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. 2024 Oct 21;102:skae318. doi: 10.1093/jas/skae318

D-aspartate stimulates growth hormone secretion in wethers

Tatsuyuki Takahashi 1,✉, Kyosuke Kidachi 2, Mikiko Yukawa 3, Tomoki Hachinohe 4, Yuina Takashima 5, Mao Fujimura 6, Atsuko Saito 7, Daichi Soga 8, Chihiro Ota 9, Eri Niizuma 10, Katsuyoshi Sato 11, Hideki Ogasawara 12, Yohei Kurose 13
PMCID: PMC11630845  PMID: 39432441

Abstract

Growth hormone (GH) is an essential factor in enhancing the productivity of animals. In ruminants, L-aspartate (L-Asp) stimulates the secretion of GH; however, the effect of D-Asp on GH remains unknown. Here, we examined the effect of D-Asp on GH secretion in wethers. Blood GH, insulin, adrenaline, noradrenaline, non-esterified fatty acid (NEFA), and glucose concentrations were evaluated in response to the intravenous infusion of a high-dose (0.1 mmol/kg/min) of D-Asp for 20 min. Further, concentrations of these biomolecules were evaluated when a low-dose (0.05 mmol/kg/min) of D-Asp was continuously infused intravenously for 20 min. Finally, the direct effect of D-Asp on GH secretion was determined using cultured sections of the anterior pituitary tissue from wethers. Infusion of the high-dose of D-Asp markedly increased blood GH concentrations (P < 0.05), resulting in an increase in the area under the curve (AUC). Plasma GH concentrations and AUC also increased in response to infusion of a low D-Asp dose. Infusion of a high and low D-Asp dose caused a prolonged reduction in plasma insulin concentrations, and the AUC was lower (P < 0.05). Plasma NEFA concentrations gradually increased after the end of D-Asp infusion, with a low D-Asp dose infusion resulting in significantly higher concentrations at 90 min (P < 0.05). Plasma adrenaline, noradrenaline, and glucose concentrations did not show significant changes despite differences in the dose of D-Asp. Although D-Asp treatments stimulated GH secretion in the cultured sections of pituitary tissues, the effect was not significant. These results suggest that D-Asp stimulates the secretion of GH in wethers through not only a direct action on the pituitary gland but also through another pathway of GH stimulation.

Keywords: D-aspartate, growth hormone, pituitary gland, ruminants, wethers

Growth hormone level is closely associated with enhanced productivity in ruminants. Hence, we investigated the beneficial effects of D-aspartate on GH secretion and its mechanism of action.

Graphical Abstract

Graphical Abstract.

Graphical Abstract

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Introduction

The secretion of growth hormone (GH) is primarily regulated by the coordinated actions of growth hormone-releasing hormone (GHRH) and somatostatin (SST), which is released from the hypothalamus. GHRH stimulates GH secretion via binding to GHRH receptors expressed in the somatotrophs of the anterior pituitary, and somatostatin suppresses the GH secretion via binding to SST receptor types 2 and 5. GH promotes body growth through the regulation of longitudinal bone growth (Ohlsson et al., 1998) and protein synthesis (muscle development) (Moller et al., 2009). It additionally facilitates glycogenesis and glycogenolysis in the liver and kidneys and lipolysis in adipose tissue (Møller and Jørgensen, 2009; Vijayakumar et al., 2010). In ruminants, GH acts directly and/or indirectly on the mammary gland to enhance milk production (Hull and Harvey, 2001; Svennersten-Sjaunja and Olsson, 2005). Therefore, the control of GH secretion is a key factor in improving productivity in ruminants.

