QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) Online Supporting Material Purchase Article View Shopping Cart Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wang, J. Articles by Wu, G. Search for Related Content PubMed PubMed Citation Articles by Wang, J. Articles by Wu, G.

---

Genomics, Proteomics, and Metabolomics

Gene Expression Is Altered in Piglet Small Intestine by Weaning and Dietary Glutamine Supplementation^1–3,

⁴ State Key Laboratory of Animal Nutrition, China Agricultural University, Beijing, China 100094; ⁵ Texas Agricultural Experiment Station, The Texas A&M University System, College Station, TX 77843; ⁶ Institute of Subtropical Agriculture, Chinese Academy of Sciences, Changsha, Hunan, China 410128; and ⁷ College of Animal Science and Technology, Hunan Agricultural University, Changsha, Hunan, China 410128

^* To whom correspondence should be addressed: g-wu{at}tamu.edu, wangfl{at}mafic.ac.cn, or yinyulong{at}isa.ac.cn.

	ABSTRACT

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
Dietary supplementation of glutamine prevents intestinal dysfunction and atrophy in weanling piglets, but the underlying mechanism(s) are largely unknown. This study was conducted to test the hypothesis that weaning or glutamine may modulate expression of genes that are crucial for intestinal metabolism and function. In Expt. 1, we obtained small intestine from 28-d-old pigs weaned at 21 d of age and from age-matched suckling piglets. In Expt. 2, piglets were weaned at 21 d of age and then had free access to diets supplemented with 1% L-glutamine (wt:wt) or isonitrogenous L-alanine (control). At d 28, we collected small intestine for biochemical and morphological measurements and microarray analysis of gene expression using the Operon Porcine Genome Oligo set. Early weaning resulted in increased (52–346%) expression of genes related to oxidative stress and immune activation but decreased (35–77%) expression of genes related to macronutrient metabolism and cell proliferation in the gut. Dietary glutamine supplementation increased intestinal expression (120–124%) of genes that are necessary for cell growth and removal of oxidants, while reducing (34–75%) expression of genes that promote oxidative stress and immune activation. Functionally, the glutamine treatment enhanced intestinal oxidative-defense capacity (indicated by a 29% increase in glutathione concentration), prevented jejunal atrophy, and promoted small intestine growth (+12%) and body weight gain (+19%) in weaned piglets. These findings reveal coordinate alterations of gene expression in response to weaning and aid in providing molecular mechanisms for the beneficial effect of dietary glutamine supplementation to improve nutrition status in young mammals.

	Introduction

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

Weaning is associated with reduced food consumption by young mammals, including piglets (8), therefore decreasing the intake of glutamine from the diet. Notably, weanling piglets often experience intestinal dysfunction and atrophy (9). Recent work has shown that dietary supplementation with glutamine prevents these gut problems (10). However, the underlying mechanism(s) are largely unknown. We hypothesized that weaning or glutamine may modulate expression of genes that are crucial for intestinal metabolism and function. This hypothesis was tested in the present study using microarray technology, which provides a powerful discovery tool to simultaneously analyze expression of thousands of genes in a tissue (11).

	Materials and Methods

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
All animals used in this study were humanely managed according to the established guidelines of the USDA. The experimental protocol was approved by the Texas A&M University Institutional Animal Care and Use Committee.

    Animals and tissue collection. Pregnant sows were fed daily a 2-kg gestating diet during the entire period of pregnancy that met the NRC-recommended requirements for nutrients (12). After farrowing, sows had free access to a corn- and soybean meal-based diet that also met NRC-recommended requirements (13). Each sow freely nursed 9 piglets before weaning at 21 d of age. All sows had free access to drinking water during gestation and lactation periods. Neonatal piglets were used in 2 series of experiments.

In Expt. 1, 24 21-d-old piglets with similar body weights from 6 litters (4 piglets per litter) were assigned randomly to 1 of the 2 groups on the basis of their litter origins (n = 12/group). Piglets in group 1 continued to be nursed by sows, whereas piglets in group 2 were weaned and housed in pens (2 piglets per pen) of the same animal facilities and had free access to a corn- and soybean meal-based diet and drinking water, as we previously described (10). Milk consumption by suckling piglets was measured with another similar group of 6 piglets over an 8-h period at d 21, 24, and 28 of age, using the weight-suckle-weight technique (14) and the mean value for the 3 measurements was used to represent nutrient intake by a piglet. An additional group of piglets was used for the measurement of milk consumption, because this technique was associated with stress on piglets. Feed intake by weanling piglets was determined during the 7-d period after weaning. On d 28, at 1 h after suckling or feeding, blood samples (3 mL) were obtained from the jugular vein and piglets were then humanely killed after anesthesia, as we previously described (3). Plasma was obtained after centrifugation at 12,000 x g; 1 min and stored at –80°C. The whole small intestine was weighed and its length was measured after careful removal of luminal contents. The luminal contents were centrifuged at 12,000 x g; 1 min. The supernatant fluid was placed in liquid nitrogen and stored at –80°C until analysis for glutamine. In neonatal pigs, the small intestine was defined as the portion of the digestive tract between the pylorus and the ileocecal valve, with the first 10-cm segment being duodenum (13). The jejunum and ileum constituted 40 and 60%, respectively, of the small intestine below the duodenum (13). A portion of mid-jejunum (3 cm each in length) was placed in 4% paraformaldehyde for subsequent analysis of morphology (14) and another set of samples (10 cm long) were obtained for glutathione analysis (15). Jejunal samples (5 g) were placed in RNAlater solution (Ambion) and stored at –80°C before use for the isolation of total RNA.

