrifuge tubes. The samples were incubated for 60 min (30°C), DNase I was then added, and the incubation continued for 15 min. RNA was immediately extracted (27), and labeled nuclear transcripts were hybridized to specific GHR cDNA probes, previously immobilized on nylon membranes by use of a slot-blot apparatus. The probes were based on the sequences of rainbow trout GHRs reported previously (37); bases 1855–2667 for GHR 1 and bases 1835–2725 for GHR 2. Blots were quantified by phosphor imaging, as described previously (12).
The stability of GHR mRNAs also was determined in isolated hepatocytes (2 × 106 cells/ml) essentially as described previously (12). Cells were preincubated with actinomycin D (5 μg/ml) for 30 min, then collected by centrifugation, washed once, and incubated in the presence or absence of 100 ng/ml SS-14-I. At various times up to 24 h, the cells were collected by centrifugation, and RNA was extracted and analyzed by quantitative real-time PCR as described above.
Quantitative data are presented as means ± SE. Ligand binding characteristics were calculated with the SigmaPlot Ligand Binding Module (SPSS, Chicago, IL) using the one-site model, which gave the best fit for the data and was used for interpretation of results. Statistical differences were estimated by Student's t-test or one-way ANOVA, as appropriate; multiple comparisons among means were evaluated by Duncan's test. A probability level of 0.05 was used to indicate significance. All statistics were performed using SigmaStat (SPSS, Chicago, IL).
Effects of SS on hepatic GH binding and internalization.
SS implantation significantly elevated plasma SS-14-I levels (7.4 ± 0.2 ng/ml) over those observed in saline-implanted fish (5.3 ± 0.3 ng/ml). In vivo exposure of fish to SS-14-I significantly decreased hepatic GH binding capacity from 331.5 ± 34.2 fmol/mg protein in control fish to 58.4 ± 6.4 fmol/mg protein in SS-14-I-treated fish (Fig. 1).
To determine whether or not the effects of SS-14-I were direct, GH binding to isolated hepatocytes was assessed. Scatchard analysis yielded a linear plot, suggesting a single class of high-affinity binding site for GH. (Fig. 2, inset). Treatment of hepatocytes with SS-14-I significantly reduced GH binding capacity from 1194 ± 104 fmol/cell to 610 ± 58 fmol/cell (Fig. 2). Treatment with SS-25-II, a predominant form of SS in the pancreas of teleost fish (11), similarly reduced GH binding capacity in hepatocytes. Neither SS-14-I nor SS-25-II affected GH binding affinity in isolated hepatocytes (Fig. 2, inset).
To determine whether or not the decrease in GH binding capacity results from internalization, the effects of SS-14-I on processing of 125I-labeled tGH by isolated hepatocytes was investigated. 125I-labeled tGH was internalized in a time-dependent manner, with appreciable internalization evident within 0.5 h and becoming saturated within 6–9 h (Fig. 3). Internalization of 125I-labeled tGH into control- and SS-14-treated hepatocytes was similar up to 1 h. After 2 h, however, SS-14-treated hepatocytes displayed significantly greater internalization of 125I-labeled tGH than control-treated cells. By 6 h, SS-14-I increased internalization of 125I-labeled tGH 58% over that observed in controls.
Effects of SS on steady-state levels of GHR mRNAs.
To determine whether or not the decreases in GH binding capacity also result from decreased GHR synthesis, the effects of SSs on steady-state levels of GHR mRNAs in isolated hepatocytes was evaluated. Rainbow trout express two distinct mRNAs encoding for separate GHRs, GHR 1 and GHR 2. SS-14-I directly inhibited steady-state levels of GHR 1 and GHR 2 mRNA in isolated hepatocytes in a dose-dependent manner (Fig. 4). The dose responsiveness of GHR 1 and GHR 2 mRNA expression to SS-14-I was similar; maximum inhibition resulted in reductions in the steady-state mRNA levels of 84% and 88%, respectively, at 1,000 ng/ml.
The time course of SS-14-I action on GHR expression is shown in Fig. 5. The effects of SS-14-I were rapid; steady-state levels of GHR mRNAs declined sharply within 3 h and remained depressed over the course of the experiment. The temporal responsiveness to SS-14-I on the expression of GHR 1 and GHR 2 mRNAs was similar. After 24 h, steady-state levels of GHR 1 mRNA were reduced by 85% and those of GHR 2 were reduced by 88% compared with controls.
The influence of various SS isoforms on GHR expression is shown in Fig. 6. While SS-14-I inhibited the expression of both GHR 1 and GHR 2 mRNAs, the NH2 terminally extended peptide, SS-28-I, had no significant effect on GHR mRNA levels. The [Tyr7,Gly10]-substituted SSs, SS-25-II and SS-28-II, inhibited the expression of GHR mRNAs, whereas, SS-14-II promoted GHR mRNA expression.
