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Nutrient Metabolism

Onset of Small Intestinal Atrophy Is Associated with Reduced Intestinal Blood Flow in TPN-Fed Neonatal Piglets^1,2

U.S. Department of Agriculture/ARS Children’s Nutrition Research Center, Department of Pediatrics, Baylor College of Medicine, Houston, TX 77030 and ^* Department of Medical Physiology, University of Copenhagen, DK-2200 Copenhagen, Denmark

³To whom correspondence should be addressed. E-mail: dburrin{at}bcm.tmc.edu.

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
Our aim was to determine the speed of onset of total parenteral nutrition (TPN)-induced mucosal atrophy, and whether this is associated with changes in intestinal blood flow and tissue metabolism in neonatal piglets. Piglets were implanted with jugular venous and duodenal catheters and either a portal venous or superior mesenteric artery (SMA) blood flow probe. At 3 wk of age, piglets were randomly assigned to receive continuous enteral formula feeding (n = 8) or TPN (n = 17) for 24 or 48 h. Blood flow was recorded continuously and piglets were given an i.v. bolus of bromodeoxyuridine and ¹³C-phenylalanine to measure crypt cell proliferation and protein synthesis, respectively. After 8 h of TPN, portal and SMA blood flow decreased 30% compared with enteral feeding (P < 0.01), and remained near levels of food-deprived piglets for the remaining 48 h of TPN. After 24 h, TPN reduced jejunal inducible nitric oxide synthase (iNOS) activity and protein abundance (P < 0.05), small intestinal weight, and villous height (P < 0.01) compared with enterally fed piglets. Cell proliferation and DNA mass were decreased (P < 0.05) and apoptosis increased (P < 0.05) after 48 h of TPN. Protein synthesis was lower (P < 0.05) after 24 h of TPN, and protein mass was lower (P < 0.05) after 48 h of TPN, compared with enteral feeding. These data indicate that the transition from enteral to parenteral nutrition induced a rapid (<8 h) decrease in intestinal blood flow, and this likely precedes villous atrophy and the suppression of protein synthesis at 24 h, and of cell proliferation and survival at 48 h.

KEY WORDS: • enteral nutrition • intestinal adaptation • nitric oxide synthase • cell proliferation • protein synthesis

Total parenteral nutrition (TPN)⁴ leads to small intestinal atrophy and diminished intestinal function (1,2). Chronic TPN in neonatal piglets reduced proximal small bowel weight by as much as 70% compared with that of enterally fed piglets (3). Chronic TPN also was shown to induce anatomical changes, including decreased villous height, crypt depth, and villous surface area, and epithelial cell numbers (4,5). TPN-induced intestinal atrophy in piglets is associated with decreased protein synthesis and cell proliferation and with increased apoptosis and proteolysis in comparison with enterally fed piglets (6).

Due to the continual processes of cell proliferation and cell loss via exfoliation into the lumen, the tissue metabolic rate (e.g., oxygen uptake and protein synthesis) is relatively high in the small intestine compared with other tissues such as muscle (7). The high metabolic requirement of the small intestine renders it susceptible to luminal nutrient deprivation. Moreover, within the small intestine, the protein synthesis rates are higher in the proximal than in the distal region, reflecting a higher rate of cell turnover (8). In rodents and neonatal piglets, the enterocyte life span averages 3–5 d, depending on the location along the small intestine (9). Thus, it is possible that the suppression of mucosal epithelial cell kinetics by TPN occurs within a few days. Indeed, TPN-induced mucosal atrophy was observed within 3 d in rodents (10). That study demonstrated the most striking decreases in jejunal mucosal wet weights, protein, and DNA content after only 3 d of TPN, but the mean values decreased only slightly in the ensuing 12 d. Consistent with this finding, a recent study in neonatal piglets demonstrated a significant reduction in mucosal mass and villous height within 3 d (11).

