1US Department of Agriculture/Agricultural Research Service, Children's Nutrition Research Center, Department of Pediatrics, Baylor College of Medicine, Houston, Texas 77030; 2Department of Animal Science, University of Kentucky, Lexington, Kentucky 40546-0215; and 3Department of Animal and Poultry Science, University of Guelph, Guelph, Ontario, Canada N1G 2W1
Submitted 22 May 2003 ; accepted in final form 23 March 2004
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ABSTRACT |
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excitatory amino acids; gut mucosa; transporter affinity; gene expression; neonates
Amino acids are key nutrients that are essential for gut mucosal growth, because they serve as metabolic fuels (6, 61) and precursors for the synthesis of protein (2, 55), nucleosides, and polyamines (6). L-Glutamate is one of the most intensively investigated amino acids in gut mucosal growth and metabolism (50, 60). Earlier studies showed that enteral glutamate was extensively metabolized by the gut mucosa in dogs and rats (42, 61). With the use of stable isotope tracer balance techniques in vivo, it was further shown that enteral glutamate was preferentially metabolized to CO2 and, specifically, used as a precursor for the biosynthesis of glutathione, arginine, and proline by the gut mucosa in neonatal pigs fed milk-based diets (51).
L-Glutamate transport by the enterocyte apical membrane is the initial step of the entire glutamate metabolism pathway in the gut mucosa. Physiological characterization of intestinal amino acid transport systems responsible for glutamate uptake in the mammalian species has been largely conducted with brush-border membrane vesicles prepared from total mucosal scrapings of mature animals (3, 49). Yet how neonatal intestinal epithelial cells effectively transport enteral glutamate across the apical membrane remains unclear.
L-Glutamate transport across the enterocyte apical membrane was shown to be through the Na+-dependent high-affinity XAG system and/or the low-affinity Bo system (3, 29, 49). Although both transport activities couple the absorption of glutamate to Na+, major differences exist between these transport activities. Whereas the Bo system functions primarily as a neutral amino acid transporter, recognizing L-glutamate only at pH less than about 5.5 (24), L-glutamate and L- and D-aspartate are the principal substrates of the XAG system, which functions as an obligate countertransporter of K+ (3, 10, 30, 49). In contrast to the Bo system, the molecular identities of four proteins [glutamate/aspartate transporter 1 (GLAST-1), glutamate transporter 1 (GLT-1), excitatory amino acid carrier (EAAC)-1, and excitatory amino acid transporters (EAAT)-4 and -5] capable of XAG system activity have been described in brain and various nonbrain tissues (12, 23, 47, 56). Recently, EAAC-1 expression has been examined in the rat (11, 53) small intestinal mucosa, EAAC-1 and GLT-1 proteins have been identified throughout the intestinal epithelium of sheep and cattle (18, 19), and GLAST-1 is known to be expressed by the cryptlike cell line IEC-17 (38, 39) and the differentiating human colorectal carcinoma Caco-2 cell line (37). With regard to the site of glutamate uptake activity along intestinal villi, early studies conducted with quantitative autoradiographic techniques suggested that the expression of various amino acid transport systems was limited to the upper third of the villi (8). Despite these advances in our understanding of intestinal glutamate uptake, overall, little is known about L-glutamate transport activity or the proteins in the apical membrane of epithelial cells, especially as these cells differentiate during their migration up the crypt-villus axis in the neonate.
Therefore, the objectives of this study were to establish the major apical membrane transport activities for L-glutamate absorption and identify specific glutamate transport proteins of epithelial cells at different stages of differentiation along the small intestinal crypt-villus axis in neonatal pigs fed a milk protein-based formula. Given the extensive metabolism of enteral glutamate in the neonate, we hypothesized that glutamate transporter activity and expression would be higher in differentiated upper villus cells than in proliferative crypt cells. Because upper villus cells are differentiated into an absorptive phenotype, we expected the activity and transporter content of glutamate transporters to be higher than in proliferative crypt cells.
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MATERIALS AND METHODS |
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Animals and tissue preparation. A total of 30 piglets, taken from different sows at the age of 7 days, were individually housed in stainless steel metabolic cages equipped with feeders in a room maintained at 28°C. The piglets were fed three times a day a liquid, milk protein-based formula containing 25% lactose, 25% protein, and 10% fat (Litter Life, Merricks, Middleton, WI). At 16 days of age, the piglets were euthanized for isolation of small intestinal epithelial cells. The experimental protocol was approved by the institutional Animal Care and Use Committees.
Piglets were anesthetized by inhalation of 5% isoflurane (Aerrane, Anaquest, WI) via a face mask. The abdomen was immediately opened, and the whole small intestine, starting at 30 cm posterior to the pyloric sphincter and 60 cm anterior to the ileocecal sphincter, was immediately flushed twice with 60 ml of an ice-cold phosphate-buffered saline (0.2 mM PMSF and 0.5 mM DTT, pH 7.4). To isolate enterocytes along the longitudinal axis, a 240-cm proximal small intestinal segment was dissected starting at 30 cm distal to the pyloric sphincter. In addition, a 240-cm distal small intestinal segment was dissected starting at 60 cm anterior to the ileocecal junction. Proximal and distal segments were divided into four separate segments (60 cm each) for the cell isolation procedure. After the intestinal segments were removed, the animals were killed with an overdose (50 mg/kg) of pentobarbital sodium (Sigma).
