Transgenic Mice Overexpressing Des-Acyl Ghrelin Show Small Phenotype
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内分泌学杂志 2005年第1期
Department of Medicine and Clinical Science (H.A., K.N.), Kyoto University Graduate School of Medicine, and Translational Research Center (K.T., H.I., H.H., T.A., K.K.), Kyoto University Hospital, Kyoto 606-8507; and Departments of Bioscience (Y.A.) and Biochemistry (K.K.), National Cardiovascular Center Research Institute, Osaka 565-8565, Japan
Address all correspondence and requests for reprints to: Kazuhiko Takaya, M.D., Ph.D., Translational Research Center, Kyoto University Hospital, Kyoto 606-8507, Japan. E-mail: ktakaya@kuhp.kyoto-u.ac.jp.
Abstract
Ghrelin, a 28-amino acid acylated peptide, displays strong GH-releasing activity in concert with GHRH. The fatty acid modification of ghrelin is essential for the actions, and des-acyl ghrelin, which lacks the modification, has been assumed to be devoid of biological effects. Some recent reports, however, indicate that des-acyl ghrelin has effects on cell proliferation and survival. In the present study, we generated two lines of transgenic mice bearing the preproghrelin gene under the control of chicken ?-actin promoter. Transgenic mice overexpressed des-acyl ghrelin in a wide variety of tissues, and plasma des-acyl ghrelin levels reached 10- and 44-fold of those in control mice. They exhibited lower body weights and shorter nose-to-anus lengths, compared with control mice. The serum GH levels tended to be lower, and the serum IGF-I levels were significantly lower in both male and female transgenic mice than control mice. The responses of GH to administered GHRH were normal, whereas those to administered ghrelin were reduced, especially in female transgenic mice, compared with control mice. These data suggest that overexpressed des-acyl ghrelin may modulate the GH-IGF-I axis and result in small phenotype in transgenic mice.
Introduction
GHRELIN, AN ACYLATED peptide of 28 amino acids, was identified as an endogenous ligand for the GH secretagogue (GHS) receptor (GHS-R) (1). The major site of production of ghrelin is the stomach and it is also expressed in the hypothalamus (2, 3, 4, 5). Plasma ghrelin levels are regulated by acute feeding states. They rise by fasting and are rapidly suppressed by feeding (3, 6, 7, 8). Secretion of ghrelin is also regulated by chronic feeding states. Plasma ghrelin levels are elevated in patients with anorexia nervosa and food-restricted animals and are reduced in obese subjects (3, 6, 7, 8, 9, 10). These data suggest the possible involvement of ghrelin in energy homeostasis. In fact, ghrelin stimulates food intake in animals and humans and exhibits anticachectic effect in cancer-bearing mice (8, 11, 12, 13).
Exogenously administered ghrelin strongly stimulates GH release in a clear dose-dependent manner in vivo (1, 2, 14, 15, 16). The site of ghrelin action on GH release is not well known to date. The GHS-R is reported to be expressed in the pituitary as well as hypothalamus (17, 18, 19). Previous studies indicate that ghrelin binds to membranes from the pituitary and stimulates GH release from cultured pituitary cells (1, 20), suggesting that the pituitary is one of the sites of ghrelin actions. The stimulatory effect of GHSs and ghrelin on GH secretion, however, is more prominent in vivo than in vitro, and intact GHRH signaling is essential for the effect (1, 21). Hexarelin, one of the potent GHSs, cannot efficiently stimulate GH release in patients with GHRH receptor deficiency (22). Moreover, as we demonstrated, ghrelin has a synergistic action with GHRH. Even a low dose of ghrelin can highly augment GH release by GHRH (23). These data indicate a critical role of the hypothalamus in the stimulatory effect of ghrelin on GH secretion. The strong potency of ghrelin suggests its role as a physiological regulator of GH secretion (1, 2, 14, 15, 16). The issue, however, is currently controversial. One recent study (24), using a GHS antagonist, revealed that circulating ghrelin in peripheral blood may not play a role in generating pulsatile GH secretion. Moreover, deletion of ghrelin impairs neither growth nor appetite, indicating that ghrelin is not critically required for GH secretion (25). Another study (26), however, demonstrated that the attenuation of the GHS-R expression in vivo results in reduction in food intake and growth, suggesting a physiological role of the ghrelin-GHS-R system in the secretory regulation of GH.
The acylation of ghrelin is assumed to be essential for its actions (1). Des-acyl ghrelin, which lacks the fatty acid modification and circulates at 10-fold higher concentration than acylated ghrelin (1, 3, 27), is devoid of any endocrine activities including GH release, based on previous studies (1, 28). Recent studies (29, 30), however, indicated that des-acyl ghrelin may share with acylated ghrelin the modulation of neoplastic cell proliferation and cardiovascular cell survival in vitro. Moreover, one study shows that des-acyl ghrelin may offset the inhibitory effect of acylated ghrelin on insulin secretion (28). Although previous studies indicated that several tissues and cell lines produce des-acyl and/or acylated ghrelin (3, 27, 31, 32), the mechanism by which ghrelin is acylated is also unknown to date.
In the present study, we generated transgenic mice bearing the preproghrelin gene under the control of a cytomegalovirus immediate early enhancer and a modified chicken ?-actin promoter, designated CAG promoter (33, 34). This promoter sequence has been demonstrated to have high activity in cultured cells and transgenic mice (33, 34). Transgenic mice in the present study overexpressed des-acyl ghrelin in plasma and a wide variety of tissues and showed small phenotype. Here we show that des-acyl ghrelin may modulate endogenous ghrelin action and alter the GH-IGF-I axis in transgenic mice.
Materials and Methods
All procedures in animal experiments were approved by the Kyoto University Graduate School of Medicine Committee on Animal Research. The procedures were performed in accordance with the principle and guidelines established by the committee.
Plasmid construction and generation of transgenic mice
The full-length mouse preproghrelin cDNA (1) and the pCAGGS expression vector including the CAG promoter (34) were kindly donated by Professor Masayasu Kojima (Division of Molecular Genetics, Institute of Life Science, Kurume University, Kurume, Japan) and Professor Jun-ichi Miyazaki (Department of Nutrition and Physiological Chemistry, Osaka University School of Medicine, Osaka, Japan), respectively. Plasmid pCAGGS-ghrelin was constructed by inserting the mouse preproghrelin cDNA into the unique EcoRI site between the CAG promoter and 3'-flanking sequence of the rabbit ?-globin gene of the pCAGGS expression vector. The DNA fragment was excised from its plasmid by digestion with SalI and HindIII and then purified and microinjected into the pronuclei of fertilized eggs obtained from BDF1 female mice (Charles River Japan, Yokohama, Japan) as reported previously (35). Founder transgenic mice were identified by PCR analysis and bred with C57BL/6 mice (Japan CLEA, Osaka, Japan). Mice were housed in air-conditioned animal quarters, with the lights on between 0800 and 2000 h and were given standard rat chow (CE-2, 352 kcal per 100 g, Japan CLEA) and water ad libitum.
Measurement of total and acylated ghrelin levels in tissue samples
Tissues such as the stomach, cerebrum, heart, and kidney were removed from 8-wk-old mice under anesthesia with diethyl ether. Each sample was diced and boiled for 7 min in a 5-fold volume of water. The solution was adjusted to 1.0 M acetic acid and 20 mM hydrogen chloride after boiling, and the tissue was homogenized. The supernatant was obtained after centrifugation at 10,000 rpm for 30 min. Tissue ghrelin levels were measured using two kinds of RIAs, C-RIA for the carboxyl terminal and N-RIA for the amino terminal of ghrelin as reported previously (9, 27). C-RIA and N-RIA recognize total (acylated plus des-acyl ghrelin) and acylated ghrelin, respectively (9, 27).
Measurement of plasma total and acylated ghrelin levels
Blood samples were collected from the inferior vena cava of mice under anesthesia with diethyl ether. The samples were immediately transferred to chilled polypropylene tubes containing Na2EDTA (1 mg/ml) and aprotinin (Ohkura Pharmaceutical, Inc., Kyoto, Japan; 1000 kallikrein inactivator U/ml) and centrifuged at 4 C. For N-RIA, hydrogen chloride was added to the samples at final concentration of 0.1 N immediately after the separation of plasma. Plasma ghrelin was measured as reported previously (1, 3, 27). Briefly, the samples were subjected to a Sep-Pak C18 cartridge and C-RIA and N-RIA were carried out.
Measurement of body weights and lengths, organ weights, and daily food intake
Body weights of control and transgenic mice were measured weekly, beginning at 3 wk of age. Body lengths of 8- and 52-wk-old mice were measured by manual immobilization and extension of mice to the nose-to-anus lengths, always by the same individual. Body mass indexes (BMIs = weight/(nose-to-anus lengths)2) were calculated in 8- and 52-wk-old control and transgenic mice (36, 37). Organs such as the pituitary, stomach, cerebrum, heart, liver, kidney, spleen, pancreas, and epididymal fat were removed from 8-wk-old mice under anesthesia with diethyl ether and weighed. Daily food intake was monitored for 3 wk, beginning at 5 wk of age.