The biological mechanisms that lead to mucosal atrophy during TPN have not been established, but are believed to involve both local nutrient-sensing cell signals (12) and humoral signals, such as gut hormones (13,14). Given that TPN-induced mucosal atrophy was observed within 3 d, it is likely that short-term physiologic mediators are involved. One such factor may be intestinal blood flow and tissue perfusion. Intestinal blood flow is strictly regulated to maintain the supply of oxygen and essential nutrients, while removing metabolic waste products. Intestinal blood flow increases rapidly with feeding in neonatal and mature animals (15,16). In neonatal piglets, we found that portal blood flow (PBF) increases up to 30–50% above values in food-deprived piglets almost immediately (within 1–3 h) after the start of enteral feeding (17). Conversely, TPN reduces intestinal blood flow as evidenced from recent studies in which piglets administered chronic TPN had lower portal blood flow rates than enterally fed piglets (18).

We hypothesized that intestinal mucosal atrophy occurs within 48 h of TPN and is associated with reduced intestinal blood flow in the neonatal piglet model. Our primary aim in this study was to quantify measurements of mucosal morphology, cell proliferation, apoptosis, and protein synthesis in neonatal piglets fed enterally or by TPN for 24 or 48 h and compare this with the rates of intestinal blood flow during this period.

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
    Animals. The study protocol was approved by the Animal Care and Use Committee of Baylor College of Medicine and was conducted in accordance with the NIH guidelines (19). Female crossbred piglets (n = 25; Large White x Hampshire x Duroc) were purchased from the Texas Department of Criminal Justice at the age of 5 d. They received an antibiotic injection (20 mg/kg enrofloxacin, Bayer) on the day of arrival at the animal facility. For the next week, they were housed at 30°C and fed a liquid milk replacer formula (Advance Liqui-Wean, Milk Specialties) at an increasing daily rate of 25 g (d 1–2) to 50 g (d 3–7) of dry matter/(kg body weight · d). The composition of the milk replacer formula was 430 g lactose, 130 g fat, and 250 g protein/1000 g dry matter. Pigs were fed liquid formula at a rate of 240 mL/(kg · d), which supplied 13.5 g protein/(kg · d) and 815 kJ/(kg · d), 90% of the NRC requirement. The formula did not contain antibiotics.

    Surgical procedure. The piglets were fed formula for 7–8 d. Surgery was performed at 11–12 d of age after overnight food deprivation. Under aseptic conditions and isoflurane anesthesia, the piglets were surgically implanted with a Tygon catheter (1.78 mm diameter) in the external jugular vein, and a silicone catheter (1.65 mm diameter, Baxter Healthcare) in the duodenum. An ultrasonic blood flow probe (Transonic Systems) was placed around the portal vein (6–10 mm probe) or the superior mesenteric artery (3–4 mm probe). The catheters were filled with saline containing heparin and exteriorized on the left flank (duodenal catheter and portal probe lead) or between the scapulae (jugular catheter). Immediately after surgery, the piglets received i.m. analgesic (0.1 mg/kg butorphenol tartrate, Fort Dodge Labs) and antibiotic (20 mg/kg enrofloxacin, Bayer). The catheters were protected with gauze pads and secured with an elastic bandage. After the surgery, the piglets were offered 60 mL/kg formula overnight, followed by 120 mL/kg on d 1 postsurgery and full feed (240 mL/kg) on d 2 postsurgery.