Amino acids (AA) are the building blocks of proteins. They also exist in a free state in intracellular fluid and blood plasma, and maintain homeostasis through regulating specific physiological functions, such as regulation of neurotransmitters (Zhou and Danbolt, 2014), immune (Li et al., 2007), and gut barrier function (Wang et al., 2009) and improvement of metabolic disease (Wu, 2009). Additionally, AA act on the endocrine organs that control hormone secretion associated with energy metabolism (Reiter et al., 1990; Villalobos et al., 1997; Li et al., 2004; Yang et al., 2012; Alamshah et al., 2017; Modvig et al., 2021). Notably, the administration of high-dose L-arginine (L-Arg) induces GH secretion via inhibiting endogenous SST release in humans and rats (Alba-Roth et al., 1988), and elevates GH mRNA expression levels via activating nitric oxide (NO)/NO synthase (NOS) signaling pathway in rats (Olinto et al., 2012). In sheep, blood GH concentrations increase by intracarotid continuous infusion of L-Arg (Davis, 1972), whereas L-aspartate (L-Asp) is the most potent stimulator of GH secretion among the 17 AA studied, including L-Arg (Kuhara et al., 1991). Ohata et al. (1997) have observed that L-Asp slightly, but significantly, causes GH secretion in goat primary anterior pituitary cells. Therefore, although types of AA that stimulate GH secretion differ among animal species, L-Arg, and L-Asp are potent stimulators of GH secretion.

The physiological function of L-type AA has been studied; however, AA also exists in a D-type form. D-AA has various sources such as intestinal bacteria, food, racemization, or biosynthesis. The physiological effects of D-type AA are being reported. Recently, D-type AA produced by intestinal bacteria was shown to regulate the host gut immune system (Suzuki et al., 2021). D-serine (D-Ser) is abundant in the central nervous system (CNS) of rodents and humans, suggesting that it plays an important role in the regulation of brain function (Kiriyama and Nochi, 2016). D-aspartate (D-Asp) has been detected in the CNS; it is also abundant in endocrine organs such as the pituitary gland, pineal gland, and testis (Imai et al., 1995; D’Aniello et al., 2000; Topo et al., 2009), and is associated with the regulation of hormone secretion. Intraperitoneal administration of D-Asp results in the elevation of GH, luteinizing hormone (LH), testosterone, and progesterone concentrations in the blood of rats (D’Aniello et al., 2000). Similar to that in rodents, D-Asp level is high in the pituitary and pineal glands of sheep, and subcutaneous administration of D-Asp leads to increased blood LH concentrations (Boni et al., 2006). As D-Asp stimulates GH secretion in rats, it is expected that D-Asp, similar to L-Asp, stimulates GH secretion in ruminants. D-Asp induced-GH secretion has, however, not been reported yet in ruminants.

We hypothesized that D-Asp induces GH secretion in sheep by acting directly on the pituitary gland. This study aimed to investigate the effects of D-Asp on GH secretion through intravenous infusion and experiments with cultured sections of the anterior pituitary gland in sheep.

Materials and Methods

All the experiments performed in this study were approved by the President of Kitasato University through the judgment of the Institutional Animal Care and Use Committee of Kitasato University (Approval number: #20-020). Wethers used in the experiment were obtained from the Field Science Center, Towada Farm, School of Veterinary Medicine, Kitasato University (Japan).

Short-term infusion test

A total of 4 wethers (aged 1 year, 49.2 ± 0.2 kg BW) were used for the infusion test. The animals were kept in metabolic cages in a temperature-controlled room at 20 ± 2 °C under a 12:12 light-dark cycle during the infusion experiment. The sheep were fed a mixture feed of hay cubes and timothy hay for 0.65 times the energy maintenance requirement based on the Japanese Feeding Standard for Sheep (1996) twice a day (total 1.3 times amount/day) (Agriculture, Forestry and Fisheries Research Council Secretariat, 1996), and water and mineral salts were consumed ad libitum. On the day of the infusion test, the wethers were fitted with indwelling catheters (SurFlash Polyurethane I.V. Catheter 18G × 2 1/2; TERUMO corporation) in the jugular vein and rested for at least 1 h before the experiment was conducted. Two different approaches were considered—high and low-dose infusion. The experimental groups for the high-dose infusion comprised saline (control), D-Asp (0.1 mmol/kg BW/min), and L-Asp (0.1 mmol/kg BW/min). The trial was designed as 4 × 3 incomplete Latin squares. After a two-month recovery period, we examined the effects of a low D-Asp dose using the same animals that had been used in the high-dose infusion experiment. The experimental groups here comprised saline (control) and D-Asp (0.05 mmol/kg BW/min). The trial was designed as a crossover study. Animals were administered the fluid continuously for 20 min using peristaltic pumps and blood samples were obtained over time from pre- to post-infusion. The injection start time was set as “0 minutes” and blood was drawn at −30, −20, −10, 0, 10, 20, 30, 40, 50, 60, and 90 min. The blood samples were then transferred to tubes containing 10 U/mL heparin and 100 KIU/mL aprotinin, mixed thoroughly by gentle inversion, and then were placed on ice. Plasma samples were obtained by centrifugation (1,970×g, 20 min, 4 °C) and stored at −30 °C until analysis. Here, 0.5 M D-Asp (FUJIFILM Wako Pure Chemical Corporation, Japan) or L-Asp (FUJIFILM Wako Pure Chemical Corporation, Japan) solution was adjusted to pH 7.4 using NaOH and HCl was prepared and used after filter sterilization (0.22 μm; Merck Millipore Ltd., USA).

