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Original Articles: Gastroenterology

Does Small Intestinal Atresia Affect Epithelial Protein Expression in Human Newborns?

Schaart, Maaike W*,†; Yamanouchi, Takeshi†,‡; van Nispen, Danielle JPM§; Raatgeep, Rolien HC§; van Goudoever, Johannes B*; de Krijger, Ronald R|,|; Tibboel, Dick; Einerhand, Alexandra WC§; Renes, Ingrid B*

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Journal of Pediatric Gastroenterology and Nutrition: November 2006 - Volume 43 - Issue 5 - p 576-583
doi: 10.1097/01.mpg.0000235755.22111.83
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Abstract

INTRODUCTION

At an estimated overall incidence of 1 in 1500 to 5000 live births, small intestinal atresia is a well-known cause of neonatal intestinal obstruction. It is believed to have its genesis as an embryopathy at the level of the duodenum in the first trimester of pregnancy or as an anomaly of ischemic origin in the second or third trimester of pregnancy (1).

The small intestine is one of the most metabolically active tissues in the body. Its best-regulated functions are epithelial proliferation, differentiation and apoptosis. Four specialized cell types in the epithelium play key roles in maintaining intestinal functions: enterocytes, goblet cells, Paneth cells and enteroendocrine cells. Enterocytes facilitate digestion, uptake and transport of nutrients by expressing, for example, the sugar-degrading enzyme sucrase-isomaltase (SI), the glucose and fructose transporters 2 and 5 (Glut2 and Glut5) and the intestinal fatty acid binding protein (i-FABP) (2,3). Enterocytes also express alkaline phosphatase (AP), which aids intestinal defense by detoxifying endotoxins (4). Goblet cells synthesize the secretory mucin 2 (MUC2), the structural component of the intestinal mucus layer that protects the epithelium from mechanical stress, bacteria, viruses and other pathogens (5). Goblet cells also synthesize and secrete trefoil factor 3 (TFF3), a bioactive peptide involved in epithelial protection and repair (6). Synthesizing antimicrobial peptides, Paneth cells are crucial to epithelial defence (7,8). Enteroendocrine cells are specialized in the mucosal secretion of hormonal peptides.

Evaluation of mucosal morphology in chick embryos (9) and fetal lambs (10,11) after experimental intestinal atresia, and in human neonates (12) with jejunal or ileal intestinal atresia revealed an abundance of shorter, flattened villi in the proximal, dilated segment. The distal, narrowed segment had tall, hypertrophic villi, often obliterating the intestinal lumen. The obstruction would prevent access of luminal components that may stimulate intestinal development, such as amniotic fluid (before birth) and/or enteral nutrition (after birth). Absence of these components may lead to an immature bowel distal to the obstruction. In the current study, we hypothesized that human newborns show mature epithelial differentiation proximal to a jejunal or ileal atresia, but immature epithelial differentiation distally. We used a variety of relevant markers to immunohistochemically investigate enterocyte differentiation: lactase, SI, sodium glucose cotransporter 1 (SGLT-1), Glut2, Glut5, i-FABP and AP. Goblet cell differentiation was determined by the expression of MUC2 and TFF3 and Paneth cell differentiation by the expression of the antibacterial enzyme lysozyme. Thus, by studying all of these parameters in conjunction, we aimed to gain more insight into intestinal epithelial protein expression and thus epithelial differentiation in human newborns, proximal and distal to jejunal and ileal atresias.

PATIENTS AND METHODS

Patients

Patients eligible for this study were newborns who had undergone surgery for jejunal or ileal atresia in the Department of Pediatric Surgery, Erasmus Medical Center (MC)–Sophia Children's Hospital (Rotterdam, The Netherlands). The diagnosis had been made by plain x-ray or ultrasound and was confirmed at surgery. We excluded cases of duodenal atresia, atresias complicating meconium ileus and/or meconium peritonitis and atresias associated with gastroschisis. Table 1 shows main clinical characteristics of the 16 patients studied.