Sequential isolation of epithelial cells along the crypt-villus axis. We recently developed a procedure for sequentially isolating enterocytes along the crypt-villus axis (14) by adapting the distended intestinal sac technique previously developed in rats (58, 59). By aid of a 50-ml syringe fitted with a feeding needle, the divided segments were then quickly filled with a preincubation buffer (27 mM sodium citrate, 0.2 mM PMSF, and 0.5 mM DTT, pH 7.4), oxygenated with 19:1 (vol/vol) O2-CO2, and warmed in a water bath (37°C). The filled intestinal segments were sealed with two pairs of hemostatic forceps, immersed in saline (154 mM NaCl) in 2-liter glass beakers, and incubated for 15 min. After preincubation, the intestinal segments were filled with an isolation buffer (1.5 mM Na2EDTA, 0.2 mM PMSF, 0.5 mM DTT, and 2 mM D-glucose, oxygenated with the O2-CO2 mixture, and warmed at 37°C) for the sequential isolation of 12 cell fractions (F1F12) from the villus tip to the bottom crypt.
For in vitro glutamate transport measurements on individual cell fractions, a total of 12 cell fractions were isolated from 24 piglets, and the isolated cell fractions were stored separately for each pig. The F1F6 cell fractions were collected through six consecutive 10-min incubations and buffer removal. The F7F12 cell fractions were collected through six consecutive 15-min incubations and buffer removal. Each cell fraction was washed twice with 150 ml of oxygenated cell suspension buffer (155 mM KCl, pH 7.4), and the cells were retained through centrifugation at 400 g for 4 min at 4°C. The washed cells were immediately frozen at 80°C for future isolation of brush-border membrane vesicles for transport measurements. Isolated cell fractions were also pooled between the proximal and the distal intestinal segments for each piglet.
For molecular identification of the putative glutamate transporters, three major cell fractions, consisting of the upper (F1F4), the middle (F5F8), and the crypt (F9F12) cell fractions, were sequentially isolated through three consecutive incubations at 40, 50, and 60 min, respectively, from six additional piglets with the use of the distended intestinal sac technique described above. Isolated cell fractions from each piglet were immediately frozen in liquid nitrogen and stored at 80°C.
Preparation of apical membrane vesicles.
Apical membrane vesicles were isolated by Mg2+ precipitation and differential centrifugation according to a previously established procedure (13, 28). Specifically, for each batch of apical membrane vesicle preparation, 5 g of frozen cell fractions collected from two piglets were thawed in ice-cold homogenate buffer (50 mM D-mannitol, pH 7.4) at 20 ml of the homogenate buffer per gram of cells and homogenized by a Polytron homogenizer. The resulting homogenate was pooled and centrifuged at 2,000 g for 15 min. After the top foam layer was removed and the pellets were discarded, the supernatant was mixed with 1 M MgCl2 to contain 10 mM MgCl2, stirred for 15 min, and then centrifuged at 2,400 g for 15 min. After the top foam layer was discarded, the supernatant was centrifuged at 19,000 g for 30 min to generate crude apical membrane pellets. The crude apical membrane pellets were then suspended in a suitable amount of a membrane suspension buffer (150 mM KSCN and 180 mM D-mannitol, pH 7.0) and centrifuged at 39,000 g for 30 min to generate the final apical membrane vesicle pellets. The final pellets were resuspended with a 25-gauge needle in a suitable volume of the same membrane suspension buffer. The final membrane vesicle suspension was assayed for protein content and diluted with the same buffer to contain 4 mg protein/ml for subsequent transport measurements. Several aliquots of the final membrane vesicle suspension were taken for the assays of marker enzyme activities.
Protein and marker enzyme assays.