Expt. 2 was conducted as for Expt. 1, except that 24 piglets were weaned and then assigned randomly to 1 of the 2 treatment groups (n = 12/group), representing supplementation with 1% L-glutamine (wt:wt) and 1.22% L-alanine (wt:wt; isonitrogenous control) to the corn- and soybean meal-based diet (10). The basal diet, which contained 21.0% crude protein (on an as-fed basis; 89.6% dry matter), was analyzed for amino acids using HPLC (1,7). The contents (percent of diet on an as-fed basis) of amino acids were as follows: arginine, 1.34; alanine, 1.27; aspartate+asparagine, 2.18; glutamate+glutamine, 3.36; glycine, 0.86; histidine, 0.56; isoleucine, 0.87; leucine, 1.73; lysine, 1.38; methionine, 0.35; phenylalanine, 0.97; proline, 1.52; serine, 0.76; threonine, 0.83; tryptophan, 0.24; tyrosine, 0.74; and valine, 0.97. The amount of supplemental glutamine was based on the results of our previous study that indicated its efficacy in preventing intestinal atrophy and enhancing growth in early-weaned piglets (10). At 28 d of age, 1 h after feeding, tissues were obtained, processed, and stored, as described in Expt. 1.

Alanine was chosen for isonitrogenous control from among nonessential amino acids on the basis of the following considerations. First, in contrast to glycine, alanine is not an antioxidant (16). Second, glycine, glutamate, aspartate, and metabolites of tyrosine (dopa and dopamine) are neurotransmitters (17); therefore, dietary supplementation of these amino acids may alter food intake by pigs. In addition, glutamate is a substrate for the synthesis of glutamine (10) and glutathione (16). Third, both glutamate and proline are major precursors for endogenous synthesis of arginine (5) and, thus, their addition to the diet may augment arginine provision in vivo (18). Fourth, in animals, serine and asparagine are readily converted to glycine and aspartate, respectively (4). Fifth, tyrosine metabolism via tyrosine hydroxylase requires tetrahydrobiopterin (17); therefore, its supplementation may reduce the availability of this essential cofactor for the generation of nitric oxide [a major vasodilator and a key regulator of smooth muscle relaxation (19)]. Sixth, cysteine is a sulfur-containing amino acid and a substrate for the synthesis of glutathione (an antioxidant) (16). The catabolism of increased amounts of cysteine leads to the production of H₂SO₄, which seriously disturbs acid-base balance in animals (17). Finally, all amino acids, except for leucine and lysine, are potential substrates for hepatic gluconeogenesis. However, alanine is not toxic and animals (including piglets) have a high capacity to catabolize this neutral amino acid (18). Thus, among nonessential amino acids, alanine is most appropriate for isonitrogenous control in this study. In our preliminary study, we found that adding 1.22% L-alanine (wt:wt) to the basal diet for 21- to 28-d-old weaned pigs did not affect food intake, small-intestinal weight and morphology, concentrations of glutamine and glutathione in jejunal mucosa, concentrations of glucose and arginine in plasma, whole-body weight gain, or expression of genes in the small intestine compared with no alanine supplementation (n = 6 pigs/treatment group; G. Wu, unpublished data).

    Histological and biochemical analyses of the small intestine. Villus height, crypt depth, and lamina propria depth in jejunum were determined with the aid of a microscope (10x magnification), as we previously described (10,13). Free glutamine in the jejunal luminal content was analyzed using the HPLC method involving precolumn derivatization with o-phthaldialdehyde (1). Jejunal reduced glutathione (GSH)⁸ and oxidized glutathione (GSSG) were measured using an HPLC method of Jones et al. (15), except that: 1) fluorescence detection (Waters 2475 Multi Fluorescence Detector) was set at 590-nm excitation and 610-nm emission (0.0–7.5 min) to eliminate the appearance of amino acid peaks and at 335-nm excitation and 610-nm emission (7.5–38 min) for GSH and GSSG detection; and 2) gain of the detection was set at 100 (0–32.2 min) for GSH detection and at 1000 (32.2–38 min) for GSSG detection. GSH and GSSG were quantified on the basis of authentic standards (Sigma Chemicals) using the Millennium-32 software and workstation.