TABLE 1
TABLE 1:
Patient characteristics

Ten of the 16 newborns had received standard nutrition for 1 or 2 days after birth, before being transferred to our hospital. Upon onset of clinical symptoms for intestinal obstruction, they had been given adequate intravenous fluid resuscitation to achieve hemodynamic stabilization. Antenatal maternal ultrasonography showing polyhydramnion or dilated intestines had pointed at intestinal obstruction in the other 6 patients. They received no enteral feeding after birth but intravenous glucose infusion.

Experimental Design

We studied intestinal tissue sections of these patients provided by the Erasmus MC Tissue Bank, with permission of the local medical ethics committee and according to the Code Proper Secondary Use of Human Tissue, thus in compliance with Dutch law and ethics regulations. Expression of the epithelial enzymes was studied in sections taken at surgery, proximal and/or distal to the jejunal or ileal atresia in close vicinity to the resection areas. For patients with multiple atresias, we preferentially used sections close to first atretic segment. If these were not available, we designated a random intestinal segment proximal or distal to one of the other atretic segments as the distal part and studied this. In addition, we studied small-intestinal tissue samples of 8 control patients. For the five patients, undergoing stoma formation or bowel resection in the neonatal period for intestinal perforation (n = 1), bowel necrosis (n = 1), volvulus (n = 1) and bowel stenosis (n = 2), these had been obtained during surgery. For the 3 patients, these had been obtained at autopsy after death from complications of congenital diaphragmatic hernia diafragmatica (n = 2) or persistent pulmonary hypertension of the newborn (n = 1). All intestinal biopsies were immediately fixed in 4% (wt/vol) paraformaldehyde in phosphate buffered saline (PBS) and prepared for light microscopy. Two investigators, blinded for the proximal and distal resection areas, independently assessed the sections on histology and epithelial protein expression.

Histology

Sections of 5-μm thickness were routinely stained with hematoxylin and eosin to study histological changes (ie, distortion of crypt/villus epithelium and lymphoid aggregates close to the jejunal or ileal atresia). Atrophy was defined as a crypt/villus ratio of 1:3 or less, and hypertrophy was defined as a crypt/villus ratio of more than 1:4.

Immunohistochemistry

Five-micrometer-thick paraffin sections were cut and deparaffinized through a graded series of xylol-ethanol as described previously (13). Briefly, endogenous peroxidase activity was inactivated with 3% (vol/vol) hydrogen peroxide in PBS for 30 minutes, followed by rinsing in PBS for 15 minutes. The sections were boiled in 0.01 mol/L citrate buffer (pH 6.0) or EDTA (5 mmol/L, pH 8.0) for 10 minutes. To reduce nonspecific binding, the sections were then incubated with TENG-T (10 mmol/L Tris-HCl, 5 mmol/L EDTA, 150 mmol/L NaCl, 0.25% (wt/vol) gelatin and 0.05% (wt/vol) Tween 20) for 30 minutes. This was followed by overnight incubation with primary antibodies. To determine enterocyte-specific protein expression, we used antihuman lactase (DR BB 2/33: 1:2000; A.Q.), antirat SI (1:6000) (14), anti-rabbit SGLT-1 (1:2000) (15), antihuman Glut2 (1:2000; B.T.), antirat Glut5 (1:1500; D.R.Y.) and antihuman i-FABP (1:4000) (16). As a marker for goblet cell–specific protein expression, we used a human MUC2-specific antibody (We9, 1:200) (17) and antihuman TFF3 (1:2000, see following paragraph for antibody preparation). Antihuman lysozyme (1:25, Dako, Glostrup, Denmark) was used to detect Paneth cell–specific protein expression. Sections were then incubated for 1 hour with biotinylated horse antimouse IgG (diluted 1:1000, Vector Laboratories) or with biotinylated goat anti-rabbit IgG (diluted 1:2000, Vector Laboratories) followed by 1 hour incubation with ABC/PO complex (Vectastain Elite Kit, Vector Laboratories) diluted 1:400. After incubation, binding was visualized in 0.5 mg/mL 3,3′-diaminobenzidine, 0.02% (vol/vol) H2O2 in 30 mmol/L imidazole and 1 mmol/L EDTA (pH 7.0).

