Skip to main content
NCBI home page
Journal of Animal Science logoLink to Journal of Animal Science
. 2019 Feb 11;97(4):1796–1805. doi: 10.1093/jas/skz059

The effects of dietary supplementation with hyodeoxycholic acid on the differentiation and function of enteroendocrine cells and the serum biochemical indices in weaned piglets1

Enyan Zong 1,#, Shanling Yan 1,#, Meiwei Wang 1, Lanmei Yin 1, Qiye Wang 1, Jia Yin 1, Jianzhong Li 1, Yali Li 1, Xueqin Ding 1, Pengfei Huang 1, Shanping He 1, Huansheng Yang 1,2,✉, Yulong Yin 1,2
PMCID: PMC6447273  PMID: 30753616

Abstract

Bile acid, a cholesterol metabolite, promotes gastrointestinal tract digestion and absorption of cholesterol, lipids, and fat-soluble vitamins. It is a signaling regulatory molecule that influences a variety of endocrinal and metabolic activities. This study investigated the effects of hyodeoxycholic acid (HDCA) as a dietary supplement on endocrine cell differentiation and function and weaned piglet serum biochemical indices. Sixteen piglets [Duroc × (Landrace × Yorkshire)] were individually housed and weaned at 21 d of age (BW of 6.14 ± 0.22 kg). Uniform weight animals were randomly assigned to 1 of 2 treatments (8 replicate pens per treatment and 1 piglet per pen). The treatments were 1) base diet (control) and 2) base diet supplemented with 2 g/kg of HDCA. Control and HDCA piglet numbers of chromogranin A (CgA)-positive cells per crypt did not differ. HDCA CgA-positive cells numbers decreased (P < 0.05) in the jejunal villi showed a tendency to decrease (P < 0.10) in the ileal villi and showed tendency toward an increase (P < 0.10) in the duodenal villi compared with the controls. The HDCA diet led to a decline in glucagon-like peptide 2 (P < 0.01) concentrations, but did not affect plasma glucagon-like peptide 1. HDCA supplementation increased (P < 0.05) the mRNA expression of jejunal Insm1, Sst, PG, and Gast, but decreased (P < 0.05) duodenal expression of Insm1, jejunal Pdx1, and ileal NeuroD1. HDCA elevated globulin and immunoglobulin A (P < 0.05) serum concentrations and decreased the albumin/globulin ratio (P < 0.05). Total protein and immunoglobulin G serum levels tended to increase compared with the control group. These results indicate that dietary HDCA at 2 g/kg may regulate enteroendocrine cell differentiation and play a role in increasing weaned piglet humoral immunity.

Keywords: cell differentiation, enteroendocrine cell, hyodeoxycholic acid, intestine, weaned piglet

INTRODUCTION

As a vital organ, the intestine digests and absorbs nutrients to maintain energy homeostasis. The gut epithelial primary types of differentiated cells are Paneth cells, goblet cells, enterocytes, and enteroendocrine cells (EECs). This cell differentiation ensures an optimal nutrient absorption. EECs have evolved in part to control nutrient absorption efficiency by secreting specialized peptide hormones that act as either autocrines, paracrines, or endocrines to transmit signals related to nutrient availability (Lee et al., 2017). Weaning piglets can suffer from transient anorexia and face severe deterioration of intestinal mucosal structures and function. This deterioration may relate to weaning stress, which affects intestinal epithelial cell proliferation, differentiation, and function (Ipharraguerre et al., 2013; Yang et al., 2016b). Early weaning can be accompanied by a notable reduction in circulating glucagon-like peptide 2 (GLP-2) levels (Burrin et al., 2003; Petersen et al., 2003). GLP-2 is a vaso-active intestinal polypeptide released by enteroendocrine L cells primarily in response to luminal nutrients. It plays an important role in intestinal nutrient absorption and mucosal enterocyte proliferation (Gunawardene et al., 2011). Previous studies showed that intestinal luminal bile acids activate guanosine protein-coupled bile acid receptors (TGR5) expressed in mucosal enteroendocrine L cells and cooperate with luminal nutrients to stimulate postprandial glucagon-like peptide 1 (GLP-1) and GLP-2 secretions (Jain et al., 2012; Hansen et al., 2016). Bile acids have emerged as potent hormonal regulators capable of stimulating GLP-1, a coproduct of proglucagon released in parallel with GLP-2, secretion (Rindi et al., 2004; Katsuma et al., 2005; De Diego-Cabero et al., 2015). GLP-2 secretion is mediated by direct nutrient stimulation of the L cells and indirect action from enteroendocrine and neural inputs (Burrin et al., 2003). The continuous enteral administration of chenodeoxycholic acid (CDC), a primary bile acid known to activate TGR5, to newborn piglets fed parenterally increased GLP-2 plasma concentrations and prevented gut atrophy which might otherwise result from a lack of enteral nutrition (Jain et al., 2012). This study investigated whether exogenous bile acids could induce a similar response in weaning piglets and whether it might influence immune-related factors including, but not limited to, aspartate aminotransferase (AST); alanine aminotransferase (ALT); total protein (TP); albumin (ALB); globulin (GLO); albumin/globulin (A/G); immunoglobulin M (IgM); and immunoglobulin A (IgA) in the serum. It was hypothesized that bile acids might influence GLP-1 and GLP-2 secretions by affecting EEC differentiation and might further affect weaning piglet serum biochemical indices. The hypothesis was tested by evaluating how dietary hyodeoxycholic acid (HDCA) might influence weaning piglet EEC differentiation and function and serum biochemical indices.

MATERIALS AND METHODS

The experimental design and procedures in this study were reviewed and approved by the Animal Care and Use Committee of Hunan Normal University, Changsha, Hunan, China.

Animals and Experimental Treatments

Sixteen piglets [Duroc × (Landrace × Yorkshire)] were individually housed and weaned at 21 d of age. Animals with fairly uniform BW (6.14 ± 0.22 kg) were then randomly assigned to 1 of 2 treatments, with 8 replicate pens per treatment. The treatments were feedings of either a base diet (control) or a base diet supplemented with 2 g/kg of HDCA (Sigma, St. Louis, MO). The experimental diets (Table 1) were mash feeds and formulated to meet the National Research Council (NRC) nutrient specifications for weaned pigs (NRC, 2012). All piglets were kept in a temperature-controlled nursery where they had ad libitum access to feed and water during the 7 d post-weaning experimental period. Feed consumption per pen was measured daily. Growth performance results as ADG along with ADFI were collected for each subject.

