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Department of Biochemistry, Memorial University of Newfoundland, St. John's, Canada A1B 3X9
* To whom correspondence should be addressed. E-mail: rbertolo{at}mun.ca.
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ABSTRACT |
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 TOP ABSTRACT IntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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28 mg/g) was continuously perfused in isolated loops for 120 min, including a 30-min 3H-phenylalanine flooding dose. We measured fractional synthesis rates of total mucosal protein and mucin by analyzing 3H-phenylalanine incorporation. Fractional rates of total mucosal protein synthesis were higher in loops perfused with solutions containing threonine at 56 mg/g (66 ± 4%/d) compared with 0 mg/g (42 ± 9%/d) and 21 mg/g (53 ± 6%/d) (P < 0.05). For mucin, fractional rates of synthesis differed between 0 mg/g (323 ± 72%/d), 21 mg/g (347 ± 49%/d), and 56 mg/g (414 ± 31%/d) (P < 0.05). In addition, total proline and threonine concentrations in the protein hydrolysates increased with luminal threonine concentration (P < 0.05), indicating an increase in threonine- and proline-rich proteins. De novo synthesis of mucosal and mucin proteins is acutely sensitive to luminal threonine concentration, which demonstrates the importance of dietary amino acid supply to gut protein metabolism.
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Introduction |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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One of the primary fates of amino acids extracted by the gut is synthesis of proteins, which are mostly secreted into the lumen (9). These endogenous secretions include large quantities of mucus, which protect the gut from pathogens and antinutritional factors. The main component of mucus is mucins, glycoproteins with a protein core that is particularly rich in threonine, proline, and serine, with threonine alone constituting 28–35% of the total amino acids (10,11). Mucin protein is very resistant to digestion and its amino acids cannot be reutilized by the animal (5,10). Small intestinal mucins, as well as colonic mucins, are readily digested by colonic microbes, but these amino acids are not available to the host because of negligible amino acid absorption in the large intestine. Therefore, intestinal mucin secretion represents a significant net loss of indispensable amino acids to the animal. Indeed, for most indispensable amino acids, gastrointestinal losses at the ileum accounted for 14–33% of the maintenance requirement, but for threonine, the contribution was 61% (12). Together, these data suggest that the gut has a high threonine requirement, which is a significant proportion of the whole body requirement. Indeed, mucin synthesis (13,14) and goblet cell morphology (15) are sensitive to dietary threonine supply. In situations of gut stress that would increase mucus turnover, it follows that threonine utilization would increase in proportion to whole body needs. Given the amino acid costs associated with stress-induced nonspecific protein synthesis, the costs for mucin-associated amino acids, such as threonine, would be much greater due to their specialized role in gut secretions. If threonine that is utilized during the first-pass metabolism for mucin synthesis is increased, then the availability of threonine in portal circulation will decrease and in turn decrease threonine that is available for peripheral tissues (16).
Because threonine is an indispensable amino acid and because the threonine requirement of the gut is disproportionately higher than other amino acids, the study of this amino acid in the gut is of particular interest nutritionally. Adegoke et al. (17,18) successfully validated a continuously perfused gut loop model using a luminal flooding dose technique that maintains first-pass nutrient exposure, limits any systemic effects by the perfused nutrients, and allows for multiple loops within an animal, which decreases experimental error. In this study, our objective was to adapt this model to demonstrate the limiting amino acid concept for luminal threonine on protein and mucin synthesis in the gut. We hypothesized that if luminal, first-pass threonine supply is deficient, then protein, and especially mucin, synthesis will be limited acutely, even in animals with adequate threonine status.