    Study protocol. One week after the surgery, at the age of 18–22 d, the piglets were randomly assigned to 3 study groups and deprived of food for 16 h before the feeding studies. After the 16 h of food deprivation, all piglets were given an oral feeding (20 mL/kg) of formula immediately followed by a duodenal infusion of formula at the rate of 10 mL/kg hourly for 8 h. After the 8-h enteral feeding period, the piglets were administered 1 of the following treatments: an enterally fed group administered continuous duodenal feeding at 10 mL/(kg · h) for 24 h (n = 4 piglets) or 48 h (n = 4 piglets) and a parenterally fed group administered TPN for 24 h (n = 9 piglets) or 48 h (n = 8 piglets). The TPN solution was infused i.v. at a rate of 10 mL/(kg · h) and consisted of glucose (104 g/L), lipid (21 g/L), a complete amino acid mixture (55 g/L, including glutamine), electrolytes, trace minerals, and vitamins to meet or exceed the requirements for neonatal pigs. To measure intestinal blood flow specifically, a separate group of piglets (n = 3), implanted with superior mesenteric arterial (SMA) probes, underwent the same procedure described above, except the period of TPN was only 20 h instead of 48 h. In all piglets, blood flow was recorded continuously starting 1 h before the oral bolus feeding, during the 8-h duodenal formula infusion, and until the end of the respective treatment periods. At these infusion rates, the enterally and TPN-fed piglets received approximately equal nutrient intakes [0.55 g protein/(kg · h) and 35 kJ/(kg · h)]. Blood samples were collected every 12 h in EDTA tubes and were frozen immediately in liquid nitrogen.

    In vivo cell proliferation and protein synthesis. An intravenous bolus of BrdU (bromodeoxyuridine, 50 mg/kg body weight; Sigma Aldrich) was given 4 h before the pigs were killed to estimate an index of crypt cell proliferation (see below). In addition, a bolus dose of ¹³C-phenylalanine (1.5 mmol/kg phenylalanine containing 0.15 mmol/kg ¹³C₆-phenylalanine, Cambridge Isotope Laboratories) was given 30 min before the pigs were killed to measure the rate of tissue protein synthesis. Pigs were killed with a venous injection of pentobarbital sodium (50 mg/kg body weight) and sodium phenytoin (5 mg/kg weight; Beuthanasia-D, Schering-Plough Animal Health). The abdomen was opened, and the whole small intestine was excised and quickly flushed with ice-cold saline, weighed, and divided into parts of equal length, designated as jejunum and ileum. Samples of jejunum were placed in formalin for morphological and BrdU analysis. The remaining intestinal tissues were frozen in liquid nitrogen and used for subsequent measurement of protein and DNA contents and isotopic enrichments.

    Blood flow measurement. In all piglets, either PBF or SMA flow was measured continuously by transit-time ultrasound via the ultrasonic transducer implanted in the respective vessels. The flow probe was coupled to a Transonic T206 flowmeter (Transonic Systems), and the blood flow rate was recorded and analyzed using the WinDaq/Pro+ software program (Dataq Instruments).

    Morphometry, cell proliferation, and apoptosis. Morphometry analysis was performed on formalin-fixed, hematoxylin and eosin–stained sections as described previously (6). In vivo crypt cell proliferation was measured as described previously (6). BrdU-labeled cells were detected by immunohistochemistry in formalin-fixed, paraffin-embedded sections and expressed as a percentage of total nuclei per crypt. Measurements of apoptosis were made based on cell morphology observed in 400X images by a single, trained observer who was unaware of the treatments. The apoptotic cells were characterized by evidence of condensed irregular chromatin, nuclear fragmentation, and intensely eosinophilic cytoplasm. Apoptotic cells were expressed as a percentage of the total epithelial cell numbers in the villus and crypt compartment of the same section.

    Tissue protein and DNA, and plasma analysis. Samples of jejunum tissue were pulverized in liquid nitrogen and assayed for protein (Pierce) and DNA (20) concentrations. Plasma amino acids were analyzed by reversed-phase HPLC for their phenylisothiocyanate derivatives (Pico Tag, Waters) (21) and plasma glucose using a glucose oxidase assay (kit 315–100; Sigma-Aldrich). Plasma glucagon-like peptide 2 (GLP-2) concentrations were quantified by RIA as described previously (17).

    Nitric oxide synthase (NOS) activity and NOS protein abundance. Intestinal tissue samples were used for determining the activities of inducible NOS (iNOS) and constitutive NOS (cNOS) by measuring the conversion of L-[¹⁴C]arginine into L-[¹⁴C]citrulline by a method similar to that described previously (21,22). Frozen tissue samples were homogenized, sonicated, and centrifuged at 12,000 x g for 20 min at 4°C. Extracts were assayed for iNOS activity in the presence of 60 mmol/L valine (an inhibitor of arginase), 0.1 mmol/L L-citrulline (to prevent the potential recycling of ¹⁴C-citrulline to arginine), and 4 mmol/L EGTA. Total NOS activity was assayed using the same conditions as iNOS, except that 2 mmol/L CaCl₂ replaced 4 mmol/L EGTA. For radioactivity blanks, 4 mmol/L N^G-nitro-L-arginine methyl ester (an inhibitor of NOS) was also included.