Trefoil Factor 3 Cloning, Expression and Antibody Preparation

The coding sequence for human (h)TFF3 was amplified from the HITF plasmid using primers: 5′-TACGTAGAGGAGTACGTCGGCCTG-3′ containing the SnaBI restriction site and 5′-TCAATGATGATGATGATGATGGAAGGTGCATTCTGCTTCCT-3′ (18). The resulting polymerase chain reaction product was cloned into pCR2.1 and verified by sequence analysis. Subsequently, the coding sequence of hTFF3 was subcloned into the yeast expression vector pPic9K using SnaBI and EcoRI. The vector was transformed into, and expressed by, the Pichia pastoris strains KM71 and GS115 by using the Pichia Multi-Copy Expression kit (Invitrogen) according to the manufacturer's instructions. Recombinant hTFF3/HIS-tag fusion proteins were isolated using a nickel column (Pharmacia, Diegem, Belgium) according to the manufacturer's instructions. The eluted recombinant hTFF3 protein was dialyzed and concentrated. Subsequently, New Zealand White rabbits (Broekman, Utrecht, The Netherlands) were immunized with recombinant hTFF3 diluted in Gerbu Adjuvant (Instruchemie, Hilversum, The Netherlands) according to the manufacturer's instructions. The polyclonal TFF3 antibody preparation was performed with approval of the Erasmus MC Animal Studies Ethics Committee.

Histochemistry

Enterocyte-specific alkaline phosphatase activity was assessed by a 1-step assay. The deparaffinized and rehydrated tissues sections were incubated with a Tris-buffer (pH 9.5) containing 50 μL 4-nitroblue tetrazolium chloride (Vector Laboratories) and 37.5 μL 5-bromo-4-chloro-3-indolyl-phosphate (Vector Laboratories) according to the manufacturer's protocol. The color reaction was performed for 1 hour in the dark and was stopped with distilled water; slides were mounted with aquamount improved (Gurr, Brunschwig, Amsterdam, The Netherlands).

Statistics

Patient characteristics are presented as the mean ± SD.

RESULTS

Patients

The 16 patients with a jejunal or ileal atresia had mean gestational age of 37 ± 2 weeks and mean birth weight of 2770 ± 551 g (Table 1). Surgery had been performed at 3 ± 1 days after birth. In 4 patients, intestinal anastomosis had not been possible during the initial surgery, and they underwent reanastomosis after 54 ± 58 days. One infant had died 38 days after birth from complications of antenatal ischemia caused by twin-to-twin transfusion syndrome.

The 8 control patients had mean gestational age of 37 ± 2 weeks, and intestinal surgery or autopsy had been performed 4 ± 4 days after birth.

Morphology

Sections both proximal and distal to jejunal and ileal atresias showed a patchy pattern of morphological changes. Changes observed were villus atrophy, flattening of crypt and villus cells, flattening of the surface epithelium and villus hypertrophy (Fig. 1). The lamina propria contained hemorrhages and some lymphoid aggregates, plasma cells, individual lymphocytes and eosinophilic granulations comparable with the normal situation in the lamina propria.

FIG. 1
FIG. 1:
Epithelial morphology. Hematoxylin and eosin staining. Atrophic (A and C) and hypertrophic (B and D) segments were observed both proximal (A and B) and distal (C and D) to jejunal and ileal atresias.

Enterocyte-specific Protein Expression

Enterocyte-specific protein expression was studied immunohistochemically, using antibodies against lactase, SI, Glut2, Glut5, i-FABP and SGLT-1. All of the enterocyte markers were normally expressed in control sections (data not shown). Lactase, SI, SGLT-1 and Glut5 protein expression is normally confined to the brush border of villus enterocytes (19,20). In the atresia patients, these proteins were expressed both proximal and distal to the atretic segment in jejunum as well as ileum (Fig. 2A and B,: SI; Fig. 2C and D, lactase; Glut5 and SGLT-1, data not shown). Glucose and fructose transporter 2 was expressed at the basolateral membrane of villus enterocytes. Similar to the proteins expressed at the brush border, Glut2 was normally expressed proximal and distal to jejunal and ileal atresias (Fig. 2E and F). Intestinal-FABP protein is normally found in the cytosol of the jejunal and ileal villus enterocytes (21). As with the membrane-bound proteins, expression of i-FABP was normal (Fig. 2G and H). The in situ AP activity was observed in the brush border of the surface enterocytes both in proximal and distal segments of jejunal and ileal atresias (data not shown). Expression of all described enterocyte-specific proteins did not differ between segments characterized by atrophy and segments characterized by hypertrophy (Fig. 3A and B,: SI; other markers, data not shown). Thus, all enterocyte-specific proteins investigated were present in the small intestine (jejunum and ileum) at 3 ± 1 days after birth. Enterocyte-specific protein expression in the small intestine was unaffected by the presence of a jejunal or ileal atresia.