Table 1.

Experimental diet ingredients

Component Content, %
Corn 37.76
Extruded corn 20.00
Soybean meal, 43% CP 8.00
Concentrated soy protein 7.00
Whey 10.00
Fish meal, 63% CP 5.00
Plasma protein powder 4.50
l-Lys HCl, 98% 0.33
dl-Met 0.08
l-Thr 0.03
l-Trp 0.01
Glucose 2.00
Soybean oil 2.00
Limestone 1.04
Monocalcium phosphate 0.50
Choline chloride, 50% 0.10
Antioxidants 0.05
Zinc oxide 0.30
Citric acid 0.30
Vitamin–mineral premix1 1.00
Total 100
Calculated composition
 CP, % 18.0
 ME, MJ/kg 14.2
 Lys,2 % 1.35
 Met,2 % 0.39
 Met + Cys,2 % 0.74
 Thr,2 % 0.79
 Trp,2 % 0.22
Open in a new tab

1Vitamin–mineral premix per kg of feed: vitamin A, 10,000 IU; vitamin D3, 1,000 IU; vitamin E, 80 IU; vitamin K3, 2.0 mg; vitamin B12, 0.03 mg; riboflavin, 12 mg; niacin, 40 mg; d-pantothenic acid, 25 mg; biotin, 0.25 mg; folic acid, 1.6 mg; thiamin, 3.0 mg; pyridoxine, 2.25 mg; choline chloride, 300 mg; Fe (FeSO4), 150 mg; Zn (ZnSO4), 100 mg; Mn (MnSO4), 30 mg; Cu (CuSO4), 25 mg; I (KIO3), 25 mg; Co (CoSO4), 0.3 mg; Se (Na2SeO3), 0.3 mg; and ethoxyquin, 4.0 mg.

2Standardized ileal-digestible.

Plasma Collection and Analysis

At the end of the trial, the piglets were 28 d old. They were fasted for 12 h (overnight) and individually weighed. A 10-mL blood sample was collected into heparin-coated tubes via jugular vein puncture and centrifuged at 3,000 × g and 4 °C for 10 min to recover plasma. Plasma samples were immediately stored at −80 °C until analyzed for total GLP-1, GLP-2, and biochemical indices.

Tissue Collection and Serum Biochemical Index Analyses

Immediately after blood sampling, piglets were given a general anesthesia and sacrificed by an intravenous injection of 4% sodium pentobarbital solution (40 mg/kg BW). The gastrointestinal (GI) tract was immediately resected and separated the duodenum, jejunum, and ileum. The first approximately 20 cm distal from the stomach was designated the duodenum. The ileal portion, about 20 cm in length, was collected beginning approximately 10 cm proximal to the ileocecal valve. The 20-cm-long jejunal portion was isolated from the middle of the intestine. The segments were flushed with normal saline (pH = 7.0) and bisected crosswise. A section, approximately 2 cm long, was fixed with a 10% formaldehyde-phosphate buffer and kept at 4 °C, whereas the other section provided mucosal samplings. Immediately after collection, the mucosal tissues were frozen in liquid nitrogen and stored at −80 °C until required for further analysis (He et al., 2013; Yan et al., 2018). Blood urea nitrogen, glucose, alkaline phosphatase, AST, ALT, AST/ALT, TP, ALB, GLO, A/G, IgM, IgA, immunoglobulin G (IgG), lactate dehydrogenase, and IGF were measured using a Beckman CX4 Chemistry Analyzer (Beckman Coulter) and commercial kits (Sino-German Beijing Leadman Biochemistry Technology Company, Beijing, China).

Intestinal Morphology

Duodenal, jejunal, and ileal samples were fixed in formalin and embedded in paraffin. Samples were dehydrated in graded alcohol baths and ade transparent with xylene. They were then soaked for an hour with soft wax and 2 h with hard wax and embedded in paraffin. Using a microtome, approximately 5-µm-thick cross-sectional samples were cut and stained with hematoxylin and eosin. Villus height (VH) and crypt depth (CD) for each section were calculated using a microscope at 50× combined magnification and an image processing and analysis system (DM3000; Leica, Germany). At least 30 well-oriented intact villi and their associated crypts were examined. The distances from the tip of the villus to the mouth of the crypt (VH) and the crypt mouth to the crypt base (CD) were taken to assess VH and CD consistently. The VH-to-CD ratio (VH:CD) was determined.

Immunohistochemical Examination

Four-µm-thick tissue sections were mounted on charged slides. After dissolving the paraffin with xylene, the tissues were rehydrated in graded alcohol baths and PBS (pH = 7.0). They were then soaked for 10 min in 3% H2O2. Antigen retrieval was performed by boiling the slides for 30 min in a sodium citrate buffer (pH = 6.0). The slides cooled to room temperature before being thrice washed in PBS. Rabbit monoclonal anti-chromogranin A (CgA) antibody (Abcam) was diluted in PBS (1:600) and incubated for 1 h at 37 °C. After secondary antibody (Santa Cruz Biotechnology Inc.), incubation, and labeling according to supplier instructions, the tissue slides were counterstained with hematoxylin and dehydrated. Cover slips were placed on the slides to prepare for further analysis. Using a light microscope (DM3000; Leica, Germany) at 100× magnification, the number of CgA-positive cells per crypt and per individual villus (15 crypts and 15 villi per field) found in 10 randomly chosen fields was tallied. Mean values for the 10 fields per sample were then calculated and reported for each piglet (Saqui-Salces et al., 2017).

RNA Extraction and cDNA Synthesis

Tissue from each sample type, approximately 100 mg, was pulverized in liquid nitrogen (Zhou et al., 2012). Total RNA was isolated from the homogenate using a TRIZOL reagent (100 mg tissue/mm of Trizol; TaKaRa, Beijing, China). RNA integrity was checked via 1% agarose gel electrophoresis stained with 10 µg/mL ethidium bromide. RNA quality and quantity were determined by a UV spectrophotometer (NanoDrop ND-1000; Thermo Fisher Scientific). For all samples, RNA had an OD260:OD280 ratio of between 1.8 and 2.0 (Yang et al., 2016a). DNA-free RNA (1 µg) was used for reverse transcription and PCR. First-strand cDNA was synthesized with an RT Reagent Kit and gDNA Eraser (TaKaRa).