NOS isoform protein abundance was determined as described previously (21). Frozen intestinal tissue samples were homogenized, sonicated, centrifuged at 12,000 x g for 15 min at 4°C. Equal amounts (100 µg) of supernatant protein extracts were separated on a 7.5% denatured SDS-PAGE gel and transferred to polyvinylidene difluoride membranes. Membranes were blocked with 10 mL Pierce Superblock solution and then incubated with a primary antibody [rabbit polyclonal antibody against human eNOS (endothelial NOS, H-159) or iNOS (inducible NOS, H-174)(Santa Cruz Biotechnology)] diluted 1:500 in Tris-buffered saline with added Tween-20 solution (0.1%). Membranes were incubated with a secondary antibody (anti-rabbit IgG conjugated with biotin, 1:25,000, Santa-Cruz Biotechnology), enhanced with Neutravidin-HRP (1:25,000, Pierce), and allowed to react with horseradish peroxidase substrate (ECL-plus, Amersham Biosciences). Membranes were exposed to X-ray film for 30–120 s, and the image was scanned and quantified by ImageQuant 5.0 software (Molecular Dynamics, Amersham Biosciences). The apparent molecular weight of the eNOS and iNOS bands was 124 and 130 kDa, respectively.

    Mass spectrometry. Jejunal tissue were homogenized and deproteinized with 2 mol/L perchloric acid and the perchloric acid–soluble (tissue-free pool) and acid–insoluble (protein-bound pool) fractions were subjected to MS analysis as described previously (8). The acid-insoluble fraction was hydrolyzed with 6 mol/L HCl for 24 h before GC-MS analysis. The isotopic enrichment of [U-¹³C]-phenylalanine (M + 6 isotopomer) in the 2 tissue pools was determined by GC-MS analysis of the n-propyl ester heptafluorobutyramide derivative using methane-negative chemical ionization. The analyses were performed with a 5890 series II GC linked to a model 5989B (Hewlett-Packard) quadrupole MS. The isotopic enrichment of phenylalanine was determined by monitoring ions at a mass-to-change ratio of 383–389.

    Calculations. Protein synthesis was calculated as described previously (8) as the fractional synthesis rate (FSR, %/d)

where IE_bound and IE_free are the isotopic enrichments (mol% excess) of [¹³C₆]phenylalanine of the perchloric acid-insoluble (protein-bound) and perchloric acid–soluble (tissue-free) pool, t is the time of labeling (min), and 1440 is the number of minutes in a day.

    Statistics. The differences in morphology as well as intestinal weights, NOS activities, and protein synthesis were first analyzed using two-way ANOVA, with diet (enteral or TPN) and time (24 or 48 h) as main effects. We also analyzed these data using one-way ANOVA comparing 3 groups, by pooling the enteral groups and comparing differences between TPN at 24 and 48 h. In this analysis, differences among the 3 groups were determined using a Tukey multiple comparison analysis. The portal blood flow was analyzed with ANOVA using the Mixed procedure of SAS (SAS Institute). The model included the effects of time (24 or 48 h) and treatment (enteral feeding or TPN) as fixed effects, and piglet was included as a random effect. The SMA blood flow was also analyzed using the Mixed procedure with only time and animal in the model. SMA flow rates during the enteral feeding period were compared with the baseline value from food-deprived piglets, and flow rates during the TPN period were compared with the mean value estimated during the enteral feeding period. The results are expressed as means and SEM. Differences of P < 0.05 were considered significant.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
    Pig weights and amino acid and glucose concentrations. Piglet body weights were not influenced by dietary treatments (5.2 ± 0.2, 4.8 ± 0.2, and 5.4 ± 0.3 kg for enterally, 24 h TPN, and 48 h TPN fed piglets, P = 0.31). In addition, there were no significant differences in the plasma concentrations of essential amino acids (1993 ± 601, 2443 ± 643, and 2435 ± 453 µmol/L), nonessential amino acids (3293 ± 731, 3748 ± 1449, and 3482 ± 373 µmol/L), and glucose (7.4 ± 3.2, 6.3 ± 1.0, 5.7 ± 0.8 mmol/L) among the enteral, 24 h TPN, and 48 h TPN groups, respectively.