Pituitary tissue culture

A total of 8 wethers (aged 1–3 years, 49.6 ± 2.9 kg BW) were used. The animals were kept in individual pens under natural photoperiodic and thermoperiodic conditions. Feeding management was the same as in experiment 1. Animals were administered xylazine (0.1 mg/kg BW) and sodium pentobarbital (20 mg/kg BW) via intravenous injection and were euthanized under deep anesthesia by exsanguination through the carotid artery. After euthanasia, the pituitary gland was immediately collected and placed in a cold artificial cerebrospinal fluid (aCSF) solution (NaCl 125 mM, KCl 2.5 mM, CaCl2 2.0 mM, MgCl2 1.0 mM, NaH2PO4 1.25 mM, NaHCO3 25 mM, glucose 5.6 mM, pH7.4) containing penicillin and streptomycin (NACALAI TESQUE, INC, Japan). The entire pituitary gland was aerated with 95% O2/5% CO2 for 15 min and subsequently 150–200-μm thick sections were obtained under cold aCSF using a microslicer (DTK-1000, DOSAKA EM CO., LTD, Japan). Each section was transferred to a netwell (Corning, USA) and pre-cultured with an aCSF solution for 1 h in a 37 °C-5% CO2 incubator. After pre-incubation, the medium was collected from each well, and then the sections were treated with D-Asp (0.1, 1.0, 10 mM), L-Asp (0.1, 1.0, 10 mM), or GHRH (1.0 μM) (LKT laboratories Inc, USA) for 1 h. After stimulation, the media in which sections were incubated were collected and stored at −80 °C until analysis.

Hormone and metabolites analysis

The concentrations of GH in plasma and culture media were determined using time-resolved fluorescence immunoassay (TR-FIA) according to a method used in a previous report with slight modification (Sugino et al., 2002). Rabbit anti-ovine growth hormone polyclonal antibody was added to wells coated with anti-rabbit IgG as primary antibody (1:50,000, NIDDK, USA) diluted in assay buffer (50 mM Tris, 145 mM NaCl, 20 mM DTPA, 0.05 % NaN3, 0.01% Tween40, 0.5% BSA, 0.05% γ-globulin, 0.01 mg/mL phenol red, pH 7.8) and incubated for 18 h at 24 °C. Samples or standards (ovine growth hormone, NIDDK, USA) were added to each well and incubated for 20 h at 24 °C. The samples were then treated with Eu-labeled ovine growth hormone for 3 h at 24 °C. Finally, enhancement solution (PerkinElmer Japan Co., Ltd, Kanagawa, Japan) was added to the wells and the plates were incubated for 5 min at 24 °C. Eu fluorescence was measured using a multi-label plate reader (ARVO, PerkinElmer Japan Co., Ltd, Kanagawa, Japan). The intra- and inter-assay coefficient of variation for the GH concentrations were 7% and 11%, respectively. All samples were analyzed in triplicate.

Plasma insulin concentrations were also determined using TR-FIA according to a protocol in a previous report with slight modification (Takahashi et al., 2006). Guinea pig anti-bovine insulin polyclonal antibody was added to wells coated with anti-guinea pig IgG as primary antibody (1:50,000, Immundiagnostik, Germany) diluted in assay buffer (composition given above) and incubated for 17 h at 24 °C. Samples or standards (bovine insulin, Sigma, USA) were loaded into each well and incubated for 20 h at 4 °C. The samples were then treated with Eu-labeled bovine insulin for 2 h at 4 °C. Enhancement solution treatment and Eu fluorescence measurement were carried out as stated above for GH assay. The intra- and inter-assay coefficient of variation for the insulin concentrations were 5% and 6%, respectively. All samples were analyzed in triplicate.