Protein was determined by using the Bio-Rad protein dye reagent and bovine serum albumin (fraction V) as a standard. All the following enzyme assays were carried out under the conditions that enzyme reactions were linear with time. Aminopeptidase N activity was assayed according to an established procedure (32). The incubations were conducted in a final volume of 0.200 ml containing membrane suspension or cell homogenate (2.520 µg protein), 50 mM Na2HPO4, and 28.0 mM L-alanine-p-nitroanilide hydrochloride at 37°C and pH 7.0 (adjusted with 0.1 M H3PO4 and 0.1 M NaOH) for 20 min. The K+-stimulated p-nitrophenyl phosphatase activity was measured according to a previously established procedure (41). The incubations were carried out in a final volume of 1.000 ml containing membrane suspension or cell homogenate (110 µg protein), 90 mM KCl or NaCl, 10 mM MgCl2, 5.0 mM Na2EDTA, 2.0 mM p-nitrophenyl phosphate Tris salt, and 10 mM Tris·HCl at 37°C and pH 7.0 (adjusted with 0.5 M Tris·base and 0.5 M Tris·HCl) for 10 min. The succinate dehydrogenase activity was measured according to King (26). The incubations were carried out in a final volume of 1.500 ml containing membrane suspension or cell homogenate (2.5 µg protein), 75 mM disodium phosphate, 0.67 mM K3Fe(CN)6, and 80.0 mM disodium succinate at 37°C and pH 7.0 (adjusted with 0.5 M Tris·base and 0.1 M Tris·HCl) for 10 min. The D-glucose-6-phosphatase activity was measured according to Hübscher and West (20). The incubations were carried out in a final volume of 0.400 ml containing membrane suspension or cell homogenate (2.5 µg protein), 29 mM D-glucose-6-phosphate barium salt, 4.0 mM Na2EDTA, 2.0 mM KF, and 200 mM sodium maleate at 37°C and pH 6.0 (adjusted with 0.5 M NaOH) for 10 min. The acid phosphatase activity was measured according to Hübscher and West. The incubations were carried out in a final volume of 0.400 ml containing membrane suspension or cell homogenate (2.5 µg protein), 15 mM -glycerol phosphate, 4.0 mM Na2EDTA, and 52.0 mM sodium acetate at 37°C and pH 5.4 (adjusted with 0.5 M acetic acid and 0.5 M NaOH) for 10 min.
In vitro transport measurements. In vitro transport experiments were carried out with the rapid filtration procedure (13), which was conducted on the day of membrane vesicle preparation. After protein assay, the final membrane vesicle suspension was equilibrated for an additional 30 min before transport measurements. Membrane vesicles were preloaded with a buffer containing 180 mM D-mannitol, 150 mM KSCN, and 10 mM Trizma·HCl, pH 7.0. The uptake buffer for measuring L-glutamate transport via the high-affinity XAG system contained 2.4 µM L-[G-3H]glutamate, 120 mM D-mannitol, 60 mM L-phenylalanine, 150 mM NaSCN, and 10 mM Trizma·HCl, pH 6.0. The uptake buffer for measuring L-glutamate transport via the Bo system contained 2.4 µM L-[G-3H]glutamate, 120 mM D-mannitol, 60 mM D-aspartate, 150 mM NaSCN, and 10 mM Trizma·HCl, pH 7.0. Fifty microliters of uptake buffer were first pipetted into the bottom of a polystyrene tube (100 x 15 mm), and then 10 µl of apical membrane vesicle suspension were spotted onto the side of the tube in two separate drops directly above the uptake buffer with a Microman pipette. After the tube was warmed for 20 s at room temperature (24°C), uptake incubation was initiated by a foot switch-activated Bibromixer, and the process was terminated by the addition of 1.125 ml of ice-cold wash solution (180 mM D-mannitol, 150 mM NaSCN, 10 mM Trizma·HCl, and 0.1 mM HgCl2 at pH 7.0 or 6.0 for measuring L-glutamate transport via the XAG system). One milliliter of the uptake mixture was then rapidly pipetted onto 0.45-µm cellulose acetate filters (presoaked with 20 mM L-glutamate, pH 7.4, to minimize nonspecific binding of these 2 amino acids to the filters) mounted in a Manifold filtration unit, which was connected to a vacuum source. On the basis of our preliminary measurements, a 6-s incubation was used to measure initial L-[G-3H]glutamate tracer transport rate at 2 µM. Timing was performed with an electronic timer-intervalometer (model 545, GraLab). The filters were immediately washed three times with 5 ml of ice-cold wash solution. The remaining solution in the incubation tubes was collected and at the end of the experiment was pooled and counted for the average initial radioactivity in the uptake medium. After a 30-min extraction in 5 ml of Ecolume scintillant, filters were counted with a liquid scintillation analyzer with automatic quench correction.
The components of L-glutamate transport via the XAG and the Bo systems were partitioned according to a previously established protocol (27). L-Glutamate transport by the XAG system was measured in the presence of 50 mM L-phenylalanine, a typical Bo system substrate, in the uptake medium to block possible L-glutamate transport by the Bo transporter (see legends to Figs. 1 and 2). L-Glutamate transport by the Bo system was measured in the presence of 50 mM D-aspartate, a typical XAG system substrate, in the uptake medium to block possible L-glutamate transport by the XAG transporter (see legends to Figs. 1 and 2).
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Each uptake experiment was conducted in triplicate. Three separate uptake experiments were conducted using three different batches of membrane vesicle suspension prepared from cell fractions pooled from two piglets. Inasmuch as isolated cell fractions were also pooled between the proximal and distal intestinal segments for each piglet, all amino acid transport results represented the whole small intestine of the neonatal piglet fed the liquid formula.
Immunoblot analysis for expression of EAAC-1 and GLT-1.