    Intestinal weights and morphology, protein contents and protein synthesis rate. The total small intestinal weight (jejunum and ileum) decreased from 43.2 ± 2.0 g/kg in enterally fed piglets to 35.9 ± 7.1 g/kg in 24 h TPN and 26.7 ± 2.7 g/kg in 48 h TPN piglets (P < 0.01). Because the ileum was largely unaffected by TPN, only the results from the jejunum are presented. Two-way ANOVA revealed significant main effect differences (P < 0.05), indicating that jejunal weight, villous height, protein mass, and protein fractional synthesis rate were lower in TPN than enterally fed piglets; however, there was neither a main effect of time nor were there significant diet by time interactions. Based on Tukey’s analysis of the 3 groups, jejunal mass was 23 and 44% lower (P < 0.05) after 24 h and 48 h of TPN, respectively, compared with enterally fed pigs (Table 1). Representative mucosal histological sections are shown in Figure 1. Jejunal villous height was higher in the enterally fed piglets (583 ± 81 µm) than in 24 h TPN (482 ± 41 µm) or 48 h TPN piglets (488 ± 73 µm) (P < 0.01). Tissue dry weight and protein concentrations (as mg/g tissue, data not shown) did not differ among enterally fed, 24 h TPN, and 48 h TPN piglets, but when expressed as mg/kg body weight (Table 1), protein masses were highest in enterally fed piglets, intermediate in 24 h TPN, and lowest in 48 h TPN piglets (P < 0.01). The protein mass decreased 41% in jejunum after 48 h of TPN. The FSR of enterally fed piglets (108%/d) was significantly higher (P < 0.05, Tukey’s test) than that in 24 h and 48 h TPN groups (Fig. 2).

View larger version (84K): [in this window] [in a new window]   FIGURE 1 Intestinal mucosal morphology of hematoxylin and eosin–stained sections of jejunum from piglets fed enterally (Top panel) or by TPN for 48 h (Bottom panel).

View larger version (26K): [in this window] [in a new window]   FIGURE 2 Protein fractional synthesis rate in jejunum of piglets fed enterally or by TPN for 24 or 48 h. Data are expressed as means ± SEM, n = 8 or 9. TPN is significantly lower than enteral (two-way ANOVA). *Different from enterally fed piglets, P < 0.05 (Tukey’s test).

View larger version (88K): [in this window] [in a new window]   FIGURE 3 BrdU-positive crypt cells (dark-brown stained) in jejunum sections from piglets fed enterally (Top panel) or by TPN (Bottom panel) for 48 h. Sections are counterstained with hematoxylin.

View larger version (23K): [in this window] [in a new window]   FIGURE 4 Proportion of BrdU-stained crypt cells in jejunum of piglets fed enterally or by TPN for 24 and 48 h. Data are expressed as means ± SEM, n = 8 or 9. TPN is significantly lower than enteral (two-way ANOVA). *Different from enterally fed piglets, P < 0.05 (Tukey’s test).

View larger version (96K): [in this window] [in a new window]   FIGURE 5 Apoptotic morphology of epithelial cells in the villus (Top panel) and crypt (Bottom panel) observed in hematoxylin and eosin–stained sections of jejunum from piglets fed by TPN for 48 h.