FIG. 2
FIG. 2:
Enterocyte-specific protein expression. Sucrase-isomaltase (A and B) and lactase (C and D) were both expressed proximal (A and C) and distal (B and D) to jejunal and ileal atresias. Glucose and fructose transporter 2 was expressed at the basolateral membrane of jejunal and ileal villus enterocytes, both proximal (E) and distal (F) to the atretic segment. Intestinal fatty acid binding protein was expressed in the cytosol of jejunal and ileal villus enterocytes both proximal (G) and distal (H) to the atresia.
FIG. 3
FIG. 3:
Enterocyte-specific SI and goblet cell–specific MUC2 expression in areas characterized by atrophy or hypertrophy. Enterocyte-specific SI expression (atrophy [A] and hypertrophy [B]) and goblet cell–specific MUC2 expression (atrophy [C] and hypertrophy [D]).

Goblet Cell– and Paneth Cell–specific Protein Expression

Goblet cell–specific protein expression patterns were studied using MUC2 and TFF3 specific antibodies. Similar to enterocyte-specific protein expression, control sections showed normal expression of MUC2 and TFF3 (data not shown).

Trefoil factor 3 specificity was determined on adult human stomach tissue obtained during gastroscopy and control small-intestinal sections. Figure 4 shows the negative and positive controls for TFF3.

FIG. 4
FIG. 4:
Trefoil factor 3 specificity determined on adult human stomach tissue (A, negative control) and human neonatal small-intestinal tissue (B, positive control).

No differences were found in MUC2 and TFF3 protein expression on either side of the jejunal or ileal atresia independent of atrophy or hypertrophy (Fig. 3C and D, MUC2; TFF3, data not shown). Goblet cell–specific MUC2 and TFF3 expression is shown in Figure 5 (A–D).

FIG. 5
FIG. 5:
Goblet cell–specific protein expression and Paneth cell–specific protein expression. Mucin 2 (A and B) and TFF3 (C and D) were expressed on either side of jejunal or ileal atresias (proximal, A and C; distal, B and D). Lysozyme was expressed in Paneth cells at the crypt base in jejunum and ileum, both proximal (E) and distal (F) to the atresias.

Lysozyme was used as a marker for Paneth cell–specific cell function. It showed a normal expression pattern in control neonatal small bowel (data not shown). In the biopsies taken from our patients, lysozyme was expressed in Paneth cells at the crypt base in jejunum and ileum. Lysozyme expression in patients' sections did not differ between proximal and distal segments (Fig. 5E and F) and neither between atrophic and hypertrophic areas (data not shown). Also, localization of Paneth cells at the bottom of the crypt did not differ between proximal and distal segments. Thus, similar to the enterocyte markers, goblet cell– and Paneth cell–specific protein expression was observed at a mean age of 3 days after birth. Expression did not differ either proximal or distal to jejunal and ileal atresias.

DISCUSSION

We investigated the effect of jejunal and ileal atresia on epithelial morphology and epithelial protein expression in human newborns, proximal and distal to the atretic segment, shortly after birth. Morphological analysis demonstrated structural alterations of villi and crypts at both sides of the atretic segment. We observed areas with hypertrophic villi at either side of the jejunal and ileal atresia. Our findings are in line with studies by Touloukian and Wright (12), and Tilson (22) showing significant villus hypertrophy, most marked in segments distal to the small-intestinal obstruction. However, next to hypertrophic villi, we found villus atrophy both proximal and distal to jejunal and ileal atresias, which was not previously reported. The natural history of the defect during the prenatal period of life (ie, vasculary insufficiency resulting in the atresia followed by patchy repair) may explain our histological findings.