Real-Time PCR

Primers for the selected genes (Table 2) were designed using Oligo 6.0 software (Molecular Biology Insights, Cascade, CO), and synthesized by TSINGKE Biological Technology (Beijing, China). The identity of PCR product was checked by 2% agarose gel electrophoresis and DNA sequencing. Real-time PCR was performed with a StepOnePlus System (QuantStudio, Thermo Fisher Scientific). Each reaction included 5 µL of SYBR Green mix (Thermo Fisher Scientific), 0.3 µL each of forward and reverse primers, and 1 µL of 5-fold-diluted cDNA (Yang et al., 2016c). After pre-denaturation (10 s at 95 °C), 40 amplification cycles were performed. Each cycle consisted of 5 s at 95 °C, the 20× at 60 °C, followed by a melting curve program which took the sample from 60 to 99 °C at a heating rate of 0.1 °C/s and included fluorescence measurement. For each sample, β-actin amplification of was used to normalize the expression of selected genes. mRNA relative expression was calculated according to the 2−ΔΔ Ct method (Xiong et al., 2015). The efficiency of real-time reverse-transcription PCR was determined by the amplification of a dilution series of cDNA according to the equation 10(−1/slope). Target mRNA and β-actin mRNA were amplified with comparable efficiencies.

Table 2.

Primer sequences used to investigate genes of interest

Gene1 Primers Sequences (5′ to 3′) Size, bp
β-actin Forward AGTTGAAGGTGGTCTCGTGG 216
Reverse TGCGGGACATCAAGGAGAAG
Intestinal endocrine-related transcription factors
Ngn3 Forward AACTTGCAGACGAAAAAGCC 90
Reverse GCGCCATCCTCGTTTTAG
NeuroD1 Forward ACAACTACAGGAGACCTAAACA 109
Reverse GAGAACTGAGACACTCGTCTG
Pdx1 Forward TTACTTCAACGTGCAACGGT 249
Reverse TCCCACACTGAGTTTTCGCT
Pax6 Forward GGGTTGCATAGGCAGGTTATT 182
Reverse TCCCCATCAGCAGTAGTTTCA
Insm1 Forward GCAACAGGAGCCAATCTCTT 195
Reverse ATTCACCCAAAACAACCCGT
Pax4 Forward ACACGGTGAGGATCTGGTTT 120
Reverse CTGGGGAAGCACTTGGTAGA
Nkx2.2 Forward GCCAGCCCATGCAGGGAGTAC 219
Reverse GCCAGACACCAACGATGAGGA
Endocrine hormones
 CCK Forward ATACTCGGCCAGAAGGTGCTT 100
Reverse AGGGCGGTGCAAAAGGTAGAC
 Sst Forward AGCTCCAGCCTCATTTCATCC 174
Reverse TCCCCGACTCCGTCAGTTTCT
 Gast Forward ATGTGGCTCTTTGCCCCTGTT 110
Reverse TGATCCATGTGCTGGCTCTGG
 GIP Forward AGAGCGACTGGAAACACAAC 250
Reverse GTGAAGGGCAGAGTCCAATC
 PYY Forward GCCTGCTCATCTGCCTGGGGA 165
Reverse CGTCTGGGCTGTCACGTTTCC
 Ghrl Forward ACACCAGAAAGTGCAGCAGAG 108
Reverse CCTTCCACCTCACCACTGTCTT
 PG Forward GACTGAAGACAAGCGCCACT 110
Reverse TCCTCTTGGTGTTCATCAGCCACT
Open in a new tab

1Names of genes: Ngn3 = neurogenin 3; NeuroD1 = neurogenic differentiation 1; Pdx1 = pancreatic-duodenal homeobox 1; Pax6 = paired box gene 6; Insm1 = insulinoma-associated 1; Pax4 = paired box gene 4; Nkx2.2 = NK2 transcription factor related = locus 2; CCK = cholecystokinin; Sst = somatostatin; Gast = gastrin; GIP = glucose-dependent insulinotropic peptide; PYY = peptide YY; Ghrl = ghrelin; PG = proglucagon.

Enzyme-Linked Immunosorbent Assay

Glucagon-like peptide 1 concentrations in serum samples were determined using one type of ELISA kit (Cusabio Biotech Co., Ltd., China), having a detection range of 2.4 to 150 pg/mL, and a minimum detectable dose of pig GLP-1 typically less than 0.6 pg/mL. Intra-assay precision CV is about 5.1%. Interassay precision CV is about 8.7%. Serum GLP-2 was quantified with another type of ELISA kit (Kanglang Biological Technology Co., Ltd., China), whose detection range is 0.2 to 8 pmol/L. The intra-assay precision CV is about 6.2%, and the interassay precision CV is about 10.4%. Both ELISA kits employ a quantitative sandwich enzyme immunoassay technique. All measurements were made according to the manufacturer instructions.

Statistical Analysis

Statistical analysis was performed using Student’s t-tests with SAS software (Version 9.2; SAS Institute Inc., Cary, NC). Data were presented as means and SEM. Differences between treatment results were considered significant at P < 0.05.

RESULTS

Growth Performance and Intestinal Morphology

No significant differences in ADG (41.07 ± 17.17 g vs. 44.64 ± 24.36 g) and ADFI (224.43 ± 25.78 g vs. 268.65 ± 22.75 g) were detected between the control and the HDCA group. There were no significant differences in VH, CD, and VH:CD between the control and HDCA group (Table 3). These results suggest that a 7-d feeding dietary HDCA period had no major effects on weaning piglet growth performance or intestinal morphology.

Table 3.