View larger version (26K): [in this window] [in a new window]   FIGURE 6 Rates of villous and crypt epithelial cell apoptosis in the jejunum of piglets fed enterally or by TPN for 24 and 48 h. Data are expressed as means ± SEM, n = 8 or 9. TPN is significantly higher than enteral (two-way ANOVA). *Different from enterally fed piglets, P < 0.05 (Tukey’s test).

View larger version (29K): [in this window] [in a new window]   FIGURE 7 Portal blood flow in piglets fed enterally or by TPN. Blood flow is expressed as the change from the mean blood flow during the period of food deprivation (h 0). All piglets were administered continuous duodenal feeding for 8 h (h 1–8); after that, the enteral group continued duodenal feeding, whereas the TPN-fed piglets were administered total parenteral nutrition for 24 or 48 h. Data points are expressed as least-squares means ± SEM; starting at 7 h after the initiation of TPN, the differences between the TPN (n = 16) and enterally (n = 8) fed groups were significant, P < 0.001 (mixed model ANOVA).

View larger version (23K): [in this window] [in a new window]   FIGURE 8 Superior mesenteric artery blood flow in piglets fed enterally and by TPN. Blood flow is expressed as the change from the mean blood flow during the period of food deprivation (h 0). All piglets received continuous duodenal feeding for 8 h (h 1–8), and then switched to TPN for 20 h. Data were analyzed by mixed model ANOVA and expressed as least-squares means ± SEM. Differences in mean flow at each time point vs. the food-deprived baseline level were determined during enteral feeding and TPN. *Different from baseline, P < 0.05, n = 3 piglets.

View larger version (25K): [in this window] [in a new window]   FIGURE 9 Total NOS (tNOS), inducible NOS (iNOS), and constitutive NOS (cNOS) activities in jejunum of piglets fed enterally or by TPN for 24 or 48 h. Data are expressed as means ± SEM, n = 8 or 9. TPN is significantly lower than enteral (two-way ANOVA). *Different from enterally fed piglets, P < 0.05 (Tukey’s test).

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

TPN-induced mucosal atrophy.

Longstanding evidence from animal studies indicates that the intestinal mucosa loses a major part of its structure and function within several days of starvation (23). Food deprivation for >24 h was shown to significantly decrease mucosal mass, but functional downregulation actually occurs within 12–18 h, which may be considered the maximal duration of food deprivation without enterally administered nutrients in normal life (24). In the case of TPN, considerable evidence showed that substantial intestinal mucosal atrophy occurs in young and mature animals after 1 wk of TPN (2,3,6,25). However, 2 studies showed that mucosal atrophy occurs within 3 d after the onset of TPN (10,11). In both of the latter studies, the most prominent decrease in intestinal mucosal weight, protein, and DNA content occurred after only 3 d of TPN, and these measures showed only a slight further decrease after 7 or 15 d of TPN. In the present study, we found a reduction in jejunal mass (23–40%) comparable to that reported in piglets after 3 d of TPN (11). However, the magnitude of intestinal villous atrophy that we observed after both 24 and 48 h (17%) was not as large as the range (30–50%) reported in piglets administered TPN for 3 d or more (6,11). Thus, based on the present study, as well as past studies in piglets, it appears that the reduction in mucosal mass occurs largely within 48 h, whereas the process of villus atrophy is complete within 72 h of TPN.

Previous studies involving chronic TPN showed that mucosal atrophy is accompanied by a suppression of protein synthesis and cell proliferation and a stimulation of proteolysis and apoptosis (6,8,25). In agreement with these studies, we found that jejunal protein synthesis was decreased within 24 h of TPN and preceded the decrease in protein mass at 48 h. Similarly, the rate of cell proliferation in the jejunum decreased in parallel with DNA mass after 48 h of TPN, and the rate of apoptosis in both the villous and crypt cells increased after 48 h of TPN. Taken together, the present results suggest that the early onset of TPN-induced mucosal atrophy is associated with reduced cellular metabolism (i.e., protein synthesis) and followed subsequently by suppression of cell proliferation and survival. The significant decrease in jejunum villous height after 24 h of TPN administration despite no change in cell proliferation, apoptosis, or cell numbers may have been due to reduced cell hydration or volume.

