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Technical Briefs |
1 Department of Biochemistry, National Cardiovascular Center Research Institute, and2 Department of Internal Medicine, National Cardiovascular Center, Osaka 565-8565, Japan;3 Translational Research Center, Kyoto University Hospital, Kyoto 606-8507, Japan
aaddress correspondence to this author at: Department of Biochemistry, National Cardiovascular Center Research Institute, National Cardiovascular Center, Osaka 565-8565, Japan; fax 81-6-6835-5402, e-mail kangawa{at}ri.ncvc.go.jp
Ghrelin is an acylated peptide with growth-hormone-releasing activity (1). It was first isolated from rat and human stomach during the search for an endogenous ligand to the "orphan" G-protein-coupled receptor, growth hormone secretagogue receptor (2). The peptide contains 28 amino acids, and n-octanoylation of the Ser-3 hydroxyl group is necessary for biological activity. Most studies have focused on the somatotropic and orexigenic roles of ghrelin; therefore, little is known about the kinetics of this peptide. Because the ester bond is both chemically and enzymatically unstable, elimination of the octanoyl modification of ghrelin can occur during storage, handling, and/or dissolution in culture medium (3). Because of increased interest in ghrelin measurements, a standardized method of sample collection is required.
In the present study, which focused on the active form of ghrelin, we investigated the effects of anticoagulants and storage conditions on ghrelin stability. To distinguish the active form of ghrelin, we established two ghrelin-specific RIAs; N-RIA recognizes the N-terminal, octanoyl-modified portion of the peptide, whereas C-RIA recognizes the C-terminal portion. Thus, the value determined by N-RIA specifically measures active ghrelin, whereas the value determined by C-RIA gives the total ghrelin immunoreactivity, including both active and desacyl ghrelin (4)(5)(6). The minimum detectable quantities in the N- and C-RIAs were 5.0 and 50 pmol/L, respectively. The respective intra- and interassay CV were 3% and 6% for the N-RIA and 6% and 9% for the C-RIA (n = 8 assays). Data are reported as the mean (SD). Comparisons of the time course of ghrelin concentrations between subgroups were made by two-way ANOVA for repeated measures, followed by the Scheffé test. P <0.05 was considered statistically significant.
All blood samples were taken from three healthy male volunteers who gave written informed consent. Blood was taken from the forearm vein and immediately divided into tubes for serum and plasma preparation using (a) disodium EDTA (1 g/L) with aprotinin (500 000 kIU/L), (b) disodium EDTA alone, (c) heparin sodium, or (d) no anticoagulant. Synthetic human ghrelin was added to each blood sample at a final concentration of 40 µg/L; each sample was then sequentially divided into two aliquots for incubation at either 4 or 37 °C. After incubation for 0, 30, and 60 min, blood samples were centrifuged, diluted 1:200 in RIA buffer, and subjected to ghrelin-specific RIAs. A comparison of the effects of different anticoagulants on the detected ghrelin concentrations is shown in Table 1A
. Although the serum and three different plasma samples tested gave comparable results for total ghrelin by C-RIA, the N-RIA gave ghrelin concentrations that were significantly decreased at 37 °C. When the ghrelin was measured by N-RIA, serum samples were highly affected by such treatment; samples stored for 60 min at 37 °C lost 
35% of the ghrelin compared with the basal values at 0 min (P <0.05). The ghrelin concentrations in samples containing heparin as an anticoagulant were also significantly decreased (P <0.05). When EDTA–aprotinin was used as the anticoagulant for plasma treatment, the decreases in ghrelin stability were smaller than for other procedures. Storage at 4 °C also improved ghrelin stability.
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To explore optimum storage conditions, we examined the effect of plasma pH on ghrelin stability. The EDTA–aprotinin-treated plasma (n = 3) was divided into five samples; the pH was then adjusted to 3, 4, 5, 6, or 7.4 with 1 mol/L HCl. Synthetic human ghrelin was then added to each sample aliquot at a final concentration of 75 µg/L. Each of the five plasma aliquots was then subdivided into two, with one stored at 4 °C and the other stored at 37 °C. The effects of acidification on ghrelin stability in plasma are summarized in Table 1B
. When stored at 37 °C, ghrelin concentrations measured by N-RIA gradually decreased at all pH values tested. However, ghrelin was most stable in highly acidified plasma samples (pH 3–4). At pH 3–5 and a storage temperature of 4 °C, the stability of ghrelin in plasma did not change significantly over a 6-h period. By C-RIA, ghrelin concentrations remained stable across the different pH and storage temperature conditions.
We then evaluated the effects of repeated freezing and thawing on the stability of ghrelin. EDTA–aprotinin-treated plasma samples were divided into two pH groups; one was acidified to pH 4, whereas the other was not acidified (pH 7.4). After the addition of synthetic human ghrelin (75 µg/L), we subjected the samples to four freeze–thaw cycles. Repeated freezing and thawing also influenced ghrelin stability (Table 1C
). As in the N-RIA, ghrelin concentrations in untreated plasma samples decreased significantly with each successive freeze–thaw cycle, whereas the ghrelin remained relatively stable after acidification. Ghrelin concentrations by C-RIA were unchanged despite repeated freeze–thaw treatments in both acidified and untreated plasma samples.
