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Invited Review

Role of Intestinal Transporters in Neonatal Nutrition: Carbohydrates, Proteins, Lipids, Minerals, and Vitamins

Boudry, Gaëlle*; David, Elmer S; Douard, Véronique; Monteiro, Iona M; Le Huërou-Luron, Isabelle*; Ferraris, Ronaldo P

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Journal of Pediatric Gastroenterology and Nutrition: October 2010 - Volume 51 - Issue 4 - p 380-401
doi: 10.1097/MPG.0b013e3181eb5ad6
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After ingestion, food is mechanically and chemically digested in the stomach and small intestine and become components that can be efficiently absorbed. In the case of water-soluble nutrients that cannot diffuse across the lipid membranes, nutrient transporters must be present to absorb these nutrients from the luminal compartment across the apical membrane and into the cytoplasm of the intestinal cells. Another class of carrier proteins must be present in the basolateral membrane, not only to transport these nutrients from the cytoplasm into the blood but also to provide nutrition to intestinal cells by transport of nutrients from the blood to the cytoplasm between meals or during periods of fasting. Even some lipid-soluble nutrients that can diffuse through the plasma membrane are now known to also be transported by carriers. A large variety of these essential transporters are synthesized by the small intestine to facilitate absorption. For many nutrients at physiological concentrations, transport across the apical membrane is rate limiting. With some exceptions, paracellular transport through the tight junction between cells contributes little to the overall flux.

Infants triple their birth weight in the first year of life, and therefore consume enormous amounts of food and nutrients relative to their body weight. Infants require ∼125 kcal · kg−1 · day−1, of which ∼50, 30, and 45 are for basal metabolism, growth, and activity, respectively (1). As described in the following sections, failure of intestinal transporters to function properly often presents symptoms as “failure to thrive” because nutrients are not absorbed and as diarrhea because unabsorbed nutrients upset luminal osmolality or become substrates of intestinal bacteria. In this review, we itemize the nutrients that constitute human milk and various infant milk formulas, then briefly describe their importance in neonatal nutrition, the daily requirements (per kilogram, and related to normal adult requirement), and their pathophysiology (eg, what happens if the nutrient is absent from the neonatal diet). We then describe for each nutrient the transporter(s) that absorbs it from the intestinal lumen into the enterocyte cytosol and from the cytosol into the portal blood. In the following section, we describe transporters responsible for absorbing digested products of the macronutrients carbohydrates, protein, and lipids, as well as those for absorbing micronutrients such as minerals and vitamins. We emphasize here an extensive approach, covering the vast array of transport systems needed by neonates for absorbing a large variety of nutrients, and how those transporters may be regulated during ontogenetic development. Descriptions are brief and the characteristics of each transporter cannot be described in depth for reasons of space. Readers are referred to reviews that offer more mechanistic insights: for carbohydrates (2,3), proteins (4–6), minerals (7–10), water-soluble vitamins (11,12), and lipids (13,14).


Infants absorb glucose and galactose after lactase digestion of lactose from mother's milk, and glucose is an important ingredient of infant formulas. Galactose enters glycolysis and is used like glucose in neonatal metabolism. Because glucose can be synthesized from other sources by gluconeogenesis, there is no minimum required of glucose from dietary sources. Carbohydrate levels can fluctuate depending on the levels of other macro- and micronutrients required by the neonate, and during proximate analysis of diets, is often reported as “other” or “carbohydrate by difference” after dry matter, ash, nitrogen, and fats are analyzed. A newborn's diet consists of lactose as the primary carbohydrate, providing about 40% of energies in infants fed mature human milk or a standard “humanized” formula (1). When carbohydrate levels are unusually high, levels of proteins and lipids will be lower, increasing the risk of protein:energy malnutrition. Neonates chronically consuming high carbohydrate may be at risk for fatty livers in later years. Fructose as corn syrup is the third monosaccharide found in milk formulas. Fructose is not found in significant levels in mother's milk but is found mainly in fruits and vegetables, and is not essential. Because fructose is the sweetest of all natural sugars and is cheaper to mass produce than glucose, fructose may be substituted for some glucose in some infant formulas because a lesser amount is required to induce the same degree of sweetness.

Human milk (in mothers with full-term babies) has 37% carbohydrate (7 g/100 mL milk) in the form of lactose, and provides 52% of total energies found in milk (15). Bovine milk protein–based infant formulas contain similar levels of lactose-based carbohydrates; soy-based formulas have slightly lower levels of carbohydrate (6.7 g/100 mL), providing 40% of total energies. In both bovine milk–based and soy-based formulas, carbohydrates provide only ∼43% of total energies, mainly because of the higher contribution of fats in formula milk to total energy content of milk. The source of the carbohydrate in formula milk is lactose, sucrose, corn syrup, or combinations of these 3 sweeteners.

Transport in the Apical Membrane

Lactase in the brush border membrane liberates glucose and galactose from lactose. The Na+-dependent glucose transporter SGLT1 (gene name Slc5a1, human chromosome 22q12.3) transports glucose and galactose across the apical membrane. It is active and depends on the Na+ gradient generated by the Na+/K+-ATPase in the basolateral membrane. SGLT1 is found along the entire length of the small intestine of most mammals, but expression is greatest in the duodenum and jejunum (16). The universality of glucose as nutrient means that mutations involving SGLT1 would likely be fatal or selected against during evolution. Indeed, glucose-galactose malabsorption (GGM) characterized by defects in SGLT1 is a rare disease and there are only 200 known cases worldwide (17). The disease arises mostly from consanguineous relationships. GGM is characterized by severe diarrhea, which is fatal within a few weeks after birth unless lactose, glucose, and galactose are removed from the diet. The diarrhea stops with fasting or withdrawal from the diet of lactose, glucose, or galactose, but can return if these sugars are reintroduced in the diet (18). Fructose absorption is unaffected. In GGM, SGLT1 defects involve 22 missense, 4 splice-site, and 3 nonsense mutations so that a truncated protein is produced. Most missense mutations involve trafficking defects so that no SGLT1 makes it to the apical membrane. Only 1 mutation actually has a defect in the transport site. Hence, GGM is mainly due to mutant SGLT1 proteins that are either truncated or not targeted properly to the cell membrane (18).

GLUT5 (Slc2a5, 1p36.2), a member of the facilitative glucose transporter family, is specific for fructose transport across the apical membrane. It is found along the entire small intestine, but expression is greatest in the most proximal regions (19). Although intestinal fructose malabsorption in children has been functionally characterized as potentially a transport defect (20), it is not due to GLUT5 mutations (21). Malabsorption may, however, be due not to structural or trafficking defects in GLUT5, but to low abundance during early development and a failure to upregulate GLUT5 when fructose is present in the diet. GLUT5 seems expressed at low levels in neonates, and it is possible that low levels of this transporter will cause malabsorption. Breath hydrogen tests indicate that infants 1 year old or younger malabsorb fructose at much higher rates than 3- or 5-year-olds, and that juices containing a large amount of fructose or a high ratio of fructose to glucose can induce greater rates of malabsorption (22). Up to 70% of infants 1 year old or younger had malabsorption after consumption of 1 to 2 g fructose per kilogram body weight (23). Luminal fructose has been shown in rodent models to induce GLUT5 expression and activity during weaning. Dietary fructose fails to induce rat GLUT5 before weaning, but this developmental limitation can be overcome by dexamethasone treatment before fructose ingestion (24,25). In adult rodents, fructose readily stimulates intestinal expression and activity of GLUT5 (26). High-fructose corn syrup (HFCS), as the name implies, contains 42%, 55%, or 90% fructose, and is the main sweetener used in soft drinks, baked goods, and even some infant formulas. The popular use of HFCS has therefore led to the introduction of fructose into the diets of neonates.