    RNA extraction. Frozen tissue (0.5 g) was homogenized in 5 mL TRIzol reagent (Invitrogen) and total RNA was isolated according to the manufacturer's recommendations. RNA integrity and quantity were analyzed using the Agilent 2100 Bioanalyzer and RNA 6000 NanoLabChip kit (Agilent catalog no. 5065–4474). The 28S ribosome:18S ribosome peak areas ratio was 1.80 for all samples, indicating little degradation of RNA.

    Generation of labeled antisense RNA for microarray. One microgram of total RNA was used as the starting material for amplification using Amino Allyl MessageAmp II aRNA Amplification kit (Ambion catalog no. 1753). The procedure consisted of reverse transcription with an oligo(dT) primer bearing a T7 promoter and arrayscript, a reverse transcriptase enzyme that catalyzes the synthesis of a full-length cDNA. The cDNA underwent second-strand synthesis and then used as template for in vitro transcription with T7 RNA polymerase and the incorporation of the modified nucleotide, 5-(3-aminoallyl)-UTP, into antisense RNA (aRNA). The aminoallyl-UTP contained a reactive primary amino group on the C5 position of uracil that was coupled to N-hydroxysuccinimidyl eater-derivatized reactive dyes (Cy3 and Cy5). In vitro transcription generated hundreds to thousands of aRNA copies of each mRNA in the sample. The quality and quantity of the amplified RNA were analyzed via capillary electrophoresis using RNA nano-assay technique (Agilent 2100 Bioanalyzer). No ribosomal RNA (rRNA) contamination was detected for all samples, indicating the high quality of the aRNA obtained.

Five micrograms of aRNA was used for the coupling reaction using the Mono-Reactive Cy3 and Cy5 dyes (Amersham Biosciences). In Expt. 1, jejunal RNA samples from each of 4 sow-reared piglets were used as controls and labeled with Cy3 (green), whereas RNA samples from each of 4 weaned piglets were labeled with Cy5 (red). In Expt. 2, jejunal RNA samples from alanine (control)- and glutamine-supplemented weaned piglets (n = 4/group) were labeled with Cy3 and Cy5, respectively. A dye swap was adopted for each experiment.

    Microarray hybridization and analysis. One microgram of each labeled aRNA was used for hybridization to the porcine oligomicroarray. The paired aRNA samples were combined and fragmented using fragmentation reagents (catalog no. 8740, Ambion) and hybridized overnight with hybridization buffer (SlideHyb catalog no. 8861, Ambion) at 48°C to glass arrays with the Operon Pig Genome Oligo Set containing 11,000 oligonucleotides (genes spotted by the Microarray Core Facility, Department of Systems Biology and Translational Medicine, Texas A&M Health Science Center, Temple, TX). Following the hybridization, the arrays were washed and scanned using a GenePix 4000A scanner (Axon Instruments).

    Real-time RT-PCR confirmation of gene expression. Real-time RT-PCR technology was employed to verify changes in the mRNA levels of select genes (Supplemental Table 1) obtained from the microarray analysis. First-strand cDNAs were synthesized from 1 µg of total RNA (10 ng for internal standard: 18S RNA) using oligo (deoxythymidine) primers, random hexamer primers, and SuperScript II Reverse Transcriptase, as we described (20). RT-PCR analysis was performed using the SYBR Green method and the ABI 7900 Sequence Detection System (Applied Biosystems). The thermal cycling parameters were as follows: 50°C for 2 min, 95°C for 10 min, followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. Primers were designed using Primer Express Software version 1.5 (Applied Biosystems) (Supplemental Table 1). Values for cycle threshold, the value of cycle at which the fluorescence achieves a predetermined threshold, were determined using the Applied Biosystems Software. The cycle threshold values were analyzed using the generalized estimating equations model and the PROC GENMOD procedure of the Statistical Analysis System, as we described (21). All of the data were normalized with the 18S rRNA gene in the same samples and are expressed as the relative values to those of piglets fed the control diet.

    Statistical analysis. Results are expressed as means ± SEM. Data on tissue metabolite concentrations and intestinal morphology were statistically analyzed using an unpaired t test. In microarray analysis, the acquired data were transformed (to accommodate the dye swap), normalized, and filtered using the GeneSpring v7.2 software package (Silicon Genetics, Agilent Technologies). Gene expression significance was assessed using multiple t tests and the Benjamini Hochberg false discovery rate multiple testing correction (20,21). P-values 0.05 were considered significant. The GeneSpring v7.2 software was used to categorize the genes that were up- and downregulated.

	Results

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
    Food intake, body weights, and small intestine weight. Food intake and daily body weight gain between 21 and 28 d of age were reduced (P < 0.01) by 36 and 47%, respectively, in weaned pigs compared with age-matched suckling piglets (Expt. 1; Table 1). At d 28, the small intestine weight was 26% lower (P < 0.01) in weaned than in sow-reared piglets (Table 1). Supplementing 1.0% L-glutamine to the diet for weaned piglets did not affect feed intake but increased (P < 0.05) daily body weight gain between 21 and 28 d of age by 19% compared with the control group (Expt. 2; Table 1).