Effects of SS on GHR mRNA transcription and stability.
To determine whether SS-induced changes in the steady-state levels of GHR mRNAs resulted from alterations in the rates of GHR transcription, nascent mRNA transcripts were evaluated by nuclear run-on assays. SS-14-I was found to inhibit both basal and stimulated GHR mRNA transcription. At a concentration of 100 ng/ml, SS-14-I significantly decreased the rates of transcription of GHR 1 and GHR 2 mRNAs to 32.4% and 56.5% of control rates, respectively, after 6 h of incubation (Fig. 7). tGH significantly increased the rates of GHR transcription after 6 h; GHR 1 increased to 315% of control rates, whereas GHR 2 increased to 255% of control rates. SS-14-I inhibited GH-stimulated GHR transcription to rates similar to those observed in control-treated cells.
To determine whether SS-induced changes in the steady-state levels of GHR mRNAs resulted from alterations in the rates of GHR mRNA degradation, the stability of mRNAs was evaluated by decay curves. The half-life of each GHR mRNA species in rainbow trout hepatocytes was estimated to be ∼9 h. The half-lives of GHR 1 and GHR 2 mRNAs in SS-14-I-treated cells were not significantly different from those in control cells (data not shown), indicating that the stability of GHR mRNAs was unaffected by the presence of SS-14-I.
Organismal growth integrates a vast array of biological processes and is coordinated by numerous chemical mediators. A central element of organismal growth control is the GH-IGF-1 axis; however, various other factors, including thyroid hormones, cortisol, insulin, IGFBPs, and local growth factors, also are important (6, 10, 17). SSs previously were shown to regulate growth by inhibiting GH release (36). The results of this study indicate that SSs also have extrapituitary actions on growth and confirm our hypothesis that SSs regulate hepatic GH sensitivity by altering GH binding capacity and expression of GHR mRNAs. The effects of SSs on GH sensitivity in rainbow trout occur at physiological levels, which range in plasma concentration from 0.2 ng/ml in fed animals to >10 ng/ml in fasted animals (11).
An important factor in determining the responsiveness of cells to hormones that act through membrane-bound receptors is the presence of receptors on the cell membrane. The present findings indicate that SSs reduced hepatic binding capacity in vivo and in vitro. The decrease in GH binding capacity on isolated hepatocytes within 6 h revealed that the actions of SS-14-I and SS-25-II were both rapid and direct. Notably, only a single class of GH binding site was observed, consistent with previous studies in salmonids (9, 15), despite the presence of two distinct GHR mRNAs reported in trout and other fish (37). Although there are some differences in the extracellular domains of the GHRs of fish, the ligand binding characteristics of the different forms are not known. The availability of GHR at the plasma membrane is linked to the intracellular pool of GHRs. The intracellular pool may contain newly synthesized GHRs, and therefore, the expression of GHR mRNAs, mRNA stability, and rate of translation may play a role in modulation of receptor availability. Posttranslational modulation of the intracellular pool, in terms of recruitment of receptors to the plasma membrane, degradation in the lysosomes, or internalization of surface receptors would provide additional means of regulating GH sensitivity and action in target cells.
Internalization results from the endocytosis of the GH-receptor complex and represents a mechanism for regulating GH sensitivity of target cells. Interestingly, unlike many other receptors, internalized GHRs are not recycled back to the surface but are degraded in the lysosomes and are only replaced by newly synthesized receptors (34). This study revealed that SS-14-I significantly increased the internalization of GH. Such pronounced effects are consistent with downregulation of cell surface GHRs and contributed, at least in part, to the observed reduction of GH binding in the presence of SSs in vivo and in vitro. Two mechanisms for GH internalization have been observed in mammals: a ubiquitin-dependent system appears to modulate internalization of receptor dimers in the presence and absence of ligand, while a ubiquitin-independent system appears to be involved with endocytosis of GHRs not part of a dimer (34). The mechanism(s) by which SS-14 influences GHR internalization and the significance of the presence of two GHRs are not known. King et al. (20) showed that although dexamethasone and phorbol ester phorbol myristate acetate both decrease cellular GH binding, independent mechanisms were employed that involved interactions with different regions of the GHR.