Transport in the Basolateral Membrane

In classical models of sugar absorption, GLUT2 (Slc2a2, 3q26.1-q26.2) is the basolateral transporter for glucose, galactose, and fructose. Levels of GLUT2 mRNA are regulated by luminal glucose and fructose concentrations, as well as by systemic factors released during feeding (27). Partly because expression seems regulated by systemic factors, GLUT2 mRNA abundance is generally similar along the length of the small intestine (28). GLUT2 may be trafficked to the apical membrane to participate in sugar absorption after consumption of high levels of carbohydrate (29) to complement the function of SGLT1 and GLUT5. However, there is no intestinal glucose transport if SGLT1 is mutated (18), and there is no intestinal fructose transport if GLUT5 is knocked out (30), so the significance of the GLUT2 pathway is unclear. There is transepithelial glucose transport in GLUT2 knockout mice (31). GLUT2 mutations can lead to a rare genetic disease called Fanconi-Bickel syndrome, which is characterized by hepatomegaly, growth retardation, and hypoglycemia typically in young infants (32,33).


Rapid growth during the neonatal period is sustained by the highest rate of protein deposition occurring within a lifetime (0.27 g · kg−1 · day−1 for the first 2 months of life as opposed to 0.07 for 16- to 18-year-old individuals (34)). Protein needs are therefore elevated during this period of life. An adequate intake (AI) is used as the goal for intake by infants not only for protein but also for other essential macro- and micronutrients.


In full-term infants up to 6 months of age, the requirement for total protein may be estimated on the basis of the protein content of human milk (Table 1) (34–37). A minimum of 1.8 g/100 kcal (12 g/L) and a maximum protein concentration of 2.8 to 4.5 g/100 kcal have been recommended by several groups (34). Formulation of infant formula must account and compensate for differences between human milk and formula milk in digestibility, bioavailability, and efficiency of utilization of dietary proteins. A higher amount of proteins is required for hydrolysate formulas and soy protein–based formulas (2.25 g/100 for both).

Protein and amino acid requirements in human neonates

The minimum requirement of each essential and semiessential amino acid must equal or exceed the concentration of amino acids in human milk, whereas the maximum allowable level is considered to be 2 times the minimum value. Although not a standard amino acid and therefore not proteinogenic, taurine is considered semiessential in infants. However, no recommendation on taurine supplementation has been reported so far, although most infant formulas in the market contain taurine.


The major source of proteins in infant formulas is bovine milk protein, with whey-based formulas dominating the market in industrialized countries since the mid-1990s. The amino acid composition of bovine whey and casein is different from that of human milk proteins. Therefore, to compensate for this difference, the amount of protein per energy content needs to be slightly higher in formula than in human milk (generally 2.5 g/100 kcal) (36), resulting in higher levels of amino acids per energy content than human milk. The present trend is to reduce protein concentration to 1.8 g/100 kcal by using specific fractions of whey proteins to closely match human milk. Other sources of protein are used in specialized infant formulas targeting specific end users. Soy protein formulas are often recommended in cases of lactose intolerance, galactosemia, or allergy to bovine milk proteins, or used because of personal preferences. About 25% of infants in the United States used soy-based formulas in the first year of life. Free amino acid–based formulas have also been developed for infants experiencing allergy to dietary proteins (38). Other sources such as rice protein still need further investigations to confirm efficacy and safety (39). Finally, technology to synthesize recombinant human milk proteins under commercial conditions is a promising tool to further improve protein quality in infant formulas and make these as similar to human milk as possible (40).

Pathophysiology of Deficiency and Excess

Protein deficiency is often referred to as protein-energy malnutrition, the major form being kwashiorkor. Although protein and amino acid deficiency is unlikely to happen in infants consuming adequate levels of formulas, there are concerns about the acute and chronic effects of a high-protein diet on as yet immature organs involved in nitrogen metabolism. Chronically elevated levels of amino acids in the blood may surpass the capacity of hepatic and renal systems to metabolize and excrete the excess nitrogen. This may lead to acidosis, diarrhea, and excessive levels of blood ammonia and urea (41). An additional concern is that excess protein intake during infancy has also been linked to obesity later in life. High protein intake may stimulate the secretion of insulin and insulin-like growth factor-1 (IGF-1), which, in turn, can enhance growth as well as adipogenic activity and adipocyte differentiation (42).

Peptide Transporter: PEPT1

Di- and tripeptides are absorbed at the apical membrane through the peptide transporter PEPT1 (Slc15a1, 13q33-q34), and then hydrolyzed by peptidases in the cytoplasm of enterocytes. Transport systems in the basolateral membrane mediate the exit of free amino acids from the cytoplasm to the portal circulation. Quantitatively, entry of peptides by PEPT1 is the predominant mode of absorption of protein digestion products in adults and infants (43,44). Small peptides resistant to hydrolysis by cytosolic peptidases do enter the blood, but these constitute a relatively minor component (45).

PEPT1 accepts a wide variety of chemically and structurally diverse di- and tripeptides as substrates; the potential physiological substrates include 400 different dipeptides and 8000 tripeptides (43). The transport of peptides by PEPT1 is dependent on an H+ gradient generated by the Na+/H+ exchanger dependent, in turn, on the Na+ gradient (43). The H+:peptide stoichiometry is the same (1:1) for differently charged peptides. To our knowledge, no inherited disorder involving PEPT1 has been reported.

Distribution in the Intestine

PEPT1 protein is expressed exclusively in the brush border membrane (46). In adult humans, rabbits, and pigs, the duodenum exhibits the greatest expression of PEPT1, and there is a decreasing gradient of PEPT1 expression toward the colon (47–49). In rats, PEPT1 expression either does not vary with intestinal region or slowly increases toward the ileum (50–53).

Regulation by Nutrients

Dipeptides but not free amino acids regulate PEPT1 expression, suggesting that PEPT1 abundance is regulated by its substrates but not by the hydrolysis products of those substrates (54). A lack of substrate (protein-free diet) also increases peptide absorption capacity (55,56). Hence, levels of PEPT1 expression and activity are responsive to changes in quantity and composition of dietary protein (6,54,57). Transporters are typically upregulated when dietary concentrations of their substrates increase, if those substrates are used for energies, and also upregulated when dietary concentrations decrease, if those substrates are essential (58). In the case of PEPT1, its di- and tripeptide substrates fulfill both criteria. Regulation of PEPT1 by dietary substrate appears to occur by 2 mechanisms: increasing gene transcription rate and increasing mRNA stability (4).

Appearance During Development

To our knowledge, there has been no report yet of PEPT1 expression in the fetal and neonatal intestine of humans, but there have been several reports of PEPT1 expression during early ontogenetic development of various mammals. PEPT1 mRNA transcript or protein is expressed in intestines of rat embryos 17 days postconception (59). During the perinatal period, PEPT1 mRNA and protein levels usually increase during the fetal period and peak either at birth in piglet or 3 to 5 days after birth in rats and then decline in both species until weaning (6,59, Boudry and Le Huërou-Luron, personal communication). A novel feature of PEPT1 development is its presence in the colon of neonatal rats (59) and pigs (Boudry and Le Huërou-Luron, personal communication). Colonic expression may ensure maximal absorption of peptides at this early stage when protein requirements are great.

