http://orcid.org/0000-0003-3320-2381
Figures
Abstract
The experiment was conducted to study the effect of the glutamate (Glu) on muscle protein loss through toll-like receptor 4 (TLR4), nucleotide-binding oligomerization domain proteins (NODs), Akt/Forkhead Box O (Akt/FOXO) and mammalian target of rapamycin (mTOR) signaling pathways in LPS-challenged piglets. Twenty-four weaned piglets were assigned into four treatments: (1) Control; (2) LPS+0% Glu; (3) LPS + 1.0% Glu; (4) LPS + 2.0% Glu. The experiment was lasted for 28 days. On d 28, the piglets in the LPS challenged groups were injected with LPS on 100 μg/kg body weight (BW), and the piglets in the control group were injected with the same volume of 0.9% NaCl solution. After 4 h LPS or saline injection, the piglets were slaughtered and the muscle samples were collected. Glu supplementation increased the protein/DNA ratio in gastrocnemius muscle, and the protein content in longissimus dorsi (LD) muscle after LPS challenge (P<0.05). In addition, Glu supplementation decreased TLR4, IL-1 receptor-associated kinase (IRAK) 1, receptor-interacting serine/threonine-protein kinase (RIPK) 2, and nuclear factor-κB (NF-κB) mRNA expression in gastrocnemius muscle (P<0.05), MyD88 mRNA expression in LD muscle, and FOXO1 mRNA expression in LD muscle (P<0.05). Moreover, Glu supplementation increased p-Akt/t-Akt ratio (P<0.05) in gastrocnemius muscle, and p-4EBP1/t-4EBP1 ratio in both gastrocnemius and LD muscles (P<0.05). Glu supplementation in the piglets’ diets might be an effective strategy to alleviate LPS-induced muscle protein loss, which might be due to suppressing the mRNA expression of TLR4 and NODs signaling-related genes, and modulating Akt/FOXO and mTOR signaling pathways.
Citation: Kang P, Wang X, Wu H, Zhu H, Hou Y, Wang L, et al. (2017) Glutamate alleviates muscle protein loss by modulating TLR4, NODs, Akt/FOXO and mTOR signaling pathways in LPS-challenged piglets. PLoS ONE 12(8): e0182246. https://doi.org/10.1371/journal.pone.0182246
Editor: Cristoforo Scavone, Universidade de Sao Paulo, BRAZIL
Received: December 24, 2016; Accepted: July 14, 2017; Published: August 4, 2017
Copyright: © 2017 Kang et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All data files are available from the Figshare database (accession number 10.6084/m9.figshare.5038910).
Funding: Funded by the National Natural Science Foundation of China (31372318 and 31422053), and the Project of the Hubei Provincial Department of Education (T201508), Yulan Liu.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Toll-like receptor (TLR) 4 as a key component of the innate immunity is present in skeletal muscle [1]. Its indispensable member, myeloid differentiation factor (MyD) 88 [2–3], interacts with TLR4 to initiate nuclear factor-κB (NF-κB), which then up-regulates the expression of the genes encoding pro-inflammatory cytokines during LPS challenge. As well as TLR4, nucleotide-binding oligomerization domain protein (NOD) 1 and NOD 2 can activate NF-κB to release pro-inflammatory cytokines [4–6]. The up-regulation of the pro-inflammatory cytokines can cause loss of lean body mass [7–9]. In addition, pro-inflammatory cytokines can lead to muscle atrophy partially via changing the Akt / Forkhead Box O (FOXO)/ubiquitin-proteasome proteolysis (UPP) pathway [10]. Akt/FOXO signaling cascade is an important signaling mechanism in the prosurvival pathway [11]). UPP is the primary proteolytic system, which can degrade most cell proteins [12] and contribute to 75% protein degradation during skeletal muscle atrophy [13–14]. Therefore, UPP serves a critical function in controlling levels of specific proteins. Moreover, many evidences have shown that mammalian target of rapamycin (mTOR) pathway also plays a very crucial role on protein synthesis.
Recent findings have revealed nonessential amino acids in diet are also crucial for animal’s growth and health [15]. Glutamate (Glu) is one of the nonessential amino acids, however, evidence has shown that Glu is a conditionally essential amino acid for weaning piglets. Glu supplementation (as in monosodium glutamate) in postweaning piglet’s diets can be up to 4% without any side-effects, even improve growth performance of piglets for 21d [16]. Nagata reported that a high intake of Glu significantly decreased the risk of total stroke mortality in women [17]. In addition, López-Colomé et al. reported that Glu activated protein synthesis at transcriptional and translational levels in retina [18]. Wu found that Glu was a functional amino acid participating in many kinds of metabolism pathways, including protein production [19].