    Intestinal morphology and glutathione concentrations. Villus height, crypt depth, and lamina-propria depth in the jejunum of 21-d-old sow-reared piglets were 345 ± 14, 203 ± 12, and 205 ± 13 µm, respectively. Villus height in the jejunum was reduced (P < 0.01) by 43% in 28-d-old weaned piglets than in age-matched suckling piglets (Expt. 1, Table 3). Crypt depth or lamina propria depth did not differ between weaned and sow-reared piglets. Dietary glutamine supplementation increased (P < 0.01) jejunal villus height by 38% but had no effect on crypt depth or lamina propria depth (Expt. 2, Table 3).

	Discussion

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
After mammalian neonates are weaned from their mothers, they undergo tremendous changes in intestinal structure and function (8,9). However, the underlying molecular and cellular mechanisms are largely unknown. Due to the invasive nature of biochemical research on intestinal development, the piglet provides a useful animal model for studying the responses of the neonatal gut to weaning (10). With the recent availability of the microarray technology, which can analyze simultaneously expression of thousands of genes in a tissue (11), we identified significant changes in intestinal expression of key regulatory genes in response to weaning and dietary glutamine supplementation (Tables 4–6). Functionally, the glutamine treatment resulted in increased oxidative-defense capacity (Table 2), prevention of intestinal atrophy (Table 3), and enhanced growth performance (Table 1) of early-weaned piglets.

Weaning was associated with reduced expression of 18 genes out of 11,000 in the piglet small intestine (Table 4). Interestingly, these genes encode for proteins that are related to: 1) the regulation of gene expression [DNA-binding protein inhibitor ID-2 (22)]; 2) protein and peptide degradation [aminopeptidase A, cathepsin F, and ubiquitin carboxyl-terminal hydrolase (23)]; 3) lipid metabolism [acyl-CoA dehydrogenase, carnitine transporter 2, and oxysterol binding protein-related protein 10 (24)]; 4) signal transduction [adenylate cyclase and ADP-ribosylation factor GTPase-activating protein I (25)]; 5) immune function [leukocyte antigen-related protein and vanin-1 (26)]; 6) growth regulation (insulin-like growth factor II precursor and somatostatin precursor); 7) aminosugar synthesis [N-acyl-D-glucosamine 2-epinerase (27)]; and 8) intestinal transport [apolipoprotein A-IV precursor, fatty acid-binding protein, sodium- and chloride-dependent creatine transporter I, and preprogalanin (28)]. Downregulation of these genes is expected to reduce oxidative defense capacity, intestinal transport and utilization of dietary nutrients (particularly lipids and proteins), immune response, and synthesis of glycoproteins (major proteins secreted by the intestinal mucosa), as well as proliferation and differentiation of intestinal epithelial cells. This is consistent with the previous report that intestinal atrophy and dysfunction often occur in early-weaned piglets (8,9) in association with an increased GSSG:GSH ratio (Table 3), an indicator of cellular oxidative stress (16).

Expression of 21 genes was enhanced in the small intestine of piglets in response to early weaning (Table 5). These genes encode for proteins that play crucial roles in regulating: 1) lipid metabolism [3-β-hydroxysteroid-⁸, ⁷-isomerase, C-4 methyl sterol oxidase, diphosphomevalonate decarboxylase, hydroxymethylglutaryl-CoA synthase, and squalene epoxidase (29,30)]; 2) water secretion [aquaporin-8 (31)]; 3) aminosugar degradation (core 2-β-16-N-acetyl-glucosaminyltransferase); 4) xenobiotic metabolism and oxidative defense [cytochrome P450, glutathione transferase -1 (32), and nudix hydrolase-5 (33)]; 5) carbohydrate metabolism (galactoside 2--L-fucosyltransferase); 6) immune function (Ig chain C, Ig chain C, Ig chain, Ig J chain, lysozyme, polymeric Ig receptor precursor); and 7) cell division [septin-5 (34)]. Emerging evidence shows that cAMP mediates expression of aquaporin-8 expression in small intestinal mucosa in response to cholera toxin (35). The enhanced expression of aquaporin-8, which stimulates water permeation across the gut (31), may help explain the increased secretion of water from intestinal mucosal cells and the malabsorption of water from the intestinal lumen and, therefore, diarrhea that frequently occurs in early-weaned piglets (36). Additionally, weaned piglets were fed a typical corn- and soybean meal-based diet that contained high levels of plant-origin antigens (37). Results from the present study indicate that a response to such a diet was increased expression of genes responsible for the synthesis of Ig in the small intestine (Table 4). Also, the increased expression of glutathione S-transferase [which catalyzes the conjugation of GSH to electrophilic substances (32)] and nudix hydrolase-5 [which eliminates toxic nucleotide derivatives from cells and regulates the levels of signaling nucleotides (33)] suggests the presence of oxidative stress in the small intestine of weaned piglets. In support of this view, the GSSG:GSH ratio in the jejunum was elevated by 59% in response to early weaning (Table 2).