Amino Acid Transporters


The terminology of amino acid transport tends to be confusing because different amino acids can be transported by 1 or more amino acid transporters, whereas 1 amino acid transporter can absorb different amino acids, and there is substantial overlap in substrate affinities. To minimize confusion, these transport activities are referred to as “systems” rather than transporters to indicate a functionally identified transport activity that appears to be similar in various cell types (60). An amino acid transport system accepts a specific group of amino acids rather than individual amino acids. They are characterized by their type of substrate, Na+ and/or Cl2− dependency, and localization in either the brush border or basolateral membrane (Table 2) (49,51,52,60–62). In the brush border membrane, the main systems are the broadly specific systems Bo,+ for neutral amino acids, bo,+ (or y+) for cationic amino acids, and X2−AG for anionic amino acids. A deficiency in amino acid transporters is associated with several inherited disorders depending on the transport system involved (Table 3) (60).

Classification of intestinal amino acid transport system
Disorders associated with amino acid transporter deficiency

Distribution in the Intestine

Each transport system has its unique type of distribution in the gut (Table 2). However, expression of most of the apical amino acid transport systems generally increases toward the ileum. This inverse distribution compared with PEPT1 probably reflects the gradual aboral increase of luminal-free amino acid concentrations at the same time as the decrease of peptide concentrations as peptide hydrolysis continues. Interestingly, systems ASC and Bo,+ are highly expressed in the human colon (49). The significance of the colonic presence of these transporters in terms of nutrition, that is, bioavailability of the amino acids transported by these systems, needs to be elucidated. In contrast to the ileum-oriented distribution of apical amino acid transport systems, the intestinal distribution of basolateral amino acid transport systems, which control the flux of amino acids toward and from the blood, is more variable and less known.

Regulation by Nutrients

Intestinal amino acid transporters are regulated by changes in levels of dietary protein or free amino acid mixtures (63,64), following the principles outlined by Ferraris and Diamond (58). Transport systems that take up solely nonessential amino acids typically are regulated monotonically by dietary levels of their substrates, and increase in activity when dietary protein levels increase. Regulation of systems that transport essential or relatively toxic amino acids is more complex; amino acids can induce their own transporters but also modulate the other transporters (65). A high-protein or a protein-deficient diet can induce a modest increase in uptake of essential amino acids (64). A recent study demonstrated that amino acid absorption at 1 intestinal site can be regulated by the intraluminal concentration of that amino acid at a more proximal or more distal intestinal site, through neural mechanisms (66). Amino acid uptake may also be indirectly modulated by PEPT1 activity. Because many amino acid transport systems function as exchangers, the entry of amino acids in the form of peptides through PEPT1 may be important for the net movement of amino acids (67), and this exchange pathway may allow absorption of certain amino acids that modulate activity of transporters for which they are not substrates.

Appearance During Development

The human small intestine seems capable of amino acid transport early during ontogenetic development. Changes in electrical activity (transepithelial potential difference) associated with alanine transport have been reported in the intestine of 14- to 16-week-old fetuses (68). Transport typical of systems Bo, X2−AG, bo,+, y+L, and IMINO was detected in intestinal brush border membrane vesicles of 17- to 20-week-old fetuses (69,70). Transport rates decreased along the proximodistal gradient of the fetal intestine, unlike those of adults that were greatest in the ileum (69). In rats, amino acid transport characteristic of systems XAG and A are described in E17 embryos; intestinal mRNA transcripts for system A appear by E14, a full week before parturition (71,72). In piglets, transport of leucine initially detected in 7-week-old fetuses increased gradually in the last 2 weeks of gestation, along with the appearance of the proximodistal gradient of transport typical of adults (73).

Amino acid uptake normalized to intestinal weight tends to decrease with postnatal age. In piglets, a sharp decrease in uptake rates of various amino acids occurs in the first 24 hours after birth. Uptake rates then return to birth values at day 7 of life and decline further until weaning (73,74). Nevertheless, the total capacity of the piglet intestine to absorb amino acids increased with age due to increased length and mass of the intestine (73). In rats and mice, age-related differences are also observed with a decrease in transport system expression and/or specific activity from birth or 1 or 2 days after birth until weaning (72,75–81). Similarly, due to the massive growth of the intestine during this suckling period, total uptake capacity increases with age (79). Expression of apical amino acid transport systems increases with age in neonatal chickens, whereas that of basolateral transporters tends to decrease perinatally (82). This difference in evolution of amino acid transporters between the apical and basolateral sides of the enterocyte probably reflects the switch from parenteral to enteral nutrition that occurs at birth in altricial species.

In summary, expression of intestinal peptide and amino acid transport systems during the perinatal period seems designed to ensure maximum protein absorption. Before birth, the intestine seems to prepare for this task by increasing peptide and amino acid transport capacity. After birth until weaning, the decline in most of the peptides and amino acid transport capacity (when expressed per kilogram of body weight) probably reflects the decrease in protein requirements (Fig. 1) (83).

Protein requirements (dotted line) and peptide and amino acid transport capacity of the entire intestine expressed per kilogram of body weight (solid line) with age in the neonatal period. Protein requirement per kilogram of body weight gradually decreases with age in the neonate until weaning. In parallel, and despite the increase of total capacity of the intestine to transport peptides and amino acids, it decreases when expressed by kilogram of body weight. Peptide and amino acid transport capacity always exceeds the requirement with a safety margin, so the organism is certain that intestinal absorptive capacity does not limit growth (256).


Various minerals are present in breast milk and are added to infant formula. In this section we review the absorption of the 3 major minerals, Ca2+, Pi, and Mg2+, which make up 98% of the body's mineral content and are essential for tissue and bone formation. We also review the absorption of 2 major trace minerals, iron and zinc. All of these minerals are absorbed both actively and passively, and, in this review, we focus on the active transport and the transporters involved. The function, recommended AI, and diseases associated with excess or deficient intake of minerals are summarized in Table 4(84).

Function, dietary intake, and clinical states of deficiency and excess of the minerals


Active transcellular transport of Ca2+ occurs mainly in the duodenum and consists of passive uptake across the apical membrane, cytoplasmic transport, and ATP-dependent extrusion across the basolateral membrane.

Apical Transport

Ca2+ transport across the apical membrane occurs through a channel-like Ca2+ transporter (CaT1, Sla7a1) that mediates intestinal Ca2+ absorption (85). CaT1 is also called TRPV6 because it belongs to the transient receptor potential vanilloid (TRPV) family of ion channels (86). In humans, CaT1 mRNA is expressed throughout the gastrointestinal (GI) tract, including the esophagus, stomach, duodenum, jejunum, ileum, and colon, and in other organs (87). The CaT1 gene was assigned to the long arm of the human chromosome 7q33-34. In mice, intestinal CaT1 mRNA levels increase 30-fold at weaning, coincident with the induction of calbindin-D9k expression, and both are strongly regulated by dietary Ca2+ intake and 1,25-dihydroxyvitamin D3(88,89). However, significant active intestinal Ca2+ transport occurs in TRPV6-null mice, thus challenging the dogma that TRPV6 is essential for vitamin D–induced active intestinal Ca2+ transport (90). Estrogens have a distinct, vitamin D–independent effect at the genomic level on upregulation of CaT1 (91). Prolactin directly stimulates active and passive Ca2+ uptake in rat duodenum (92). The synthetic glucocorticoid prednisolone reduces the intestinal Ca2+ absorption capacity through diminished duodenal expression of CaT1 independent of systemic vitamin D, particularly calcitriol (93). This may be relevant to the premature neonate who receives steroids for lung maturation or bronchopulmonary dysplasia.