Skeletal muscle is an important site to regulate the whole body metabolism. Lipopolysaccharide (LPS) challenge can decrease protein synthesis and increase protein degradation in muscle [20], and further induce muscle protein loss or muscle atrophy. However, whether Glu has an effect on the protein loss in the muscle after LPS challenge has not been systematically investigated. Therefore, in this study, we targeted to investigate the effect of Glu inclusion in the weaning piglet diet on muscle protein loss via TLR4, NODs, Akt/FOXO/ and mTOR signaling pathways in skeletal muscle after LPS challenge.
Materials and methods
Experimental design
Twenty four healthy weaned piglets (Duroc × Large White × Landrace, 7.02 ± 0.21 kg) were purchased from Hubei Oden Agriculture and Animal Husbandry Technology Co., Ltd and randomly assigned into four treatments: (1) Control [control diet (0% Glu) and saline injection]; (2) LPS + 0% Glu [control diet and LPS (Escherichia coli serotype 055: B5; Sigma Chemical, St. Louis, MO) challenge]; (3) LPS + 1.0% Glu (1.0% Glu-supplemented diet and LPS challenge); (4) LPS + 2.0% Glu (2.0% Glu-supplemented diet and LPS challenge). Each treatment had six replicates, and each replicate had one piglet. In this experiment, we focused on determining the response to dietary Glu supplementation among LPS-challenged pigs, so we did not include the effect of Glu without LPS challenge. However, previous studies showed that without LPS challenge, dietary supplementation of 1.0% and 2.0% Glu did not affect growth performance (unpublished data). The diets were isonitrogenous with alanine (purity > 99.5%, Wuhan Amino Acid Bio-Chemical Co., Ltd), and the nutrient composition was required according to NRC (1998) [21], which is shown in Table 1. The experiment was lasted for 28 days. On d 28, the piglets in the LPS-challenge groups were injected intraperitoneally with LPS on 100 μg/kg body weight (BW), and the piglets in the control group were injected with the same volume of 0.9% NaCl solution. The L-Glu we used was purchased from Wuhan Amino Acid Bio-Chemical Co., Ltd (purity > 99.1%).
Animal feeding
The animal use protocol for this research was approved by the Animal Care and Use Committee of Wuhan Polytechnic University. All piglets were housed in a stainless pens with 1.20 × 1.10 m2, and the temperature was 25~27°C. During the whole experimental period, the tested diets and water were provided ad libitum.
Muscle samples collection
In our previous studies, we found that tissue was damaged at 4 h after LPS injection [22–23]. Therefore, on day 28, all the pigs were slaughtered under anesthesia with an intravenous injection of sodium pentobarbital (80 mg/kg BW) at 4 h after LPS or saline injection. Then the gastrocnemius muscle was harvested under the semitendinosus and biceps flexor cruris (left hind leg), and the longissimus dorsi (LD) muscle was collected from the 10th rib (right side of the carcass), respectively. All the muscle samples were frozen in liquid nitrogen immediately, and then stored at -80°C for further analysis.
Muscle protein, DNA and RNA measurement
Muscle DNA, RNA and protein contents were determined according to the published methods. Briefly, the mucosa (about 0.5g) was homogenized in ice-cold PBS (0·05M-Na3PO4, 2·0M NaCl, 2 ×10−3 M-EDTA, pH 7·4). Mucosal protein content was measured using a detergent-compatible protein assay (Bio-Rad Laboratories, Hercules, CA, USA) [24]. DNA content was assayed by a fluorometric method [25]. RNA levels was determined by ultraviolet absorption at 232·nm and 260·nm [26].