Previous studies have shown that glutamine regulates MAPK activation and C-Jun signaling in enterocytes (38). In addition, this amino acid enhances intestinal oxidative metabolism, polyamine synthesis, ion transport, and cell proliferation (39,40) as well as cytoprotection via heat shock proteins (41,42). Consistent with these reports, we found that, in response to dietary glutamine supplementation, expression of 8 genes was downregulated in the small intestine (Table 6). These genes are related to cellular signaling transduction [casein kinase 1 epsilon and MAPK-6 (43)], the cell cycle [Rho-related GTP-binding protein (44)], apoptosis [KLF-10 (45)], immune activation (La antoantigen homolog and ICAM-1), protein modification [peptidyl-prolyl isomerase G (46)], and gene expression (pre-mRNA cleavage complex II). Enhanced expression of antigens and activation of leukocytes (e.g. lymphocytes and macrophages) in the intestinal epithelium because of activation of the MAPK-6 signaling may contribute to intestinal dysfunction and diarrhea in weanling pigs (36). In addition, peptidyl-prolyl isomerase G plays a role in posttranslational protein modifications and cell proliferation (46), which is likely unfavorable for intestinal integrity in weanling piglets. Likewise, ICAM-1 associates with receptors of the integrin family proteins, thereby playing an important role in immune activation (47). Also, Rho GTPase inhibits cell cycle progression by decreasing Ras (a signal transduction protein with an abbreviation that originated from rat sarcoma)- and Raf (an oncogene that encodes protein kinase with an abbreviation from rat fibrosarcoma)-induced fibroblast transformation (44). Furthermore, KLF-10 induces and promotes apoptosis through the mitochondrial apoptotic pathway (48). Therefore, downregulation of these genes provides an additional explanation for the effect of dietary glutamine supplementation on reducing intestinal damage and enhancing intestinal cell proliferation in early-weaned piglets (10).

Another beneficial effect of glutamine supplementation is increased expression of 6 genes (Table 6) related to transcription regulation [AF-9 protein (49)], lipid metabolism (endozepine), iron absorption (heme-binding protein), cytoskeletal structure and function (myosin), defense against pathological microorganisms [IL-13R--1 (50) and endozepine (51)], and regulation of nutrient metabolism (signal recognition particle 72K chain). AF-9 is a transcription factor that regulates gene expression and cell growth (52). IL-13 stimulates contractility of intestinal smooth muscle (50) and, thus, the movement of luminal digesta along the small intestine, which facilitates the digestion of macronutrients by various digestive enzymes and the absorption of resultant smaller molecules by enterocytes. Signal recognition particle (a cytoplasmic ribonucleoprotein), which consists of 1 RNA and 6 proteins, regulates gene expression and protein function (53). Additionally, endozepine, which is expressed in intestinal mucosal cells (54), has multiple functions in intestinal metabolism, physiology, and immunology. This polypeptide acts like acyl CoA-binding protein, therefore regulating lipid metabolism, assembly, and trafficking across the small intestine (55). Also, porcine endozepine has a potent antibacterial activity in the porcine gut (56). Because early weaning is often associated with immunological challenges in the intestine (36), elevated expression of endozepine in glutamine-supplemented piglets may protect the gut from infections during weaning. Collectively, these findings provide a mechanism for the effect of dietary glutamine supplementation on preventing intestinal atrophy and improving nutrient digestion and utilization in early-weaned piglets (10).

In conclusion, results of the microarray analysis reveal that early weaning resulted in increased expression of genes that promote oxidative stress and immune activation but decreased expression of genes related to nutrient utilization and cell proliferation in the piglet small intestine. In contrast, dietary supplementation of glutamine to weanling piglets enhanced expression of genes that prevent oxidative stress, improve antibacterial activity, enhance nutrient absorption, and stimulate cell growth. These novel findings reveal coordinate alterations of gene expression in response to weaning and aid in providing molecular mechanisms for explaining the previous observation that dietary glutamine supplementation prevents intestinal dysfunction and enhances the growth performance of early-weaned piglets. Further work is necessary to determine how these changes in gene expression translate into enhanced gut growth and function.

	ACKNOWLEDGMENTS

 
We thank Haijun Gao for technical assistance, Kenton Lillie for help with pig management, and Frances Mutscher for office support.

	FOOTNOTES

 
¹ Supported by the National Basic Research Program of China (no. 2004CB117503), Natural Science Foundation of China (nos. 30525029, 30528006, 30600434, 30671517, 30671522, 30700581, and 30771558), Outstanding Overseas Chinese Scholar Fund of the Chinese Academy of Sciences (no. 2005-1-4), Beijing Municipal Natural Science Foundation (no. 6082017), Texas Agricultural Experiment Station (no. H-8200), and National Research Initiative Competitive Grant (2008-35206-18764) from the USDA Cooperative State Research, Education, and Extension Service.

² Author disclosures: J. J. Wang, L. X. Chen, P. Li, X. L. Li, H. J. Zhou, F. L. Wang, D. F. Li, Y. L. Yin, and G. Wu, no conflicts of interest.

³ Supplemental Table 1 is available with the online posting of this paper at jn.nutrition.org.