Measurement of blood glucose, serum total protein, total cholesterol, and hormones
To examine the nutritional conditions, blood glucose and serum total protein and total cholesterol levels were measured. Eight-week-old control and transgenic mice were used. Four hundred microliters of blood samples were collected from the tail vein of mice for blood glucose levels at 1000 h after 12 h fasting. Then the mice were anesthetized with diethyl ether, and 400 μl of blood samples were collected from the inferior vena cava for serum total protein, total cholesterol, and hormone levels. Blood glucose, serum total protein, and total cholesterol levels were measured by the glucose oxidase method with a reflectance glucometer (One Touch II; Lifescan, Milpitas, CA), BCA protein assay reagent kit (Pierce, Rockford, IL), and Amplex red cholesterol assay kit (Molecular Probes, Eugene, OR), respectively. Serum GH and IGF-I levels were measured with EIA kits (SPI-BIO, Bonde, France, and Diagnostic Systems Laboratories Inc., Webster, TX, respectively). Serum insulin and plasma ACTH levels were measured with EIA kits (Morinaga, Tokyo, Japan) and ACTH-RIA kit (Nichols Institute Diagnostics, San Juan Capistrano, CA), respectively. Serum TSH, LH, and FSH levels were measured with EIA kits (Amersham Biosciences, Buckinghamshire, UK).
Effects of GHRH and ghrelin on serum GH levels
Human GHRH and rat ghrelin were purchased from Sumitomo Pharmaceuticals Co., Ltd. (Osaka, Japan) and Peptide Institute, Inc. (Osaka, Japan), respectively. Male and female 8-wk-old control and Tg 10–1 mice were used under no anesthesia. Control and transgenic mice were housed in the same cage and tested on the same day. Forty mice were divided into five groups for blood sampling. Eight mice in the same group were used for each blood sampling. Control and transgenic mice were iv injected with human GHRH (60 μg/kg) or rat ghrelin (40 μg/kg). Four hundred microliters of blood samples were collected from the inferior vena cava of mice 0, 10, 20, 30, and 60 min after the injection. Serum GH levels were measured with an EIA kit (SPI-BIO).
Real-time PCR analysis of preproghrelin, GH, GHRH, somatostatin, and GHS-R mRNAs
Total RNAs from tissues, such as the stomach, small intestine, cerebrum, hypothalamus, pituitary, liver, kidney, lung, heart, and skeletal muscle, were extracted using the acid guanidinium thiocyanate-phenol-chloroform method (38). First-strand cDNA was synthesized from 1 μg of total RNA using Superscript II RT (Life Technologies, Inc., St. Louis, MO) with random hexamers according to the manufacturer’s instructions. Taqman-PCR was performed with the ABI Prism 7700 sequence detection system (Applied Biosystems, Foster City, CA) using VIC-labeled fluorogenic probes specific for preproghrelin, GH, GHRH, somatostatin, or GHS-R transcript, or the internal standard glyceraldehyde-3-phosphate dehydrogenase. Oligo primers and probes (Table 1) were chosen using the Primer Express software (Applied Biosystems). The PCR was performed using Taqman Universal PCR Mastermix (Applied Biosystems) to which primers and probes were added (final concentrations 400 and 200 nM, respectively). All samples were run in triplicate in 96-well plates in the ABI Prism 7700 sequence detector according to the manufacturer’s standard protocol. For the primer sets, serial dilutions were conducted with different cDNA preparations to confirm the kinetics of the PCR. There was no significant difference in glyceraldehyde-3-phosphate dehydrogenase mRNA levels among experimental groups.
TABLE 1. Primer and probe sequences for real-time PCR analysis for preproghrelin, GH, GHRH, somatostatin, and GHS-R mRNAs
Effects of continuous infusion of des-acyl ghrelin on the GH-IGF-I axis and body weights
Rat des-acyl ghrelin was purchased from Peptide Institute, Inc. Des-acyl ghrelin was dissolved in saline at a concentration of 700 μg/ml and stored in osmotic minipumps (DURECT Corp., Cupertino, CA). The minipumps were implanted into the peritoneum. Des-acyl ghrelin or saline was infused continuously through the minipumps into 4-wk-old C57/BL6 mice (Japan CLEA) for 10 d. The minipumps were continuously delivering saline or 250 μg/kg·d of des-acyl ghrelin for 10 d at a speed of 0.22 μl/h. Body weights were measured daily for 10 d. Four hundred microliters of blood samples for the measurement of serum GH and IGF-1 levels were collected from the inferior vena cava of mice under anesthesia with diethyl ether 10 d after the implantation.
Hematoxylin eosin and immunohistochemical staining for total ghrelin, acylated ghrelin, and GH of the pituitary
The pituitaries were removed from male 8-wk-old mice under anesthesia with diethyl ether and fixed with 4% paraformaldehyde and 0.2% picric acid and embedded in paraffin. The tissues were cut in 3-μm-thick slices. Samples were subjected to immunohistochemical staining for total and acylated ghrelin as well as hematoxylin eosin staining. After pretreatment with 0.3% hydrogen peroxide and incubation with normal goat serum, all slices were incubated overnight at 4 C with ghrelin(13–28) antiserum recognizing total (des-acyl plus acylated) ghrelin (final dilution, 1:5000), antighrelin(1–11) antiserum specifically recognizing acylated ghrelin (final dilution, 1:5000), or anti-GH antiserum (Biogenesis, Poole, UK) (final dilution, 1:200). All of the sections were stained by the avidin-biotin complex method and counterstained with hematoxylin as reported previously (39).
Statistical analysis
Results are expressed as the mean ± SEM. ANOVA followed by the t test was used to assess differences between control and transgenic mice. P < 0.05 was considered to be statistically significant.
Results
Generation of transgenic mice and preproghrelin mRNA levels
Two lines of transgenic mice with six (Tg 10–1) and 12 (Tg 9–2) copy numbers were identified by PCR and Southern blot analysis. Preproghrelin mRNAs were detected only in the stomach, small intestine, lung, pituitary, and hypothalamus of control mice, and the amounts were 100, 4, 2.1, 1.5, and 0.5 in arbitrary units (AU), respectively (Fig. 1). On the other hand, they were detected in all tissues examined in Tg 9–2 and Tg 10–1 mice, and the amounts in the stomach of Tg 9–2 and Tg 10–1 mice reached 1100 and 5200 AU, respectively (Fig. 1). Preproghrelin mRNA levels in other tissues of Tg 9–2 and Tg 10–1 mice also exceeded those of control mice.
FIG. 1. Preproghrelin mRNA levels in the tissues of control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice quantified by real-time PCR analysis. Lanes 1, stomach; 2, small intestine; 3, cerebrum; 4, hypothalamus; 5, pituitary; 6, liver; 7, kidney; 8, lung; 9, heart; and 10, skeletal muscle.
Total and acylated ghrelin levels in tissues and plasma
Eight-week-old control, Tg 9–2, and Tg 10–1 mice were used (Table 2). Although high total ghrelin levels were detected in the stomach, only very low levels were detected in other tissues of control mice. Tg 9–2 and Tg 10–1 mice showed significantly higher total ghrelin levels in the stomach than control mice (P < 0.01 for each). Tg 9–2 and Tg 10–1 mice also showed total ghrelin levels in all of the other tissues significantly higher than control mice. High levels of acylated ghrelin were also detected in the stomach of control, Tg 9–2, and Tg 10–1 mice. There was, however, no significant difference between control and Tg 9–2 mice and between control and Tg 10–1 mice. Only very low acylated ghrelin levels if any were detected in other tissues of control, Tg 9–2, and Tg 10–1 mice. Plasma total ghrelin levels in control, Tg 9–2, and Tg 10–1 mice were 1104.5 ± 94.4, 11230.6 ± 1147.1, and 48565.5 ± 9291.5 fmol/ml, respectively. Those in Tg 9–2 and Tg 10–1 mice were significantly higher than those in control mice (P < 0.01 for each). Plasma acylated ghrelin levels in control, Tg 9–2, and Tg 10–1 mice were 83.7 ± 11.9, 79.7 ± 10.1, and 86.3 ± 21.1 fmol/ml, respectively. The differences between control and Tg 9–2 mice and control and Tg 10–1 mice were not significant.
TABLE 2. Total and acylated ghrelin levels in plasma and tissues of 8-wk-old control and transgenic mice (n = 8/group)
Body weights and lengths, relative organ weights, and BMIs
Body weights of control, Tg 9–2, and Tg 10–1 mice are shown in Table 3 and Fig. 2A. Male Tg 9–2 and Tg 10–1 mice were significantly lighter in the body weight than control mice (P < 0.05 and P < 0.01, respectively). Female Tg 10–1 mice were also significantly lighter than control mice (P < 0.01). The difference between female control and Tg 9–2 mice was not significant. Fifteen-week-old male and female Tg 10–1 and male Tg 9–2 mice were still significantly lighter than control mice (P < 0.05, P < 0.01, and P < 0.01, respectively). Body lengths (nose-to-anus lengths) of control and transgenic mice are shown in Table 3. Eight-week-old male Tg 9–2 and Tg 10–1 mice were significantly shorter in the body length than control mice (P < 0.05 and P < 0.01, respectively). Female Tg 10–1 mice were significantly shorter than control mice (P < 0.01). The difference between female control and Tg 9–2 mice was not significant. BMIs were calculated from the body weights and lengths. No significant difference was noted between control and Tg 9–2 mice and control and Tg 10–1 mice (Table 3). Fifty-two-week-old male Tg 9–2 and Tg10–1 mice were still significantly lighter and shorter, compared with control mice (Table 3), and no significant difference was noted in BMIs between control and transgenic mice. Relative organ weights of 8-wk-old male control and Tg 10–1 mice were calculated from the organ and body weights. No significant difference was noted between control and Tg 10–1 mice (Fig. 2B). No significant difference was noted in the pituitary size between control and Tg 10–1 mice (0.058 ± 0.002 and 0.055 ± 0.003 mg/body weight (grams), respectively).