Muscle mRNA expression analysis
The genes expression, including TLR4, MyD88, IL-1 receptor-associated kinase (IRAK) 1, TNF receptor-associated factor (TRAF) 6, NOD1, NOD2, receptor-interacting serine/threonine-protein kinase (RIPK) 2, NF-κB, TNF-α (TLR4 and NOD signaling pathways), Akt1, FOXO1, FOXO4, muscle atrophy F-box (MAFbx) and muscle RING finger 1 (MuRF1) (Akt/FOXO/UPP signaling pathway) were measured by real-time PCR according to Liu et al [27]. Briefly, total RNA was isolated by the Trizol reagent (#9108, TaKaRa Biotechnology (Dalian) Co., Ltd., Dalian, China), and cDNA synthesis was conducted by PrimeScript® RT reagent kit (#RR047A, TaKaRa Biotechnology (Dalian) Co., Ltd., Dalian, China) according to the manufactures instruction. Quantitative results of the mRNA expression for the target genes was carried out on a ABI 7500 Real-Time PCR System (Applied Biosystems, Life Technologies) using a SYBR® Premix Ex TaqTM (Tli RNaseH Plus) qPCR kit (#RR420A, TaKaRa Biotechnology (Dalian) Co., Ltd., Dalian, China). The PCR cycling conditions were 95°C × 30 s, followed by 40 cycles of 95°C × 5 s and 60°C × 34 s. The primers were designed with Primer 6.0 software and synthesized by TAKARA Biotechnology (Dalian) Co., LTD, China. The primers sequences are shown in Table 2, and the 2-ΔΔCT method was used to calculate the mRNA expression of the target genes relative to GAPDH [28].
Muscle protein expression analysis
Protein expression in muscles was analyzed by Western blot according to our previous studies [29–30]. Briefly, proteins in supernatant fluids (65 μg) were separated on 12% sodium dodecyl sulfate-polyacrylamide gel, and then transferred onto polyvinylidene difluoride membranes for immunoblotting. The first antibodies including total Akt (t-Akt, #9272), phosphorylated Akt (p-Akt, Ser473, #9271), total FOXO1 (t-FOXO1, #9454), phosphorylated FOXO1 (p-FOXO1, Ser256, #9461), total mTOR (t-mTOR, #2972), phosphorylated mTOR (p-mTOR, Ser2448, #2971), total 4EBP1 (t-4EBP1, #9452) and phosphorylated 4EBP1 (p-4EBP1, Thr70, #9455) were purchased from Cell Signaling (Danvers, MA), and GAPDH (V-18) (#sc-20357) were purchased from Santa Cruz Biotechnology (Texas, USA). All first antibodies were diluted into 1:1000. The secondary antibodies including goat anti-rabbit IgG-HRP (#ANT020) and rabbit anti-goat IgG-HRP (#ANT021) were purchased from Antgene Biotech (Wuhan, China) and were diluted into 1:5000. The bands were analyzed by densitometry using GeneTools software (Syngene), and the abundance of the phosphorylated proteins was normalized to the total protein contents.
Statistical analysis
The experimental data were analyzed by ANOVA with SPSS 17.0 software. Differences among treatments were determined by Duncan’s multiple range tests. All data were expressed as means ± SE. The statistical significance level for all analyses was set at p<0.05. A one-way MANOVA was used to determine the effect size (partial eta squared) and the statistical power of the model.
Results
Contents of protein, DNA and RNA in muscles
As shown in Fig 1, Glu supplementation increased the protein content (Fig 1b) in LD muscle (P<0.05), and the protein/DNA ratio (Fig 1e) in gastrocnemius muscle (P<0.05) after LPS challenge.
Mean ± SE; n = 6, one piglet per pen. a,b,c Mean values with unlike letters were significantly different (P<0.05). A one-way MANOVA revealed the partial eta squared was 0.349, and the power to detect the effect was 0.711. Control (non-challenged control), piglets fed a control diet and injected with sterile saline; LPS (LPS-challenged control), piglets fed the same control diet and injected with LPS; LPS + 1.0% Glu, piglets fed a 1.0% Glu diet and injected with LPS; LPS + 2.0% Glu, piglets fed a 2.0% Glu diet and injected with LPS.
Key genes mRNA expression in TLR4 and NOD signaling pathways in muscles
As shown in Table 3, compared to the control group, LPS challenge increased the mRNA expression of TLR4, MyD88, NOD2 and RIPK2 in gastrocnemius muscle, and the mRNA expression of MyD88 in LD muscle (P<0.05), and decreased the mRNA expression of NOD1 in gastrocnemius muscle and IRAK1 in LD muscle (P<0.05). In addition, as for the LPS-challenged piglets, Glu supplementation up to 2.0% decreased TLR4, IRAK1, RIPK2 and NF-κB mRNA expression in gastrocnemius muscle (P<0.05), and MyD88 mRNA expression in LD muscle (P<0.05).
Key genes mRNA expression in Akt/FOXO/UPP signaling pathway in muscles
As shown in Table 4, compared to the control group, LPS challenge increased the mRNA expression of FOXO1 and MuRF1 in both gastrocnemius and LD muscles, and decreased the mRNA expression of FOXO4 in LD muscle (P<0.05). Glu supplementation up to 2.0% decreased the FOXO1 mRNA expression in LD muscle (P<0.05) in LPS-challenged piglets.