TPN-induced suppression of intestinal blood flow.

Intestinal blood flow is strictly regulated to maintain the supply of oxygen and essential nutrients and to remove waste products. Intestinal blood flow typically represents 15–20% of cardiac output in the unfed state (15). However, during feeding and nutrient absorption, blood flow is increased sequentially as the chyme passes over the mucosal surface. In human infants, fasting mesenteric artery blood flow, measured noninvasively with Doppler ultrasound, increased up to >50% with feeding, and reached a peak within 1 h after feeding (27,28). Within 2 h after feeding, the splanchnic blood flow reverted to the prefeeding level (29). As in human infants, the rapid stimulation of intestinal mucosal blood flow also was demonstrated in formula- and milk-fed neonatal piglets (16,26). Van Goudoever and co-workers (17) measured PBF continuously and also observed a marked increase (30–50%) above food-deprived levels within 1 h after enteral feeding, which reached the maximum within 3–6 h in continuously fed piglets. Thus, the changes in PBF are largely indicative of intestinal blood flow because the small intestine represents the largest proportion of portal-drained visceral tissues. This was evident in the similar patterns of response in both PBF and SMA flow to enteral and parenteral nutrition.

Given the rapid responsiveness of intestinal blood flow to enteral feeding, we predicted that TPN would suppress PBF relative to the rate observed with enteral feeding. To our knowledge, the present results are the first to demonstrate that TPN acutely reduces intestinal blood flow relative to enteral nutrition. We previously found that PBF was reduced after 1 wk in TPN-fed pigs (18); however, the temporal nature of this response was unknown. In this study, we found that both PBF and SMA flow increased rapidly after the initiation of enteral feeding, reached a maximum of 40% above food-deprived levels, and remained constant during the 48 h of continuous duodenal feeding. However, after the onset of TPN, both PBF and SMA flow decreased to approximately the food-deprived values within only 8 h and, more importantly, persisted at that level throughout the 20–48 h of TPN. Thus, TPN was insufficient to maintain intestinal blood flow at the level in enterally fed piglets, even though the nutrients were provided in equal amounts via both the parenteral and enteral routes. This was supported by evidence that the circulating concentrations of glucose and amino acids were similar among the 3 groups. The results suggest but do not prove that the early decrease in intestinal blood flow within 6–12 h may precede the appearance of mucosal atrophy observed after 24 h of TPN; earlier mucosal measurements are required to prove this point. Furthermore, the TPN-induced suppression of SMA flow supports the idea that perfusion was reduced in the small intestine where mucosal atrophy was localized. These results suggest that reduced tissue perfusion and substrate availability may be an early event that leads to local suppression of epithelial cell protein synthesis, cell proliferation, cell survival, and eventually mucosal atrophy.

TPN induced suppression of NOS activity.

The TPN-induced changes in PBF and SMA flow prompted us to examine whether NO production could be involved in the local regulation of intestinal blood flow. Nitric oxide is an important regulator of vascular perfusion and is produced by 3 enzyme systems: 2 constitutive forms, endothelial (eNOS) and neuronal (nNOS), and 1 inducible form (iNOS). Low levels of NO production, typically ascribed to the constitutive NOS forms, are regarded as good for cell function, whereas excessive NO production, associated with iNOS expression, is considered detrimental for tissue well being (30). Generally, eNOS activity is considered to be the NOS isoform involved in regulation of vasodilatation and blood flow rate, whereas iNOS is induced during inflammation and ischemic injury (31). Indeed, iNOS production was increased in premature piglets with necrotizing enterocolitis (32), but may be a key mediator of early villous repair after acute injury (33). Moreover, iNOS is expressed in the gut under normal physiologic conditions because the mucosal surface is constantly exposed to foreign antibodies in food; nevertheless, its role in modulating blood flow has not been established (30).