TABLE 3. Body weights, lengths, and BMIs of 8-wk-old and 52-wk-old control and transgenic mice (n = 8/group)
FIG. 2. Body weights (BW) and relative organ weights. A, Body weights of male (left panel) and female (right panel) control (triangles), Tg 9–2 (circles), and Tg 10–1 (squares) mice (n = 8/group). B, Relative organ weights of 8-wk-old control (closed bars) and Tg 10–1 (open bars) mice calculated from the organ and body weights (n = 8/group). 1, stomach; 2, cerebrum; 3, heart; 4, liver; 5, kidney; 6, spleen; 7, pancreas; 8, epididymal fat. a, P < 0.05; b, P < 0.01 (vs. control mice).
Immunohistochemical staining for total and acylated ghrelin of the pituitary
Immunohistochemical staining for total and acylated ghrelin is shown in Fig. 3, A–D. None of total ghrelin-positive cells were observed in the pituitary of control mice (Fig. 3A). On the other hand, many total ghrelin-immunoreactive pituitary cells were observed in Tg 10–1 mice (Fig. 3B). Approximately 30% of the anterior pituitary cells in all sections examined were total ghrelin immunoreactive. None of acylated ghrelin-positive cells were observed in the pituitary of either control or Tg 10–1 mice (Fig. 3, C and D).
FIG. 3. The localization of total and acylated ghrelin-immunoreactive cells in the pituitary of 8-wk-old male control (A and C) and Tg 10–1 (B and D) mice. A and B, An antiserum raised to ghrelin(13–28) recognizing total (acylated plus des-acyl) ghrelin was used. C and D, An antiserum raised to ghrelin(1–11) specifically recognizing acylated ghrelin was used. Original magnification, x40. The immunoreactive cells are stained brown by the avidin-biotin complex methods.
Food intake and biochemical parameters in the blood
Although absolute amounts of daily food intake were reduced in Tg 9–2 and Tg 10–1 mice, the amounts per body weight were not significantly changed in either male or female Tg 9–2 or Tg 10–1 mice, compared with control mice (Table 4). No significant differences in blood glucose, serum total protein, total cholesterol, and insulin levels were noted between 8-wk-old control and Tg 9–2 mice and control and Tg 10–1 mice (Table 4).
TABLE 4. Daily food intake, blood glucose, serum total protein, total cholesterol, and insulin levels in 8-wk-old control and transgenic mice (n = 8/group)
Serum GH, IGF-1, and pituitary GH mRNA levels
Serum GH levels in male control, Tg 9–2, and Tg 10–1 mice were 5.5 ± 1.9, 3.7 ± 0.7, and 2.3 ± 0.9 ng/ml, respectively (Fig. 4A). Those in female control, Tg 9–2, and Tg 10–1 mice were 4.7 ± 1.7, 2.5 ± 0.9, and 1.7 ± 0.8 ng/ml, respectively (Fig. 4A). There were tendencies for decline in serum GH levels in male and female Tg 9–2 and Tg 10–1 mice, compared with control mice, although the differences between them were not significant. Serum IGF-I levels in male control, Tg 9–2, and Tg 10–1 mice were 522 ± 23.6, 413.2 ± 49.0, and 364.1 ± 25.6 ng/ml, respectively (Fig. 4B). Those in male Tg 9–2 and Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.01 for each). Serum IGF-I levels in female control, Tg 9–2, and Tg 10–1 mice were 509.7 ± 43.1, 545.5 ± 64.1, and 253.7 ± 36.4 ng/ml, respectively (Fig. 4B). Those in female Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.01). The difference between female control and Tg 9–2 mice was not significant.
FIG. 4. Serum GH, IGF-I, and pituitary GH mRNA levels in 8-wk-old control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice (n = 8/group). A, Serum GH levels. B, Serum IGF-I levels. C, Pituitary GH mRNA levels. a, P < 0.05; b, P < 0.01 (vs. control mice).
Pituitary GH mRNA levels in male control, Tg 9–2, and Tg 10–1 mice were 1.00, 0.62, and 0.42 AU, respectively. Those in Tg 9–2 and Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.05 and P < 0.01, respectively). Pituitary GH mRNA levels in female control, Tg 9–2, and Tg 10–1 mice were 1.00, 0.97, and 0.71 AU. Those in female Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.05). The difference between those in female control and Tg 9–2 mice was not significant (Fig. 4C).
Plasma ACTH, serum TSH, LH, and FSH levels
Plasma ACTH, serum TSH, LH, and FSH levels in 8-wk-old in male control and transgenic mice are shown in Table 5. No significant difference was noted in the levels between control and transgenic mice.
TABLE 5. Plasma ACTH, serum TSH, LH, and FSH levels of 8-wk-old control and transgenic mice (n = 8/group)
Hematoxylin eosin and immunohistochemical staining for GH of the pituitary
Hematoxylin eosin staining is shown in Fig. 5, A and B. The pituitary morphology of Tg 10–1 mice was not different from that of the control mice. Immunohistochemical staining for GH is shown in Fig. 5, C and D. The distribution of GH-immunoreactive cells in the pituitary of Tg mice was similar to that of control mice.
FIG. 5. Morphology of the pituitary and the localization of GH-immunoreactive cells in the pituitary of 8-wk-old male control (A and C) and Tg 10–1 (B and D) mice. A and B, Hematoxylin eosin (HE) staining. C and D, The localization of total and GH-immunoreactive cells in the pituitary. Original magnification, x40. The immunoreactive cells are stained brown by the avidin-biotin complex methods.
Effects of GHRH and ghrelin on GH release
Control and Tg 10–1 mice were used. Serum GH levels after GHRH administration in male and female Tg 10–1 mice were similar to those of control mice throughout the course of the experiment (Fig. 6A). There was no significant difference in serum GH level at each time point between both male and female Tg-10 and control mice. Serum GH levels 10 min after ghrelin administration in male Tg10–1 and control mice were 63.1 ± 6.8 and 72.6 ± 12.0 ng/ml, respectively (Fig. 6B, left panel). The difference was not significant. Serum GH levels 20 min after ghrelin administration in male Tg10–1 and control mice were 30.2 ± 6.7 and 61.2 ± 15.5 ng/ml, respectively, and levels after 30 min were 11.8 ± 1.4 and 21.9 ± 4.1 ng/ml, respectively (Fig. 6B, left panel). Both differences were significant (P < 0.01). Serum GH levels 10 min after ghrelin administration in female Tg10–1 and control mice were 8.7 ± 3.7 and 52.8 ± 8.2 ng/ml, respectively, and those after 20 min were 29.8 ± 6.3 and 78.5 ± 14.3 ng/ml, respectively (Fig. 6B, right panel). Both differences were significant (P < 0.01). Serum GH levels 30 min after ghrelin administration in female Tg10–1 and control mice were 22.8 ± 6.3 and 22.3 ± 8.8 ng/ml, respectively (Fig. 6B, right panel). The difference was not significant.
FIG. 6. The responses of GH to GHRH and ghrelin in 8-wk-old control (closed circles) and Tg 10–1 (open triangles) mice. A, Time course of serum GH levels after iv injection of 60 μg/kg GHRH (n = 8/each point). B, Time course of serum GH levels after iv injection of 40 μg/kg ghrelin (n = 8/each point). a, P < 0.01 (vs. control mice).
Expression of GHS-R in the pituitary
GHS-R mRNA levels of male control, Tg 9–2, and Tg 10–1 mice were 1.00, 1.56, and 3.46 AU, respectively (Fig. 7). The difference between control and Tg 10–1 mice was significant (P < 0.01).
FIG. 7. Pituitary GHS-R mRNA levels in 8-wk-old control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice quantified by real-time PCR analysis (n = 8/group). a, P < 0.01 (vs. control mice).
Expression of hypothalamic neuropeptides that regulate GH secretion
GHRH mRNA levels of male control, Tg 9–2, and Tg 10–1 mice were 1.00, 0.88, and, 0.80 AU, respectively (Fig. 8A). The differences between control and Tg 9–2 mice and control and Tg 10–1 mice were not significant. Somatostatin mRNA levels of male control, Tg 9–2, and Tg 10–1 mice were 1.00, 1.08, and 0.97 AU, respectively (Fig. 8B). The differences between control and Tg 9–2 mice and control and Tg 10–1 mice were not significant.
FIG. 8. Hypothalamic GHRH and somatostatin mRNA levels in 8-wk-old control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice quantified by real-time PCR analysis. A, GHRH mRNA levels (n = 8/group). B, Somatostatin mRNA levels (n = 8/group).