Protein expression of t-Akt, p-Akt, t-FOXO1 and p-FOXO1 in muscles
As shown in Figs 2 and 3, compared to the control group, LPS challenge decreased p-FOXO1/t-FOXO1 ratio (Fig 3c) in gastrocnemius muscle (P<0.05). Glu supplementation up to 1.0% increased p-Akt/t-Akt ratio (Fig 2c) in gastrocnemius muscle after LPS challenge (P<0.05).
Mean ± SE; n = 6, one piglet per pen. a,b Mean values with unlike letters were significantly different (P<0.05). A one-way MANOVA revealed the partial eta squared was 0.550, and the power to detect the effect was 0.938. Control (non-challenged control), piglets fed a control diet and injected with sterile saline; LPS (LPS-challenged control), piglets fed the same control diet and injected with LPS; LPS + 1.0% Glu, piglets fed a 1.0% Glu diet and injected with LPS; LPS + 2.0% Glu, piglets fed a 2.0% Glu diet and injected with LPS.
Mean ± SE; n = 6, one piglet per pen. a,b Mean values with unlike letters were significantly different (P<0.05). A one-way MANOVA revealed the partial eta squared was 0.550, and the power to detect the effect was 0.938. Control (non-challenged control), piglets fed a control diet and injected with sterile saline; LPS (LPS-challenged control), piglets fed the same control diet and injected with LPS; LPS + 1.0% Glu, piglets fed a 1.0% Glu diet and injected with LPS; LPS + 2.0% Glu, piglets fed a 2.0% Glu diet and injected with LPS.
Protein expression of t-mTOR, p-mTOR, t-4EBP1 and p-4EBP1 in muscles
As shown in Fig 4, both LPS challenge and Glu inclusion had no effect on the protein expression of t-mTOR and p-mTOR: t-mTOR ratio (P>0.05). As shown in Fig 5, compared to the control group, LPS challenge decreased p-4EBP1/t-4EBP1 in LD muscle (Fig 5c) (P<0.05). Glu inclusion up to 1.0% increased the p-4EBP1/t-4EBP1 ratio in both gastrocnemius (Fig 5c) and LD muscles (Fig 5d) after LPS challenge (P<0.05).
Mean ± SE; n = 6, one piglet per pen. A one-way MANOVA revealed the partial eta squared was 0.481, and the power to detect the effect was 0.949. CONTR (non-challenged control), piglets fed a control diet and injected with sterile saline; LPS (LPS-challenged control), piglets fed the same control diet and injected with LPS; LPS + 1.0% Glu, piglets fed a 1.0% Glu diet and injected with LPS; LPS + 2.0% Glu, piglets fed a 2.0% Glu diet and injected with LPS.
Mean ± SE; n = 6, one piglet per pen. a,b,c Mean values with unlike letters were significantly different (P<0.05). A one-way MANOVA revealed the partial eta squared was 0.481, and the power to detect the effect was 0.949. CONTR (non-challenged control), piglets fed a control diet and injected with sterile saline; LPS (LPS-challenged control), piglets fed the same control diet and injected with LPS; LPS + 1.0% Glu, piglets fed a 1.0% Glu diet and injected with LPS; LPS + 2.0% Glu, piglets fed a 2.0% Glu diet and injected with LPS.
Discussion
Skeletal muscle is the most abundant tissue of the body. LPS can trigger skeletal muscle damage [31], mainly characterized by a decrease in total protein [32], accordingly, the damaged muscle has an ability to regenerate new muscle fibers [33], including the increased protein synthesis. Protein/DNA ratio can be a measure of muscle protein concentration [34] and cell size [35]. In the current study, we found Glu inclusion increased protein/DNA ratio in gastrocnemius muscle, as well as protein content in LD muscle after LPS challenge, which indicates that Glu supplementation stimulated protein synthetic capacity and inhibited protein degradation in muscle cells after LPS challenge.