Cell fractions were homogenized in a buffer consisting of 50 mM D-mannitol, 10 mM HEPES, 1 mM Na2EDTA, 0.2 mM PMSF, and 2 µg/ml each of N--p-tosyl-L-lysine ketone, N-tosyl-L-phenylalanine chloromethyl-ketone, leupeptin hemisulfate, aprotinin, and pepstatin A and stored at 80°C until immunoblot analyses or further processed to isolate apical membrane and then stored at 80°C until immunoblot analysis. Analysis of transporter protein expression by enterocyte populations was conducted by immunoblot analysis using 50 µg of homogenate or membrane protein and antibodies and procedures of Matthews et al. (34). The amount of transporter immunoreactivity measured among cell populations was normalized to crypt values within an animal to control for differences in hybridization intensity among blots.
Extraction of RNA and determination of EAAC-1, GLT-1, and 18S ribosomal mRNA expression by RT-PCR and Southern blot analyses. For RNA analyses, 6002,000 mg of each cell fraction were homogenized in 5 ml of TRIzol according to instructions of the manufacturer (GIBCO-BRL, Grand Island, NY), and recovered total RNA was suspended in RNase-free, distilled and deionized water and stored at 80°C. Five micrograms of total RNA were reverse-transcribed using Superscript II reverse transcriptase and random and oligo(dT) primers according to instructions of the manufacturer (GIBCO-BRL). The PCR primers were as follows: 5'-GGGACAGATTCTGGTGGATT-3' and 5'-GTGATCCTCTTGTCCAC-3' (EAAC-1) and 5'-GAAAAAACCCATTCTCCTTTTT-3' and 5'-CCGACTGGGAGGACGAATC-3' (GLT-1). The expected size of the EAAC-1 RT-PCR product was 369 bp (corresponding to bp 7351,104 of the human EAAC-1 sequence; GenBank accession no. U06469), whereas the expected size of the GLT-1 RT-PCR product was 159 bp (corresponding to bp 1,0541,213 of the human GLT-1 sequence; GenBank accession no. U03505). These primers were originally reported by O'Kane et al. (43) and previously documented to be successful in our laboratory work (18). The PCR primers used for amplification of a potential 398-bp 18S ribosomal protein cDNA were 5'-CCGCGGTTCTATTTTGTTGGTTTT-3' and 5'-CGGGCCGGGTGAGGTTTC-3'. These primers corresponded to bp 118141 and 499516, respectively, of the previously reported porcine 18S ribosomal protein (GenBank accession no. AF102857).
For EAAC-1 and 18S ribosomal RNA, PCR conditions were 25 cycles at 94°C for 30 s, 51°C for 30 s, and 68°C for 1 min with a final extension at 68°C. For GLT-1, 35 cycles at 94°C for 1 min, 50°C for 45 s, and 72°C for 1 min with a final extension at 72°C for 7 min were performed. The cycles were preceded by 5- and 10-min denaturing (94°C) of cDNA in the presence of MgCl2 for EAAC-1 and GLT-1, respectively. For all reactions, thermal cycling was performed using Taq DNA polymerase (GIBCO-BRL) in 50 µl and 2 mM MgCl2, and cycle times were optimal for product formation. Within a cell fraction of an individual animal, the same RT product was used for EAAC-1, GLT-1, and 18S ribosomal RNA PCR, but as separate PCR.
For ethidium bromide detection and Southern blot analyses, RT-PCR products were separated by electrophoresis through a 1.2% agarose gel in Tris-acetic acid-EDTA buffer and stained with ethidium bromide. The sizes of the RT-PCR products were estimated by regression of migration distance compared with a 100-bp DNA standard ladder (GIBCO-BRL). Subsequently, for Southern analysis, the same gels were denatured (0.4 M NaOH-1.5 M NaCl) for 30 min, rinsed twice with distilled water, neutralized (0.5 M Tris·HCl-1.5 M NaCl, pH 7.5) for 30 min, and transferred by downward capillary action onto a 0.45-µm positively charged nylon membrane (Hybond N+, Amersham Pharmacia Biotech, Arlington Heights, IL). After they were blotted, the membranes were air dried and cross-linked by ultraviolet light.
