Full-length cDNA of the mouse GH secretagogue receptor (GHS-R1a) (GenBank accession number NM_177330.4; residues 201–1377) was obtained via RT-PCR using mouse hypothalamus cDNA as the template. The sense and antisense primers were 5′-caccctcctcaggggaccagattt-3′ and 5′-aatgagcgatgacggagagat-3,’ respectively. The amplified cDNA was ligated into the pcDNA3.2/V5/GW/D-TOPO Expression Kit (Invitrogen, Tokyo, Japan). The expression vector, mouse GHS-R-pcDNA3.2, was transfected into Chinese hamster ovary (CHO) cells. Stably expressing cells were selected using 1 mg/ml G418 (Nacalai Tesque, Kyoto, Japan). The selected cell line, CHO-mouse GHS-R1a-line 24–5, showed the highest expression of mouse GHS-R1a mRNA. Cells were cultured in a humidified environment with 95% air and 5% CO₂. Changes in intracellular Ca²⁺ concentrations ([Ca²⁺]_i) were measured using a FlexStation 3 fluorometric imaging plate reader (Molecular Devices, CA, USA). CHO-mouse GHS-R1a cells (3 × 10⁴ cells) were plated in 96-well black-walled microplates (Corning, NY, USA) for 20 h before each assay. The cells were incubated with 100 µL of Calcium 4 assay kit (Molecular Devices) for 1 h, and then 50 µL of each sample was added to the CHO-mouse GHS-R1a cells. The maximum [Ca²⁺]_i change was determined as the response.

Bovine (Japanese black breed, 6 months old, male) and porcine (three-crossbred pig <LWD>, one month old, male) stomachs were provided by the Department of Veterinary Anatomy at the University of Miyazaki (Miyazaki, Japan). Equine (thoroughbred, 7 years old, male) stomach samples were obtained from the Equine Research Institute of the Japan Racing Association (Tochigi, Japan). All procedures were performed in accordance with the guidelines of the Japanese Physiological Society for animal care, and the experiments were approved by the University of Miyazaki Animal Experiment Committee (2015–006). Written informed consent was obtained from the owners for the participation of their animals in this study.

Frozen tissues (20 g, 25 g, and 100 g of bovine, porcine, and equine stomachs, respectively) were pulverized and boiled for 10 min in 10 volumes of water to inactivate intrinsic proteases, as described previously (32). The samples were then chilled on ice and adjusted to 1 M acetic acid (AcOH) by adding glacial AcOH. Boiled stomach tissue was homogenized using a Polytron mixer (Kinematica Inc., Lucerne, Switzerland). The crude acid extracts were then centrifuged for 30 min at 10,000 xg. The supernatant was diluted with an equal volume of distilled water and concentrated to about 1/3 amount using an evaporator. The residual concentrate was subjected to acetone precipitation at a 66% acetone concentration. After removing the precipitate, the acetone supernatant was evaporated. It was loaded onto a 10 g Sep-Pak C18 cartridge (Waters), which was pre-equilibrated with 0.1% trifluoroacetic acid (TFA). The cartridge was washed with a 10% acetonitrile (ACN)/0.1% TFA solution and eluted with 100 mL of 60% ACN/0.1% TFA. The eluate was evaporated and lyophilized. The residual materials were redissolved in 1 M AcOH and then adsorbed on an SP-Sephadex C-25 (H + form) chromatography column (Amersham Pharmacia Biotech, Buckinghamshire, UK) pre-equilibrated with 1 M AcOH. Successive elutions with 1 M AcOH, 2 M pyridine, and 2 M pyridine-AcOH (pH 5.0) yielded three fractions: SP-I, SP-II, and SP-III. The lyophilized SP-III fraction, which contained strong basic peptides and ghrelin activity, was separated via CM-ion-exchange HPLC on a TSK CM-2SW column (4.6 x 250 mm; Tosoh) at pH 6.5. The active fractions were further purified using the same column at pH 4.8. The active fractions were diluted in equal volumes of 0.1% TFA and subjected to Sep-Pak Light C18 cartridge (Waters) purification to remove excess salt. The sample was eluted with 60% ACN/0.1% TFA, evaporated ACN, and then separated via reverse-phase (RP)-HPLC using a Symmetry 300 TM C18 column (3.9 x150 mm, Waters) at a flow rate of 1 mL/min on a linear gradient from 10% to 60% ACN/0.1% TFA for 80 min. The eluate was collected in 0.5-mL fractions. An aliquot of each fraction was assayed for ghrelin activity using FlexStation 3. The active fractions were further purified via RP-HPLC using a diphenyl column, 219TP5215 (2.1 x150 mm; Vydac, Hesperia, CA) for 80 min on a linear gradient from 10% to 60% ACN/0.1% TFA at a flow rate of 0.2 mL/min. Each absorption peak was collected, and an aliquot of each fraction was assayed for activity using FlexStation 3. Some peaks were further purified on a Chemcosorb 3-ODS-H column (2.1 x 75 mm, Chemco Scientific, Osaka, Japan) for 80 min. Approximately 20 pmol of the final purified peptide from the main peak was analyzed using a protein sequencer (PPSQ-33A; Shimazu, Kyoto, Japan). One picomole was used for molecular weight determination using MALDI-TOF mass spectrometry (Autoflex III, Bruker).