The plasma glucose and NEFA concentrations were measured using commercially available kits (Glucose C-II Test-Wako; Wako Pure Chemical Co., Tokyo, Japan) (NEFA C Test-Wako; Wako Pure Chemical Co., Tokyo, Japan). All samples were analyzed in triplicate.

Preparation for catecholamine analysis

To determine plasma catecholamine concentrations, the samples were adjusted as described previously (Shimizu et al., 2006). Briefly, a total of 100 μL of 0.1 M EDTA-2Na solution, 1 mL of 1.5 M Tris buffer (pH8.6), 30 mg of activated alumina, and 1 ng of 2, 5-dihydroxybenzoic acid (DHBA) were added as internal control to 500 μL of plasma. The samples were shaken for 10 min using a direct mixer. They were centrifuged at 3,000 rpm, 3 min, 4 °C, and the supernatant was removed. The pellet was then washed with ultra-pure water and centrifuged at 3,000 rpm, 3 min, 4 °C. Again, the supernatant was removed, and the washing process was repeated 3 times. The samples were then mixed with 200 μL of 2% acetate solution containing 100 μM EDTA-2Na and shaken for 10 s. The samples were then incubated for 10 min at 24 °C, the supernatants were obtained by centrifugation at 4,300 rpm, 5 min, 4 °C, and were stored at −80 °C until analysis.

HPLC

Catecholamine (adrenaline and noradrenaline) concentrations were measured using high-performance liquid chromatography (HPLC) and electrochemical detection (HTEC-500, Eicom, Japan) according to our report with slight modification (Kurose and Terashima., 1999). The analytical conditions were as follows: detector, +450 mV potential against an Ag/AgCl reference electrode; column, Eicompak SC-5ODS (2.1 × 150 mm); mobile phase, 0.1 M phosphate buffer (pH 6.0) containing 50 mg/L disodium EDTA, 600 mg/L sodium 1-octanesulfonate, and 12% methanol; flow rate 0.23 mL/min. The specific retention time for each compound was determined using adrenaline and noradrenaline standards. The amount of catecholamine in each sample was calculated using the peak height ratio relative to that of 3, 4-dihydroxybenzylamine (DHBA) using PowerChrom v2.3 software (eDAQ Pty Ltd, NSW Australia).

Statistical analysis

All data are presented as mean ± SE. Plasma GH, insulin, adrenaline, and noradrenaline are expressed as a percentage rate of change from the baseline (−10 min) values. Statistical significance was defined as P < 0.05 and tendency was defined as 0.05 < P < 0.1. For the infusion experiment, data were analyzed using two-way repeated measure ANOVA. When there was a significant interaction, we conducted a Tukey-honestly significant difference (HSD) test among 3 groups or a paired t-test between 2 groups. For the total area under the curve (AUC) analysis, data were calculated using the trapezoidal method (−10 to 90 min) and evaluated using paired t-tests between 2 groups. For the anterior pituitary slice culture experiment, data were analyzed by using Dunnett’s test. All the statistical analyses were performed using the SPSS software (version 28, SPSS Institute, Inc., Chicago, IL, USA).

Results

Effects of short-term infusion of aspartate on blood GH, insulin, metabolites, and catecholamine concentrations