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Materials and Methods |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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The present experiment was conducted in accordance with the Canadian Council on Animal Care Guidelines and was authorized by the institutional Animal Care Committee. Six food-deprived, 10-kg Yorkshire pigs (Stuart King Farms) were anesthetized using an initial flow of 4% halothane mixed with oxygen and maintained with 2% halothane and oxygen. Under anesthesia, a midline incision was made to expose the small intestine. The ligament of Treitz was identified and 15 cm distal of the ligament, an inlet polyethylene cannula (i.d., 1.6 mm; o.d., 3.2 mm; Watson Marlow) was inserted into a hole made with an 18-gauge needle through the intestinal wall into the lumen. The cannula was secured with suture tied around the intestine and tubing. A 10-cm section of intestine was isolated and an outlet cannula was inserted similarly at the opposite end of the loop. A total of 4 loops was prepared similarly with 50 cm of intestine between loops. The loops were flushed with PBS (125 mmol/L NaCl, 15.9 mmol/L Na2HPO4, 1.2 mmol/L NaH2PO4, pH 7.4, 37°C) to remove any chyme that may be present. The intestines were then rinsed with warm 0.9% saline and placed gently back into the abdominal cavity for the duration of the experiment. To prevent adhesion between the handled intestines, adequate moisture levels were maintained by spraying the intestine with warm 0.9% saline and covering the abdomen with plastic wrap. Heart rate, respiration rate, body temperature, and oxygen saturation were monitored throughout the experiment.
Amino acid solution perfusions.
The perfusion solutions were continuously warmed in a water bath at 37°C. The inlet and outlet cannulae were connected to a multichannel peristaltic pump (Watson Marlow) that pumped respective perfusion solutions through the loops (5 mL luminal volume) at 3 mL/min in a closed loop system using a total perfusate volume of 50 mL. This flow rate approximates that estimated for chyme in human jejunum (i.e. 2–4 mL/min) (19) and ileum (i.e. 2.4 mL/min) (20). The various solutions and loops were randomly allocated. After a 90-min perfusion, each amino acid solution was replaced with an equivalent solution containing 50 µCi 3H-phenylalanine (Amersham Biosciences) in a 2-mmol/L phenylalanine solution. After this flooding dose of phenylalanine was perfused for an additional 30 min, the loop was excised using cautery to prevent bleeding, flushed with cold PBS, slit lengthwise, and scraped with a microscope slide on ice to remove the mucosa. Mucosal tissue was immediately frozen in liquid nitrogen and stored at –70°C until further analysis. To estimate the amount of tracer that was absorbed from the loops and then arterially extracted during the flooding dose, samples of intestinal tissue were removed from midway between the treated loops and analyzed for amino acids and radioactivity.
The amino acid profile of the basal perfusion solution was based on the profile in sow milk (21) with glutamine added (Table 1) with a total amino acid concentration of 27.1 mmol/L, which compares to the human jejunal digesta concentrations of total amino acids in the food-deprived state (i.e. 20 mmol/L) and of free amino acids in the fed state (i.e. 29 mmol/L) (22). In a similar loop model, total amino acid concentrations of the perfusate between 3.75 and 30 mmol/L did not affect protein synthesis rates (18). The osmolarity of the perfusates was 325 mosmol/L, which is similar to rat small intestinal osmolarity, ranging between 316 and 384 mosmol/L (23); in this model, it has been previously demonstrated in pigs that protein synthesis rates do not change using perfusates with 250–380 mosmol/L (17). The crystalline amino acids (Sigma-Aldrich) were dissolved in deionized water heated to 55–65°C under a blanket of nitrogen gas to prevent oxidation. A total of 151 mg/L of phenylalanine was added to one-half of the volume for the basal 90-min perfusion. To the other one-half, 330 mg/L (2 mmol/L) of phenylalanine was added and used as the flooding dose solution (17). PBS salts and test amino acids, threonine and serine, were then added according to the threonine level required (Table 2). Although serine is also enriched in mucin proteins, we decided to use this dispensable amino acid to maintain isonitrogenicity, because its plasma concentrations are particularly stable during fasting and refeeding in piglets, indicating its synthesis is not limited in young pigs, especially compared with other dispensable amino acids such as alanine, glycine, glutamate, and aspartate (24). Bertolo et al. (6) previously estimated the requirement of threonine in orally fed, 4-kg piglets as 28 mg/g amino acids. Three levels of threonine were tested in this study: 0, 21, and 56 mg threonine/g total amino acids, which reflect a range of threonine to amino acid ratios above and below the ratio required by the whole body (6). The test amino acids were added to both the basal and flooding dose perfusion solutions.