Ghrelin (Human) was purchased from Peptide Institute, Inc. (Osaka, Japan).

The interaction between ghrelin and CHO-mouse GHS-R1a-line 24–5 was examined using synthetic peptides. Human ghrelin induced dose-dependent, robust increases in [Ca²⁺]_i in CHO-mouse GHS-R1a-line 24–5, with half-maximal response concentrations (EC₅₀) of 1.15 × 10–⁹ M ( Figure 1 ). Ghrelin did not induce a response in CHO cells transfected with vector alone (data not shown).

Three groups with ghrelin activity were identified via CM ion-exchange HPLC (pH 6.5) of the SP-III fraction ( Figure 2A ). Each active group was purified via CM ion-exchange HPLC (pH 4.8) and three separate rounds of RP-HPLC. The final purification of the group B fractions ( Figure 2A ), which gave the highest ghrelin yield, is shown in Figure 2B . Four active peptides were isolated from the three groups using CM-HPLC ( Table 1 ). The amino acid sequence of the group B peptide showing the highest activity was determined to be GSXFLSPEHQKLQRKEAKKPSGRLKPR (X was unidentified by the protein sequencer because of an acyl modification). We predicted that X was serine based on the NCBI accession no. NM_174067, and the isolated peptide was identified as a ghrelin peptide based on its sequence homology with other ghrelins. The isolated peptides were analyzed using MALDI-TOF MS to determine the molecular weight of the bovine ghrelins. Table 1 shows the measured molecular masses, isolation yields, and deduced molecular forms of the isolated peptides. In addition to peptide and genetic information, molecular masses helped in the prediction of two peptide sequences: bovine ghrelin 1–27 (GSSFLSPEHQKLQRKEAKKPSGRLKPR) and bovine ghrelin 1–26 (GSSFLSPEHQKLQRKEAKKPSGRLKP), which lack the last arginine from bovine ghrelin 1–27. The major form of bovine ghrelin was isolated from group B, peak 2, which yielded approximately 367 pmol of peptide. The expected sequence of this peptide was ghrelin 1–27 with n-octanoic acid (C8:0). For some peptides, we could not determine fatty acid modifications because of the inconsistency between the expected and measured masses.

Two groups with ghrelin activity were identified via CM ion-exchange HPLC (pH 6.5) of the SP-III fraction ( Figure 3A ). Each active group was purified via CM ion-exchange HPLC (pH 4.8) and three separate rounds of RP-HPLC. The final purification of the group B fractions ( Figure 3A ), which gave the highest ghrelin yield, is shown in Figure 3B . Four active peptides were isolated from the two groups using CM-HPLC ( Table 2 ). The amino acid sequence of the group B peptide showing the highest activity was determined to be GSXFLSPEHQKVQQRKESKKPAAKLKPR. We predicted that X was serine based on the NCBI accession no. NM_213807.2 The isolated peptide was found to be a ghrelin peptide based on its sequence homology with other ghrelins. To determine the molecular weight of porcine ghrelins, isolated peptides were analyzed using MALDI-TOF mass spectrometry. Table 2 shows the measured molecular masses, isolation yields, and deduced molecular forms of the isolated peptides. In addition to peptide and genetic information, the molecular masses helped predict two peptide sequences: porcine ghrelin 1–28 (GSSFLSPEHQKVQQRKESKKPAAKLKPR) and porcine ghrelin 1–27 (GSSFLSPEHQKVQQRKESKKPAAKLKP), which lack the last arginine from porcine ghrelin 1–28. The major form of porcine ghrelin was isolated from Group B, peak 2, and these fractions yielded approximately 713 pmol of the peptide. The expected sequence of this peptide was porcine ghrelin 1–28 with n-octanoic acid (C8:0). For some peptides, we could not determine fatty acid modifications because of the inconsistency between the expected and measured masses.

Two groups with ghrelin activity were identified via CM ion-exchange HPLC (pH 6.5) of the SP-III fraction ( Figure 4A ). Each active group was purified via CM ion-exchange HPLC (pH 4.8) and three separate rounds of RP-HPLC. The final purification of the group A fractions ( Figure 4A ), which gave the highest ghrelin yield, is shown in Figure 4B . Six active peptides were isolated from the two groups using CM-HPLC ( Table 3 ). The amino acid sequence of the group A peptide showing the highest activity was determined to be GSXFLSPEHHKVQHRKESKKPPAKLKPR. We predicted that X was serine based on the NCBI accession no. XM_023619973: The isolated peptide was a ghrelin peptide based on its sequence homology with other ghrelins. Isolated peptides were analyzed using MALDI-TOF MS to determine the molecular weight of equine ghrelins. Table 3 shows the measured molecular masses, isolation yields, and deduced molecular forms of the isolated peptides. In addition to peptide and genetic information, the molecular masses helped predict two peptide sequences: equine ghrelin 1–28 (GSSFLSPEHHKVQHRKESKKPPAKLKPR) and ghrelin 1–27 (GSSFLSPEHHKVQHRKESKKPPAKLKPR), which lack the last arginine from equine ghrelin 1–28. The major form of equine ghrelin was isolated from group A, peak 1, and these fractions yielded an approximately 523 pmol of the peptide. The expected sequence of this peptide is equine ghrelin 1–28 with n-butanoic acid (C4:0). For some peptides, we could not determine fatty acid modifications because of the inconsistency between the expected and measured masses.