Effects of continuous infusion of des-acyl ghrelin on GH-IGF-I axis and body weights
Male and female C57BL/6 mice were used. Serum GH levels after 10 d treatment with saline and des-acyl ghrelin in male mice were 5.8 ± 1.1 and 7.5 ± 2.0 ng/ml, respectively. The difference was not significant. Those with saline and des-acyl ghrelin in female mice were 9.2 ± 2.2 and 9.5 ± 1.8 ng/ml, respectively. The difference was not significant either. Serum IGF-I levels after 10 d treatment with saline and des-acyl ghrelin in male mice were 769.3 ± 16.6 and 768.7 ± 21.6 ng/ml, respectively. The difference was not significant. Those with saline and des-acyl ghrelin in female mice were 766.2 ± 13.4 and 719.4 ± 49.1 ng/ml, respectively. The difference was not significant either. Body weights and lengths in des-acyl ghrelin-injected mice were not significantly different from those in saline-injected mice in either males or females (data not shown).
Discussion
We have generated transgenic mouse lines that overexpress preproghrelin mRNA in a wide variety of tissues. The wide tissue distribution of preproghrelin mRNA in transgenic mice was consistent with previous reports on transgenic mice using the CAG promoter (33, 34). Preproghrelin mRNA expression was increased, especially in Tg 10–1 mice, and its amount in the stomach reached 52-fold of that in control mice. Consistent with the elevated mRNA expression, peptide levels of total ghrelin (des-acyl plus acylated ghrelin) in various tissues were also elevated in transgenic mice. Plasma total ghrelin levels in transgenic mice showed marked results. Those in transgenic mice showed 10- and 44-fold of those in control mice. We originally intended to generate mice overexpressing biologically active ghrelin. Unexpectedly, acylated ghrelin levels were not changed in all tissues examined and plasma of transgenic mice, compared with those of control mice, indicating that transgenic mice overexpress only des-acyl ghrelin. The expression of acylated ghrelin has been reported in a small number of tissues, such as the stomach (X/A cells), duodenum, hypothalamus, and pancreatic -cells (1, 31, 39, 40). These reports and our present data suggest that only a limited number of cell lineages may able to process proghrelin or acylate ghrelin. The underlying mechanism by which ghrelin is acylated is unknown to date. Further study is needed to clarify the mechanism of the acylation.
The acylation of ghrelin is assumed to be essential for its actions, and des-acyl ghrelin, which lacks the modification, is devoid of endocrine actions, based on previous studies (1, 41). However, recent studies indicated that des-acyl ghrelin may have some actions. Des-acyl ghrelin as well as acylated ghrelin causes a significant inhibition of cell proliferation in human breast carcinoma cell lines (29) and inhibits cell death in cardiomyocytes and endothelial cells through ERK1/2 and phosphatidylinositol 3-kinase/AKT (30). In addition, one study (42) reported that acylated and des-acyl ghrelin promote adipogenesis directly in vivo by a mechanism independent of known GHS-Rs. Moreover, another study (28) indicated that des-acyl ghrelin may offset the action of acylated ghrelin on insulin secretion. Ghrelin has been shown to induce a reduction in serum insulin levels. In the study, coadministration of acylated plus des-acyl ghrelin did not result in any changes in serum insulin levels in humans, suggesting that ghrelin action on insulin is modulated by des-acyl ghrelin.
The present study indicates that transgenic mice overexpressing des-acyl ghrelin show small phenotype. Longitudinal growth was the most reduced in female Tg 10–1 mice (20% reduction from control mice). The phenotype was not associated with changes in BMIs. These mice did not show decreased food intake or decreased body fat mass. In addition, they showed normal nutritional condition, based on their biochemical parameters, including blood glucose, serum total protein, and total cholesterol levels. These data indicate that the small phenotype of transgenic mice is not attributed to poor nutritional condition.
Serum IGF-I levels were significantly reduced in male and female transgenic mice, compared with control mice. Female Tg 10–1 mice had no less than 50% reduction in serum IGF-I levels, compared with control mice. Although the differences in serum GH levels between control and transgenic mice were not statistically significant, probably because of the pulsatile character of GH secretion, the levels tended to be reduced in transgenic mice, compared with control mice, and the mean GH level of Tg10–1 mice was only 50% of that of control mice. It should be emphasized that Tg 10–1 mice showed lower serum GH levels than Tg 9–2 mice. Body weights and lengths of the former were more reduced than the latter. It should be also noted that the former showed higher des-acyl ghrelin expression than the latter. Reduced pituitary GH mRNA levels in transgenic mice support the observation. The GH-IGF-I axis-specific alteration in transgenic mice was also indicated by the measurement of other anterior pituitary hormones than GH. Plasma ACTH, serum TSH, LH, and FSH levels were not altered.
The size and morphology of the pituitary including the somatotrope populations of transgenic mice were similar to those of control mice. These data indicated that there is no apparent change, suggesting developmental problems in the pituitary of transgenic mice.
Responses of GH to GHRH and ghrelin in transgenic mice exhibited intriguing results. Transgenic mice showed normal response of GH to GHRH. Alternatively, if we consider that the basal GH levels are lower in transgenic mice, the similar maximal response might indicate that they are hyperresponsive to GHRH. It is not likely that an insufficient dose of GHRH induced submaximal response of GH in both control and transgenic mice, judging from previous reports (43). On the other hand, the responses of GH to ghrelin were reduced in transgenic mice. It is noteworthy that the reduction was much greater in female transgenic mice than in male mice, if we take their serum IGF-I levels into account. Taken together our results and these reports indicate that overexpression of des-acyl ghrelin in our mice may result in reduction of GH response to endogenous ghrelin, and it may result in the reduced serum IGF-I levels in transgenic mice.
The reduced GH response to ghrelin in transgenic mice could be due to down-regulated the GHS-R. However, the pituitary GHS-R mRNA levels in the transgenic mice were rather elevated. It is not likely that overexpressed des-acyl ghrelin acts as a blocking agent to the GHS-R because 125I-labeled acylated ghrelin bound to the GHS-R cannot be displaced by des-acyl ghrelin (20). Overexpressed des-acyl ghrelin may have some effects on endogenous GH secretion, modifying the action of endogenous ghrelin in transgenic mice via, for instance, another receptor or modulation of the signal transduction pathway after the GHS-R.
Previous reports indicated that the hypothalamus plays a critical role in the stimulatory effect of ghrelin on GH secretion as well as the pituitary (21, 22, 23). Because GH secretion is regulated chiefly by two hypothalamic hormones, GHRH and somatostatin, the expression of these hormones could be altered in transgenic mice. We could not find any significant difference in either GHRH or somatostatin mRNA levels between control and transgenic mice. These data might suggest that overexpressed des-acyl ghrelin acts on not only the pituitary but also the hypothalamus in the transgenic mice, judging from the fact that hypothalamus GHRH mRNA were not elevated, and somatostatin mRNA levels were not decreased despite the decreased serum GH levels.
We could not show, unfortunately, that continuous ip infusion of des-acyl ghrelin has some effect on serum GH and IGF-I levels or body weights. It should be noted, however, that plasma des-acyl ghrelin levels in transgenic mice reached 10- and 50-fold of those in control mice. Administration of a higher dose of des-acyl ghrelin, or longer administration, might result in alteration in the GH-IGF-I axis. On the other hand, the phenotype of transgenic mice might reflect direct effects of ubiquitous expression of des-acyl ghrelin. It should also be noted that high levels of des-acyl ghrelin were detected in a various tissues, especially in the pituitary, as well as in plasma of transgenic mice. The des-acyl ghrelin immunoreactive pituitary cells might play an important role in the mechanism for the altered GH-IGF-I axis in a paracrine or autocrine manner. It should be pointed out that preproghrelin mRNA is reported to be expressed in the normal pituitary (44), as we showed in the present study, suggesting its physiological role in GH secretion. The phenotype of transgenic mice may reflect the role. Further study is needed for this issue.
The mechanism underlying the sexual dimorphism in the responses of GH to ghrelin in transgenic mice is not fully understood. It might be due to the gender difference in the secretory regulation of GH. Female mice have been reported to be different from male mice in that they have noncyclical and rather low somatostatin output and that GHRH plays a dominant role in it (45). There might be a GHRH-dependent mechanism for the reduced response in transgenic mice. Indeed, one recent report (26) indicated that transgenic rats expressing an antisense GHS-R mRNA in the hypothalamic arcuate nucleus show marked gender difference in GH secretion. Although there was no significant difference in pulse frequency and baseline levels of GH between male control and transgenic rats, female transgenic rats showed lower baseline levels and fewer pulses of GH than female control rats (26).
The 94-amino acid proghrelin is cleaved to yield ghrelin. One previous study (46) demonstrated that C-terminal proghrelin peptides are present in the human circulation. Transgenic mice in the present study would also overexpress these peptides. We have not excluded the possibility that the phenotype of transgenic mice might be due to the effects of these peptides.
In conclusion, the present study demonstrates that transgenic mice overexpressing des-acyl ghrelin show small phenotype and altered GH-IGF-I axis. These observations may indicate a role of des-acyl ghrelin in the regulation of GH secretion.