SSs regulate the expression of GHRs. The present findings indicate that SS-14-I rapidly and directly inhibited steady-state levels of GHR mRNAs. The responsiveness of GHR 1 and GHR 2 expression in liver cells to SS-14-I appeared to be the similar, both in terms of potency and in terms of the temporal development of the response. Two additional isoforms of SS, SS-25-II and SS-28-II, both containing [Tyr7,Gly10]-SS-14 at their COOH terminus, also inhibited the expression of GHR mRNAs in liver cells. Two other SS isoforms, SS-28-I (an NH2-terminal extension of SS-14-I) and SS-14-II ([Tyr7,Gly10]-SS-14), either stimulated or had no effect on GHR expression. Functional differences among SS isoforms have been reported previously. Interestingly, for example, several isoforms of SS (e.g., SS-25-II, catfish SS-22) do not possess GH-inhibiting bioactivity (36). The bases of the structure-function differences may stem from any combination of the following: 1) differences in ligand-receptor interaction, 2) the existence of multiple SS receptors that display tissue-specific patterns of expression and ligand-specific binding, and 3) differential linkage of SS receptor subtypes to various cellular effector systems (29). Notably, rainbow trout do possess multiple forms of SS receptors (SSTRs) that display ligand-selective binding characteristics (13). Regardless, the regulation of GHR synthesis will remain an important aspect of growth regulation. The regulation of GHR expression also has been shown in mammals under certain conditions and by selected hormones, including pregnancy, nutritional state, and steroid hormones, such as estrogen and testosterone (2, 18, 21, 28).
The inhibitory effects of SS-14-I on steady-state levels of trout GHR mRNAs resulted from decreased rates of transcription and not from altered mRNA degradation. This conclusion is supported by several observations. First, nuclear run-on assays indicated that the number of nascent transcripts was significantly lower in nuclei isolated from SS-14-I-treated hepatocytes compared with control cells. Second, SS-14-I prevented the increase in the number of nascent transcripts stimulated by tGH. GH has been shown to stimulate GHR transcription in a variety of mammalian and cell line systems (28). Last, the inability of SS-14-I to affect GHR mRNA degradation supports the notion that SS-inhibited expression of GHR mRNAs does not involve alterations in GHR mRNA stability. The mechanism(s) by which SS alters the transcription of GHR genes remains to be determined. It is possible that one or more of the signaling pathways known to link to SSTRs (e.g., MAPK) (20) may interact with regulatory elements in the promoter region of the GHR gene (e.g., C/EBP-β) (28). Given that internalization of SSTRs affects GH expression (26) and that SSTRs previously have been shown to be internalized in trout liver (24), it also is possible that SSTR internalization reduces GHR expression.
The present findings of the effects of SS on the regulation GH sensitivity extend our knowledge of the role of GH in growth regulation. Growth of rainbow trout was inhibited by implantation with SS-14-I for 15 days; such growth retardation was accompanied by reduced plasma GH and was consistent with previous reports on SS effects on GH production and release (35, 36). Interestingly, SS-induced growth retardation also was accompanied by reduced plasma IGF-1 (35). While reduced IGF-1 production and release can be explained by reduced availability of GH (36), the present findings suggest that SSs also have a direct extrapituitary action on regulating GH sensitivity. This latter notion is supported by studies in mammals and fish in which reduced growth was accompanied by elevated plasma SS and reduced hepatic GH binding in the face of normal or elevated GH (4, 15, 33, 36). In addition, reduced growth was observed in mice lacking GHR (8). The model that emerges, therefore, suggests that SSs may regulate growth and the GH-IGF-1 axis at several levels: at the level of the pituitary, involving the inhibition of GH, and at extrapituitary levels, involving regulation of GH sensitivity. Extrapituitary or local actions by SS would add a fine control to growth regulation, possibly enabling the “uncoupling” of the growth-promoting actions of GH (e.g., stimulation of IGF-1 synthesis) (6, 10) from other actions (e.g., lipolytic) (30) that would be adaptive under certain physiological conditions (e.g., nutrient limiting, sexual maturation).
This research was supported by National Science Foundation Grant IOB-0444860 (to M. A. Sheridan). N. M. Very was supported by a fellowship from the North Dakota Experimental Program to Stimulate Competitive Research.
We thank Leslie Alexander, Whitney Fleck, Jun-Yang Gong, Jeffrey Kittilson, Felicia Lamb, Greg Melroe, Laura Nelson, Lindsey Norbeck, Jayson Poppinga, Bart Slagter, and Nathan Weiderholt for their assistance with conducting these studies and preparation of the manuscript. We also thank Dr. David Smith, Creighton University, Omaha, NE, for the custom synthesis of selected somatostatin isoforms.
Present address of N. M. Very: Institute for Molecular Virology, University of Minnesota, Minneapolis, MN 55455.
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- Copyright © 2007 the American Physiological Society