TLR4 play an important role to trigger mammalian innate immune response by recognizing bacterial LPS [36]. TLR4 can transfer signal to MyD88 and IRAK1, which then stimulate NF-κB to produce pro-inflammatory cytokines, such as TNF-α. Inhibiting pro-inflammatory cytokine production by blocking TLR4/NF-κB pathway can alleviate LPS-induced inflammatory responses [37]. As well as TLR4, NOD1 and NOD2 can activate NF-κB to induce the production of pro-inflammatory cytokines [4–6]. RIPK2 is an essential downstream adaptor protein in NODs signaling pathway, which can stimulate NF-κB pathway [38–39]. Moreover, previous study has reported that pro-inflammatory cytokines such as TNF-α promoted the loss of muscle protein in skeletal muscle [40]. In the current study, in consistent with Frisard et al (2015) [41] who found that LPS challenge stimulated TLR4 activation in skeletal muscle, we also found LPS injection up-regulated TLR4 mRNA expression, which further increased MyD88 mRNA expression in gastrocnemius muscle. Glu is an important precursor for glutamine (Gln). Kessel et al (2008) reported that oral Gln decreased the mRNA expression of TLR4 and MyD88, and resultantly alleviated intestinal mucosal injury caused by LPS in rat [42]. In this study, we found Glu inclusion reduced TLR4, IRAK1, and NOD2 mRNA expression in gastrocnemius muscle, and MyD88 mRNA expression in LD muscle after LPS challenge. In addition, Glu inclusion also decreased NF-κB and RIPK2 mRNA expression in gastrocnemius muscle. These results indicates that Glu supplementation had a positive effect on alleviating muscle protein degradation partially by suppressing the mRNA expression of TLR4 and NODs signaling-related genes after LPS challenge.
In addition, LPS interacts with TLR4/MD2 complex to initiate the inflammatory response [36, 43] and leads to muscle atrophy, which might be partially mediated by the Akt/FOXO/UPP pathway [10]. Phosphorylation of Akt inhibits proteolytic transcription factors [44]. Full activity of Akt requires phosphorylation, which stimulates protein synthesis and induces FOXO1 phosphorylating and inactivating to inhibit protein degradation [45]. FOXO initially stimulates protein degradation [46], and participates in MuRF1 and MAFbx transcription during muscle atrophy [45, 47–48]. Both MuRF1 and MAFbx are relied by UPP to degrade specific proteins within the cells. Crossland et al. found that LPS challenge droved Akt1 and FOXO to stimulate muscle protein degradation [10]. In the current study, we found that Glu inclusion decreased FOXO1 mRNA expression in LD muscle, and increased p-Akt/t-Akt ratios in gastrocnemius muscle after LPS challenge, which suggests that Glu alleviated muscle protein degradation through influencing Akt-FOXO signaling pathway, however, its inclusion could have no effect on UPP pathway evidenced by the unchanged MuRF1 and MAFbx mRNA expression.
Amino acids regulate protein synthesis and cell growth mainly through the mTOR signaling pathway [49–50]. Glu, as an efficient energy resource for the gut of animals, is also an important precursor for arginine and glutamine, which have been reported to stimulate mTOR and eukaryotic initiation factor (eIF) 4E-binding protein-1 (4EBP1) phosphorylation to increase protein synthesis [51]. LPS challenge could impair muscle protein synthesis partially resulting from the reduced activity of mTOR kinase, which was supported by the decreased phosphorylation of 4EBP1 [52–53]. 4EBP1 is one of the downstream targets in mTOR signaling pathway, which is mainly to control the rate of protein synthesis [54]. In addition, the Akt-dependent signaling pathway can inhibit proteolysis and stimulate protein synthesis in muscle via activating 4EBP1 [55]. The excitatory amino acid transporters 3 (EAAT3), which was the glutamate transporter, exists in many tissues, including skeletal muscle [56]. Almilaji et al. reported that EAAT3 could be powerfully up-regulated by mTOR, the later could augment carrier protein concentration in the cell membrane [57]. Therefore, the increased EAAT3 induced by the mTOR could theoretically augment Glu transposition. In the present study, we found Glu supplementation increased p-4EBP1/t-4EBP1 ratio both in gastrocnemius muscle and LD muscle after LPS challenge, which indicates that Glu supplementation could stimulate protein synthesis via activating mTOR signaling pathway, not only by increasing 4EBP1 phosphorylation, but probably by up-regulating Glu transposition.
Conclusion
The results from this study showed that Glu supplementation alleviated muscle protein degradation after LPS challenge not only by suppressing the mRNA expression of TLR4 and NODs signaling-related genes, but by modulating Akt-FOXO signaling pathway to decrease protein degradation and activating mTOR signaling pathway to increase protein synthesis. These results indicated that Glu supplementation might be an effective strategy to alleviate LPS-induced protein loss in muscle after LPS challenge.