Acknowledgments
The authors gratefully acknowledge the excellent technical support of Chieko Ishimoto and Hitomi Hiratani.
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Address all correspondence and requests for reprints to: Kazuhiko Takaya, M.D., Ph.D., Translational Research Center, Kyoto University Hospital, Kyoto 606-8507, Japan. E-mail: ktakaya@kuhp.kyoto-u.ac.jp.
Abstract
Ghrelin, a 28-amino acid acylated peptide, displays strong GH-releasing activity in concert with GHRH. The fatty acid modification of ghrelin is essential for the actions, and des-acyl ghrelin, which lacks the modification, has been assumed to be devoid of biological effects. Some recent reports, however, indicate that des-acyl ghrelin has effects on cell proliferation and survival. In the present study, we generated two lines of transgenic mice bearing the preproghrelin gene under the control of chicken ?-actin promoter. Transgenic mice overexpressed des-acyl ghrelin in a wide variety of tissues, and plasma des-acyl ghrelin levels reached 10- and 44-fold of those in control mice. They exhibited lower body weights and shorter nose-to-anus lengths, compared with control mice. The serum GH levels tended to be lower, and the serum IGF-I levels were significantly lower in both male and female transgenic mice than control mice. The responses of GH to administered GHRH were normal, whereas those to administered ghrelin were reduced, especially in female transgenic mice, compared with control mice. These data suggest that overexpressed des-acyl ghrelin may modulate the GH-IGF-I axis and result in small phenotype in transgenic mice.
Introduction
GHRELIN, AN ACYLATED peptide of 28 amino acids, was identified as an endogenous ligand for the GH secretagogue (GHS) receptor (GHS-R) (1). The major site of production of ghrelin is the stomach and it is also expressed in the hypothalamus (2, 3, 4, 5). Plasma ghrelin levels are regulated by acute feeding states. They rise by fasting and are rapidly suppressed by feeding (3, 6, 7, 8). Secretion of ghrelin is also regulated by chronic feeding states. Plasma ghrelin levels are elevated in patients with anorexia nervosa and food-restricted animals and are reduced in obese subjects (3, 6, 7, 8, 9, 10). These data suggest the possible involvement of ghrelin in energy homeostasis. In fact, ghrelin stimulates food intake in animals and humans and exhibits anticachectic effect in cancer-bearing mice (8, 11, 12, 13).
Exogenously administered ghrelin strongly stimulates GH release in a clear dose-dependent manner in vivo (1, 2, 14, 15, 16). The site of ghrelin action on GH release is not well known to date. The GHS-R is reported to be expressed in the pituitary as well as hypothalamus (17, 18, 19). Previous studies indicate that ghrelin binds to membranes from the pituitary and stimulates GH release from cultured pituitary cells (1, 20), suggesting that the pituitary is one of the sites of ghrelin actions. The stimulatory effect of GHSs and ghrelin on GH secretion, however, is more prominent in vivo than in vitro, and intact GHRH signaling is essential for the effect (1, 21). Hexarelin, one of the potent GHSs, cannot efficiently stimulate GH release in patients with GHRH receptor deficiency (22). Moreover, as we demonstrated, ghrelin has a synergistic action with GHRH. Even a low dose of ghrelin can highly augment GH release by GHRH (23). These data indicate a critical role of the hypothalamus in the stimulatory effect of ghrelin on GH secretion. The strong potency of ghrelin suggests its role as a physiological regulator of GH secretion (1, 2, 14, 15, 16). The issue, however, is currently controversial. One recent study (24), using a GHS antagonist, revealed that circulating ghrelin in peripheral blood may not play a role in generating pulsatile GH secretion. Moreover, deletion of ghrelin impairs neither growth nor appetite, indicating that ghrelin is not critically required for GH secretion (25). Another study (26), however, demonstrated that the attenuation of the GHS-R expression in vivo results in reduction in food intake and growth, suggesting a physiological role of the ghrelin-GHS-R system in the secretory regulation of GH.
The acylation of ghrelin is assumed to be essential for its actions (1). Des-acyl ghrelin, which lacks the fatty acid modification and circulates at 10-fold higher concentration than acylated ghrelin (1, 3, 27), is devoid of any endocrine activities including GH release, based on previous studies (1, 28). Recent studies (29, 30), however, indicated that des-acyl ghrelin may share with acylated ghrelin the modulation of neoplastic cell proliferation and cardiovascular cell survival in vitro. Moreover, one study shows that des-acyl ghrelin may offset the inhibitory effect of acylated ghrelin on insulin secretion (28). Although previous studies indicated that several tissues and cell lines produce des-acyl and/or acylated ghrelin (3, 27, 31, 32), the mechanism by which ghrelin is acylated is also unknown to date.
In the present study, we generated transgenic mice bearing the preproghrelin gene under the control of a cytomegalovirus immediate early enhancer and a modified chicken ?-actin promoter, designated CAG promoter (33, 34). This promoter sequence has been demonstrated to have high activity in cultured cells and transgenic mice (33, 34). Transgenic mice in the present study overexpressed des-acyl ghrelin in plasma and a wide variety of tissues and showed small phenotype. Here we show that des-acyl ghrelin may modulate endogenous ghrelin action and alter the GH-IGF-I axis in transgenic mice.
Materials and Methods
All procedures in animal experiments were approved by the Kyoto University Graduate School of Medicine Committee on Animal Research. The procedures were performed in accordance with the principle and guidelines established by the committee.
Plasmid construction and generation of transgenic mice
The full-length mouse preproghrelin cDNA (1) and the pCAGGS expression vector including the CAG promoter (34) were kindly donated by Professor Masayasu Kojima (Division of Molecular Genetics, Institute of Life Science, Kurume University, Kurume, Japan) and Professor Jun-ichi Miyazaki (Department of Nutrition and Physiological Chemistry, Osaka University School of Medicine, Osaka, Japan), respectively. Plasmid pCAGGS-ghrelin was constructed by inserting the mouse preproghrelin cDNA into the unique EcoRI site between the CAG promoter and 3'-flanking sequence of the rabbit ?-globin gene of the pCAGGS expression vector. The DNA fragment was excised from its plasmid by digestion with SalI and HindIII and then purified and microinjected into the pronuclei of fertilized eggs obtained from BDF1 female mice (Charles River Japan, Yokohama, Japan) as reported previously (35). Founder transgenic mice were identified by PCR analysis and bred with C57BL/6 mice (Japan CLEA, Osaka, Japan). Mice were housed in air-conditioned animal quarters, with the lights on between 0800 and 2000 h and were given standard rat chow (CE-2, 352 kcal per 100 g, Japan CLEA) and water ad libitum.
Measurement of total and acylated ghrelin levels in tissue samples
Tissues such as the stomach, cerebrum, heart, and kidney were removed from 8-wk-old mice under anesthesia with diethyl ether. Each sample was diced and boiled for 7 min in a 5-fold volume of water. The solution was adjusted to 1.0 M acetic acid and 20 mM hydrogen chloride after boiling, and the tissue was homogenized. The supernatant was obtained after centrifugation at 10,000 rpm for 30 min. Tissue ghrelin levels were measured using two kinds of RIAs, C-RIA for the carboxyl terminal and N-RIA for the amino terminal of ghrelin as reported previously (9, 27). C-RIA and N-RIA recognize total (acylated plus des-acyl ghrelin) and acylated ghrelin, respectively (9, 27).
Measurement of plasma total and acylated ghrelin levels
Blood samples were collected from the inferior vena cava of mice under anesthesia with diethyl ether. The samples were immediately transferred to chilled polypropylene tubes containing Na2EDTA (1 mg/ml) and aprotinin (Ohkura Pharmaceutical, Inc., Kyoto, Japan; 1000 kallikrein inactivator U/ml) and centrifuged at 4 C. For N-RIA, hydrogen chloride was added to the samples at final concentration of 0.1 N immediately after the separation of plasma. Plasma ghrelin was measured as reported previously (1, 3, 27). Briefly, the samples were subjected to a Sep-Pak C18 cartridge and C-RIA and N-RIA were carried out.
Measurement of body weights and lengths, organ weights, and daily food intake
Body weights of control and transgenic mice were measured weekly, beginning at 3 wk of age. Body lengths of 8- and 52-wk-old mice were measured by manual immobilization and extension of mice to the nose-to-anus lengths, always by the same individual. Body mass indexes (BMIs = weight/(nose-to-anus lengths)2) were calculated in 8- and 52-wk-old control and transgenic mice (36, 37). Organs such as the pituitary, stomach, cerebrum, heart, liver, kidney, spleen, pancreas, and epididymal fat were removed from 8-wk-old mice under anesthesia with diethyl ether and weighed. Daily food intake was monitored for 3 wk, beginning at 5 wk of age.