Oligonucleotides with sequences internal to our partial-length porcine EAAC-1 sequence and previously reported 18S ribosomal RNA protein were designed and used for Southern analysis. The sequences of these "nested" probes were 5'-ATCCTCTTGTCCACCTGGTTCTTCTCTTCA-3' and 5'-ACTCCTGGTGGTGCCCTTCCGTCAATTCCTTTAA-3', respectively. After synthesis (Invitrogen, San Diego, CA), the oligonucleotides were end labeled with [-32P]ATP (3,000 Ci/mmol; Amersham) using a forward 5'-terminal cDNA end-labeling reaction kit (GIBCO-BRL). Briefly, 5 pmol of oligonucleotide were incubated with 25 µCi of [
-32P]ATP and 10 U of T4 polynucleotide kinase for 10 min at 37°C. The reaction was terminated by addition of Na2EDTA to a final concentration of 5 mM. Radiolabeled oligonucleotides were separated from unincorporated [
-32P]ATP molecules using a G-50 Sephadex spin column (Amersham Pharmacia, Piscataway, NJ), after phenol-chloroform separation of the aqueous phase using standard molecular biology techniques. In serial hybridization experiments with the blots, recovered 32P-labeled oligonucleotides were boiled for 5 min, mixed with 5 ml of 50°C hybridization buffer (34), and used immediately in Southern blot analysis. After a 2-h prehybridization period, blots were hybridized for 4 h with the appropriate 32P-labeled oligonucleotide at 50°C in hybridization buffer. All blots were initially washed at 50°C for 2 min in a 5x SSC solution (0.75 M NaCl-0.075 M sodium citrate, pH 7.0). Subsequent washes were also carried out for 2 min but varied in SSC concentrations. Typically, blots were serially washed once in 2.5x SSC and then with 1.25x SSC. After an appropriate exposure period of the membrane to autoradiographic film (Amersham Pharmacia), digital images of the audioradiographic film were recorded, and relative intensities of bands were determined as previously described (57). The ratios of EAAC-1 to 18S ribosomal RNA protein cDNA intensities were computed to account for potential variation of RT-PCR product generation, loading, and transfer and, thus, allowed for the relative comparison of EAAC-1 expression among the epithelial cell fractions.
Generation of partial-length porcine EAAC-1 cDNA. The EAAC-1 RT-PCR product was cloned into the EcoRI multiple cloning site of pCRII (Invitrogen) plasmid vector according to instructions of the manufacturer. The identity of the cDNA was determined by dideoxy-mediated chain termination sequencing by the University of Florida DNA Sequencing Laboratory (Gainesville, FL). The nucleotide sequence reported here resides in the GenBank/EMBL data bank (accession no. AY195622). Sequence alignments and comparisons to other GenBank entries were performed using BLAST 2.0 software (blast@ncbi.nlm.nih.gov).
Calculations and kinetic and statistical analyses of transport data.
The initial rates of L-[G-3H]glutamate tracer and/or total L-glutamate uptake under various experimental conditions were calculated according to our previously established procedure (13)
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Kinetic parameter estimates of the L-glutamate transporter affinity, the maximal transport activity, and the transmembrane diffusion were determined according to a previously established tracer inhibitory kinetic model (28, 31)
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Kinetic parameter estimates of the L-glutamate transporter affinity, the maximal transport activity, and transmembrane permeability constant also were determined according to a previously established classical kinetic model (62)
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All kinetic parameter estimates of L-glutamate transport activities were analyzed according to Eqs. 2 and 3 using the Fig.P curve-fitting program (1993, Biosoft, Cambridge, UK). Comparison of kinetic parameter estimates was conducted by using the pooled two-tailed Student's t-test (7).
The effect of cell population on relative expression of XAG system transporter protein and mRNA was evaluated using the general linear model procedure of SAS (SAS Institute, Cary, NC). When appropriate, protected Fisher's least significant difference procedures of the SAS were used to compare cell population means.
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RESULTS |
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The 16-day-old neonatal piglets in this study were associated with a mean ± SE (n = 30) fresh weight of 209 ± 9.8 g and length of 7.02 ± 0.2 m for the small intestine at 4.7 ± 0.4 kg body wt. We also observed that 50% of the total fresh weight of the small intestine was epithelial cells enriched in enterocytes. Furthermore, the upper villus, middle villus, and crypt region accounted for
28.4 ± 0.6, 45.5 ± 0.7, and 26.3 ± 0.8% (mean ± SE, n = 30) of the total epithelial cells, respectively, on a wet weight basis.
Characterization of the apical membrane prepared from neonatal porcine enterocytes isolated along the crypt-villus axis. Our previous work showed that more aminopeptidase N than other apical membrane hydrolases, such as lactase, sucrase, and alkaline phosphatase, was expressed in the crypt epithelial cells (14). We therefore chose to use this enzyme- specific activity as a marker for the enterocyte apical membrane. The apical membrane vesicles prepared from all 12 cell fractions (F1F12) were enriched 6- to 10-fold in the aminopeptidase N specific activity compared with their corresponding cell homogenates.
Enrichment of the enzyme marker for the enterocyte basolateral membrane, K+-stimulated phosphatase-specific activity was less than onefold in the apical membrane prepared from all cell fractions compared with their corresponding cell homogenates, indicating little contamination of the apical membrane preparation with the enterocyte basolateral membrane.
Enrichments of other enzyme markers, including the mitochondria membrane marker succinate dehydrogenase specific activity, the endoplasmic reticulum membrane marker D-glucose-6-phosphatase specific activity, and the lysosome membrane marker acid phosphatase specific activity, were <1.7-fold in the apical membrane prepared from all cell fractions compared with their corresponding cell homogenates, suggesting little contamination of the apical membrane preparation with the other intracellular membrane fractions.