Acknowledgments
This study was supported by the National Natural Science Foundation of China (31372318 and 31422053), and the Project of the Hubei Provincial Department of Education (T201508).
References
- 1. Radin MS, Sinha S, Bhatt BA, Dedousis N, O'Doherty RM. Inhibition or deletion of the lipopolysaccharide receptor Toll-like receptor-4 confers partial protection against lipid-induced insulin resistance in rodent skeletal muscle. Diabetologia. 2008; 51: 336–346. pmid:18060381
- 2. Kawai T, Adahi O, Ogawa T, Takeda K, Akira S. Unresponsiveness of MyD88-deficient mice to endotoxin. Immunity. 1999; 11:115–122. pmid:10435584
- 3. Muzio M, Ni J, Feng P, Dixit VM. IRAK (Pelle) family member IRAK-2 and MyD88 as proximal mediators of IL-1 signaling. Science. 1997; 278:1612–1615. pmid:9374458
- 4. Correa RG, Milutinovic S, Reed JC. Roles of NOD1 (NLRC1) and NOD2 (NLRC2) in innate immunity and inflammatory diseases. Biosci Rep. 2012; 32:597–608. pmid:22908883
- 5. Moreira LO, Zamboni DS. NOD1 and NOD2 signaling in infection and inflammation. Front Immunol. 2012; 3:328–339. pmid:23162548
- 6. Philpott DJ, Sorbara MT, Robertson SJ, Croitoru K, Girardin SE. NOD proteins: regulators of inflammation in health and disease. Nat Rev Immunol. 2014; 14:9–23. pmid:24336102
- 7. Kimmel PL, Phillips TM, Simmens SJ, Peterson RA, Weihs KL, Alleyne S, et al. Immunologic function and survival in hemodialysis patients. Kidney Int. 1998; 54:236–244. pmid:9648084
- 8. Bruunsgaard H, Pedersen M, Pedersen BK. Aging and proinflammatory cytokines. Curr Opin Hematol. 2001; 8:131–136. pmid:11303144
- 9. Stentz FB, Umpierrez GE, Cuervo R, Kitabchi AE. Proinflammatory cytokines, markers of cardiovascular risks, oxidative stress, and lipid peroxidation in patients with Hyperglycemic Crises. Diabetes. 2004; 53:2079–2086. pmid:15277389
- 10. Crossland H, Constantin-Teodosiu D, Gardiner SM, Constantin D, Greenhaff PL. A potential role for Akt/FOXO signalling in both protein loss and the impairment of muscle carbohydrate oxidation during sepsis in rodent skeletal muscle. J Physiol. 2008; 586:5589–5600. pmid:18818241
- 11. Juhasz B, Thirunavukkarasu M, Pant R, Zhan L, Penumathsa SV, Secor ER Jr, et al. Bromelain induces cardioprotection against ischemia-reperfusion injury through Akt/FOXO pathway in rat myocardium. Am J Physiol Heart Circ Physiol. 2008; 294: H1365–H1370. pmid:18192224
- 12. Hershko A, Ciechanover A. The ubiquitin system. Annu Rev Biochem. 1998; 67: 425–479. pmid:9759494
- 13. Kandarian SC, Jackman RW. Intracellular signaling during skeletal muscle atrophy. Muscle Nerve. 2006; 33:155–165. pmid:16228971
- 14. Franch HA, Price SR. Molecular signaling pathways regulating muscle proteolysis during atrophy. Curr Opin Clin Nutr Metab Care. 2005; 8:271–275. pmid:15809529
- 15. Wu G, Wu Z, Dai Z, Yang Y, Wang W, Liu C, et al. Dietary requirements of “nutritionally non-essential amino acids”by animals and humans. Amino Acids. 2013; 44:1107–1113. pmid:23247926
- 16. Rezaei R, Knabe DA, Tekwe CD, Dahanayaka S, Ficken MD, Fielder SE, et al. Dietary supplementation with monosodium glutamate is safe and improves growth performance in postweaning pigs. Amino Acids. 2013; 44:911–923. pmid:23117836
- 17. Nagata C, Wada K, Tamura T, Kawachi T, Konishi K, Tsuji M, et al. Dietary intakes of glutamic acid and glycine are associated with stroke mortality in Japanese adults. J Nutr. 2015; 145:720–728. pmid:25833775
- 18. María López-Colomé A, Martínez-Lozada Z, Guillem AM, López E, Ortega A. Glutamate transporter-dependent mTOR phosphorylation in Müller glia cells. ASN Neuro. 2012; 4:331–342.