⁸ Abbreviations used: aRNA, antisense RNA; GSH, reduced glutathione; GSSG, oxidized glutathione; ICAM-1, intercellular adhesion molecule-1; IL, interleukin; KLF-10, transforming growth factor β-inducible early growth response protein I krueppel-like factor 10; MAPK, mitogen-activated protein kinase; rRNA, ribosomal RNA.

Manuscript received 13 February 2008. Initial review completed 10 March 2008. Revision accepted 13 March 2008.

	LITERATURE CITED

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 

1. Wu G, Knabe DA. Free and protein-bound amino acids in sow's colostrums and milk. J Nutr. 1994;124:415–24.[Abstract/Free Full Text]

2. Davis TA, Nguyen HV, Garciaa-Bravo R, Fiorotto ML, Jackson EM, Lewis DS, Lee DR, Reeds PJ. Amino acid composition of human milk is not unique. J Nutr. 1994;124:1126–32.[Abstract/Free Full Text]

3. Reeds PJ, Burrin DG. The gut and amino acid homeostasis. Nutrition. 2000;16:666–8.[Medline]

4. Wu G. Intestinal mucosal amino acid catabolism. J Nutr. 1998;128:1249–52.[Abstract/Free Full Text]

5. Wu G, Morris SM. Arginine metabolism: nitric oxide and beyond. Biochem J. 1998;336:1–17.[Medline]

6. Flynn NE, Wu G. An important role for endogenous synthesis of arginine in maintaining arginine homeostasis in neonatal pigs. Am J Physiol Regul Integr Comp Physiol. 1996;271:R1149–55.[Abstract/Free Full Text]

7. Wang X, Qiao SY, Yin YL, Yue LY, Wang ZY, Wu G. Deficiency or excess of dietary threonine reduces protein synthesis in jejunum and skeletal muscle of young pigs. J Nutr. 2007;137:1442–6.[Abstract/Free Full Text]

8. Miller BG, James PS, Smith MW. Effect of weaning on the capacity of pig intestinal villi to digest and absorb nutrients. J Sci Food Agric. 1986;107:579–89.

9. Gu X, Li D, She R. Effect of weaning on small intestinal structure and function in the piglet. Arch Anim Nutr. 2002;56:275–86.

10. Wu G, Meier SA, Knabe DA. Dietary glutamine supplementation prevents jejunal atrophy in weaned pigs. J Nutr. 1996;126:2578–84.[Abstract/Free Full Text]

11. Nguyen DV, Arpat AB, Wang N, Carroll RJ. DNA microarray experiments: biological and technological aspects. Biometrics. 2002;58:701–17.[CrossRef][Medline]

12. Wu G, Bazer FW, Tuo W. Developmental changes of free amino acid concentrations in fetal fluids of pigs. J Nutr. 1995;125:2859–68.[Abstract/Free Full Text]

13. Wang JJ, Chen LX, Li DF, Yin YL, Wang XQ, Li P, Dangott LJ, Hu WX, Wu G. Intrauterine growth restriction affects the proteomes of the small intestine, liver and skeletal muscle in newborn pigs. J Nutr. 2008;138:60–6.[Abstract/Free Full Text]

14. Wu G, Flynn NE, Knabe DA. Enhanced intestinal synthesis of polyamines from proline in cortisol-treated piglets. Am J Physiol Endocrinol Metab. 2000;279:E395–402.[Abstract/Free Full Text]

15. Jones DP, Carlson JL, Samiec PS, Sternberg P, Mody VC, Reed RL, Brown LAS. Glutathione measurement is human plasma: evaluation of sample collection, storage and derivatization conditions for analysis of dansyl derivatives by HPLC. Clin Chim Acta. 1998;275:175–84.[CrossRef][Medline]

16. Fang YZ, Yang G, Wu G. Free radicals, antioxidants, and nutrition. Nutrition. 2002;8:872–9.

17. Li P, Yin YL, Li D, Kim SW, Wu G. Amino acids and immune function. Br J Nutr. 2007;98:237–52.[CrossRef][Medline]

18. Kim SW, Wu G. Dietary arginine supplementation enhances the growth of milk-fed young pigs. J Nutr. 2004;134:625–30.[Abstract/Free Full Text]

19. Jobgen WS, Fried SK, Fu WJ, Meininger CJ, Wu G. Regulatory role for the arginine-nitric oxide pathway in metabolism of energy substrates. J Nutr Biochem. 2006;17:571–88.[CrossRef][Medline]

20. Fu WJ, Hayne TE, Kohl R, Hu JB, Shi WJ, Spencer TE, Carroll RJ, Meininger CJ, Wu G. Dietary L-arginine supplementation reduces fat mass in Zucker diabetic fatty rats. J Nutr. 2005;135:714–21.[Abstract/Free Full Text]

21. Fu WJ, Hu J, Spencer T, Carroll R, Wu G. Statistical models in assessing fold changes of gene expression in real-time RT-PCR experiments. Comput Biol Chem. 2006;30:21–6.[CrossRef][Medline]