In agreement with our findings in normal neonatal intestinal tissue, several studies have clearly demonstrated expression of lactase, SI, SGLT-1, Glut2, Glut5 and AP; the goblet cell markers MUC2 and TFF3 and the Paneth cell marker lysozyme already at messenger RNA (mRNA) and/or protein level in healthy fetal small intestine and persisting after birth (6,23,24). More specifically, mRNAs encoding the glucose transporter proteins Glut2 and Glut5 are detectable in human fetal intestine as early as 11 weeks postconception (23,24). Sodium glucose cotransporter 1 mRNA is detectable at 17 weeks of gestation and sucrose-isomaltase mRNA at 13 weeks of gestation (23). Lactase is expressed early in gestation, but its activity increases markedly during the third trimester, probably to meet the needs of full-term newborns (24). Dahlqvist and Lindberg detected jejunal AP activity in 11-week-old fetuses (25) detected jejunal AP activity in 11-week-old fetuses. Intestinal fatty acid binding protein has not been studied in fetal human tissue but is expressed in adult intestinal tissue (26). Lin and colleagues (6) detected the goblet cell marker TFF3 by immunohistochemistry in human intestine as early as 12 weeks of gestation. Goblet cell–specific MUC2 mRNA is expressed throughout the intestinal epithelium at 9 weeks postconception, therefore concomitantly with endodermal cytodifferentiation associated with the formation of premature crypts and villi (27). Paneth cell–specific lysozyme specifically stains at about 20 weeks of gestation (28). In the present study, we found for each patient enterocyte-, goblet cell– and Paneth cell–specific marker expression proximal to the jejunal or ileal atresia. Taken together, these data indicate that the epithelial-specific proteins are expressed in early neonatal life (within 3 days after birth) suggesting that enterocytes, goblet cells and Paneth cells can express a wide variety of proteins very early in life. More important, the expression of these proteins is not affected by the jejunal or ileal atresia itself.

The presence of luminal components such as amniotic fluid and biliary-pancreatic secretion during fetal intestinal development may be crucial to epithelial protein induction in utero and after birth. Surana and Puri (29), for example, showed that amniotic fluid has a nutritive role for the fetus and that intestinal obstruction blocking this pathway causes intrauterine growth retardation. Furthermore, animal studies found experimental elimination of fetal ingestion to result in retarded growth of the gastrointestinal tract and changes in the intestinal epithelial architecture (30,31). Our present findings in human material nevertheless suggest that absence of luminal factors does not affect induction and/or maintenance of epithelial-specific protein expression. Furthermore, these data imply that an intrinsic program encoded in the epithelial cells determines enterocyte-, goblet cell– and Paneth cell–specific protein expression. Our findings are in line with isograft studies demonstrating normal induction of intestinal liver–FABP in fetal intestinal tissues implanted subcutaneously into nude mice. Rubin and colleagues (32) therefore suggest that the gut stem cell is multipotent, has great capacity for self-renewal and can to be programmed/imprinted with positional information. Recent mouse studies show that mesenchymal proteins such as Wnt and bone morphogenetic proteins can direct intestinal epithelial differentiation and thus epithial-specific gene expression (33–35), implying a pivotal role for these factors in the epithelial differentiation program. Studies in pig and sheep found systemic and luminal factors (hormones and growth factors) influence intestinal development and differentiation (36).

Although we detected no differences in SI and lactase protein expression proximal and distal to jejunal and ileal atresias, we cannot rule out differences in enterocyte-specific enzyme activity because we did not quantify disaccharidase activity. Serrano et al. (37) did, however, and measured lower activity of SI and lactase proximal to a small-intestinal obstruction. Distally, only lactase activity was significantly reduced. Reduced lactase activity may affect lactose digestion and thus may indirectly be responsible for these patients' failure to thrive. Surgery for small intestinal atresia regularly is associated with symptoms of malabsorption and growth retardation (38). Even if intestinal surgery is successful and food intake is adequate, these symptoms may persist for several months (39). However, we still feel that factors such as villus morphology, gut caliber, motility and remaining intestinal length have greater impact on delayed postoperative recovery than the reduced protein expression levels observed by Serrano et al. (37).