Initial rates of L-[G-3H]glutamate tracer transport in the apical membrane of neonatal porcine enterocytes along the crypt-villus axis. It has been well documented that high-affinity XAG and low-affinity Bo systems are responsible for transporting L-glutamate across the mammalian enterocyte apical membrane (3, 49). It is thus logical to first compare the relative importance of the high-affinity XAG and the low-affinity Bo systems in transporting glutamate across the apical membrane of proliferating, differentiating, and differentiated neonatal porcine epithelial cells. Inasmuch as Na+-dependent L-glutamate transport via the Bo system across the enterocyte apical membrane is an electrogenic process, membrane potential across the cell membrane is an important driving force in transporting amino acids under normal physiological conditions (3, 10). Our previous work revealed a sharply decreasing gradient in ouabain-sensitive Na+-K+-ATPase and ouabain-insensitive Na+-ATPase activities from the tip villus to the bottom crypt cells (14), suggesting a decreasing membrane potential gradient associated with the neonatal porcine epithelial cells during cell proliferation and differentiation. To eliminate possible contributing effects from differences in membrane potential and the apical membrane permeability among the cell fractions, the membrane-permeable anion SCN was used in the membrane suspension buffer and also in amino acid uptake buffers to clamp the membrane potential (15).
Initial rates of L-[G-3H]glutamate tracer transport across the apical membrane via the high-affinity XAG system showed a cubic pattern of progressive increases (P < 0.05) from the tip villus (F1) to the bottom crypt epithelial cells (F12) at the tracer substrate concentration of 2.0 µM (Fig. 1). At the same substrate concentration, initial rates of L-[G-3H]glutamate tracer transport across the apical membrane via the low-affinity Bo system showed no differences (P > 0.05) from the tip villus (F1) to the bottom crypt (F12) epithelial cells (Fig. 1). Furthermore, initial rates of L-[G-3H]glutamate tracer transport via the high-affinity XAG system were much higher (P < 0.05) than those via the low-affinity Bo system in all cell fractions, indicating that glutamate transport via the high-affinity XAG system is the major pathway for absorbing the enteral glutamate.
Kinetics of L-glutamate transport via the XAG transporter in the apical membrane prepared from neonatal porcine enterocytes along the crypt-villus axis. Inasmuch as initial rates of L-[G-3H]glutamate tracer transport across the apical membrane via the Bo system were very low in the presence of increasing levels of unlabeled L-glutamate, we were unable to determine the kinetics of glutamate transport via the Bo system in the apical membrane vesicles. Kinetics of glutamate transport via the XAG system were determined with apical membrane vesicles prepared from the upper villus (F1F4), middle villus (F5F8), and crypt (F9F12) cells under the condition that the membrane potential in the apical membrane vesicles was clamped with SCN.
We compared two well-defined kinetic models to quantify glutamate transport kinetics, inasmuch as both models allowed simultaneous determination of the component of transmembrane diffusion (28, 31, 62), and very few previous literature reports simultaneously compared both models in the same studies. The tracer inhibitory kinetic model uses initial tracer L-[G-3H]glutamate transport rate as the independent variable, the concentrations of unlabeled L-glutamate as the dependent variable, and the L-[G-3H]glutamate tracer concentration (2 µM) as an input parameter. Thus the kinetic curve associated with this model (Eq. 2) visually resembles competitive inhibition kinetics, namely, the inhibition of tracer L-[G-3H]glutamate uptake by unlabeled L-glutamate (Figs. 2A, 3A, and 4A). Kinetics of L-glutamate transport were also analyzed with the classical model (Eq. 3) after partition of the transmembrane diffusion component, and the saturable component was found to be within 00.22 mM L-glutamate (Figs. 2B, 3B, and 4B). The kinetics of glutamate transport were determined with the apical membrane vesicles prepared from the pooled upper villus (Fig. 2), the pooled middle villus (Fig. 3), and the pooled crypt (Fig. 4) cells. The kinetics of L-glutamate transport in the apical membrane vesicles of the upper, middle, and crypt enterocyte fractions were best fitted by a single transport system (Figs. 24). The relation between initial tracer L-[G-3H]glutamate transport rates and the logarithmically transformed, unlabeled L-glutamine concentrations is presented in Fig. 5.
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Five mammalian glutamate transporters featuring the high-affinity XAG transport system have been cloned and characterized (22, 37). Four of the five cloned glutamate transporters, GLAST, GLT-1, EAAC-1, and EAAT-4, are defined in animals, whereas the respective glutamate transporters are named EAAT-1, EAAT-2, EAAT-3, and EAAT-4 in humans (22).