Cytoplasmic and Basolateral Transport

In the cytoplasm, dietary Ca2+ binds to calbindin-D9k (94) and then traverses the cytoplasm. Ca2+ can also be transported through the cytoplasm via vesicles or endosome-mediated tunnelling transport via membrane-bound organelles (95). Ca2+ transport in the small intestine of the calbindin-D9k knockout mouse, however, is normal (96). Calbindin-D9k decreases with age and increases with increasing concentrations of vitamin D and Pi (97,98). Ca2+ is extruded through the basolateral membrane via the plasma membrane Ca2+-ATPase 1b (PMCA 1b, Atp2b1, 12q21.3). A smaller amount is transported by the Na/Ca exchanger1 (NCX1, Slc8a1, 2p23-22) transporter (94).


Phosphorus is found as inorganic phosphate (Pi) in the body and is involved in the maintenance of pH, storage and transfer of energy, synthesis of nucleotides, and growth. It is found mostly in bones and teeth.

The predominant mode of Pi uptake is transcellular via the apical Na+-dependent phosphate transporter NaPi2b (Slc34a2, 4p15.3-1) (99). The gene encoding this cDNA was mapped to human chromosome 4p15.1-p15.3 (100). NaPi2b mRNA and protein expression in neonatal goats increases up to 8 to 11 weeks of age, and in the ruminating goat, at 5 months of life, the protein expression is comparable to 8 to 11 weeks of age, but the mRNA decreases (101). Pi uptake, NaPi2b mRNA, and protein expression were markedly higher in neonatal compared with weaning and adult rats (102), as would be expected because the demand for phosphate is greatest in rapidly growing young mammals. In fact, transepithelial Pi uptake in adult rats is mostly Na+ independent and may be paracellular, suggesting that NaPi2b is no longer involved (103,104). The NaPi2b cotransporter is highly abundant in mouse ileum and is almost absent in the duodenum and jejunum (105). In the rat, NaPi2b is mostly in the duodenum and jejunum, with negligible expression in the ileum (106). Kirchner et al (102) demonstrated a higher concentration in the middle small intestine as compared with the distal small intestine in neonatal rats. High levels of dietary fructose may inhibit expression of rat NaPi2b, particularly in neonates when expression is the greatest (102). A low-phosphate diet increases transport activity as well as protein and mRNA expression of NaPi2b (107,108). The cytosolic and basolateral transport mechanisms for Pi have yet to be elucidated.


Mg2+ is a cofactor for numerous enzymes and plays a role in contraction and relaxation of muscles, production and transport of energy, and production of proteins. The majority of Mg2+ in the body resides in bone (109). Intestinal absorption of Mg2+ occurs by both a passive paracellular and an active transcellular process (110). The mode of Mg2+ transport changes at the time of weaning from mainly a passive mechanism in the suckling period to a carrier-mediated mechanism in adolescent rats (111). Mg2+ absorption is several-fold higher in the suckling than in the weaning rats. The synthetic glucocorticoid methylprednisolone decreases Mg2+ absorption in the suckling but not in the weaning rats (112). Transport of Mg2+ across the apical brush border is via TRPM6, which is a member of the transient receptor potential melastatin channel (TRPM) family and is expressed in intestinal epithelia and kidney tubules (113). The TRPM6 gene is mapped to chromosome 9q22. Mutations in this gene have been identified in autosomal-recessive hypomagnesemia with secondary hypocalcemia (114). Patients present with convulsions in early infancy and respond well to intravenous administration of Mg2+, followed by lifelong supplementation of dietary Mg2+(115). Mg2+-enriched diets increase TRPM6 mRNA levels in the colon of mice (113). How the Mg2+ is extruded through the basolateral membrane remains unknown at this time. There is a specific Mg2+ transporter recently discovered in the kidney, but an intestinal homologue has not yet been discovered (10).


Iron (Fe) is a component of hemoglobin and numerous enzymes. It is added in infant formulas as ferrous (Fe2+) sulfate. The transport of dietary Fe2+ from the intestinal lumen across the epithelial apical membrane is by the divalent metal transporter 1 (DMT1 [also known as DCT1]). In the cytosol, dietary iron is bound to iron-binding proteins and exits the cell via the basolateral transporter called ferroportin (FPN1, Slc40a1, 2q32) (116,117). FPN1 is thought to associate with hephaestin, a multicopper ferroxidase protein required for the export of iron across the basolateral membrane. Mice without a functional hephaestin absorb iron normally but are unable to export it out of the intestinal cells so that the iron trapped in the cytosol is lost during intestinal cell turnover (118).

DMT1 (Slc11a2, 12q13) is highly expressed in the duodenum and has an unusually broad substrate range that includes Fe2+, Zn2+, Mn2+, Co2+, Cd2+, Cu2+, Ni2+, and Pb2+(117). Fe2+ absorption is tightly regulated by dietary intake and stage of ontogenetic development. Intestinal iron absorption rates are high in the perinatal and weaning periods and then decrease to rates exhibited by adults. Expression of the iron transporters (DMT1 and FPN1) in the duodenum of rats is developmentally regulated and markedly increases postnatally, until sexual maturity (119). The genes encoding DMT1 and FPN1 are also high in the distal intestine in the neonate and decrease to adult levels at the time of weaning, accounting for the high iron absorption in the neonate (120). However, iron absorption in the colon of neonatal rat pups is significantly higher than at the time of weaning.

Regulation of rates of iron absorption is complicated. Hepcidin is a peptide hormone whose secretion by the liver varies with iron deficiency or overload. Under conditions of overload, increased hepcidin secretion into the blood binds to ferroportin causing it to internalize and degrade, thus preventing export of iron out of the cell (121). This internalization and degradation of the ferroportin involves activation of Janus kinase 2 (Jak 2) (122). Hepcidin levels are low in patients with hemochromatosis, an iron overload disorder (123). The important ability to regulate iron absorption depends on the developmental stage. Human infants cannot regulate iron absorption at 6 months of age, but older infants at 9 months of age can significantly increase iron absorption when dietary iron intake is low (124). Likewise, neonatal rat pups are unable to downregulate DMT1 and FPN1 expression in response to iron supplementation until the time of weaning (119). FPN1 expression increases in iron-deficient states (116).