- 19. Wu G. Intestinal mucosal amino acid catabolism. J Nutr. 1998; 128:1249–1252. pmid:9687539
- 20. Jepson MM, Pell JM, Bates PC, Millward DJ. The effects of endotoxaemia on protein metabolism in skeletal muscle and liver of fed and fasted rats. Biochem J. 1986; 235:329–336. pmid:3527153
- 21.
NRC. Nutrient Requirements of Swine (10th Ed.). National Academic Press, Washington, DC. 1998
- 22. Liu Y, Huang J, Hou Y, Zhu H, Zhao S, Ding B, et al. Dietary arginine supplementation alleviates intestinal mucosal disruption induced by Escherichia coli lipopolysaccharide in weaned pigs. Br J Nutr. 2008; 100: 552–560. pmid:18275628
- 23. Li Q, Liu Y, Che Z, Zhu H, Meng G, Hou Y, et al. Dietary L-arginine supplementation alleviates liver injury caused by Escherichia coli LPS in weaned pigs. Innate Immun. 2012; 18:804–814. pmid:22441699
- 24. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ. Protein measurement with Folin phenol reagent. J Biol Chem. 1951; 193: 265–275. pmid:14907713
- 25. Labarca C, Paigen K. A simple, rapid, and sensitive DNA assay procedure. Anal Biochem 1. 1980; 344–352.
- 26. Storch D, Heilmayer O, Hardewig I, Pörtner HO. In vitro protein synthesis capacities in a cold stenothermal and a temperate eurythermal pectinid. J Comp Physiol. 2003; 173B: 611–620.
- 27. Liu Y, Wang X, Wu H, Chen S, Zhu H, Zhang J, et al. Glycine enhances muscle protein mass associated with maintaining Akt-mTOR-FOXO1 signaling and suppressing TLR4 and NOD2 signaling in piglets challenged with LPS. Am J Physiol Regul Integr Comp Physiol. 2016; 11:R365–R373.
- 28. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and 2-ΔΔCT method. Methods. 2001; 25: 402–408. pmid:11846609
- 29. Liu Y, Chen F, Odle J, Lin X, Jacobi SK, Zhu H, et al. Fish oil enhances intestinal integrity and inhibits TLR4 and NOD2 signaling pathways in weaned pigs after LPS challenge. J Nutr. 2012; 142: 2017–2024. pmid:23014495
- 30. Liu Y, Chen F, Odle J, Lin X, Zhu H, Shi H, et al. Fish oil increases muscle protein mass and modulates Akt/FOXO, TLR4, and NOD signaling in weanling piglets after lipopolysaccharide challenge. J Nutr. 2013; 143:1331–1339. pmid:23739309
- 31. Jortay J, Senou M, Delaigle A, Noel L, Funahashi T, Maeda N, et al. Local induction of adiponectin reduces lipopolysaccharide-triggered skeletal muscle damage. Endocrinology. 2010; 151: 4840–4851. pmid:20702578
- 32. McKinnell IW, Rudnicki MA. Molecular mechanisms of muscle atrophy. Cell. 2004; 119:907–910. pmid:15620349
- 33. Carlson BM. The regeneration of skeletal muscle. A review. Am J Anat. 1973; 137:119–149. pmid:4350147
- 34. Petersson B, Hultman E, Andersson K, Wernerman J. Human skeletal muscle protein: effect of malnutrition, elective surgery and total parenteral nutrition. Clin Sci (Lond). 1995; 88:479–484.
- 35. Enesco M, and Leblond CP. Increase in cell number as a factor in the growth of the young male rat. J Embryol Exp Morphol. 1962; 10:530–562.