Infusion of a high-dose of D- and L-Asp (0.1 mmol/kg/min) presented a significant time × treatment interaction (P < 0.001); however, neither treatment altered blood GH concentrations at 10 min after infusion (Figure 1a). GH concentrations subsequently increased, with a significant increase in GH concentrations at 20- and 30-min after the start of the infusion compared with those in control (P < 0.05). Moreover, blood GH concentrations in the D-Asp treatment group were higher at 40 min (P = 0.092), whereas those in the L-Asp treatment group remained significantly elevated at 40 min compared with those in the control group (P = 0.035). Blood GH concentrations gradually declined thereafter and then returned to basal levels at 90 min. A significant time × treatment interaction was also observed in the low D-Asp dose (0.05 mmol/kg/min) group (P < 0.001) (Figure 1b). As with the high-dose infusion group, blood GH concentrations peaked at 20 min, and the GH concentrations in the D-Asp group tended to be higher at 30 min compared with those in the control group (P = 0.088). The concentrations declined and returned to basal concentrations at 50 min. The AUC of the high dose D- and L-Asp infusion was higher than that of the control group (P = 0.054 and P = 0.050, respectively) (Figure 1c). The AUC at the low-dose treatment was also higher for the D-Asp group than for the control group, but the difference was not significant. (Figure 1d).

Figure 1.

Figure 1.

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Change in plasma GH concentrations from the baseline with intravenous D-Asp infusion at high or low doses. The variation of plasma GH concentrations (a) and (b) from the baseline, and their AUC values (c) and (d) are shown. (a) and (c) Wethers were received with D-Asp (circle), L-Asp (triangle) at 0.1 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). (b) and (d) After a two-month recovery period, the same wethers received D-Asp (circle) at 0.05 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). Plasma GH concentrations are presented as a percentage rate of change from the −10 min values. (c) and (d) AUC for GH indicates the total GH change rate from −10 to 90 min based on the rate of change value. Gray layers indicate infusion time. Data are presented as mean ± SD. *: Control vs D-Asp (P < 0.05), #: Control vs L-Asp (P < 0.05), †: Control vs D-Asp (P < 0.10). The number of replications per treatment (n = 4).

The infusion of high-dose D-Asp did not show a significant time × treatment interaction (P = 0.084) for plasma insulin concentrations (Figure 2a). However, the infusion of D-Asp suppressed plasma insulin concentrations more rapidly and strongly than the infusion of L-Asp. D-Asp treatment maintained the suppression until the end of the experiment, whereas L-Asp treatment returned to approximately basal values in 40 min. The result demonstrated that the AUC of insulin in a high-dose D-Asp treatment group was significantly lower than those in the control group (P = 0.038) and tended to be lower than those in the L-Asp treatment group (P = 0.057) (Figure 2c). In the low-dose treatment group, the infusion of low D-Asp dose did not show a significant time × treatment interaction (P = 0.131) (Figure 2b). Additionally, the AUC was also lower than that of the control group, but the difference was not significant (Figure 2d).

Figure 2.

Figure 2.

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Change in plasma insulin concentrations from the baseline with intravenous D-Asp infusion at high or low doses. The variation of plasma insulin concentrations (a) and (b) from the baseline, and their AUC values (c) and (d) are shown. (a) and (c) Wethers received D-Asp (circle), L-Asp (triangle) at 0.1 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). (b) and (d) After a two-month recovery period, the same wethers received D-Asp (circle) at 0.05 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). Plasma insulin concentrations are presented as a percentage rate of change from the −10 min values. (c) and (d) AUC for insulin indicates the total insulin change rate from −10 to 90 min based on the rate of change value. Gray layers indicate infusion time. Data are presented as mean ± SD. *: Control vs D-Asp (P < 0.05). The number of replications per treatment (n = 4).

In terms of plasma adrenaline, there was no significant time × treatment interaction in the high-dose infusion tests (Figure 3a). AUC also showed no significant difference among D-Asp, L-Asp, and saline groups (Figure 3c). Even in the low-dose infusion tests, there was no significant time × treatment interaction in plasma adrenaline (Figure 3b). Additionally, AUC did not differ significantly from the control, similar to the high-dose treatment (Figure 3d). The results of the high-dose infusion tests in terms of plasma noradrenaline did not show a significant time × treatment interactions (Figure 4a). The AUC was not significantly different among D-Asp, L-Asp, and saline groups (Figure 4c). There was also no significant time × treatment interaction in plasma noradrenaline even in the low-dose treatment (Figure 4b). Although the AUC was significantly higher for D-Asp than for control (P = 0.013) (Figure 4d), noradrenaline concentrations did not exhibit any dose-dependent change in response to D-Asp treatment.

Figure 3.

Figure 3.