Like the enterocyte markers, the goblet cell markers MUC2 and TFF3 and the Paneth cell marker lysozyme were also expressed at both sides of the jejunal and ileal atresia. Thus, our findings suggest that defense and repair functions of the small-intestinal mucosa are maintained by goblet cell and Paneth cell expression of MUC2, TFF3 and lysozyme, respectively, close to the atretic segment.

In conclusion, in contrast to observations in animals the present study demonstrates that the small-intestinal epithelium in humans is mature at birth. Luminal components, such as amniotic fluid (before birth) and enteral nutrition (after birth), are not essential for the epithelial maturation of the intestine. Taken together, the results highlight that epithelial protein expression, which is crucial to nutrient absorption, epithelial defense and repair in the small intestine, is genetically imprinted and that indicates the presence of indispensable ontogenetic factors.

REFERENCES

1. Miller AJW, Rode H, Cywes S. Intestinal atresia and stenosis. In: Ashcraft KW, Holcomb GW, Murphy JP, eds. Pediatric Surgery. Philadelphia, PA: Elsevier Saunders, 2005:416–34.
2. Van Beers EH, Buller HA, Grand RJ, et al. Intestinal brush border glycohydrolases: structure, function, and development. Crit Rev Biochem Mol Biol 1995; 30:197–262.
3. Cohn SM, Simon TC, Roth KA, et al. Use of transgenic mice to map cis-acting elements in the intestinal fatty acid binding protein gene (Fabpi) that control its cell lineage–specific and regional patterns of expression along the duodenal-colonic and crypt-villus axes of the gut epithelium. J Cell Biol 1992; 119:27–44.
4. Poelstra K, Bakker WW, Klok PA, et al. Dephosphorylation of endotoxin by alkaline phosphatase in vivo. Am J Pathol 1997; 151:1163–1169.
5. Van Klinken BJ, Dekker J, Buller HA, et al. Mucin gene structure and expression: protection vs. adhesion. Am J Physiol 1995; 269:G613–G627.
6. Lin J, Nadroo AM, Chen W, et al. Ontogeny and prenatal expression of trefoil factor 3/ITF in the human intestine. Early Hum Dev 2003; 71:103–109.
7. Hancock RE. Peptide antibiotics. Lancet 1997; 349:418–422.
8. Mahida YR, Rose F, Chan WC. Antimicrobial peptides in the gastrointestinal tract. Gut 1997; 40:161–163.
9. Tovar JA, Sunol M, Lopez de Torre B, et al. Mucosal morphology in experimental intestinal atresia: studies in the chick embryo. J Pediatr Surg 1991; 26:184–189.
10. Touloukian RJ. Antenatal intestinal adaptation with experimental jejunoileal atresia. J Pediatr Surg 1978; 13:468–474.
11. Trahair JF, Harding R. Ultrastructural anomalies in the fetal small intestine indicate that fetal swallowing is important for normal development: an experimental study. Virchows Arch A Pathol Anat Histopathol 1992; 420:305–312.
12. Touloukian RJ, Wright HK. Intrauterine villus hypertrophy with jejunoileal atresia. J Pediatr Surg 1973; 8:779–784.
13. Verburg M, Renes IB, Meijer HP, et al. Selective sparing of goblet cells and paneth cells in the intestine of methotrexate-treated rats. Am J Physiol Gastrointest Liver Physiol 2000; 279:G1037–G1047.
14. Yeh KY, Yeh M, Holt PR. Thyroxine and cortisone cooperate to modulate postnatal intestinal enzyme differentiation in the rat. Am J Physiol 1991; 260:G371–G378.
15. Hirayama BA, Lostao MP, Panayotova-Heiermann M, et al. Kinetic and specificity differences between rat, human, and rabbit Na+-glucose cotransporters (SGLT-1). Am J Physiol 1996; 270:G919–G926.
16. Kanda T, Fujii H, Fujita M, et al. Intestinal fatty acid binding protein is available for diagnosis of intestinal ischaemia: immunochemical analysis of two patients with ischaemic intestinal diseases. Gut 1995; 36:788–791.
17. Tytgat KM, Bovelander FJ, Opdam FJ, et al. Biosynthesis of rat MUC2 in colon and its analogy with human MUC2. Biochem J 1995; 309:221–229.