As well as differences in assay methodologies, differences in sample handling, such as the method of storage, effects of anticoagulants, or previous freezing and thawing of the samples, could influence the reported values (7)(8)(9)(10). Instability of peptides and proteins can be divided into two forms: chemical and physical instability (11)(12). The chemical degradation of peptides is influenced by the pH of the aqueous solution; human parathyroid hormone and luteinizing-hormone-releasing hormone derivatives are examples (13)(14)(15). We demonstrated that in whole blood and plasma, ghrelin is unstable. The degradation of octanoylated ghrelin was shown to be attributable to hydrolysis to desacyl ghrelin (see Fig. 1
in the Data Supplement that accompanies the online version of this Technical Brief at http://www.clinchem.org/content/vol50/issue6/). Acidification is a simple, reliable procedure that protected against degradation of the acylated modification and dramatically improved stability at pH 4. On the other hand, the stability of the octanoyl modification of ghrelin was markedly decreased in strongly acidic (below pH 2), neutral, and alkaline solutions (data not shown).
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We evaluated the effectiveness of measuring active ghrelin compared with total ghrelin in response to oral glucose tolerance tests (OGTTs). Four healthy male volunteers (age range, 28–35 years; body mass index, 21.5–23.7 kg/m2) were examined on 2 separate days (100 g of glucose administered on 1 day, and 50 g of glucose administered on the other day) at least 2 weeks apart in a randomized, crossover study. After the volunteers fasted overnight, 50 or 100 g of glucose was administered orally between 0930–1000. Blood samples were obtained at 0, 1, 2, 3, and 4 h after glucose ingestion. To each plasma sample was added 1 mol/L HCl (10% of plasma volume), which acidified the sample to pH 4; samples were then treated with Sep-Pak C18 cartridges for ghrelin RIAs. After glucose ingestion, the mean plasma ghrelin concentrations as determined by N-RIA and C-RIA decreased to a nadir at 1 h (Fig. 1
). At this point, 60.3% and 73.0% of the basal concentration was detected by the N-RIA and C-RIA, respectively, after the 100-g OGTT, and 64.2% and 78.7% of the basal concentration was detected after the 50-g OGTT. Plasma ghrelin values increased thereafter, although plasma ghrelin concentrations measured by the C-RIA were significantly lower for up to 2 h after the 100-g glucose load. The N-RIA for ghrelin could detect differences in the changes in ghrelin concentrations between the 50-g and 100-g OGTTs at 3 h. The ghrelin values observed with the C-RIA exhibited changes similar those in the N-RIA, but the changes were small and delayed. These effects may be attributable to the differential rates of metabolic turnover for octanoylated and desacylated ghrelin in circulating blood (see Fig. 2 in the online Date Supplement).
The results for the plasma ghrelin response to the OGTTs show that measuring the concentration of active ghrelin is useful for studying plasma ghrelin changes over short time periods. Plasma concentrations of active ghrelin changed more rapidly and dynamically than those of total ghrelin immunoreactivity. Fasting led to markedly increased plasma ghrelin values as measured by N-RIA, and the values decreased in a clearer dose-dependent manner in rats after glucose injection compared with those measured by C-RIA (16). The proportion of active ghrelin in plasma was 2–5% of total ghrelin in rodents. In this study, the quantity of active ghrelin was 10% of the total ghrelin in human plasma (data not shown). These findings imply that inactive desacyl ghrelin circulates in the bloodstream at much higher concentrations than active ghrelin. Similar to previous studies in which ghrelin concentrations were measured by C-RIA (17), desacyl ghrelin is relatively stable, and its stability is not altered by different storage conditions. An analogous situation has been reported for the activity of pancreatic beta cells, which secrete insulin and C-peptide in a 1:1 molar ratio. However, the half-life of C-peptide is much longer than that of insulin, leaving more C-peptide available in the circulation for quantification (18)(19). Measurement of C-peptide provides an assessment of ß-cell secretory activity. Similarly, desacyl ghrelin concentrations may serve as an indicator of ghrelin secretory function (20).
To acquire accurate data on ghrelin concentrations, this study recommends a standard procedure for the collection of blood samples: (a) the collection of blood samples with EDTA–aprotinin is preferred; (b) blood samples should be chilled and centrifuged as soon as possible, at least within 30 min after collection; and (c) because acidification is the best method for the preservation of plasma ghrelin, 1 mol/L HCl (10% of sample volume) can be added to the plasma sample for adjustment to pH 4.
Acknowledgments
We thank H. Mondo and M. Miyazaki for technical assistance. This work was supported by grants from the Ministry of Education, Science, Sports and Culture of Japan; the Ministry of Health, Labor and Welfare of Japan; the Promotion of Fundamental Studies in Health Science from the Organization for Pharmaceutical Safety and Research of Japan; and the Takeda Science Foundation.
References
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| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
) and 100-g (•) OGTTs in four healthy individuals.