Effects of dietary supplementation with HDCA on intestinal morphology in weaned piglets1

Dietary treatment
Item2 Control HDCA P
Villus height, µm
 Duodenum 325.86 ± 20.19 344.76 ± 25.02 0.566
 Jejunum 316.69 ± 23.77 305.45 ± 15.11 0.696
 Ileum 261.77 ± 11.29 297.90 ± 24.22 0.198
Crypt depth, µm
 Duodenum 390.23 ± 28.38 459.59 ± 31.58 0.125
 Jejunum 295.15 ± 13.69 280.48 ± 13.62 0.460
 Ileum 258.66 ± 14.55 276.40 ± 22.92 0.524
VH:CD
 Duodenum 0.84 ± 0.03 0.77 ± 0.07 0.396
 Jejunum 1.11 ± 0.12 1.11 ± 0.09 0.958
 Ileum 1.03 ± 0.06 1.16 ± 0.16 0.460
Open in a new tab

1Eight piglets per treatment. Control = base diet; HDCA = base diet supplemented with 2 g of HDCA per kg of diet.

2VH = villus height; CD = crypt depth.

HDCA Supplement Effects on EECs and Plasma GLP

Immunohistochemical staining was performed using chromogranin A as the marker to measure intestinal EEC. The number of endocrine CgA-positive cells per crypt in the duodenum, jejunum, or ileum (Fig. 1) did not differ between dietary treatments. Piglets fed the base diet had more cells (P < 0.05) in the jejunal villi compared with the HDCA piglets. HDCA supplementation decreased (P < 0.01) plasma GLP-2 concentrations, but had no effect on plasma GLP-1 concentrations compared with the control group (Fig. 2).

Figure 1.

Figure 1.

Open in a new tab

Dietary HDCA supplement effects on the number of endocrine chromogranin A-positive cells per villus (A) and crypt (B). Values are least squares means ± SEM, n = 8 per treatment. Bars indicate 95% confidence intervals (fold-change up or down). Control = base diet; HDCA = base diet supplemented with 2 g of HDCA/kg of diet. For jejunum samples in (A), difference between treatments was significant at P < 0.05.

Figure 2.

Figure 2.

Open in a new tab

HDCA supplement effects on weaned piglet mean GLP-1 and GLP-2 plasma concentrations. Values are least squares means ± SEM, n = 8 per treatment. Control = base diet; HDCA = base diet supplemented with 2 g of HDCA/diet kg. For GLP-2, difference between treatments was significant at P < 0.05.

Gene Expression Related to EEC Differentiation in the Small Intestine

mRNA expression of numerous transcription factors (TFs; Table 4) and hormone-related genes (Table 5) was measured in the duodenum, jejunum, and ileum. The HDCA diet resulted in decreased duodenal expression of insulinoma-associated 1 (Insm1) but increased jejunal expression. The mRNA abundance of the genes for somatostatin (Sst), proglucagon (PG), and gastrin (Gast) in the jejunum was greater in the HDCA group (P < 0.05) than in the control group. Expression of pancreatic-duodenal homeobox 1 (Pdx1) decreased, paired box gene 4 (Pax4) tended to decrease, and NK2 TF-related locus 2 (NKx2.2) expression tended to increase in the HDCA group compared with the control group. In the ileum, the HDCA diet was associated with a tendency for neurogenic differentiation 1 (NeuroD1) expression to decrease, a tendency for peptide YY (PYY) to decrease, and a tendency for Gast expression to increase compared with the control group.

Table 4.

Effects of dietary supplementation with HDCA on relative abundance of mRNA for transcription factors in duodenum, jejunum, and ileum of weaned piglets1

Dietary treatment
Item2 Control HDCA P
Ngn3
 Duodenum 1.21 ± 0.30 1.22 ± 0.18 0.989
 Jejunum 1.10 ± 0.21 1.29 ± 0.34 0.660
 Ileum 1.08 ± 0.17 0.74 ± 0.15 0.151
NeuroD1
 Duodenum 1.08 ± 0.14 0.99 ± 0.14 0.663
 Jejunum 1.07 ± 0.15 1.49 ± 0.29 0.220
 Ileum 1.06 ± 0.14 0.56 ± 0.08 0.009
Pdx1
 Duodenum 1.04 ± 0.11 0.88 ± 0.15 0.251
 Jejunum 0.99 ± 0.20 0.50 ± 0.06 0.050
 Ileum 1.06 ± 0.17 1.43 ± 0.20 0.189
Pax6
 Duodenum 1.02 ± 0.08 1.22 ± 0.18 0.331
 Jejunum 1.02 ± 0.07 0.82 ± 0.10 0.131
 Ileum 1.08 ± 0.14 0.74 ± 0.19 0.180
Insm1
 Duodenum 1.07 ± 0.16 0.68 ± 0.08 0.049
 Jejunum 1.07 ± 0.16 1.95 ± 0.25 0.006
 Ileum 1.09 ± 0.17 0.70 ± 0.16 0.115
Pax4
 Duodenum 0.91 ± 0.08 0.82 ± 0.12 0.560
 Jejunum 1.10 ± 0.19 0.65 ± 0.07 0.056
 Ileum 1.03 ± 0.09 1.03 ± 0.10 0.992
NKx2.2
 Duodenum 1.09 ± 0.18 1.39 ± 0.20 0.280
 Jejunum 1.16 ± 0.25 1.89 ± 0.31 0.099
 Ileum 1.04 ± 0.11 0.97 ± 0.09 0.623
Open in a new tab

1Values represent fold-changes due to supplement relative to control (n = 8 per treatment). Control = base diet; HDCA = base diet supplemented with 2 g of HDCA/kg of diet.

2 Ngn3 = neurogenin 3; NeuroD1 = neurogenic differentiation 1; Pdx1 = pancreatic-duodenal homeobox 1; Pax6 = paired box gene 6; Insm1 = insulinoma-associated 1; Pax4 = paired box gene 4; Nkx2.2 = NK2 transcription factor related = locus 2.

Table 5.