Measurement of blood glucose, serum total protein, total cholesterol, and hormones
To examine the nutritional conditions, blood glucose and serum total protein and total cholesterol levels were measured. Eight-week-old control and transgenic mice were used. Four hundred microliters of blood samples were collected from the tail vein of mice for blood glucose levels at 1000 h after 12 h fasting. Then the mice were anesthetized with diethyl ether, and 400 μl of blood samples were collected from the inferior vena cava for serum total protein, total cholesterol, and hormone levels. Blood glucose, serum total protein, and total cholesterol levels were measured by the glucose oxidase method with a reflectance glucometer (One Touch II; Lifescan, Milpitas, CA), BCA protein assay reagent kit (Pierce, Rockford, IL), and Amplex red cholesterol assay kit (Molecular Probes, Eugene, OR), respectively. Serum GH and IGF-I levels were measured with EIA kits (SPI-BIO, Bonde, France, and Diagnostic Systems Laboratories Inc., Webster, TX, respectively). Serum insulin and plasma ACTH levels were measured with EIA kits (Morinaga, Tokyo, Japan) and ACTH-RIA kit (Nichols Institute Diagnostics, San Juan Capistrano, CA), respectively. Serum TSH, LH, and FSH levels were measured with EIA kits (Amersham Biosciences, Buckinghamshire, UK).
Effects of GHRH and ghrelin on serum GH levels
Human GHRH and rat ghrelin were purchased from Sumitomo Pharmaceuticals Co., Ltd. (Osaka, Japan) and Peptide Institute, Inc. (Osaka, Japan), respectively. Male and female 8-wk-old control and Tg 10–1 mice were used under no anesthesia. Control and transgenic mice were housed in the same cage and tested on the same day. Forty mice were divided into five groups for blood sampling. Eight mice in the same group were used for each blood sampling. Control and transgenic mice were iv injected with human GHRH (60 μg/kg) or rat ghrelin (40 μg/kg). Four hundred microliters of blood samples were collected from the inferior vena cava of mice 0, 10, 20, 30, and 60 min after the injection. Serum GH levels were measured with an EIA kit (SPI-BIO).
Real-time PCR analysis of preproghrelin, GH, GHRH, somatostatin, and GHS-R mRNAs
Total RNAs from tissues, such as the stomach, small intestine, cerebrum, hypothalamus, pituitary, liver, kidney, lung, heart, and skeletal muscle, were extracted using the acid guanidinium thiocyanate-phenol-chloroform method (38). First-strand cDNA was synthesized from 1 μg of total RNA using Superscript II RT (Life Technologies, Inc., St. Louis, MO) with random hexamers according to the manufacturer’s instructions. Taqman-PCR was performed with the ABI Prism 7700 sequence detection system (Applied Biosystems, Foster City, CA) using VIC-labeled fluorogenic probes specific for preproghrelin, GH, GHRH, somatostatin, or GHS-R transcript, or the internal standard glyceraldehyde-3-phosphate dehydrogenase. Oligo primers and probes (Table 1) were chosen using the Primer Express software (Applied Biosystems). The PCR was performed using Taqman Universal PCR Mastermix (Applied Biosystems) to which primers and probes were added (final concentrations 400 and 200 nM, respectively). All samples were run in triplicate in 96-well plates in the ABI Prism 7700 sequence detector according to the manufacturer’s standard protocol. For the primer sets, serial dilutions were conducted with different cDNA preparations to confirm the kinetics of the PCR. There was no significant difference in glyceraldehyde-3-phosphate dehydrogenase mRNA levels among experimental groups.
TABLE 1. Primer and probe sequences for real-time PCR analysis for preproghrelin, GH, GHRH, somatostatin, and GHS-R mRNAs
Effects of continuous infusion of des-acyl ghrelin on the GH-IGF-I axis and body weights
Rat des-acyl ghrelin was purchased from Peptide Institute, Inc. Des-acyl ghrelin was dissolved in saline at a concentration of 700 μg/ml and stored in osmotic minipumps (DURECT Corp., Cupertino, CA). The minipumps were implanted into the peritoneum. Des-acyl ghrelin or saline was infused continuously through the minipumps into 4-wk-old C57/BL6 mice (Japan CLEA) for 10 d. The minipumps were continuously delivering saline or 250 μg/kg·d of des-acyl ghrelin for 10 d at a speed of 0.22 μl/h. Body weights were measured daily for 10 d. Four hundred microliters of blood samples for the measurement of serum GH and IGF-1 levels were collected from the inferior vena cava of mice under anesthesia with diethyl ether 10 d after the implantation.
Hematoxylin eosin and immunohistochemical staining for total ghrelin, acylated ghrelin, and GH of the pituitary
The pituitaries were removed from male 8-wk-old mice under anesthesia with diethyl ether and fixed with 4% paraformaldehyde and 0.2% picric acid and embedded in paraffin. The tissues were cut in 3-μm-thick slices. Samples were subjected to immunohistochemical staining for total and acylated ghrelin as well as hematoxylin eosin staining. After pretreatment with 0.3% hydrogen peroxide and incubation with normal goat serum, all slices were incubated overnight at 4 C with ghrelin(13–28) antiserum recognizing total (des-acyl plus acylated) ghrelin (final dilution, 1:5000), antighrelin(1–11) antiserum specifically recognizing acylated ghrelin (final dilution, 1:5000), or anti-GH antiserum (Biogenesis, Poole, UK) (final dilution, 1:200). All of the sections were stained by the avidin-biotin complex method and counterstained with hematoxylin as reported previously (39).
Statistical analysis
Results are expressed as the mean ± SEM. ANOVA followed by the t test was used to assess differences between control and transgenic mice. P < 0.05 was considered to be statistically significant.
Results
Generation of transgenic mice and preproghrelin mRNA levels
Two lines of transgenic mice with six (Tg 10–1) and 12 (Tg 9–2) copy numbers were identified by PCR and Southern blot analysis. Preproghrelin mRNAs were detected only in the stomach, small intestine, lung, pituitary, and hypothalamus of control mice, and the amounts were 100, 4, 2.1, 1.5, and 0.5 in arbitrary units (AU), respectively (Fig. 1). On the other hand, they were detected in all tissues examined in Tg 9–2 and Tg 10–1 mice, and the amounts in the stomach of Tg 9–2 and Tg 10–1 mice reached 1100 and 5200 AU, respectively (Fig. 1). Preproghrelin mRNA levels in other tissues of Tg 9–2 and Tg 10–1 mice also exceeded those of control mice.
FIG. 1. Preproghrelin mRNA levels in the tissues of control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice quantified by real-time PCR analysis. Lanes 1, stomach; 2, small intestine; 3, cerebrum; 4, hypothalamus; 5, pituitary; 6, liver; 7, kidney; 8, lung; 9, heart; and 10, skeletal muscle.
Total and acylated ghrelin levels in tissues and plasma
Eight-week-old control, Tg 9–2, and Tg 10–1 mice were used (Table 2). Although high total ghrelin levels were detected in the stomach, only very low levels were detected in other tissues of control mice. Tg 9–2 and Tg 10–1 mice showed significantly higher total ghrelin levels in the stomach than control mice (P < 0.01 for each). Tg 9–2 and Tg 10–1 mice also showed total ghrelin levels in all of the other tissues significantly higher than control mice. High levels of acylated ghrelin were also detected in the stomach of control, Tg 9–2, and Tg 10–1 mice. There was, however, no significant difference between control and Tg 9–2 mice and between control and Tg 10–1 mice. Only very low acylated ghrelin levels if any were detected in other tissues of control, Tg 9–2, and Tg 10–1 mice. Plasma total ghrelin levels in control, Tg 9–2, and Tg 10–1 mice were 1104.5 ± 94.4, 11230.6 ± 1147.1, and 48565.5 ± 9291.5 fmol/ml, respectively. Those in Tg 9–2 and Tg 10–1 mice were significantly higher than those in control mice (P < 0.01 for each). Plasma acylated ghrelin levels in control, Tg 9–2, and Tg 10–1 mice were 83.7 ± 11.9, 79.7 ± 10.1, and 86.3 ± 21.1 fmol/ml, respectively. The differences between control and Tg 9–2 mice and control and Tg 10–1 mice were not significant.
TABLE 2. Total and acylated ghrelin levels in plasma and tissues of 8-wk-old control and transgenic mice (n = 8/group)
Body weights and lengths, relative organ weights, and BMIs
Body weights of control, Tg 9–2, and Tg 10–1 mice are shown in Table 3 and Fig. 2A. Male Tg 9–2 and Tg 10–1 mice were significantly lighter in the body weight than control mice (P < 0.05 and P < 0.01, respectively). Female Tg 10–1 mice were also significantly lighter than control mice (P < 0.01). The difference between female control and Tg 9–2 mice was not significant. Fifteen-week-old male and female Tg 10–1 and male Tg 9–2 mice were still significantly lighter than control mice (P < 0.05, P < 0.01, and P < 0.01, respectively). Body lengths (nose-to-anus lengths) of control and transgenic mice are shown in Table 3. Eight-week-old male Tg 9–2 and Tg 10–1 mice were significantly shorter in the body length than control mice (P < 0.05 and P < 0.01, respectively). Female Tg 10–1 mice were significantly shorter than control mice (P < 0.01). The difference between female control and Tg 9–2 mice was not significant. BMIs were calculated from the body weights and lengths. No significant difference was noted between control and Tg 9–2 mice and control and Tg 10–1 mice (Table 3). Fifty-two-week-old male Tg 9–2 and Tg10–1 mice were still significantly lighter and shorter, compared with control mice (Table 3), and no significant difference was noted in BMIs between control and transgenic mice. Relative organ weights of 8-wk-old male control and Tg 10–1 mice were calculated from the organ and body weights. No significant difference was noted between control and Tg 10–1 mice (Fig. 2B). No significant difference was noted in the pituitary size between control and Tg 10–1 mice (0.058 ± 0.002 and 0.055 ± 0.003 mg/body weight (grams), respectively).