18. Podolsky DK, Lynch-Devaney K, Stow JL, et al. Identification of human intestinal trefoil factor. Goblet cell–specific expression of a peptide targeted for apical secretion. J Biol Chem 1993; 268:6694–6702.
19. Alpers DH. Digestion and absorption of carbohydrates and proteins. In: Johnson LR, ed. Physiology of the Gastrointestinal Tract. New York, NY: Raven Press, 1994:1723–49.
20. Wright EM, Hirayama BA, Loo DD, et al. Intestinal sugar transport. In: Johnson LR, ed. Physiology of the Gastrointestinal Tract. New York, NY: Raven Press, 1994.
21. Alpers DH, Bass NM, Engle MJ, et al. Intestinal fatty acid binding protein may favor differential apical fatty acid binding in the intestine. Biochim Biophys Acta 2000; 1483:352–362.
22. Tilson MD. Compensatory hypertrophy of the gut in an infant with intestinal atresia. Am J Surg 1972; 123:733–734.
23. Davidson NO, Hausman AM, Ifkovits CA, et al. Human intestinal glucose transporter expression and localization of GLUT5. Am J Physiol 1992; 262:C795–C800.
24. Montgomery RK, Mulberg AE, Grand RJ. Development of the human gastrointestinal tract: twenty years of progress. Gastroenterology 1999; 116:702–731.
25. Dahlqvist A, Lindberg T. Development of the intestinal disaccharidase and alkaline phosphatase activities in the human foetus. Clin Sci (Lond) 1966; 30:517–528.
26. Pelsers MM, Namiot Z, Kisielewski W, et al. Intestinal-type and liver-type fatty acid-binding protein in the intestine. Tissue distribution and clinical utility. Clin Biochem 2003; 36:529–535.
27. Buisine MP, Devisme L, Savidge TC, et al. Mucin gene expression in human embryonic and fetal intestine. Gut 1998; 43:519–524.
28. Klockars M, Reitamo S, Adinolfi M. Ontogeny of human lysozyme. Distribution in fetal tissues. Biol Neonate 1977; 32:243–249.
29. Surana R, Puri P. Small intestinal atresia: effect on fetal nutrition. J Pediatr Surg 1994; 29:1250–1252.
30. Trahair JF. Is fetal enteral nutrition important for normal gastrointestinal growth? a discussion. JPEN J Parenter Enteral Nutr 1993; 17:82–85.
31. Trahair JF, Rodgers HF, Cool JC, et al. Altered intestinal development after jejunal ligation in fetal sheep. Virchows Arch A Pathol Anat Histopathol 1993; 423:45–50.
32. Rubin DC, Swietlicki E, Roth KA, et al. Use of fetal intestinal isografts from normal and transgenic mice to study the programming of positional information along the duodenal-to-colonic axis. J Biol Chem 1992; 267:15122–15133.
33. Blache P, van de Wetering M, Duluc I, et al. SOX9 is an intestine crypt transcription factor, is regulated by the Wnt pathway, and represses the CDX2 and MUC2 genes. J Cell Biol 2004; 166:37–47.
34. Pinto D, Clevers H. Wnt control of stem cells and differentiation in the intestinal epithelium. Exp Cell Res 2005; 306:357–363.
35. van Es JH, Jay P, Gregorieff A, et al. Wnt signalling induces maturation of Paneth cells in intestinal crypts. Nat Cell Biol 2005; 7:381–386.
36. Trahair JF, Sangild PT. Systemic and luminal influences on the perinatal development of the gut. Equine Vet J Suppl 1997; 24:40–50.
37. Serrano J, Zetterstrom R. Disaccharidase activities and intestinal absorption in infants with congenital intestinal obstruction. J Pediatr Gastroenterol Nutr 1987; 6:238–243.
38. Goulet O, Baglin-Gobet S, Talbotec C, et al. Outcome and long-term growth after extensive small bowel resection in the neonatal period: a survey of 87 children. Eur J Pediatr Surg 2005; 15:95–101.
39. Cohen IT, Greecher CP. Nutritional status following surgical correction of congenital gastrointestinal anomalies. J Pediatr Surg 1979; 14:289–386.
Keywords:

Small intestinal atresia; Epithelium; Luminal components; Neonates; Protein expression

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