Effects of dietary supplementation with HDCA on relative abundance of mRNA for hormone-related genes in duodenum, jejunum, and ileum of weaned piglets1

Dietary treatment
Item2 Control HDCA P
CCK
 Duodenum 1.02 ± 0.08 1.06 ± 0.17 0.821
 Jejunum 1.03 ± 0.11 1.19 ± 0.07 0.185
 Ileum 1.00 ± 0.18 1.31 ± 0.17 0.223
Sst
 Duodenum 1.08 ± 0.16 0.90 ± 0.11 0.349
 Jejunum 1.03 ± 0.09 1.71 ± 0.24 0.028
 Ileum 1.08 ± 0.18 1.04 ± 0.14 0.877
Gast
 Duodenum 1.05 ± 0.13 0.89 ± 0.15 0.429
 Jejunum 1.08 ± 0.14 2.75 ± 0.32 0.000
 Ileum 1.12 ± 0.20 2.07 ± 0.44 0.080
GIP
 Duodenum 1.06 ± 0.12 0.98 ± 0.13 0.684
 Jejunum 1.08 ± 0.17 1.37 ± 0.09 0.135
 Ileum 1.05 ± 0.12 0.68 ± 0.17 0.104
PYY
 Duodenum 1.07 ± 0.13 1.07 ± 0.16 0.988
 Jejunum 1.08 ± 0.16 0.93 ± 0.18 0.526
 Ileum 1.08 ± 0.16 0.72 ± 0.07 0.055
Ghrl
 Duodenum 1.21 ± 0.29 0.67 ± 0.11 0.117
 Jejunum 1.20 ± 0.24 1.20 ± 0.28 1.000
 Ileum 1.10 ± 0.19 1.07 ± 0.33 0.929
PG
 Duodenum 1.09 ± 0.17 1.41 ± 0.30 0.378
 Jejunum 1.02 ± 0.09 1.98 ± 0.28 0.006
 Ileum 1.06 ± 0.12 1.35 ± 0.14 0.134
Open in a new tab

1Values represent fold-changes due to supplement relative to control (n = 8 per treatment). Control = base diet; HDCA = base diet supplemented with 2 g of HDCA/kg of diet.

2 CCK = cholecystokinin; Sst = somatostatin; Gast = gastrin; GIP = glucose-dependent insulinotropic peptide; PYY = peptide YY; Ghrl = ghrelin; PG = proglucagon.

HDCA Dietary Supplement Effects on Weaned Piglet Serum Biochemical Indices

HDCA supplementation increased GLO (P < 0.05) and IgA (P < 0.05) serum concentrations and decreased the A/G ratio (P < 0.05). TP and IgG serum levels tended to increase (P < 0.10) compared with the control group (Table 6). These results suggest that HDCA may reduce immune function impairment and increase weaned piglets’ resistance to weaning stress.

Table 6.

Effects of dietary supplementation with HDCA on serum biochemical indexes in weaned piglets1

Dietary treatment
Component2 Control HDCA P
BUN, mmol/L 3.24 ± 1.31 3.15 ± 1.16 0.906
GLU, mmol/L 4.03 ± 0.53 4.46 ± 0.37 0.112
ALP, U/L 200.13 ± 33.24 243.00 ± 63.17 0.124
AST, U/L 38.25 ± 10.84 47.33 ± 3.56 0.561
ALT, U/L 21.50 ± 7.48 27.33 ± 10.03 0.119
AST/ALT 1.82 ± 0.36 1.75 ± 0.68 0.789
TP, g/L 33.48 ± 5.76 40.46 ± 7.96 0.080
ALB, g/L 27.46 ± 4.11 31.38 ± 5.08 0.124
GLO, g/L 6.02 ± 1.98 9.08 ± 3.05 0.042
A/G 4.83 ± 1.11 3.63 ± 0.71 0.042
IgM, mg/dL 11.70 ± 3.77 13.03 ± 5.35 0.593
IgA, mg/dL 1.23 ± 0.07 1.633 ± 0.121 0.002
IgG, mg/dL 123.47 ± 18.23 143.433 ± 23.581 0.098
LDH, U/L 404.87 ± 75.96 456.500 ± 130.304 0.376
IGF 40.71 ± 10.19 50.700 ± 13.356 0.171
Open in a new tab

1Values are least squares means ± SD, n = 8 per treatment. Control = base diet; HDCA = base diet supplemented with 2 g of HDCA/kg of diet.

2BUN = blood urea nitrogen; GLU = glucose; ALP = alkaline phosphatase; AST = aspartate aminotransferase; ALT = alanine aminotransferase; AST/ALT = aspartate aminotransferase/alanine aminotransferase; TP = total protein; ALB = albumin; GLO = globulin; A/G = albumin/globulin; IgM = immunoglobulin M; IgA = immunoglobulin A; IgG = immunoglobulin G; LDH = lactate dehydrogenase.

DISCUSSION

The effects of dietary HDCA supplementation on weaned piglet endocrine differentiation and function and serum biochemical indices were investigated. The data indicated that HDCA had little effect on weaning piglet growth or intestinal morphology. It did regulate intestinal EEC differentiation by affecting mRNA expression of TFs and genes for hormones related to differentiation. Plasma GLP-2 concentrations were reduced by supplementation. Serum biochemical indices measurements showed that dietary HDCA might affect humoral immunity in weaning piglets. These results strongly suggest that adding HDCA influences intestinal EEC differentiation and functioning and may lessen weaning stress damage to the immune functions and enhance resistance.

CgA, a member of the granin family, is located in neuron and endocrine cell vesicles (Taupenot et al., 2003). It is a common EEC marker. Any change in CgA cell densities reflects a change in the total EEC number (Deftos, 1991; El-Salhy et al., 2010; Mazzawi and El-Salhy, 2016). This HDCA supplement was found to decrease the number of CgA-positive cells in the jejunal villi. It tended to decrease in the ileal villi but tended to increase in the duodenal villi. Descending from the duodenum to the rectum, EEC frequency is highest proximally and declines steadily to a nadir in the colon before again increasing in the rectum (Gunawardene et al., 2011). Postprandially, bile acids pass down the intestinal tract, whereas small amounts of unconjugated bile acids are reabsorbed in the upper intestine via passive diffusion. Most bile acids (95%) are reabsorbed in the brush border membrane of the terminal ileum (Chiang, 2013). HDCA concentration would gradually decrease along the digestive tract. This suggests that HDCA may be associated with EEC differentiation. We would expect to see an opposite trend in the number of CgA-positive cells.