22. Kurooka H, Yokota Y. Nucleo-cytoplasmic shuttling of Id2, a negative regulator of basic helix-loop-helix transcription factors. J Biol Chem. 2005;280:4313–20.[Abstract/Free Full Text]

23. Wilkinson KD. Regulation of ubiquitin-dependent processes by deubiquitinating enzymes. FASEB J. 1997;11:1245–56.[Abstract]

24. Lessmann E, Ngob M, Leitgesc M, Mingueta S, Ridgwayb ND, Hubera M. Oxysterol-binding protein-related protein (ORP) 9 is a PDK-2 substrate and regulates Akt phosphorylation. Cell Signal. 2007;19:384–92.[CrossRef][Medline]

25. Gillingham AK, Munro S. The small G proteins of the arf family and their regulators. Annu Rev Cell Dev Biol. 2007;23:579–611.[CrossRef][Medline]

26. Berruyer C, Pouyet L, Millet V, Martin FM, LeGoffic A, Canonici A, Garcia S, Bagnis C, Naquet P, et al. Vanin-1 licenses inflammatory mediator production by gut epithelial cells and controls colitis by antagonizing peroxisome proliferators-activated receptor activity. J Exp Med. 2006;203:2817–27.[Abstract/Free Full Text]

27. Maru I, Ohta Y, Murata K, Tsukada Y. Molecular cloning and identification of N-acyl-D-glucosamine 2-epimerase from porcine kidney as a rennin-binding protein. J Biol Chem. 1996;271:16294–9.[Abstract/Free Full Text]

28. Anselmi L, Stella SL, Lakhter A, Hirano A, Tonini M, Sternini C. Galanin receptors in the rat gastrointestinal tract. Neuropeptides. 2005;39:349–52.[CrossRef][Medline]

29. Cuthbert JA, Lipsky PE. Regulation of proliferation and Ras localization in transformed cells by products of mevalonate metabolism. Cancer Res. 1997;57:3498–505.[Abstract/Free Full Text]

30. Shibata N, Arita M, Misaki Y, Dohmae N, Takio K, Ono T, Inoue K, Arai H. Supernatant protein factor, which stimulates the conversion of squalene to lanosterol, is a cytosolic squalene transfer protein and enhances cholesterol biosynthesis. Proc Natl Acad Sci USA. 2001;98:2244–9.[Abstract/Free Full Text]

31. Loo DDF, Wright EM, Zeuthen T. Water pumps. J Physiol. 2002;542:53–60.[Abstract/Free Full Text]

32. Wu GY, Fang YZ, Yang S, Lupton JR, Turner ND. Glutathione metabolism and its implications for health. J Nutr. 2004;134:489–92.[Abstract/Free Full Text]

33. McLennan AG. The Nudix hydrolase superfamily. Cell Mol Life Sci. 2006;63:123–43.[CrossRef][Medline]

34. Kinoshita M. The septins. Genome Biol. 2003;4:236.[CrossRef][Medline]

35. Flach CF, Lange S, Jennische E, Lonnroth I. Cholera toxin induces expression of ion channels and carriers in rat small intestinal mucosa. FEBS Lett. 2004;561:122–6.[CrossRef][Medline]

36. Ou DY, Li DF, Cao YH, Li XL, Yin JD, Qiao SY, Wu G. Dietary supplementation with zinc oxide decreases expression of the stem cell factor in the small intestine of weanling pigs. J Nutr Biochem. 2007;18:820–6.[CrossRef][Medline]

37. Li DF, Nelssen JL, Reddy PG, Blecha F, Hancock JD, Allee GL, Goodband RD, Klemm RD. Transient hypersensitivity to soybean-meal in the early-weaned pig. J Anim Sci. 1990;68:1790–9.[Abstract]

38. Rhoads M. Glutamine signaling in intestinal cells. JPEN J Parenter Enteral Nutr. 1999;23:S38–40.[Medline]

39. Rhoads JM, Keku EO, Woodard JP, Bangdiwala SI, Lecce JG, Gatzy JT. L-Glutamine with D-glucose stimulates oxidative metabolism and NaCl absorption in piglet jejunum. Am J Physiol Gastrointest Liver Physiol. 1992;263:G960–6.[Abstract/Free Full Text]

40. Rhoads JM, Argenzio RA, Chen WN, Rippe RA, Westwick JK, Cox AD, Berschneider HM, Brenner DA. L-Glutamine stimulates intestinal cell proliferation and activates mitogen-activated protein kinases. Am J Physiol Gastrointest Liver Physiol. 1997;272:G943–53.[Abstract/Free Full Text]

41. Wischmeyer PE, Musch MW, Madonna MB, Thisted R, Chang EB. Glutamine protects intestinal epithelial cells: role of inducible HSP70. Am J Physiol Gastrointest Liver Physiol. 1997;272:G879–84.[Abstract/Free Full Text]