- 36. Beutler B. Inferences, questions and possibilities in Toll-like receptor signalling. Nature. 2004; 430:257–263. pmid:15241424
- 37. Xiang P, Chen T, Mou Y, Wu H, Xie P, Lu G, et al. NZ suppresses TLR4/NF-κB signalings and NLRP3 inflammasome activation in LPS-induced RAW264.7 macrophages. Inflamm Res. 2015; 64:799–808. pmid:26298161
- 38. Franchi L, Warner N, Viani K, Nunez G. Function of Nod-like receptors in microbial recognition and host defense. Immunol Rev. 2009; 227:106–128. pmid:19120480
- 39. Sartor RB. Microbial influences in inflammatory bowel diseases. Gastroenterology. 2008; 134:577–594. pmid:18242222
- 40. Lang CH, Frost RA, Nairn AC, MacLean DA, Vary TC. TNF-αimpairs heart and skeletal muscle protein synthesis by altering translation initiation. Am J Physiol Endocrinol Metab. 2002; 282:E336–E347. pmid:11788365
- 41. Frisard MI, Wu Y, McMillan RP, Voelker KA, Wahlberg KA, Anderson AS, et al. Low levels of lipopolysaccharide modulate mitochondrial oxygen consumption in skeletal muscle. Metabolism. 2015; 64:416–427. pmid:25528444
- 42. Kessel A, Toubi E, Pavlotzky E, Mogilner J, Coran AG, Lurie M, et al. Treatment with glutamine is associated with down-regulation of Toll-like receptor-4 and myeloid differentiation factor 88 expression and decrease in intestinal mucosal injury caused by lipopolysaccharide endotoxaemia in a rat. Clin Exp Immunol. 2008; 151:341–347. pmid:18070149
- 43. Akira S, Takeda K. Toll-like receptor signalling. Nat Rev Immunol. 2004; 4:499–511. pmid:15229469
- 44. Bodine SC, Stitt TN, Gonzalez M, Kline WO, Stover GL, Bauerlein R, et al. Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat Cell Biol. 2001a; 3:1014–1019.
- 45. Stitt TN, Drujan D, Clarke BA, Panaro F, Timofeyva Y, Kline WO, et al. The IGF-1/PI3K/Akt pathway prevents expression of muscle atrophy-induced ubiquitin ligases by inhibiting FOXO transcription factors. Mol Cell. 2004; 14:395–403. pmid:15125842
- 46. Sandri M, Sandri C, Gilbert A, Skurk C, Calabria E, Picard A, et al. Foxo transcription factors induce the atrophy-related ubiquitin ligase atrogin-1 and cause skeletal muscle atrophy. Cell. 2004; 117: 399–412. pmid:15109499
- 47. Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA, et al. Identification of ubiquitin ligases required for skeletal muscle atrophy. Science. 2001b; 294: 1704–1708.
- 48. Wang X, Hu Z, Hu J, Du J, Mitch WE. Insulin resistance accelerates muscle protein degradation: Activation of the ubiquitin-proteasome pathway by defects in muscle cell signaling. Endocrinology. 2006; 147:4160–4168. pmid:16777975
- 49. Efeyan A, Sabatini DM. mTOR and cancer: many loops in one pathway. Curr Opin Cell Biol. 2010; 22:169–176. pmid:19945836
- 50. Inoki K, Ouyang H, Zhu T, Lindvall C, Wang Y, Zhang X, et al. TSC2 integrates Wnt and energy signals via a coordinated phosphorylation by AMPK and GSK3 to regulate cell growth. Cell. 2006; 126:955–968. pmid:16959574
- 51. Kim J, Song G, Wu G, Gao H, Johnson GA, Bazer FW. (2013) Arginine, leucine, and glutamine stimulate proliferation of porcine trophectoderm cells through the MTOR-RPS6K-RPS6-EIF4EBP1 signal transduction pathway. Biol Reprod. 2013; 88:113, 1–9. pmid:23486913
- 52. Lang CH, Frost RA, Vary TC. Regulation of muscle protein synthesis during sepsis and inflammation. Am J Physiol Endocrinol Metab. 2007; 293:E453–E459. pmid:17505052
- 53. Lang CH, Frost RA, Bronson SK, Lynch CJ, Vary TC. Skeletal muscle protein balance in mTOR heterozygous mice in response to inflammation and leucine. Am J Physiol Endocrinol Metab. 2010; 298:E1283–E1294. pmid:20388826
- 54. Ma XM, Blenis J. Molecular mechanisms of mTOR mediated translational control. Nat Rev Mol Cell Biol. 2009; 10:307–318. pmid:19339977
- 55. Zlotchenko E, Scrimgeour A, Lawrence JC, Glass DJ, Yancopoulos GD. Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat Cell Biol.2001; 3:1014–1019. pmid:11715023
- 56. Velaz-Faircloth M, McGraw TS, alandro MS, Fremeau RT Jr, Kilberg MS, Anderson KJ. Characterization and distribution of the neuronal glutamate transporter EAAC1 in rat brain. Am J Physiol. 1996; 270: C67–C75. pmid:8772431
- 57. Almilaji A, Pakladok T, Guo A, Munoz C, Föller M, Lang F. Regulation of the glutamate transporter EAAT3 by mammalian target of rapamycin mTOR. Biochem Biophys Res Commun. 2012; 421:159–163. pmid:22483750