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Change in plasma adrenaline concentrations with intravenous D-Asp infusion at high or low doses. The variation of plasma adrenaline concentrations (a) and (b) from the baseline, and their AUC values (c) and (d) are shown. (a) and (c) Wethers received D-Asp (circle), L-Asp (triangle) at 0.1 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). (b) and (d) After a two-month recovery period, the same wethers received D-Asp (circle) at 0.05 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). Plasma adrenaline concentrations are presented as a percentage rate of change from the −10 min values. (c) and (d) AUC for adrenaline indicates total adrenaline change rate from −10 to 90 min based on the rate of change value. Gray layers indicate infusion time. Data are presented as mean ± SD. The number of replications per treatment (n = 4).

Figure 4.

Figure 4.

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Change in plasma noradrenaline concentrations from the baseline with intravenous D-Asp infusion at high or low doses. The variation of plasma noradrenaline concentrations (a) and (b) from the baseline, and their AUC values (c) and (d) are shown. (a) and (c) Wethers received D-Asp (circle), L-Asp (triangle) at 0.1 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). (b) and (d) After a two-month recovery period, the same wethers received D-Asp (circle) at 0.05 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). Plasma noradrenaline concentrations are presented as a percentage rate of change from the −10 min values. (c) and (d) AUC for noradrenaline indicates the total noradrenaline change rate from −10 to 90 min based on the rate of change value. Gray layers indicate infusion time. Data are presented as mean ± SD. *: Control vs D-Asp (P < 0.05). The number of replications per a treatment (n = 4).

There was no significant time × treatment interaction in blood glucose concentrations with either the high or low D-Asp dose treatment (Figure 5a and b). As with glucose concentrations, blood NEFA concentrations in the high-dose infusion group did not show a significant time × treatment interaction (Figure 5c). However, the low-dose infusion group demonstrated a significant time × treatment interaction (P < 0.001) (Figure 5d). Blood NEFA concentrations in the low-dose D-Asp treatment tended to be lower than those in the saline treatment at 20 min after the start of infusion (P = 0.073). However, they then gradually increased and continued to increase until the end of the experiment. The low-dose D-Asp group showed an increase in NEFA concentrations at 60 min (P = 0.072), whereas they were significantly higher at 90 min compared with those in the control (P = 0.048).

Figure 5.

Figure 5.

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Change in plasma glucose and NEFA concentrations with intravenous D-Asp infusion at high or low doses. The variation of plasma glucose (a) and (b) and NEFA (c) and (d) concentrations are shown. (a) and (c) Wethers received D-Asp (circle), L-Asp (triangle) at 0.1 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). (b) and (d) After a two-month recovery period, the same wethers received D-Asp (circle) at 0.05 mmol/kg/min, or saline (square) for 20 min by intravenous infusion (n = 4 in each group). Gray layers indicate infusion time. Data are presented as mean ± SD. *: Control vs D-Asp (P < 0.05). The number of replications per treatment (n = 4).

Effect of aspartate on the release of GH from the anterior pituitary.

Growth hormone-releasing hormone (GHRH) stimulated the release of GH from the anterior pituitary tissue sections, resulting in a significant increase of approximately 8-fold greater levels than those in the control (P < 0.01) (Figure 6). Further, the addition of 10 mM D-Asp induced an approximately 3-fold increase in GH release compared with that in control, but did not show a difference with 1 mM and 0.1 mM D-Asp addition. L-Asp had a marginal increase in GH release (approximately 1.5-fold compared with that of the control) in response to 1 mM and 10 mM addition.

Figure 6.

Figure 6.

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GH secretion in response to Asp stimulation in the cultured sections of pituitary glands from wethers. Sectioned sheep pituitary glands (n = 3–8) were cultured to examine GH secretion upon stimulation with 3 concentrations (0.1, 1.0, and 10 mM) of D- or L-Asp. Ovine GHRH (1.0 μM) was used as a positive control. Data are presented as mean ± SD. **: vs Control (P < 0.01). The number of replications per treatment (n = 3–8).