42. Li N, Liboni K, Fang MZ, Samuelson D, Lewis P, Patel R, Neu J. Glutamine decreases lipopolysaccharide-induced intestinal inflammation in infant rats. Am J Physiol Gastrointest Liver Physiol. 2004;286:G914–21.[Abstract/Free Full Text]

43. Fish KJ, Cegielska A, Getman ME, Landes GM, Virshup DM. Isolation and characterization of human casein kinase I epsilon (CKI), a novel member of the CKI gene family. J Biol Chem. 1995;270:14875–83.[Abstract/Free Full Text]

44. Villalonga P, Guasch RM, Riento K, Ridley AJ. RhoE inhibits cell cycle progression and ras-Induced transformation. Mol Cell Biol. 2004;24:7829–40.[Abstract/Free Full Text]

45. Subramaniam M, Hawse JR, Johnsen SA, Spelsberg TC. Role of TIEG1 in biological processes and disease. J Cell Biochem. 2007;102:539–48.[CrossRef][Medline]

46. Lu KP, Hanes SD, Hunter T. A human peptidyl-prolyl isomerase essential for regulation of mitosis. Nature. 1996;380:544–7.[CrossRef][Medline]

47. Luo BH, Carman CV, Springer TA. Structural basis of integrin regulation and signaling. Annu Rev Immunol. 2007;25:619–47.[CrossRef][Medline]

48. Jin W, Di GH, Li JJ, Chen Y, Li WF, Wu J, Cheng TW, Yao M, Shao ZM. TIEG1 induces apoptosis through mitochondrial apoptotic pathway and promotes apoptosis induced by homoharringtonine and velcade. FEBS Lett. 2007;581:3826–32.[Medline]

49. Fuse N, Yasumoto K, Takeda K, Amae S, Yoshizawa M, Udono T, Takahashi K, Tamai M, Tomita Y, et al. Molecular cloning of cDNA encoding a novel microphthalmia-associated transcription factor isoforms with a distinct amino-terminus. J Biochem. 1999;126:1043–51.[Abstract/Free Full Text]

50. Morimoto M, Morimoto M, Zhao AP, Madden KB, Dawson H, Finkelman FD, Mentink-Kane M, Urban JF, Wynn TA, et al. Functional importance of regional differences in localized gene expression of receptors for IL-13 in murine gut. J Immunol. 2006;176:491–5.[Abstract/Free Full Text]

51. Pusch W, Jahner D, Spiess AN, Ivell R. Rat endozepine-like peptide (ELP): cDNA cloning, genomic organization and tissue-specific expression. Gene. 1999;235:51–7.[CrossRef][Medline]

52. Fischer U, Heckel D, Michel A, Janka M, Hulsebos T, Meese E. Cloning of a novel transcription factor-like gene amplified in human glioma including astrocytoma grade I. Hum Mol Genet. 1997;6:1817–22.[Abstract/Free Full Text]

53. Utz PJ, Hottelet M, Lei TM, Kim SJ. Geiger me, van Venrooij WJ, Anderson P. The 72-kDa component of signal recognition particle is cleaved during apoptosis. J Biol Chem. 1998;273:35362–70.[Abstract/Free Full Text]

54. Steyaert H, Tonon MC, Tong YA, Smihrouet F, Testart J, Pellefier G, Vaudry H. Distribution and characterization of endogenous benzodiazepine receptor ligand (endozepine)-like peptides in the rat gastrointestinal tract. Endocrinology. 1991;129:2101–9.[Abstract/Free Full Text]

55. Chang JL, Tsai HJ. Carp cDNA sequence encoding a putative diazepam-binding inhibitor/endozepine/acyl-CoA-binding protein. Biochim Biophys Acta. 1996;1298:9–11.[CrossRef][Medline]

56. Agerberth B, Boman A, Andersson M, Jornvall H, Mutt V, Boman HG. Isolation of 3 antibacterial peptides from pig intestine: gastric-inhibitory polypeptide(7042), diazepam-binding inhibitor(32–86) and a novel factor, peptide-3910. Eur J Biochem. 1993;216:623–9.[Medline]

This article has been cited by other articles:

				W. Jobgen, C. J. Meininger, S. C. Jobgen, P. Li, M.-J. Lee, S. B. Smith, T. E. Spencer, S. K. Fried, and G. Wu Dietary L-Arginine Supplementation Reduces White Fat Gain and Enhances Skeletal Muscle and Brown Fat Masses in Diet-Induced Obese Rats J. Nutr., February 1, 2009; 139(2): 230 - 237. [Abstract] [Full Text] [PDF]

				H. Gao, G. Wu, T. E. Spencer, G. A. Johnson, X. Li, and F. W. Bazer Select Nutrients in the Ovine Uterine Lumen. I. Amino Acids, Glucose, and Ions in Uterine Lumenal Flushings of Cyclic and Pregnant Ewes Biol Reprod, January 1, 2009; 80(1): 86 - 93. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) Online Supporting Material Purchase Article View Shopping Cart Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wang, J. Articles by Wu, G. Search for Related Content PubMed PubMed Citation Articles by Wang, J. Articles by Wu, G.