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The pellet containing the tissue bound amino acids was disrupted and washed 4 times with 4 mL of ice-cold 2% (wt:v) perchloric acid. The final pellet was resuspended in 1.3 mL of 1 mol/L NaOH by disrupting the pellet with a Teflon pestle; NaOH was added to a total volume of 2.4 mL. The samples were then left to sit in a 37°C water bath for 1.5 h to solubilize the protein for total protein analysis. To the remaining solution, 1.2 mL of cold 20% (wt:v) perchloric acid was added and the mixture was placed on ice for 20 min to reprecipitate the protein. The samples were then centrifuged at 2800 x g; 15 min and standards (norleucine and 14C-leucine) were then added to the pellet, which was disrupted with a glass rod in 1.2 mL of 6 mol/L HCl. This acid mixture was then transferred to a 25-mL digestion tube with cap; the tube was rinsed with additional 6 mol/L HCl to a final volume of 5 mL. The samples were digested at 110°C for 24 h for protein hydrolysis. The cooled hydrolysates were transferred to an Erlenmeyer flask and brought to a final volume of 25 mL with deionized water. The contents were filtered with a 0.45-µm filter and stored until further analysis.
Mucin purification and hydrolysis. Our method to purify mucin glycoprotein was a modified version of that developed by Faure et al. (25) in rats that exploits mucin's resistance to protease activity. Approximately 300 mg of mucosal scrapings was homogenized for 45 s in 2 mL of ice-cold 50 mmol/L Tris (pH 7.5) using a Polytron homogenizer at 30%. A solution containing 20 mg of Flavourzyme (a fungal complex of exopeptidases and endoproteases; Novozymes) was added to the homogenate and left to incubate under agitation in a 37°C water bath overnight. After 18 h, the enzymatic activity of Flavourzyme was stopped by placing the homogenate on ice for 10 min. Guanidium hydrochloride and dithiothreitol were added to 4 mol/L and 10 mmol/L concentrations, respectively, and the mixture was left at room temperature for 2 h under agitation. Finally, 25 mmol/L iodoacetamide was added to the homogenate and left overnight at room temperature under agitation. The mucins in this homogenate were then purified using gravity gel filtration chromatography using 10 mL PD-10 columns (Amersham Biosciences) filled with Sephacryl S-300 High Resolution resin (Sigma-Aldrich). The resin was equilibrated with 20 mL of 50 mmol/L Tris (pH 7.5) containing 2 mol/L guanidium HCl. The column was loaded with 1 mL of the homogenate followed by 12 mL of the equilibration buffer. Twelve fractions of 1 mL were collected, placed in dialysis tubing (Spectra/Por, MWCO 12–14,000; VWR International), and dialyzed against deionized water for 48 h. Following dialysis, the fractions were freeze-dried and resuspended in 0.1 mL of deionized water. The mucin-containing fractions were then identified by SDS-PAGE (4% stacking/7.5% migrating, at 20 mA) with periodic acid-Schiff's base staining for glycoproteins and Coomassie Blue staining for contaminating proteins. The mucin-containing fractions were identified by SDS-PAGE results and then combined in a digestion tube with 5 mL of 6 mol/L HCl and internal standards (norleucine and 14C-leucine) and hydrolyzed at 110°C for 24 h and stored for HPLC analysis.
Amino acid analysis. Samples were analyzed for amino acids using the Waters Pico-tag method, which uses phenylisothiocyanate derivatization and separation on a C18 reverse-phase column (Waters) (26). Leucine and phenylalanine fractions were collected and fraction radioactivity was determined using a liquid scintillation counter and Biodegradable Counting Scintillant (Amersham Biosciences).
Calculations and statistical analysis.
Protein and mucin fractional synthesis rates were calculated using the formula developed by Garlick et al. (27), using the following equation:
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where Ks is the fractional rate of protein or mucin synthesis in percentage of the respective protein pool synthesized per day; SRAb is the specific radioactivity of protein-bound phenylalanine; SRAf is the specific radioactivity of phenylalanine in the precursor pool (tissue free amino acids); and t is the time allowed for incorporation of the radioactive phenylalanine into protein.