Bile acids are amphipathic molecules that facilitate lipid uptake. Their levels fluctuate in the intestine as well as in the circulation depending on food intake. In addition to their role in dietary lipid absorption, bile acids are signaling molecules that activate specific bile acid receptors and trigger downstream signaling cascades (Houten et al., 2006; Thomas et al., 2008). Ipharraguerre (2013) found that the enteral administration of bile acid CDC did not result in improved early-weaned pig intestinal growth, morphology, or inflammation. De Diego-Cabero et al. (2015) found that supplementing early-weaned pigs diets with 60 mg of CDC/kg of initial BW increased the expression of the genes involved in the distal small intestine mucosal protection and barrier function. It did not affect feed intake. These additives had less effect on growth performance or intestinal morphology, but did change endogenous GLP-2 plasma concentrations. A finding that is consistent with our findings.

Glucagon-like peptides are secreted from enteroendocrine L cells in response to nutrients and bile acids (Cottrell et al., 2006; Estall and Drucker, 2006). They control metabolism by acting on structurally related, yet distinct, G protein-coupled receptors. GLP-1 regulates gut motility, appetite, and glucose homeostasis, whereas GLP-2 improves intestinal nutrient absorption while stimulating mucosal enterocyte proliferation (Gunawardene et al., 2011). This experiment demonstrated control and HDCA groups had low GLP-1 and GLP-2 plasma concentrations. Supplementing the base diet with HDCA reduced GLP-2 concentrations, which was consistent with our observation that the total intestinal number of EECs decreased. In a previous study, intragastric administration of a single dose of CDC to piglets during the first 6 d post-early weaning markedly increased endogenous GLP-2 plasma concentrations (Ipharraguerre et al., 2013). That result contrasts with our findings. CDC is a primary bile acid (De Diego-Cabero et al., 2015). HDCA is a secondary bile acid. Primary bile acids are biotransformed by gut microbiota via 2 enzymatic reactions—deconjugation and dihydroxylation—into secondary bile acids (Theriot et al., 2016), how they function may differ. HDCA and CDC doses, as applied and administered, in those 2 studies, differ. We speculate that the particular form of bile acid, its dosage, and how it is administered may explain GLP-2 secretion.

In gut epithelium, the primary differentiated cells are Paneth cells, goblet cells, enterocytes, and EECs. Neurogenin 3 (Ngn3) is required by all intestinal endocrine cells to develop (Jenny et al., 2002; Mellitzer et al., 2010). TFs, acting downstream from Ngn3, are necessary to establish a functional endocrine program or specific progenitors for endocrine lineage progenitors. TFs include NeuroD1, Insm1, Pax4, paired box gene 6 (Pax6), and Nkx2.2 (Naya et al., 1997; Larsson et al., 1998; Jenny et al., 2002; Lee et al., 2002). TF need varies according to endocrine subtype. For example, NeuroD1 is required for cholecystokinin (CCK) and S cells (Naya et al., 1997). Insm1 is required for N cells (Mellitzer et al., 2006) CCK, and PYY cells (Gierl et al., 2006). Nkx2.2 is required for CCK, glucose-dependent insulinotropic peptide, L, D, and N cells (Desai et al., 2008). Pax6 and Pax4 are necessary to control enteroendocrine specifications of intestinal subtypes (Beucher et al., 2012; Du et al., 2012). Experimental results demonstrate that, compared with controls, ileal mRNA NeuroD1 expression decreases. Duodenal Insm1 expression decreased while jejunal HDCA group expression increased. This suggests that the supplement causes EEC duodenal and ileal differentiation reduction, while increasing the jejunal endocrine N, CCK, D, and enterochromaffin (EC) cell differentiation.

Different enteroendocrine subtypes secrete specific hormones, or hormone-like peptides, which, in concert, modulate multiple physiological responses, including GI motility and secretion, glucose homeostasis, and appetite (Psichas et al., 2015). We found that, in the jejunum, Sst and Gast abundances were higher for the HDCA group than for controls. Sst is secreted by D cells and is an inhibitory hormone that reduces all other gut hormone secretions and GI tract exocrine and pancreas functions. Gast is secreted by G cells and stimulates gastric acid secretion (Dhanwate et al., 2015). Experimental results indicate that the HDCA supplements influence EEC functions by altering various hormonal secretions.

The serum biochemical indices reflect piglet health and nutritional status as well as adaptability to the environment, as they undergo physiological and/or pathological changes. The serum immunoglobulin titer is an indicator of humoral immunity (Kong et al., 2007). By protecting the intestinal gut surface against bacterial damage, plasma IgG helps to maintain optimal intestinal functioning and GI development, which, in turn, benefits piglet health and growth (Gomez et al., 1998). Elevated serum IgA and IgG concentrations, as measured in weaned piglets within the HDCA group, evidence that this particular supplement effectively boosts humoral immunity.

In summary, our data show that dietary supplementation with 2 g/kg HDCA could regulate the differentiation of intestinal EECs by affecting mRNA expression of TFs related to the differentiation and may play a role in improving humoral immunity in weaned piglets.

Footnotes

1

This work was supported by Key Programs of Frontier Scientific Research of the Chinese Academy of Sciences (QYZDY-SSW-SMC008), National Natural Science Foundation of China (31330075, 31402089), Natural Science Foundation of Hunan Province (2017JJ1020), and Young Elite Scientists Sponsorship Program by CAST (YESS20160086).