Zn2+ is a component of multiple enzymes and proteins. There are 2 families of zinc transporters encoded by the solute-linked carrier (SLC) gene families, Zip and ZnT. There are multiple transporters in each of these families. Zinc absorption occurs mainly in the small intestine. Zip transporters increase cytoplasmic zinc by promoting extracellular and, perhaps, vesicular zinc transport into cytoplasm (125). In mice, Zip4 expression is greatest in the small intestine, less in the stomach and liver, and absent in the kidneys. Intestinal Zip4 but not Zip1-3 mRNA abundance is regulated by dietary zinc (126). ZIP1 mRNA increases modestly after 16 days of gestation and then remains unchanged in the neonatal period; however, ZIP4 mRNA increases after 18 days of gestation, decreases within 5 days postpartum, and rebounds by 15 days of life (127). Under conditions of dietary deficiency, ZIP4 protein is increased on the apical membranes of enterocytes from the neonatal mice (127). The apical localization of ZIP4 at birth suggests that this protein plays a critical role in absorption of zinc from the milk in the neonatal period. An autosomal-recessive disorder, acrodermatitis enteropathica, is secondary to point mutations in the Zip4 gene, which has been mapped to human chromosome 8q24 (128). ZnT transporters are likely involved in basolateral transport. ZnT1 was the first zinc transporter identified and is mapped to chromosome 1 in humans. ZnT1, ZnT2, and ZnT4 transporters are highly expressed in the neonatal small intestine, principally near the apical surface, and then ZnT1 and 4 increase in abundance at the basolateral surface during development (129). ZnT4 is localized in the membrane of intracellular vesicles, the majority of which concentrate in the basal cytoplasmic region of mouse enterocytes (130). Zinc may also be transported as a [Gly-Gly-His-Zn] complex, using the peptide carrier system (131), most likely PEPT1 already present in the neonatal small intestine as discussed above. Metallothionein proteins act as a regulatory reservoir, binding excess zinc under conditions of toxicity and increasing availability of zinc under conditions of deficiency (132).


Water-soluble vitamins are essential for development and must be provided in the neonatal diet, with the possible exception of niacin, a vitamin, which can be synthesized endogenously from tryptophan. The amounts of vitamins in infant formulas based on bovine milk normally meet the newborn's needs. In contrast, the concentration of water-soluble vitamins in breast milk is highly dependent on maternal status and dietary intake. Generally, after birth, the blood concentration of most of these vitamins is relatively greater in the neonate compared with that of the mother's (133). Subsequently, inadequate intake by the neonate of these essential micronutrients due to inadequate dietary sources or, more important, the inability of the neonatal gut to absorb these compounds will lead to deficiency states. The recommended daily intake (RDI) of these micronutrients increases proportionately as the neonate grows and matures. Hence, the intestine is critical in maintaining and regulating their homeostases in the body, and it is imperative that the ability of the neonatal gut to assimilate these nutrients from the diet be optimal.

Requirements and Disease States

The level considered as AI reflects the average intake of vitamin B for infants consuming human milk produced by well-nourished breast-feeding mothers (Table 5) (134). Generally, AI values for each vitamin B meet infant needs. There are no reports of full-term infants exclusively fed human milk from healthy mothers and later manifested signs of vitamin B deficiency. Because breast milk from mothers who are vegetarian or who have untreated pernicious anemia has lower amounts of vitamin B12, their infants may begin to show clinical signs of B12 deficiency at about 4 months of age (135).

Water-soluble vitamins: function, adequate intake, and clinical states of deficiency and excess

In the following section, the mechanisms as well as regulation of intestinal absorption for each water-soluble vitamin are discussed briefly. Because most of the B vitamins are bound to proteins and other compounds in the diet, each vitamin needs to be initially liberated from its bound form and then absorbed by its specific transport system from the intestinal lumen. With the exception of cobalamin, whose intestinal absorption is via a receptor-mediated mechanism, the rest of the water-soluble vitamins are transported by carrier-mediated systems (11). For almost all water-soluble vitamins, there are 2 significant sources: diet and colonic bacteria (136,137).

Thiamine (Vitamin B1)

Intestinal absorption of dietary thiamine takes place mainly in the jejunum by both active (at lower concentrations) and passive (at higher concentrations) mechanisms. For a review of thiamine transport, refer to Rindi and Laforenza's article (138). Dietary thiamine is first hydrolyzed into its phosphorylated form, coenzyme thiamine pyrophosphate, in the intestinal lumen and enters the mucosal cell via an Na+-independent, H+-dependent, carrier-mediated mechanism involving the thiamine transporters THTR-1 and THTR-2, which are products of Slc19a2 (1q23.3) and Slc19a3 (2q37), respectively (139–143). In humans, both transporters are expressed along the length of the intestinal tract (144,145), but THTR-1 is expressed in both the apical and basolateral membranes whereas THTR-2 is expressed only in the apical (145,146). Intestinal thiamine uptake decreases with maturity. The mechanism underlying colonic absorption of bacterial thiamine is similar to the carrier-mediated process of thiamine uptake in the small intestine (136).

Riboflavin (Vitamin B2)

Most dietary riboflavin is consumed as a complex of food protein with flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD) (147). Gastric acidification in the stomach releases most of the coenzyme forms of riboflavin (FAD and FMN) from the protein. The noncovalently bound coenzymes are then hydrolyzed to riboflavin by nonspecific pyrophosphatases and phosphatases in the upper gut (147,148). Dietary riboflavin absorption occurs in the proximal small intestine via a rapid, saturable, Na+-independent, carrier-mediated transport system (147–149), which predominates at lower dietary concentrations. At higher concentrations, the rate of absorption in adult humans is proportional to the amount of intake, and absorption increases when riboflavin is ingested along with other foods (150) and in the presence of bile salts (150,151). A small amount of riboflavin circulates via the enterohepatic system (148). There is an adaptive regulation induced by extracellular riboflavin levels wherein a deficiency or an oversupply of riboflavin leads to an up- or downregulation, respectively, in intestinal riboflavin uptake (149,152). Riboflavin may exit the intestinal cell by another carrier located in the basolateral membrane (152). A small amount of colonic riboflavin absorption occurs via a carrier-mediated, Na+-independent transport system similar to that in the small intestine (153,154). The molecular identities of the apical and basolateral riboflavin transporters are still not known. Moreover, developmental and dietary regulation of riboflavin absorption in infants and children have not been studied.

Niacin (Vitamin B3)

Niacin requirement is met not only by dietary nicotinic acid and nicotinamide but also by the metabolic conversion of the amino acid tryptophan to niacin. The relative contribution of tryptophan is estimated to be 60 mg of tryptophan to 1 mg of niacin or 1 mg of niacin equivalents (155). Dietary niacin is absorbed via an Na+-independent, H+-dependent, carrier-mediated mechanism (156). Intestinal absorption is rapid (157). At higher concentrations, passive diffusion predominates. Although the molecular identity and regulation of this transport system is still not known, some evidence suggests that the transport system may be under the regulation of an intracellular protein tyrosine kinase–mediated pathway (156).

Pantothenic Acid (Vitamin B5)

Most tissues transport pantothenic acid into cells for the synthesis of coenzyme A (CoA). CoA in the diet is hydrolyzed in the intestinal lumen to dephospho-CoA, phosphopantetheine, and pantetheine, with the pantetheine subsequently hydrolyzed to pantothenic acid (158). Active transport occurs at low vitamin concentrations and by passive transport at higher concentrations (159). Active transport is via the SMVT (Slc5a6, 2p23) transport system, which is a carrier-mediated system shared with another vitamin, biotin (160). It is unclear how this shared transport system is regulated by the substrate levels of these micronutrients. Intestinal microflora have been observed to synthesize pantothenic acid in mice (161), but the physiological contribution of bacterial synthesis to systemic pantothenic acid levels or fecal losses in humans has not been quantified. Colonic absorption of bacterially produced pantothenic acid is via the same system found in the small intestine (162).