Discussion

In this study, we investigated the effect of D-Asp on the secretion of GH through in vivo infusion and ex vivo anterior pituitary section culture experiments. Consistent with the results of a previous study (Kuhara et al., 1991), the short-term intravenous infusion of L-Asp increased blood GH concentrations. The short-term infusion of D-Asp at the same dose resulted in clearly elevated blood GH level that peaked more rapidly than that in response to L-Asp. D’Aniello et al (2000) reported that a single bolus intraperitoneal injection of D-Asp induced an approximately 2.0-fold increase in GH and LH release in rats after 1 h of injection. Due to the short-term intravenous infusion and usage of higher doses of D-Asp in our experiments, a greater GH release, and a more rapid response were observed than those reported by D’Aniello et al (2000). Moreover, the elevation in GH level occurred in a dose-dependent manner. Taken together, these findings suggest that D-Asp has a potent stimulatory effect on GH secretion.

Growth hormone (GH) promotes body growth and increases milk production by increasing the secretion of insulin-like growth factor-I (IGF-I) via GH receptors expressed in the liver. GH and IGF-I are closely related to each other. However, in a previous report, L-Asp increased plasma GH concentrations but not plasma IGF-I concentrations (Kuhara et al., 1991). Therefore, in this experiment, the increase in plasma GH concentrations due to D-Asp did not likely induce an increase in plasma IGF-I concentrations.

To understand the mechanism of GH secretion after D-Asp infusion, we focused on the anterior pituitary gland, which is the primary source of GH secretion. We examined the direct effect of D-Asp on GH release from the pituitary gland. Contrary to expectations, D-Asp treatment only marginally stimulated GH release. Ohata et al. reported a marginal effect of L-Asp treatment on GH release from the primary pituitary cells of goat (Ohata et al., 1997). If D-Asp acts directly on the pituitary gland, GH secretion would increase immediately after the intravenous infusion. However, GH secretion was not elevated at 10 min after infusion, suggesting an indirect effect of Asp infusion on GH secretion. Moreover, D-Asp treatment of the hypothalamus and pituitary gland co-culture stimulates more GH secretion than that obtained with the treatment of the pituitary gland culture alone (D’Aniello et al., 2000). Therefore, these findings suggest that D-Asp is a potent facilitator of GH secretion through both direct action on the pituitary gland and indirect action on the hypothalamus via GHRH secretion. In addition to these pathways, regulation of GH secretion occurs via an adrenergic pathway, in which the intravenous infusion of noradrenaline suppresses GH secretion in ewes (Thomas et al., 1994). However, the adrenergic pathway may not be involved in Asp-induced increase in GH secretion, given that D- and L-Asp treatments did not alter plasma noradrenaline concentrations in our experiment.

In cattle, GH and insulin secretion are closely related, and GH acts on the pancreatic β-cells to stimulate insulin secretion (Feng et al., 2009), although their actions depend on nutritional status. We, therefore, proposed that the Asp-induced augmentation of GH secretion would result in more insulin secretion. However, insulin concentrations were sustainably low after D-Asp infusion, and the AUC of the D-Asp group was significantly lower than that of the control group. L-Asp infusion also reduced insulin concentrations, but the reduction was smaller for L-Asp than for D-Asp, and the AUC was not significantly different in L-Asp treatment compared with that for the control group. This suggests that the suppression of insulin concentrations by D-Asp acts independently of the elevation of GH secretion. In human follow-up studies, phenylalanine, tyrosine, alanine, aspartate, and glutamate have been reported to decrease insulin secretion and sensitivity (Vangipurapu et al., 2019). In contrast, intravenous and oral administration of glutamate elevated insulin secretion in rats (Bertrand et al., 1995) and glutamate stimulated insulin secretion in mouse pancreatic β-cell line MIN6 cells (Oya et al., 2011). It has recently been suggested that chronic supplementation of D-Ser diminishes insulin secretion from pancreatic β-cell via activation of the sympathetic nervous system (Suwandhi et al., 2018). In the present study, plasma adrenaline, and noradrenaline concentrations were not altered by D-Asp infusion. However, unaltered concentrations of these catecholamines do not rule out the possibility that the sympathetic nervous system is not associated with the suppression of insulin secretion in response to D-Asp. Alternatively, other inhibitory pathways for insulin secretion may have contributed. Further studies are needed to elucidate these pathways.