Associated with the induction of mucosal villous atrophy, we found a significant decrease in iNOS activity and protein abundance after 24 h of TPN, with no change in eNOS activity or protein abundance. The finding of reduced NOS activity and intestinal blood flow in TPN- vs. enterally fed piglets agrees with the reports that NOS is involved in the regulation of local blood flow. However, our finding of reduced iNOS activity is in contrast with the general theory that changes in vasodilation and blood flow are mediated by expression of the endothelial NOS isoform. In addition, Reber and co-workers recently reported increased eNOS abundance with enteral feeding in neonatal piglets (34). Our results are also in contrast with the finding of Hsu et al. (35), who found increased iNOS activity in rats administered a TPN solution orally vs. a standard diet. If NOS activity were causally linked to blood flow in this situation, the changes in NOS activity would be expected either to precede or occur simultaneously with blood flow. As discussed above with respect to villous morphology, it will be important to establish whether the temporal decrease in intestinal blood flow is paralleled by a decrease in NOS activity in the small intestine.

The results from this study demonstrate that intestinal mucosal atrophy is evident after 48 h of TPN in neonatal piglets. Moreover, the loss of mucosal protein and DNA mass after 48 h of TPN was accompanied by a reduction in protein synthesis and cell proliferation and an increase in apoptosis. A potentially important finding of this study was the fact that TPN rapidly (within 8 h) suppressed portal venous and SMA blood flow and that within 24 h of TPN, jejunal mucosal iNOS activity decreased and mucosal atrophy occurred. Whether the suppression of mucosal blood flow is causally linked to TPN-induced mucosal atrophy warrants further study. However, we showed recently that systemic infusion of GLP-2 prevents mucosal atrophy in TPN-fed piglets (6) and does so via an NO-dependent stimulation of intestinal blood flow and substrate utilization (21). Although we did observe a decrease in plasma GLP-2 concentration after 24 h of TPN, consistent with previous studies (6), further research is warranted to establish whether the early decline in PBF within 8 h of TPN is strictly correlated with a reduction in plasma GLP-2. Taken together, the findings of this study may provide a putative mechanism to explain the induction of mucosal atrophy that occurs with TPN. The rapid reduction in intestinal blood flow after initiating TPN could result in an insufficient supply of oxygen and nutrients to villous epithelial cells and lead to hypoxia, oxidant stress, and eventually cell death and villous atrophy. The cellular signal whereby TPN or the absence of luminal nutrition triggers the reduction in intestinal blood flow is unknown. However, studies have shown that increased osmolarity and glucose within the villous interstitial tissue, which occurs during the apical absorption of luminal glucose, may trigger the release of nitric oxide or serotonin and in turn affect vasodilation (36,37). Further studies are warranted to establish how the presence of nutrients on the luminal vs. basolateral surface (i.e., enteral vs. parenteral nutrition) of the mucosal epithelium triggers the stimulation of intestinal blood flow.

	ACKNOWLEDGMENTS

 
The authors thank Xiaoyan Chang and Xiaofeng Lu for technical assistance, Guoyao Wu for assistance developing the NOS enzyme assays, Michael Helmrath for surgical assistance, and Leslie Loddeke for editorial assistance.

	FOOTNOTES

 
¹ This is a publication of the USDA/ARS Children’s Nutrition Research Center, Department of Pediatrics, Baylor College of Medicine and Texas Children’s Hospital, Houston, TX. The contents of this publication do not necessarily reflect the views or policies of the U.S. Department of Agriculture, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. government.

² Supported by federal funds from the U.S. Department of Agriculture, Agricultural Research Service under Cooperative Agreement Number 58–6250-6–001, and by National Institutes of Health grants HD33920 (D.G.B.). H.N. was supported in part by grants from the Sigrid Juselius Foundation and Foundation for Pediatric Research, Finland.

⁴ Abbreviations used: BrdU, bromodeoxyuridine; GLP-2, glucagon-like peptide 2; NOS, nitric oxide synthase; PBF, portal blood flow; SMA, superior mesenteric artery; TPN, total parenteral nutrition.

Manuscript received 18 February 2004. Initial review completed 8 March 2004. Revision accepted 11 March 2004.

	LITERATURE CITED

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 

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