Pyridoxine (Vitamin B6)

Vitamin B6 comprises a group of 6 related compounds: pyridoxal (PL), pyridoxine (PN), pyridoxamine (PM), and their respective 5′-phosphates (PLP, PNP, and PMP). Because of PLP's role as a coenzyme for many enzymes involved in amino acid metabolism, vitamin B6 requirements are influenced by protein intake. In fact, increased protein intake causes a relative decrease in vitamin B6 status (163–165).

Pyridoxine absorption involves phosphatase-mediated hydrolysis followed by transport of the nonphosphorylated form into the mucosal cell. PN glucoside is absorbed less effectively than are PLP and PMP and, in humans, is deconjugated by a mucosal glucosidase (166). Intestinal pyridoxine absorption is via an Na+-independent, H+-dependent, carrier-mediated mechanism and that appears to be under the regulation of an intracellular protein kinase A (PKA)–mediated pathway (167). Colonocyte pyridoxine transport is via the same specific and regulatable carrier-mediated process as in the small intestine (168). To date, the molecular identity of the intestinal vitamin B6 uptake system and its gene has not been elucidated.

Drugs that can react with carbonyl groups have the potential to interact with PLP. Isoniazid, which is used in the treatment of tuberculosis, and L-DOPA, which is metabolized to dopamine and used in the treatment of Parkinson disease and dopamine-responsive dystonia, have been reported to reduce plasma PLP concentrations (169). Thus, vitamin B6 supplementation is routinely recommended for infants receiving isoniazid and infants breast-fed by mothers receiving isoniazid or L-DOPA.

Cyanocobalamin (Vitamin B12)

Small amounts of B12 (cobalamin) are absorbed via a complicated, active process that requires an intact stomach, intrinsic factor (a glycoprotein that the parietal cells of the stomach secrete after being stimulated by food), pancreatic sufficiency, and a normally functioning terminal ileum. In the stomach, dietary B12 is dissociated from proteins in the presence of acid and pepsin (Fig. 2). The dissociated B12 then binds to R proteins (haptocorrins or transcobalamin I) secreted by the salivary glands and the gastric mucosa. In the small intestine, pancreatic proteases partially degrade the R proteins, releasing B12 to bind with intrinsic factor. The resulting complex of intrinsic factor and B12 attaches to a specific receptor, cubilin, in the ileal mucosa; after internalization of the complex, B12 enters the enterocytes (133). This complex binds with the intestinal brush border membrane receptor cubilin which is endocytosed, processed via the endosomal-lysosomal pathway, then finds its way into the circulation via the basolateral membrane of the absorptive cells (170–172). If there is a lack of intrinsic factor (as is the case of pernicious anemia), then malabsorption of B12 occurs. If untreated, then potentially irreversible neurological damage and life-threatening anemia develop. B12 is continually secreted in the bile. In healthy individuals most of this B12 is reabsorbed and available for metabolic functions. The protein and the gene encoding the receptor for the cellular uptake of transcobalamin-bound cobalamin have recently been identified (171). Developmental regulation is not known.

Vitamin B12 processing in the body. B12 = free vitamin B12; TC I = transcobalamin I (also known as R-protein, cobalophilin, haptocorrin); IF = intrinsic factor; B12-IF = vitamin B12-intrinsic factor complex; TC II = transcobalamin II (carries vitamin B12 in blood circulation); TC III = transcobalamin III (vitamin B12 storage complex).

Biotin (Vitamin H)

A biotin carrier located in the intestinal brush border membrane transports biotin against an Na+ concentration gradient. The carrier is structurally specific, temperature dependent, and electroneutral. At pharmacological concentrations of dietary biotin, diffusion predominates (173). Human intestinal biotin uptake is via the SMVT (Slc5a6, 2p23) transport system that is also shared by pantothenic acid (162,174). Human intestinal biotin uptake is adaptively upregulated during biotin deficiency, which occurs via an increase in SMVT protein and mRNA levels (175). Regulation is mediated by transcriptional mechanisms involving binding sites for the transcriptional factor gut-protein Kruppel-like factor (GKLF) in the human SMVT promoter (176). Colonic biotin uptake occurs via the same Na+-dependent, carrier-mediated mechanism that operates in the small intestine (162).

Folic Acid (Folate)

Folic acid (pteroylmonoglutamic acid), which is the most oxidized and stable form of folate, is the form used in vitamin supplements and in fortified food products. Folate deficiency has been identified in small-for-gestational-age infants. Use of drugs, such as phenobarbital, phenytoin, and sulfasalazine, may increase the need for folate (133). The premature infant is at particular risk for folate deficiency because of insufficient hepatic stores, rapid growth, increased erythropoiesis, use of antibiotics and anticonvulsants, and inherent fat malabsorption states (133). Iron deficiency may lead to a decrease in folate utilization. Folic acid therapy may lead to zinc deficiency (177).

Intestinal folate transport is carried out by 2 transport systems: reduced folate carrier (RFC, Slc19a1, 21q22.3) and proton-coupled folate transporter (PCFT, Slc46a1, 17q11.2). Dietary folates (polyglutamate derivatives) are hydrolyzed to monoglutamate forms in the gut before absorption across the intestinal mucosa. This cleavage is accomplished by a γ-glutamylhydrolase, more commonly called folate conjugase. The monoglutamate form of folate is actively transported across the brush border membrane of proximal small intestinal cells by both RFC and PCFT, which are both saturable H+-dependent processes (178,179). Although PCFT transports folic acid at more acidic pH compared with RFC, both transporters have similar affinities for reduced and oxidized folates (179,180). When pharmacological doses of the monoglutamate form of folate are consumed, some are also absorbed by a nonsaturable mechanism involving passive diffusion (133). A product of Slc19a1, RFC, also known as RFT (reduced folate transporter) is responsible for intestinal folate transport (141) at the basolateral membrane domains of polarized enterocytes. Dietary deficiency of folate leads to an upregulation in its intestinal uptake. Intestinal folate transport process is ontogenetically regulated and decreases with age (181). Colonic folate absorption is similar to the carrier-mediated process in the small intestine (182). Coexisting iron or vitamin B12 deficiency may interfere with the diagnosis of folate deficiency (183).

Vitamin C (Ascorbic Acid)

Vitamin C acts as a cofactor in a number of metabolic reactions and as a free radical scavenger (184). It exists in reduced (ascorbic acid) or oxidized (dehydro-L-ascorbic acid [DHAA]) forms. Intestinal transport of vitamin C by passive diffusion is negligible. Other known transport mechanisms are facilitated diffusion or secondary active transport (184). There are 2 human isoforms of ascorbic acid transporters: Na+-dependent vitamin C transporters 1 and 2 (SVCT1 and SVCT2), which share the same homology with one another (185). In the small intestine, SVCT1 (5q31.2–31.3), the product of the Slc23a1 gene, is expressed at the apical membrane (185), whereas the expression of SVCT2 (Slc23a2, 20p13) is at the basolateral membrane of the enterocytes (186). Absorption sites occur throughout the entire length of the small intestine. Intestinal ascorbic acid transport is regulated by extracellular substrate levels and by an intracellular PKC-mediated pathway (187).