Data were analyzed using a 1-way ANOVA with repeated measures (to account for blocking within pigs) and Student-Newman-Keuls multiple comparisons using Graph Pad Prism 4. Means ± SEM were considered significantly different at P < 0.05.
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Results |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Mucin purification. We found the mucin purification method of Faure et al. (25) to be just as effective as the classical method using CsCl density gradient centrifugation (28) but with better efficiency in time and labor (data not shown). Our initial results following the procedures of Faure et al. (25) indicated the presence of contaminating protein bands in the glycoprotein-containing fractions when stained with Coomassie Blue. These results suggested inadequate proteolysis by Flavourzyme, so we increased the amount of Flavourzyme from 10 mg to 20 mg and increased the incubation period from 2 h to overnight (18 h). These modifications did not change periodic acid-Schiff's-staining patterns for glycoproteins but did eliminate Coomassie Blue-staining bands in the mucin-containing fractions (Supplemental Fig. 1).
Protein and mucin fractional synthesis rates. The fractional synthesis rates of mucosal protein and mucin are presented in Figures 1 and 2, respectively. Blocking within each pig (by repeated measures analysis) was significant (P < 0.05) and so data are presented as means for each threonine level (Figs. 1A,2A) and as connected data within individual pigs (Figs. 1B,2B) to demonstrate the effects of blocking. With respect to total mucosal protein, fractional rates of synthesis were higher in loops perfused with solutions containing threonine at 56 mg/g compared with 0 and 21 mg/g (n = 6; P < 0.05) (Fig. 1). For mucin glycoprotein, fractional rates of synthesis differed between 0, 21, and 56 mg/g (n = 5; P < 0.05) (Fig. 2); we did not have enough mucosa for mucin purification in one of the pigs. The control loops that were perfused with nutrient-free PBS had protein synthesis rates of 113 ± 11%/d and mucin synthesis rates of 461 ± 65%/d. Untreated adjacent intestinal tissue not directly exposed to an amino acid mixture was also extracted and analyzed to determine the extent of arterial extraction of the luminal flooding dose and to correct raw data for protein synthesis rate calculations. The protein-bound and free phenylalanine specific radioactivity in this untreated tissue was 3–11% and 6–8%, respectively, of that found in the luminally flooded loops; these data were similar to those found by Adegoke et al. (17).
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Discussion |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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It is also possible that threonine is directly stimulating protein synthesis independent of its role as a limiting precursor. Indispensable amino acids, particularly leucine, are now known to act as nutrient signals to stimulate translation initiation and induce protein synthesis via an mammalian target of rapamycin (mTOR)-regulated pathway (29). The absence of individual amino acids inhibits p70 S6 kinase activity, a key enzyme facilitating the translation of capped mRNA (30,31). Although more focus has been placed on the potency of branched-chain amino acids (30) and arginine (32), deletion of threonine alone has been shown to moderately diminish p70 S6 kinase activity (31). In our model, intracellular threonine concentrations were not diminished (Table 3), probably given the acute treatment and the fact that arterial threonine supply was available to the intestinal mucosa. Whether higher than normal luminal threonine concentrations led to intracellular activation of protein synthesis is unknown but may be a contributing factor in our observations. Regardless of whether threonine is directly stimulating protein synthesis or simply providing precursors for limiting protein synthesis, the finding that increasing luminal threonine supply can lead to enhanced mucosal protein and mucin synthesis rates is profound and nutritionally relevant.