To test the hypothesis that XAG transporter systems were responsible for the observed D-aspartate-sensitive glutamate uptake by neonatal porcine epithelial cell populations, the putative presence of EAAT-4, GLAST-1, GLT-1, and/or EAAC-1 in homogenates of upper, middle, or crypt enterocytes was evaluated by immunoblot analysis. The presence of EAAT-4 and GLAST-1 was not detected in any cell populations (data not shown). Immunoreactive bands of 167 and 181 kDa were detected in the homogenate and the apical membrane preparations from all cell populations isolated from two of six pigs (Fig. 6). To further evaluate the potential expression of GLT-1 protein by porcine small intestinal epithelia, RT-PCR analysis for GLT-1 was performed on the animal with the highest expression of GLT-1 protein (pig 2). Expression of GLT-1 mRNA was not detected by RT-PCR followed by ethidium bromide visualization (Fig. 7A). In contrast, GLT-1 was expressed by a control tissue (sheep liver) (17). Repetition of this analysis for pig 2 and an initial analysis for the other five pigs also failed to detect expression of GLT-1 mRNA (Fig. 7B).
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DISCUSSION |
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Several previous studies used different techniques to examine the expression of amino acid transport activity during the enterocyte proliferation and differentiation (8, 37, 4446). Very early work using a quantitative autoradiographic technique in studies with adult animals demonstrated that amino acid and peptide transporter activities were limited to the differentiated upper villus enterocytes (8, 25). Results of those studies contradict recent observations from differentiating Caco-2 cells (37, 4446) which show that the uptake of dipolar-neutral and cationic amino acids was higher in proliferating than in differentiated cells. On the other hand, the studies of amino acid transport activity in association with cell differentiation in cell line models are usually limited by the fact that it is difficult to differentiate the measured amino acid transport activities between the apical and basolateral membranes. Furthermore, Caco-2 cells are of colorectal origin and may exhibit certain differences from enterocytes with regard to in vivo conditions, including the different expression pattern of amino acid transport systems.
We have shown that the apical membrane maximal L-glutamate transport activity via the XAG system was high in proliferating and differentiating midvillus epithelial cells and low in differentiated upper villus cells, a pattern distinctly different from that reported in differentiating Caco-2 cells (37). Moreover, it is possible that the pattern of expressing L-glutamate activity in proliferating, differentiating, and differentiated enterocytes we observed in the neonatal pig changes with stage of development, an issue not addressed in this study. Two major factors affecting the maximal amino acid transport activity (Jmax) are the number of transporters expressed and transporter affinity. Expression of a large number of transporter proteins and a high glutamate transporter affinity could potentially result in a large Jmax. Our data showed that XAG glutamate transporter affinity was significantly lower in crypt and middle villus than in upper villus cells. It is well established that transporters with a low transporter affinity (a large Km value) are usually associated with a large Jmax value. In this situation, changes in Jmax alone could not truly reflect differences in glutamate transporter efficiency among the cell populations at different stages of proliferation and differentiation. Therefore, significant changes in the transporter affinity are believed to be an additional mechanism regulating transport capacity and efficiency. We suggest that the efficiency of L-glutamate transport across the apical membrane is likely affected by changes in transporter maximal activity and affinity along the neonatal small intestinal crypt-villus axis.
The XAG transporter affinity for L-glutamate (Km) determined from this study (4268 µM) for the neonatal pig was generally within the range of the values (4150 µM) reported in the literature (10, 27, 49). Km among upper villus, middle villus, and crypt enterocytes determined from this study was more variable than the values (107118 µM) reported in differentiating Caco-2 cells (37, 40). Differences in species, developmental stages, and experimental conditions were likely responsible for differences in the Km values between studies (13). In this study, the differences in Km may be attributed to the following factors. First, apical membrane-bound amino acid transporters are glycoproteins, and studies indicate that significant villus-crypt changes occur in the glycosylation of the apical membrane-bound proteins (35). Changes in the glycosylation might have been, in part, responsible for the different Km values among the different enterocyte fractions. Yet changes in glycosylation were not responsible for the apical membrane hydrolase lactase affinity (5). However, all of the apical membrane proteins are regulated independently; thus we could not rule out this possibility for this high-affinity XAG transporter protein at this time. Second, changes in the microvillus lipid composition and membrane fluidity, although not determined in this study, as was demonstrated in several previous studies (1, 4), were also possibly responsible for the different Km in the different cell fractions. On the other hand, a significant component of transmembrane diffusion observed with both kinetic models of analyses was consistent with previous studies with the membrane vesicle approach (13, 31, 62), suggesting the leakiness of membrane vesicles under the in vitro preparation condition as well as the intrinsic leaky nature of cell membranes (Table 1). In addition, the data obtained with both kinetic models pointed to the fact that the apical membrane of the upper villus and the crypt cells was more permeable than that of the middle villus cells, probably because of their differential combined effects of cell proliferation, differentiation, and luminal factors.
Transmembrane uptake of the enteral L-glutamate is the initial step of its entire first-pass metabolism pathway in the gut mucosa. Enteral L-glutamate metabolism in the gut mucosa has been extensively investigated with in vivo animal studies (50, 52, 54, 60). However, the metabolic fate of glutamate at the cellular level remains unclear, especially within enterocytes. Inasmuch as proliferating crypt epithelial cells are likely to be metabolically different from differentiated upper villus enterocytes, the metabolic fate of glutamate in proliferating and differentiating crypt cells warrants further investigation.