The lipid content in human milk is ∼38 g/L and represents 45% to 55% of the newborn infant's energy requirements. Lipid requirements decrease to 30% for infants 6 to 12 months of age (Table 6) (188–190). Lipids in breast milk consist mostly of triacylglycerols (∼99%) with some contributions from cholesterol esters (<1%, 10–15 mg/dL) and phospholipids (<1%, 15–20 mg/dL). A small proportion (<0.1%) is found as diacylglycerols and free fatty acids. The lipid fraction of human milk provides not only energy but also cholesterol and essential fatty acids or long-chain polyunsaturated fatty acids (LC-PUFA), which are the precursors of eicosanoids, endocannabinoids, other fat-soluble hormones, as well as liposoluble vitamins. Hence, the amount of triacylglycerols and the composition of their fatty acids have health relevance for the neonate.

Neonatal requirements for lipids and essential unsaturated fatty acids

Source of Energy and of Essential Lipids

Because the triacylglycerols provide ∼9 kcal/g, they are the best dietary source of energy. During the first 6 months of life, the energy required for growth alone represents 20% to 30% of the total energy needs, whereas at 12 months of age, the total energy cost for growth falls to 5%—under both conditions, the energy released from lipid metabolism alone would be sufficient to meet these needs (188) (Table 6).

Linoleic (LA, C18:2n-6) and α-linolenic (LNA, C18:3n-3) acids are the essential fatty acid precursors of LC-PUFA of n-6 and n-3 families. The major metabolite arising from successive elongation and desaturation of LA is arachidonic acid (ARA, C20:4 n-6) and those of LNA are eicosapentaenic (EPA, C20:5n-3) and docosahexaenoic (DHA, C22:6 n-3) acids. The n-6 and n-3 LC-PUFA play a major role in the early development of the skin, brain, and retina. In neonates, there is a rapid accretion of ARA in the whole body as well as of ARA and DHA in various organs such as the brain and retina. This indicates that specific PUFAs play important unique roles in the development of certain organ systems. Even if the preterm and full-term neonates are capable of de novo synthesis of LC-PUFA (191), most of the required LC-PUFA would have to be present in breast milk to ensure that sufficient levels of these nutrients would be available to the infant (Tables 6 and 7). It is not clear whether preterm infants have lipid and LC-PUFA requirements different from full-term babies (189,192).

Lipid composition of human term milk* and infant term formula

The n-6 and n-3 LC-PUFA also contribute to synthesis of plasma membranes and to immune, visual, cognitive, and motor functions (193–198). High levels of LC-PUFA in early life may be beneficial as they diminish the incidence of insulin resistance, obesity, and cardiovascular problems in later life (199). However, LC-PUFA like ARA may also be detrimental because they may induce precocious development of adiposity (200,201).

Although the amounts of cholesterol are especially high in breast milk (6 to 18 mg/100 mL) (202), its role in early tissue development or later adult cholesterol metabolism remains unclear. On the one hand, cholesterol is the major lipid component of some nervous cell membranes such as myelin, and an increase in neonatal plasma cholesterol concentrations is positively associated with enhancement of cerebrum weight gain and development of normal behavior and reflexes (203,204). On the other hand, adult rat offsprings display a negative correlation between serum cholesterol concentrations and the cholesterol content of their mothers' milk, suggesting the potential “protective” role of early dietary cholesterol exposure to hypercholesterolemia in adulthood (205). However, other studies do not support the hypothesis (206); thus, the influence of infant high cholesterol intake on cholesterol metabolism in adulthood remains unclear.

Lipid Composition of Breast and Formula Milk

The lipid content and composition of breast milk vary depending on stage of lactation (early, mid, or late lactation), time of feeding (foremilk vs hindmilk), time of day (morning vs evening), and mother's diet (207,208), making it difficult to simulate a complete lipid profile for use in infant formulas. Despite efforts to replicate human milk, numerous qualitative and quantitative differences persist between breast and formula milk (Table 7). Currently, vegetable oils are added to infant formula to improve lipid absorption, to increase the level of essential unsaturated fatty acids (LA and LNA), and to decrease the LA/LNA ratio (188). Whereas LA is abundant in most of the vegetable oils, LNA and other n-3 PUFA are only found in rapeseed, canola, and soy oil (188). In addition, the positions of fatty acids esterified to the glycerol backbone in various oils used to supplement the infant formulae may differ from those found in human milk. As discussed later below, these fatty acid positions are relevant because the vulnerability of esterified fatty acid to lipase digestion is dependent in part on its position in the glycerol backbone. Indeed, fatty acids in triglycerides from breast milk are mostly long-chain fatty acids (LC-FA), for example palmitic (C16:0, ∼20%–25% of triglyceride fatty acids) and stearic acids (C18:0, ∼10%) for the saturated fatty acids, and oleic acids (C18:1, ∼30%–35%) for the monounsaturated ones (209). Palmitic acid constitutes the highest proportion (53% to 70%) of saturated fatty acids at the sn-2 position of the triacylglycerol backbone (210), whereas oleic acids are mainly localized on the sn-3 and sn-1 positions. The location of palmitic acid at the sn-2 position is critical because that increases its absorption in the lumen of infants (211). Consequently, to establish in infant formula an optimal amount of palmitic acid esterified at the sn-2 position, triacylglycerols must be modified by enzymatic interesterification of tripalmitin with vegetable oil mixes and fish oil to change the positional distribution of fatty acids in the glycerol backbone. Fish, algal, and fungal oils as well as lipids from eggs are mainly used for supplementation of infant milk formulae to balance the n-3 and n-6 LC-PUFAs.

Processing of Dietary Lipids in the Gastrointestinal Tract


The process of digestion begins in the stomach where ∼15% of the triacylglycerols are released by lingual and gastric lipases, but most of the hydrolysis occurs in the duodenum by pancreatic lipases. Fatty acids in sn-1 and sn-3 positions are mostly hydrolyzed by pancreatic lipases and later on become incorporated in micelles. The fatty acids in the sn-2 position remain as monoglycerides and are mainly absorbed because these monoglycerides are polar and later easily solubilized. Pancreatic lipases are secreted by the pancreas from approximately 30 weeks of gestation but remain at low concentrations until the first year of life (212). Because lipid digestion is critical for neonatal development, it is hypothesized that other lipases such as pancreatic lipase–related protein 2 (213,214) or carboxyl ester lipase from breast milk (215) may compensate for this deficiency in levels of pancreatic lipases. Increases in luminal concentration of triacyglycerols in the duodenum stimulates the release of bile acids, whose detergent properties solubilize the products of lipid hydrolysis (sn-2-monoacylglcyerol and free fatty acids) and form mixed micelles. Micelles whose surfaces are hydrophilic diffuse through the aqueous luminal contents and eventually come into close proximity to the brush border of enterocytes and would release free fatty acids and acylglycerols, which subsequently either are taken by specific carrier molecules or diffuse into the mucosal cell.


Intestinal lipid absorption is a multistep process, traditionally divided into 3 components: apical absorption into the enterocyte, intracellular processing, and subsequent release into the lymphatic and portal circulation (216).