LITERATURE CITED

  1. Beucher A., Gjernes E., Collin C., Courtney M., Meunier A., Collombat P., and Gradwohl G.. 2012. The homeodomain-containing transcription factors Arx and Pax4 control enteroendocrine subtype specification in mice. PLoS One 7:e36449. doi:10.1371/journal.pone.0036449 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Burrin D. G. Stoll B., and Guan X.. 2003. Glucagon-like peptide 2 function in domestic animals. Domest. Anim. Endocrinol. 24:103–122. doi:10.1016/S0739-7240(02)00210-2 [DOI] [PubMed] [Google Scholar]
  3. Chiang J. Y. L. 2013. Bile acid metabolism and signaling. Compr Physiol. 3:1191–1212. doi:10.1002/cphy.c120023 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Cottrell J. J. Stoll B. Buddington R. K. Stephens J. E. Cui L. Chang X., and Burrin D. G.. 2006. Glucagon-like peptide-2 protects against TPN-induced intestinal hexose malabsorption in enterally refed piglets. Am. J. Physiol. Gastrointest. Liver Physiol. 290:G293–G300. doi:10.1152/ajpgi.00275.2005 [DOI] [PubMed] [Google Scholar]
  5. De Diego-Cabero N. Mereu A. Menoyo D. Holst J. J., and Ipharraguerre I. R.. 2015. Bile acid mediated effects on gut integrity and performance of early-weaned piglets. BMC Vet. Res. 11:111. doi:10.1186/s12917-015-0425-6 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Deftos L. J. 1991. Chromogranin A: Its role in endocrine function and as an endocrine and neuroendocrine tumor marker. Endocr. Rev. 12:181–187. doi:10.1210/edrv-12-2-181 [DOI] [PubMed] [Google Scholar]
  7. Desai S. Loomis Z. Pugh-Bernard A. Schrunk J. Doyle M. J. Minic A. McCoy E., and Sussel L.. 2008. Nkx2.2 regulates cell fate choice in the enteroendocrine cell lineages of the intestine. Dev. Biol. 313:58–66. doi:10.1016/j.ydbio.2007.09.047 [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Dhanwate A. D., Ambekar S. A., and Gaikwad M. D.. 2015. Enteroendocrine cells unfolding the mystery. Int. J. Curr. Res. 7:16246–16251. [Google Scholar]
  9. Du A. McCracken K. W. Walp E. R. Terry N. A. Klein T. J. Han A. Wells J. M., and May C. L.. 2012. Arx is required for normal enteroendocrine cell development in mice and humans. Dev. Biol. 365:175–188. doi:10.1016/j.ydbio.2012.02.024 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. El-Salhy M. Lomholt-Beck B., and Hausken T.. 2010. Chromogranin A as a possible tool in the diagnosis of irritable bowel syndrome. Scand. J. Gastroenterol. 45:1435–1439. doi:10.3109/00365521.2010.503965 [DOI] [PubMed] [Google Scholar]
  11. Estall J. L., and Drucker D. J.. 2006. Glucagon-like peptide-2. Annu. Rev. Nutr. 26:391–411. doi:10.1146/annurev.nutr.26.061505.111223 [DOI] [PubMed] [Google Scholar]
  12. Gierl M. S. Karoulias N. Wende H. Strehle M., and Birchmeier C.. 2006. The zinc-finger factor insm1 (IA-1) is essential for the development of pancreatic beta cells and intestinal endocrine cells. Genes Dev. 20:2465–2478. doi:10.1101/gad.381806 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Gomez G. G. Phillips O., and Goforth R. A.. 1998. Effect of immunoglobulin source on survival, growth, and hematological and immunological variables in pigs. J. Anim. Sci. 76:1–7. doi:10.2527/1998.7611 [DOI] [PubMed] [Google Scholar]
  14. Gunawardene B. M. Corfe B. M., and Staton C. A.. 2011. Classification and functions of enteroendocrine cells of the lower gastrointestinal tract. Int. J. Exp. Pathol. 92:219–231. doi:10.1111/j.1365-2613.2011.00767.x [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Hansen M. Scheltema M. J. Sonne D. P. Hansen J. S. Sperling M. Rehfeld J. F. Holst J. J. Vilsbøll T., and Knop F. K.. 2016. Effect of chenodeoxycholic acid and the bile acid sequestrant colesevelam on glucagon-like peptide-1 secretion. Diabetes. Obes. Metab. 18:571–580. doi:10.1111/dom.12648 [DOI] [PubMed] [Google Scholar]
  16. He L. Yang H. Hou Y. Li T. Fang J. Zhou X. Yin Y. Wu L. Nyachoti M., and Wu G.. 2013. Effects of dietary l-lysine intake on the intestinal mucosa and expression of CAT genes in weaned piglets. Amino Acids 45:383–391. doi:10.1007/s00726-013-1514-0 [DOI] [PubMed] [Google Scholar]
  17. Houten S. M. Watanabe M., and Auwerx J.. 2006. Endocrine functions of bile acids. Embo J. 25:1419–1425. doi:10.1038/sj.emboj.7601049 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Ipharraguerre I. R. Tedó G. Menoyo D. de Diego-Cabero N. Holst J. J. Nofrarías M. Mereu A., and Burrin D. G.. 2013. Bile acids induce glucagon-like peptide 2 secretion with limited effects on intestinal adaptation in early weaned pigs. J. Nutr. 143:1899–1905. doi:10.3945/jn.113.177865 [DOI] [PubMed] [Google Scholar]
  19. Jain A. K. Stoll B. Burrin D. G. Holst J. J., and Moore D. D.. 2012. Enteral bile acid treatment improves parenteral nutrition-related liver disease and intestinal mucosal atrophy in neonatal pigs. Am. J. Physiol. Gastrointest. Liver Physiol. 302:G218–G224. doi:10.1152/ajpgi.00280.2011 [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Jenny M. Uhl C. Roche C. Duluc I. Guillermin V. Guillemot F. Jensen J. Kedinger M., and Gradwohl G.. 2002. Neurogenin3 is differentially required for endocrine cell fate specification in the intestinal and gastric epithelium. Embo J. 21:6338–6347. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Katsuma S. Hirasawa A., and Tsujimoto G.. 2005. Bile acids promote glucagon-like peptide-1 secretion through TGR5 in a murine enteroendocrine cell line STC-1. Biochem. Biophys. Res. Commun. 329:386–390. doi:10.1016/j.bbrc.2005.01.139 [DOI] [PubMed] [Google Scholar]
  22. Kong X. F., Yin Y., Wu G., Liu H., Yin F., Li T., Huang R., Ruan Z., Xiong H., Deng Z., Xie M., Liao Y., and Kim S.. 2007. Dietary supplementation with Acanthopanax senticosus extract modulates cellular and humoral immunity in weaned piglets. Asian-Aust. J. Anim. Sci. 20:1453–1461. doi:10.5713/ajas.2007.1453 [Google Scholar]