To test the hypothesis that the XAG transporter system was responsible for the observed D-aspartate-sensitive glutamate uptake by cell membrane preparations, the putative presence of EAAT-4, GLAST-1, GLT-1, and/or EAAC-1 in homogenates of the upper, middle, or crypt cells was evaluated by immunoblot analysis. That EAAT-4 and GLAST-1 proteins were not detected in small intestinal epithelia of neonatal pigs, but EAAC-1 was, is consistent with that observed for developing rats (53), mature ruminant animal species (18), and growing lambs (19). In contrast, the sporadic expression (2 of 6 animals evaluated) of GLT-1 by neonatal pigs differed from that by the ruminant animal species (18, 19), inasmuch as GLT-1 was detected in small intestinal epithelium by all tested animals. Whether the relatively sporadic expression of GLT-1 by neonatal pigs reflects species-specific or ontogenic regulation of GLT-1 remains to be determined.
Immunoblot analysis of cell homogenates also demonstrated that more EAAC-1 were expressed in differentiated upper and middle villus cells than in proliferating crypt cells (Fig. 9). When the kinetic parameter estimates for XAG glutamate transport system activities among cell populations, as summarized in Table 1, are considered in terms of EAAC-1 content, it is clear that a greater EAAC-1 protein content is associated with a lower maximal transport activity and changing transporter affinity. Collectively, these data suggest that another, D-aspartate-insensitive, glutamate transport activity (other than the XAG system) may be expressed in less-differentiated cells or the functional properties of EAAC-1 change with differentiation. With regard to the first possibility, the apical membrane of rat placenta possesses such an Na+-dependent glutamate uptake activity (34). Conversely, a cytosolic protein [GTRAP-318] that binds and decreases EAAC-1 affinity for glutamate has been identified (Genbank accession no. AF240182). When GTRAP-318 and EAAC-1 were overexpressed in 293C18 cells, a loss of glutamate affinity was achieved without a change in uptake velocity. In the present study, however, decreases in glutamate uptake affinity as cell populations differentiated were accompanied by increases in the maximal velocity. Our observations that EAAC-1 is the main identifiable XAG transporter system expressed in all enterocyte populations coupled with kinetic analysis suggest that there is a single transporter system in these cell fractions. It is most likely that the XAG system transporter EAAC-1 is the only Na+-dependent glutamate transporter expressed by these epithelial cells. However, the existence of additional transporters cannot be excluded.
With regard to how the differential expression of EAAC-1 is regulated, the data of Figs. 9 and 11 indicate that the increase in EAAC-1 protein content of differentiated cells likely resulted from a combination of increased gene transcription and posttranscriptional events. That is, whereas the relative content of EAAC-1 protein and mRNA was proportional in the upper villus and crypt cell populations, the amount of EAAC-1 mRNA in the middle villus cells was low compared with protein content. This discordance between relative abundance of steady-state EAAC-1 mRNA and protein in neonatal pig enterocyte populations is consistent with that observed for EAAC-1 regulation in other tissues and developmental models (34, 48).
In conclusion, this study demonstrated that the high-affinity XAG system was the major pathway for transporting luminal L-glutamate across the apical membrane of proliferating, differentiating, and differentiated neonatal porcine epithelial cells. There were significant differences in the apical membrane glutamate maximal uptake activity and transporter affinity and the in the neonatal porcine enterocyte along the crypt-villus axis. EAAC-1 was the major XAG glutamate transporter identified, and its expression was increased with cell differentiation and regulated at the transcription and translation levels in the neonatal porcine enterocyte along the crypt-villus axis. The finding of a reduced maximal glutamate transport activity in association with a significantly increased transporter affinity associated with the XAG system in the upper villus cells supports the concept that efficiency of luminal L-glutamate uptake across the apical membrane is different along the intestinal crypt-villus axis in the neonate. However, the changes in EAAC-1 expression in isolated cells are consistent with the idea that glutamate transport is upregulated in differentiated villus enterocytes. Further study is warranted to establish whether the metabolic fate of glutamate is indeed different in undifferentiated crypt cells compared with differentiated enterocytes.
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ACKNOWLEDGMENTS |
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This work is a publication of the US Department of Agriculture/Agricultural Research Service (USDA/ARS) Children's Nutrition Research Center, Department of Pediatrics, Baylor College of Medicine. The contents of this publication do not necessarily reflect the views or policies of the USDA, nor does mention of trade names, commercial products, or organizations imply endorsement by the US Government.
Present address of M. Z. Fan: Dept. of Animal and Poultry Science, Rm. 250, #70 Animal Science/Nutrition Bldg., University of Guelph, 50 Stone Rd. East, Guelph, ON, Canada N1G 2W1 (E-mail: mfan@uoguelph.ca).
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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