At both high and low infusion doses, NEFA concentrations gradually increased after the end of intravenous D-Asp infusion and were significantly higher at 90 min after infusion of a low D-Asp dose. GH can induce lipolysis, resulting in an elevation of blood NEFA concentrations (Eisemann et al., 1986); however, L-Asp treatment did not stimulate an increase in plasma NEFA concentrations, even though GH concentrations in the L-Asp treatment group were comparable to those in the D-Asp treatment group. This suggests that the mechanism of D-Asp induced-elevation of NEFA concentrations may be independent of the increase in GH secretion. As with insulin secretion, D- and L-types of aspartic acid may affect lipid metabolism differently. Further research is needed to elucidate the lipolytic effects (direct or indirect) of D-Asp.

In conclusion, D-Asp, similar to L-Asp, stimulates GH secretion and exerts its effects via pathways acting directly on the pituitary gland and via other pathways, such as its effect on GHRH neurons in the hypothalamus. Furthermore, D-Asp not only stimulates GH secretion, but also suppresses insulin secretion and increases NEFA release, and these effects are stronger for D-Asp than for L-Asp. However, the mechanisms underlying the inhibitory effects of D-As on insulin secretion and its facilitatory effects on NEFA release could not be elucidated in detail. Further research is thus needed to gain a comprehensive understanding of the effects of D-Asp. These findings suggest that D-Asp could be a novel GH-promoting factor to improve ruminant production.

Acknowledgments

We are very grateful to T. Katsuoka, T. Kado, R. Fujimoto, Y. Harashima, C. Tokigawa (Kitasato University School of Veterinary Medicine) for assistance with animal care, management, and sampling and statistical analysis. We would also like to thank Dr. K. Takagishi (Kitasato University School of Veterinary Medicine) for his advice on statistical analysis. This work was supported by JSPS KAKENHI Grant Number JP25892027.

Glossary

Abbreviations:

GH

growth hormone

D-Asp

D-aspartate

L-Arg

L-arginine

D-Ser

D-serine

AA

amino acids

NEFA

non-esterified fatty acid

AUC

area under the curve

GHRH

growth hormone-releasing hormone

SST

somatostatin

IGF-I

insulin like growth factor-I

NO

nitric oxide

NOS

NO synthase

CNS

central nervous system

LH

luteinizing hormone

TR-FIA

time-resolved fluorescence immunoassay

DHBA

dihydroxybenzoic acid

HPLC

high-performance liquid chromatography

HSD

honestly significant difference

Contributor Information

Tatsuyuki Takahashi, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Kyosuke Kidachi, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Mikiko Yukawa, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Tomoki Hachinohe, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Yuina Takashima, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Mao Fujimura, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Atsuko Saito, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Daichi Soga, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Chihiro Ota, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Eri Niizuma, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Katsuyoshi Sato, Faculty of Bioresource Sciences, Akita Prefectural University, Akita, Japan.

Hideki Ogasawara, Field Science Center, School of Veterinary Medicine, Kitasato University, Hokkaido, Japan.

Yohei Kurose, Department of Animal Science, School of Veterinary Medicine, Kitasato University, Aomori, Japan.

Author contributions

Tatsuyuki Takahashi (Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Resources, Software, Visualization, Writing—original draft, Writing—review & editing), Kyosuke Kidachi (Data curation, Formal analysis, Investigation, Visualization), Mikiko Yukawa (Data curation, Formal analysis, Investigation), Tomoki Hachinohe (Data curation, Formal analysis, Investigation), Yuina Takashima (Data curation, Formal analysis, Investigation), Mao Fujimura (Data curation, Formal analysis, Investigation), Atsuko Saito (Formal analysis, Investigation), Daichi Soga (Investigation), Chihiro Ota (Investigation), Eri Niizuma (Investigation), Katsuyoshi Sato (Investigation, Methodology, Writing—review & editing), Hideki Ogasawara (Investigation, Methodology, Writing—review & editing), and Yohei Kurose (Conceptualization, Formal analysis, Writing—original draft, Writing—review & editing)

Conflict of interest statement

The authors declare that no conflict of interest could be perceived as prejudicing the impartiality of the research.

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