  23. Larsson L. I St-Onge L. Hougaard D. M. Sosa-Pineda B., and Gruss P.. 1998. Pax 4 and 6 regulate gastrointestinal endocrine cell development. Mech. Dev. 79:153–159. doi:S0925-4773(98)00182-8 [DOI] [PubMed] [Google Scholar]
  24. Lee J. Koehler J. Yusta B. Bahrami J. Matthews D. Rafii M. Pencharz P. B., and Drucker D. J.. 2017. Enteroendocrine-derived glucagon-like peptide-2 controls intestinal amino acid transport. Mol. Metab. 6:245–255. doi:10.1016/j.molmet.2017.01.005 [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Lee C.S., Perreault N., Brestelli J. E., and Kaestner K. H.. 2002. Neurogenin 3 is essential for the proper specification of gastric enteroendocrine cells and the maintenance of gastric epithelial cell identity. Genome Res. 12:47–56. doi:10.1101/gad.985002 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Mazzawi T., and El-Salhy M.. 2016. Changes in small intestinal chromogranin A-immunoreactive cell densities in patients with irritable bowel syndrome after receiving dietary guidance. Int. J. Mol. Med. 37:1247–1253. doi:10.3892/ijmm.2016.2523 [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Mellitzer G. Beucher A. Lobstein V. Michel P. Robine S. Kedinger M., and Gradwohl G.. 2010. Loss of enteroendocrine cells in mice alters lipid absorption and glucose homeostasis and impairs postnatal survival. J. Clin. Invest. 120:1708–1721. doi:10.1172/JCI40794 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Mellitzer G., Bonné S., Luco R. F., Van De Casteele M., Lenne-Samuel N., Collombat P., Mansouri A., Lee J., Lan M., Pipeleers D., et al. 2006. IA1 is NGN3-dependent and essential for differentiation of the endocrine pancreas. Embo J. 25:1344–1352. doi:10.1038/sj.emboj.7601011 [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. National Research Council 2012. Nutrient requirements of swine. 11th ed. Natl. Acad. Press, Washington, DC. [Google Scholar]
  30. Naya F. J. Huang H. P. Qiu Y. Mutoh H. DeMayo F. J. Leiter A. B., and Tsai M. J.. 1997. Diabetes, defective pancreatic morphogenesis, and abnormal enteroendocrine differentiation in BETA2/neurod-deficient mice. Genes Dev. 11:2323–2334. doi:10.1101/gad.11.18.2323 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Petersen Y. M. Hartmann B. Holst J. J. Le Huerou-Luron I. Bjørnvad C. R., and Sangild P. T.. 2003. Introduction of enteral food increases plasma GLP-2 and decreases GLP-2 receptor mRNA abundance during pig development. J. Nutr. 133:1781–1786. doi:10.1093/jn/133.6.1781 [DOI] [PubMed] [Google Scholar]
  32. Psichas A. Reimann F., and Gribble F. M.. 2015. Gut chemosensing mechanisms. J. Clin. Invest. 125:908–917. doi:10.1172/JCI76309 [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Rindi G. Leiter A. B. Kopin A. S. Bordi C., and Solcia E.. 2004. The “normal” endocrine cell of the gut: Changing concepts and new evidences. Ann. N. Y. Acad. Sci. 1014:1–12. doi:10.1196/annals.1294.001 [DOI] [PubMed] [Google Scholar]
  34. Saqui-Salces M., Huang Z., Ferrandis Vila M., Li J., Mielke J. A., Urriola P. E., and Shurson G. C.. 2017. Modulation of intestinal cell differentiation in growing pigs is dependent on the fiber source in the diet. J. Anim. Sci. 95:1179–1190. doi:10.2527/jas2016.0947 [DOI] [PubMed] [Google Scholar]
  35. Taupenot L., Harper K. L., and O’Connor D. T.. 2003. The chromogranin–secretogranin family. New Engl. J. Med. 348:1134–1149. doi:10.1056/NEJMra02405 [DOI] [PubMed] [Google Scholar]
  36. Theriot C. M., Bowman A. A., and Young V. B.. 2016. Antibiotic-induced alterations of the gut microbiota alter secondary bile acid production and allow for Clostridium difficile spore germination and outgrowth in the large intestine. mSphere 1:e00045–e00015. doi:10.1128/mSphere.00045-15 [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Thomas C. Pellicciari R. Pruzanski M. Auwerx J., and Schoonjans K.. 2008. Targeting bile-acid signalling for metabolic diseases. Nat. Rev. Drug Discov. 7:678–693. doi:10.1038/nrd2619 [DOI] [PubMed] [Google Scholar]
  38. Xiong X., Yang H. S., Wang X. C., Hu Q., Liu C. X., Wu X., Deng D., Hou Y. Q., Nyachoti C. M., Xiao D. F., et al. 2015. Effect of low dosage of chito-oligosaccharide supplementation on intestinal morphology, immune response, antioxidant capacity, and barrier function in weaned piglets. J. Anim. Sci. 93:1089–1097. doi:10.2527/jas.2014-7851 [DOI] [PubMed] [Google Scholar]
  39. Yan S. Long L. Zong E. Huang P. Li J. Li Y. Ding X. Xiong X. Yin Y., and Yang H.. 2018. Dietary sulfur amino acids affect jejunal cell proliferation and functions by affecting antioxidant capacity, wnt/β-catenin, and the mechanistic target of rapamycin signaling pathways in weaning piglets. J. Anim. Sci. 96:5124–5133. doi:10.1093/jas/sky349 [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Yang H. Wang X. Xiong X., and Yin Y.. 2016a. Energy metabolism in intestinal epithelial cells during maturation along the crypt-villus axis. Sci. Rep. 6:31917. doi:10.1038/srep31917 [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Yang H. S., Wu F., Long L. N., Li T. J., Xiong X., Liao P., Liu H. N., and Yin Y. L.. 2016b. Effects of yeast products on the intestinal morphology, barrier function, cytokine expression, and antioxidant system of weaned piglets. J. Zhejiang Univ. Sci. B 17:752–762. doi:10.1631/jzus.B1500192 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Yang H., Xiong X., Wang X., Li T., and Yin Y.. 2016c. Effects of weaning on intestinal crypt epithelial cells in piglets. Sci. Rep. 6:36939. doi:10.1038/srep36939 [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Zhou X., Wu X., Yin Y., Zhang C., and He L.. 2012. Preventive oral supplementation with glutamine and arginine has beneficial effects on the intestinal mucosa and inflammatory cytokines in endotoxemic rats. Amino Acids 43:813–821. doi:10.1007/s00726-011-1137-2 [DOI] [PubMed] [Google Scholar]

Articles from Journal of Animal Science are provided here courtesy of Oxford University Press

RESOURCES