TABLE 3. Body weights, lengths, and BMIs of 8-wk-old and 52-wk-old control and transgenic mice (n = 8/group)
FIG. 2. Body weights (BW) and relative organ weights. A, Body weights of male (left panel) and female (right panel) control (triangles), Tg 9–2 (circles), and Tg 10–1 (squares) mice (n = 8/group). B, Relative organ weights of 8-wk-old control (closed bars) and Tg 10–1 (open bars) mice calculated from the organ and body weights (n = 8/group). 1, stomach; 2, cerebrum; 3, heart; 4, liver; 5, kidney; 6, spleen; 7, pancreas; 8, epididymal fat. a, P < 0.05; b, P < 0.01 (vs. control mice).
Immunohistochemical staining for total and acylated ghrelin of the pituitary
Immunohistochemical staining for total and acylated ghrelin is shown in Fig. 3, A–D. None of total ghrelin-positive cells were observed in the pituitary of control mice (Fig. 3A). On the other hand, many total ghrelin-immunoreactive pituitary cells were observed in Tg 10–1 mice (Fig. 3B). Approximately 30% of the anterior pituitary cells in all sections examined were total ghrelin immunoreactive. None of acylated ghrelin-positive cells were observed in the pituitary of either control or Tg 10–1 mice (Fig. 3, C and D).
FIG. 3. The localization of total and acylated ghrelin-immunoreactive cells in the pituitary of 8-wk-old male control (A and C) and Tg 10–1 (B and D) mice. A and B, An antiserum raised to ghrelin(13–28) recognizing total (acylated plus des-acyl) ghrelin was used. C and D, An antiserum raised to ghrelin(1–11) specifically recognizing acylated ghrelin was used. Original magnification, x40. The immunoreactive cells are stained brown by the avidin-biotin complex methods.
Food intake and biochemical parameters in the blood
Although absolute amounts of daily food intake were reduced in Tg 9–2 and Tg 10–1 mice, the amounts per body weight were not significantly changed in either male or female Tg 9–2 or Tg 10–1 mice, compared with control mice (Table 4). No significant differences in blood glucose, serum total protein, total cholesterol, and insulin levels were noted between 8-wk-old control and Tg 9–2 mice and control and Tg 10–1 mice (Table 4).
TABLE 4. Daily food intake, blood glucose, serum total protein, total cholesterol, and insulin levels in 8-wk-old control and transgenic mice (n = 8/group)
Serum GH, IGF-1, and pituitary GH mRNA levels
Serum GH levels in male control, Tg 9–2, and Tg 10–1 mice were 5.5 ± 1.9, 3.7 ± 0.7, and 2.3 ± 0.9 ng/ml, respectively (Fig. 4A). Those in female control, Tg 9–2, and Tg 10–1 mice were 4.7 ± 1.7, 2.5 ± 0.9, and 1.7 ± 0.8 ng/ml, respectively (Fig. 4A). There were tendencies for decline in serum GH levels in male and female Tg 9–2 and Tg 10–1 mice, compared with control mice, although the differences between them were not significant. Serum IGF-I levels in male control, Tg 9–2, and Tg 10–1 mice were 522 ± 23.6, 413.2 ± 49.0, and 364.1 ± 25.6 ng/ml, respectively (Fig. 4B). Those in male Tg 9–2 and Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.01 for each). Serum IGF-I levels in female control, Tg 9–2, and Tg 10–1 mice were 509.7 ± 43.1, 545.5 ± 64.1, and 253.7 ± 36.4 ng/ml, respectively (Fig. 4B). Those in female Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.01). The difference between female control and Tg 9–2 mice was not significant.
FIG. 4. Serum GH, IGF-I, and pituitary GH mRNA levels in 8-wk-old control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice (n = 8/group). A, Serum GH levels. B, Serum IGF-I levels. C, Pituitary GH mRNA levels. a, P < 0.05; b, P < 0.01 (vs. control mice).
Pituitary GH mRNA levels in male control, Tg 9–2, and Tg 10–1 mice were 1.00, 0.62, and 0.42 AU, respectively. Those in Tg 9–2 and Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.05 and P < 0.01, respectively). Pituitary GH mRNA levels in female control, Tg 9–2, and Tg 10–1 mice were 1.00, 0.97, and 0.71 AU. Those in female Tg 10–1 mice were significantly reduced, compared with those in control mice (P < 0.05). The difference between those in female control and Tg 9–2 mice was not significant (Fig. 4C).
Plasma ACTH, serum TSH, LH, and FSH levels
Plasma ACTH, serum TSH, LH, and FSH levels in 8-wk-old in male control and transgenic mice are shown in Table 5. No significant difference was noted in the levels between control and transgenic mice.
TABLE 5. Plasma ACTH, serum TSH, LH, and FSH levels of 8-wk-old control and transgenic mice (n = 8/group)
Hematoxylin eosin and immunohistochemical staining for GH of the pituitary
Hematoxylin eosin staining is shown in Fig. 5, A and B. The pituitary morphology of Tg 10–1 mice was not different from that of the control mice. Immunohistochemical staining for GH is shown in Fig. 5, C and D. The distribution of GH-immunoreactive cells in the pituitary of Tg mice was similar to that of control mice.
FIG. 5. Morphology of the pituitary and the localization of GH-immunoreactive cells in the pituitary of 8-wk-old male control (A and C) and Tg 10–1 (B and D) mice. A and B, Hematoxylin eosin (HE) staining. C and D, The localization of total and GH-immunoreactive cells in the pituitary. Original magnification, x40. The immunoreactive cells are stained brown by the avidin-biotin complex methods.
Effects of GHRH and ghrelin on GH release
Control and Tg 10–1 mice were used. Serum GH levels after GHRH administration in male and female Tg 10–1 mice were similar to those of control mice throughout the course of the experiment (Fig. 6A). There was no significant difference in serum GH level at each time point between both male and female Tg-10 and control mice. Serum GH levels 10 min after ghrelin administration in male Tg10–1 and control mice were 63.1 ± 6.8 and 72.6 ± 12.0 ng/ml, respectively (Fig. 6B, left panel). The difference was not significant. Serum GH levels 20 min after ghrelin administration in male Tg10–1 and control mice were 30.2 ± 6.7 and 61.2 ± 15.5 ng/ml, respectively, and levels after 30 min were 11.8 ± 1.4 and 21.9 ± 4.1 ng/ml, respectively (Fig. 6B, left panel). Both differences were significant (P < 0.01). Serum GH levels 10 min after ghrelin administration in female Tg10–1 and control mice were 8.7 ± 3.7 and 52.8 ± 8.2 ng/ml, respectively, and those after 20 min were 29.8 ± 6.3 and 78.5 ± 14.3 ng/ml, respectively (Fig. 6B, right panel). Both differences were significant (P < 0.01). Serum GH levels 30 min after ghrelin administration in female Tg10–1 and control mice were 22.8 ± 6.3 and 22.3 ± 8.8 ng/ml, respectively (Fig. 6B, right panel). The difference was not significant.
FIG. 6. The responses of GH to GHRH and ghrelin in 8-wk-old control (closed circles) and Tg 10–1 (open triangles) mice. A, Time course of serum GH levels after iv injection of 60 μg/kg GHRH (n = 8/each point). B, Time course of serum GH levels after iv injection of 40 μg/kg ghrelin (n = 8/each point). a, P < 0.01 (vs. control mice).
Expression of GHS-R in the pituitary
GHS-R mRNA levels of male control, Tg 9–2, and Tg 10–1 mice were 1.00, 1.56, and 3.46 AU, respectively (Fig. 7). The difference between control and Tg 10–1 mice was significant (P < 0.01).
FIG. 7. Pituitary GHS-R mRNA levels in 8-wk-old control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice quantified by real-time PCR analysis (n = 8/group). a, P < 0.01 (vs. control mice).
Expression of hypothalamic neuropeptides that regulate GH secretion
GHRH mRNA levels of male control, Tg 9–2, and Tg 10–1 mice were 1.00, 0.88, and, 0.80 AU, respectively (Fig. 8A). The differences between control and Tg 9–2 mice and control and Tg 10–1 mice were not significant. Somatostatin mRNA levels of male control, Tg 9–2, and Tg 10–1 mice were 1.00, 1.08, and 0.97 AU, respectively (Fig. 8B). The differences between control and Tg 9–2 mice and control and Tg 10–1 mice were not significant.
FIG. 8. Hypothalamic GHRH and somatostatin mRNA levels in 8-wk-old control (closed bars), Tg 9–2 (shaded bars), and Tg 10–1 (open bars) mice quantified by real-time PCR analysis. A, GHRH mRNA levels (n = 8/group). B, Somatostatin mRNA levels (n = 8/group).