Although other recent studies in adult rats (13) and young pigs (14) have examined the effects of chronic whole body threonine deprivation on intestinal protein and mucin synthesis rates, our study is the first, to our knowledge, to examine the acute effects of varying dietary threonine levels. In rats, long-term feeding of diets with decreasing threonine levels did not affect total protein synthesis rates in mucosal tissues but did decrease fractional rates of mucin synthesis (13). In contrast, young pigs fed a threonine-deficient diet (i.e. 23 mg/g crude protein) for 14 d experienced both lower mucosal protein and mucin synthesis rates compared with a threonine-adequate diet (i.e. 46 mg/g crude protein) (14); however, this group also found a similar decrease in intestinal and mucin synthesis rates when pigs were fed a threonine-supplemented diet to 150% of whole body requirement (i.e. 69 mg/g crude protein). It is possible that high luminal threonine may result in transporter competition with other indispensable amino acids, as proposed by Wang et al. (14), although our study did not suggest such competition when threonine was fed at 56 mg/g. The 14-d feeding period used by both Faure et al. (13) and Wang et al. (14) may have allowed adaptation by intestinal mucosa, whereby threonine extraction from systemic circulation to the small intestinal tissue may have been upregulated to compensate for the chronic dietary deficit. In our study, the loop model and short adaptation perfusion time (90 min) was designed to preclude such compensation. These profound acute changes to protein synthesis also demonstrate the concept that intestinal cells primarily depend on luminal, first-pass nutrients in the fed state (33). Our data are not surprising considering recent data by Schaart et al. (8), who demonstrated that about three-quarters of the threonine incorporated into intestinal proteins come from dietary sources whether with a low- or high-protein diet.
Mucin synthesis rates are specifically sensitive to dietary threonine supply, both acutely in this study and chronically in studies by Faure et al. (13) in rats and Wang et al. (14) in pigs. However, it is difficult to make conclusions on the relative rate of mucin synthesis between these 3 studies. In the adult rat, it is possible that mucin synthesis does not contribute as large a proportion of total mucosal protein synthesis, which may explain why changes in mucin synthesis did not lead to measurable changes in total protein synthesis (13). Indeed, our mucin synthesis rates were double those observed by Faure et al. (13). In both studies, synthesis rates for mucin were higher than those observed for total mucosal protein, which would indicate that the mucin pool is being renewed at least every 24 h in adult rats and every 8 h in young pigs. In contrast, Wang et al. (14) found slightly lower rates of mucin synthesis (52–97%/d) compared with mucosal proteins (75–115%/d) despite using a similar young pig model and the same mucin isolation technique as our study. Our data indicate that intestinal mucosa turns over total protein (primarily in enterocytes) at a slower rate than mucin, indicating the importance of continuously renewing the mucus layer in the small intestine. Indeed, the quantitative significance of the amount of mucin synthesized was further demonstrated by the increasing proportions of threonine and proline in total mucosal protein hydrolysates as luminal threonine concentration increased (Table 3).
Adegoke et al. (17) carried out an initial study to validate the flooding dose method in the gut loop model. An interesting finding in both our study and that of Adegoke et al. (17) was that when the lumen of the small intestine was exposed to PBS only, without any nutrients, higher rates of protein synthesis were observed compared with those observed with amino acids present. Adegoke et al. (18) hypothesized that because protein synthesis utilizes high amounts of ATP to assemble a single peptide bond, energy preservation may be occurring with nutrient exposure to ensure that adequate amounts of energy are available for nutrient absorption into the systemic circulation. Therefore, protein synthesis may be suppressed when exposed to luminal nutrients to spare ATP.
In conclusion, this study demonstrated that during acute restriction of luminal threonine supply, the small intestinal mucosal and mucin protein synthesis rates are limited in young, well-nourished pigs. This finding is particularly profound considering that the arterial supply of threonine was adequate, suggesting that in the fed state, intestinal protein synthesis is primarily dependent on luminal, first-pass amino acids. These results have key nutritional implications and may help explain why a significant supply of enteral nutrients is necessary to maintain intestinal mass and integrity, even when parenteral nutrients are adequate (34).
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FOOTNOTES |
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2 Author disclosures: N. L. Nichols and R. F. Bertolo, no conflicts of interest.
3 Supplemental Figure 1 is available with the online posting of this paper at jn.nutrition.org.
Manuscript received 22 October 2007. Initial review completed 20 November 2007. Revision accepted 16 April 2008.
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