Effects of continuous infusion of des-acyl ghrelin on GH-IGF-I axis and body weights
Male and female C57BL/6 mice were used. Serum GH levels after 10 d treatment with saline and des-acyl ghrelin in male mice were 5.8 ± 1.1 and 7.5 ± 2.0 ng/ml, respectively. The difference was not significant. Those with saline and des-acyl ghrelin in female mice were 9.2 ± 2.2 and 9.5 ± 1.8 ng/ml, respectively. The difference was not significant either. Serum IGF-I levels after 10 d treatment with saline and des-acyl ghrelin in male mice were 769.3 ± 16.6 and 768.7 ± 21.6 ng/ml, respectively. The difference was not significant. Those with saline and des-acyl ghrelin in female mice were 766.2 ± 13.4 and 719.4 ± 49.1 ng/ml, respectively. The difference was not significant either. Body weights and lengths in des-acyl ghrelin-injected mice were not significantly different from those in saline-injected mice in either males or females (data not shown).
Discussion
We have generated transgenic mouse lines that overexpress preproghrelin mRNA in a wide variety of tissues. The wide tissue distribution of preproghrelin mRNA in transgenic mice was consistent with previous reports on transgenic mice using the CAG promoter (33, 34). Preproghrelin mRNA expression was increased, especially in Tg 10–1 mice, and its amount in the stomach reached 52-fold of that in control mice. Consistent with the elevated mRNA expression, peptide levels of total ghrelin (des-acyl plus acylated ghrelin) in various tissues were also elevated in transgenic mice. Plasma total ghrelin levels in transgenic mice showed marked results. Those in transgenic mice showed 10- and 44-fold of those in control mice. We originally intended to generate mice overexpressing biologically active ghrelin. Unexpectedly, acylated ghrelin levels were not changed in all tissues examined and plasma of transgenic mice, compared with those of control mice, indicating that transgenic mice overexpress only des-acyl ghrelin. The expression of acylated ghrelin has been reported in a small number of tissues, such as the stomach (X/A cells), duodenum, hypothalamus, and pancreatic -cells (1, 31, 39, 40). These reports and our present data suggest that only a limited number of cell lineages may able to process proghrelin or acylate ghrelin. The underlying mechanism by which ghrelin is acylated is unknown to date. Further study is needed to clarify the mechanism of the acylation.
The acylation of ghrelin is assumed to be essential for its actions, and des-acyl ghrelin, which lacks the modification, is devoid of endocrine actions, based on previous studies (1, 41). However, recent studies indicated that des-acyl ghrelin may have some actions. Des-acyl ghrelin as well as acylated ghrelin causes a significant inhibition of cell proliferation in human breast carcinoma cell lines (29) and inhibits cell death in cardiomyocytes and endothelial cells through ERK1/2 and phosphatidylinositol 3-kinase/AKT (30). In addition, one study (42) reported that acylated and des-acyl ghrelin promote adipogenesis directly in vivo by a mechanism independent of known GHS-Rs. Moreover, another study (28) indicated that des-acyl ghrelin may offset the action of acylated ghrelin on insulin secretion. Ghrelin has been shown to induce a reduction in serum insulin levels. In the study, coadministration of acylated plus des-acyl ghrelin did not result in any changes in serum insulin levels in humans, suggesting that ghrelin action on insulin is modulated by des-acyl ghrelin.
The present study indicates that transgenic mice overexpressing des-acyl ghrelin show small phenotype. Longitudinal growth was the most reduced in female Tg 10–1 mice (20% reduction from control mice). The phenotype was not associated with changes in BMIs. These mice did not show decreased food intake or decreased body fat mass. In addition, they showed normal nutritional condition, based on their biochemical parameters, including blood glucose, serum total protein, and total cholesterol levels. These data indicate that the small phenotype of transgenic mice is not attributed to poor nutritional condition.
Serum IGF-I levels were significantly reduced in male and female transgenic mice, compared with control mice. Female Tg 10–1 mice had no less than 50% reduction in serum IGF-I levels, compared with control mice. Although the differences in serum GH levels between control and transgenic mice were not statistically significant, probably because of the pulsatile character of GH secretion, the levels tended to be reduced in transgenic mice, compared with control mice, and the mean GH level of Tg10–1 mice was only 50% of that of control mice. It should be emphasized that Tg 10–1 mice showed lower serum GH levels than Tg 9–2 mice. Body weights and lengths of the former were more reduced than the latter. It should be also noted that the former showed higher des-acyl ghrelin expression than the latter. Reduced pituitary GH mRNA levels in transgenic mice support the observation. The GH-IGF-I axis-specific alteration in transgenic mice was also indicated by the measurement of other anterior pituitary hormones than GH. Plasma ACTH, serum TSH, LH, and FSH levels were not altered.
The size and morphology of the pituitary including the somatotrope populations of transgenic mice were similar to those of control mice. These data indicated that there is no apparent change, suggesting developmental problems in the pituitary of transgenic mice.
Responses of GH to GHRH and ghrelin in transgenic mice exhibited intriguing results. Transgenic mice showed normal response of GH to GHRH. Alternatively, if we consider that the basal GH levels are lower in transgenic mice, the similar maximal response might indicate that they are hyperresponsive to GHRH. It is not likely that an insufficient dose of GHRH induced submaximal response of GH in both control and transgenic mice, judging from previous reports (43). On the other hand, the responses of GH to ghrelin were reduced in transgenic mice. It is noteworthy that the reduction was much greater in female transgenic mice than in male mice, if we take their serum IGF-I levels into account. Taken together our results and these reports indicate that overexpression of des-acyl ghrelin in our mice may result in reduction of GH response to endogenous ghrelin, and it may result in the reduced serum IGF-I levels in transgenic mice.
The reduced GH response to ghrelin in transgenic mice could be due to down-regulated the GHS-R. However, the pituitary GHS-R mRNA levels in the transgenic mice were rather elevated. It is not likely that overexpressed des-acyl ghrelin acts as a blocking agent to the GHS-R because 125I-labeled acylated ghrelin bound to the GHS-R cannot be displaced by des-acyl ghrelin (20). Overexpressed des-acyl ghrelin may have some effects on endogenous GH secretion, modifying the action of endogenous ghrelin in transgenic mice via, for instance, another receptor or modulation of the signal transduction pathway after the GHS-R.
Previous reports indicated that the hypothalamus plays a critical role in the stimulatory effect of ghrelin on GH secretion as well as the pituitary (21, 22, 23). Because GH secretion is regulated chiefly by two hypothalamic hormones, GHRH and somatostatin, the expression of these hormones could be altered in transgenic mice. We could not find any significant difference in either GHRH or somatostatin mRNA levels between control and transgenic mice. These data might suggest that overexpressed des-acyl ghrelin acts on not only the pituitary but also the hypothalamus in the transgenic mice, judging from the fact that hypothalamus GHRH mRNA were not elevated, and somatostatin mRNA levels were not decreased despite the decreased serum GH levels.
We could not show, unfortunately, that continuous ip infusion of des-acyl ghrelin has some effect on serum GH and IGF-I levels or body weights. It should be noted, however, that plasma des-acyl ghrelin levels in transgenic mice reached 10- and 50-fold of those in control mice. Administration of a higher dose of des-acyl ghrelin, or longer administration, might result in alteration in the GH-IGF-I axis. On the other hand, the phenotype of transgenic mice might reflect direct effects of ubiquitous expression of des-acyl ghrelin. It should also be noted that high levels of des-acyl ghrelin were detected in a various tissues, especially in the pituitary, as well as in plasma of transgenic mice. The des-acyl ghrelin immunoreactive pituitary cells might play an important role in the mechanism for the altered GH-IGF-I axis in a paracrine or autocrine manner. It should be pointed out that preproghrelin mRNA is reported to be expressed in the normal pituitary (44), as we showed in the present study, suggesting its physiological role in GH secretion. The phenotype of transgenic mice may reflect the role. Further study is needed for this issue.
The mechanism underlying the sexual dimorphism in the responses of GH to ghrelin in transgenic mice is not fully understood. It might be due to the gender difference in the secretory regulation of GH. Female mice have been reported to be different from male mice in that they have noncyclical and rather low somatostatin output and that GHRH plays a dominant role in it (45). There might be a GHRH-dependent mechanism for the reduced response in transgenic mice. Indeed, one recent report (26) indicated that transgenic rats expressing an antisense GHS-R mRNA in the hypothalamic arcuate nucleus show marked gender difference in GH secretion. Although there was no significant difference in pulse frequency and baseline levels of GH between male control and transgenic rats, female transgenic rats showed lower baseline levels and fewer pulses of GH than female control rats (26).
The 94-amino acid proghrelin is cleaved to yield ghrelin. One previous study (46) demonstrated that C-terminal proghrelin peptides are present in the human circulation. Transgenic mice in the present study would also overexpress these peptides. We have not excluded the possibility that the phenotype of transgenic mice might be due to the effects of these peptides.
In conclusion, the present study demonstrates that transgenic mice overexpressing des-acyl ghrelin show small phenotype and altered GH-IGF-I axis. These observations may indicate a role of des-acyl ghrelin in the regulation of GH secretion.
Acknowledgments
The authors gratefully acknowledge the excellent technical support of Chieko Ishimoto and Hitomi Hiratani.
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