Apical Transport

Although the short-chain fatty acids diffuse passively across the enterocyte membrane, the relative importance of simple diffusion as opposed to carrier-mediated absorption in the apical absorption of LC-FA and cholesterol remains unclear. Because of their lipophilic nature, LC-FA were once thought to diffuse freely through the plasma membrane (217,218). Several studies in the last 10 years have shown evidence for carrier-mediated lipid absorption (Fig. 3). These transporters for lipids have, however, been studied mainly in adults, and little is known about their relative contribution to intestinal lipid absorption in the neonate. Several transporters for LC-FA have initially been discovered in other tissues, and 3 of these were later found expressed in the small intestine: the plasma membrane fatty acid–binding protein (FABPpm, Got2, 16q21) involved mainly in the intestinal uptake of LC-PUFA (219–222), the fatty acid transport protein 4 (FATP4, Slc27a4, 9q34.11) (223), and the rat homologue of human fatty acid translocase (FAT/CD36, Cd36, 7q11.2). FAT/CD36 is not a specific transporter of fatty acid and is also involved in the apical uptake of cholesterol (224). Intestinal cholesterol absorption was also reported to be facilitated by the scavenger receptor SR-BI (Scarb1, 12q24.31–32) (225–227) and by Niemann-Pick C-1-like 1 (NPC1L1, Npc1l1, 7p13) (228,229). Lipid transport may be bidirectional, and some lipids may be secreted by intestinal cells. For example, the ABCG5–ABCG8 (Abcg5-Abcg8, 2p21) heterodimeric transporter localized on the apical side is thought to transport intracellular cholesterol back across the apical membrane to the intestinal lumen (230).

Mechanisms of intestinal apical, intracellular, and basolateral transport of medium- and long-chain polyunsaturated fatty acids (MC/LC-FA). LC-FA cross the apical membrane of enterocytes by a number of facilitated transporters including the plasma membrane fatty acid–binding protein (FABPpm), fatty acid transport protein 4 (FATP4), fatty acid translocase (FAT/CD36), and scavenger receptor SR-BI. FAT/CD36 and SR-BI are less specific and also able to mediate the transport of cholesterol (CHO). The apical transporter Niemann-Pick C-1-like 1 (NPC1L1) is more specialized for CHO transport. The ATP-binding cassette transporters G5 and G8 (ABCG5/8) mediate an efflux of CHO to the lumen. The cytoplasmic transport of LC-PUFA involves intestinal and liver fatty acid–binding protain (L- and I-FABP). LC-FA are brought to the endoplasmic reticulum (ER) where metabolism of fatty acids requires monoacylglycerotransferase 1 (MGAT1) as well as diacyglyceroltransferase 1 and 2 (DGAT1 and 2) for the synthesis of the triglycerides (TG). Metabolism of CHO in the ER requires the acyl CoA cholesterol acyltransferase 2 (ACAT2) for the etherification of CHO into cholesterol ester (CHO-E). TG and CHO-E would be assembled by membrane transport protein (MTP) to apolipoprotein B-48 and A-IV in the ER. Chylomicron (postprandial) or very low-density lipoprotein (between meals) are processed in the Golgi and exported by exocytosis into the blood. CHO may also be release to the blood through ATP-binding cassette transporters A1 (ABCA1).

Intracellular Processing and Basolateral Release

After absorption, free fatty acids and sn-2 monoacylglycerols are bound to the intestinal fatty acid–binding protein (I-FABP, Fabp2, 4q28–31) and the liver FABP (L-FABP, Fabp1, 2p11) (231–233) to prevent their transport back into the intestinal lumen, and to facilitate their transport to the endoplasmic reticulum (ER) where they are used for the de novo synthesis of triglycerides and phospholipids (Fig. 3). There are 2 main biochemical pathways for triglyceride synthesis in the enterocytes: the monoacylglycerol (MGAT) pathway (contributing to 80% of triglyceride incorporation in the chylomicrons and involving the enzymes monoacyl-glycerol transferase 1 and 2 as well as diacyl-glycerol transferase 2) and the glycerol-3-phosphate (G3P) pathway (234). Then, in the ER, the newly synthesized TG will be transported by microsomal triglyceride transfer protein (MTP, Mttp, 4q24) to link to the apolipoproteins B-48 and A-IV (apo B-48 and apo A-IV) and to form first the primordial-chylomicrons and subsequently the prechylomicrons. In the same way, cholesterol would be esterified by acyl-CoA:cholesterol acyltransferase 2 (ACAT2, Acat2, 6q25.3) into cholesterol esters and then fused to the primordial chylomicron by MTP. The prechylomicrons are transferred to the Golgi where they eventually form chylomicrons. Chylomicrons consist of ∼90% triglycerides, ∼1% cholesterol or cholesterol esters, and ∼1% proteins, and are typically synthesized during postprandial periods. During interprandial and fasting periods, intestinal very low-density lipoprotein (VLDLs) are synthesized and consist of ∼60% triglycerides and ∼10% proteins (235). Chylomicrons as well as VLDLs leave the enterocyte by exocytosis, fusing with the lateral membrane of the enterocyte and subsequently reaching the lymphatic circulation through the lateral intracellular spaces (236).

Another independent pathway for cholesterol involves the transporter ABCA1 (Abca1, 9q31.1) present on the basolateral side (237,238), which exports cholesterol to the blood (Fig. 3).

Neonatal Regulation of the Absorptive Process

Although the regulation of FATP and FABPpm in the placenta during gestation is well documented (239), little is known about the developmental regulation of these 2 transporters as well as of the transporters NPC1L1, SR-B1, and FAT/CD36 in the intestine of the fetus and the neonate. The expression level of FATP2 (Scl27a2, 15q21.2) mRNA appears to decrease by 60% between 10 and 20 days of age. SR-B1 expression increases after 20 weeks of gestation in humans (240), but postnatal regulation was not studied.

In contrast, regulation of intracellular processing during neonatal development has received some attention, and evidence suggests that the major players of lipid processing are synthesized during gestation and that the entire process may be operational at birth. Rat fetal explants (17–20 weeks of gestation) are able to synthesize and secrete chylomicrons, VLDL, and HDL. I-FABP and L-FABP gene transcription begins in late fetal life (∼17 days of gestation in rats) (241). In humans and pigs, the expression of Apo B-48 and MTP increases drastically at birth (242,243) and Apo IV is already present in the intestine of neonatal pigs (244). During the suckling phase, the intestine is capable of a better adaptation to a high-lipid diet by increasing uptake of lipids than during the weaning period (245). Greater lipid uptake during the suckling phase may result from an increased fluidity of the brush border membrane (246,247) and/or from greater rates of facilitated, carrier-mediated lipid transport. After weaning, alterations in lipid composition of brush border membrane decrease its fluidity (247), which may result in decreased rates of facilitated lipid transport. During suckling but not in weaning rats and in adults, jejunal apo B-48 and apo A-IV protein levels are stimulated by dietary fatty acids (242,244,248–250). In piglets, jejunal and ileal MTP expressions are also enhanced by high level of lipids in the lumen during the suckling phase (242).


Liposoluble vitamins include vitamins A, D, E, and K (Table 8) (251–256). Vitamin A exists as retinol, retinal, and retinoic acid, and absorption of dietary vitamin A increases the mucosal and plasma levels of all 3 forms of vitamin A (257). These retinoids are essential for vision, normal embryonic development, and control of cellular growth and differentiation (258). Vitamin E refers to a family of closely related compounds, the 4 tocopherols (α-, β-, δ-, γ-) and the 4 tocotrienols (α-, β-, δ-, γ-). Vitamin E displays antioxidant properties but is also involved in cell signaling and proliferation as well as regulation of gene expression. Vitamin D, whose active form is 1,25-dihydroxy vitamin D (calcitriol), is essential for Ca2+ and Pi metabolism, bone health, and cell growth and development (259). Vitamin K, whose 2 major isoforms are K1 and K3, is involved in the synthesis of coagulation factors, particularly that of prothrombin.

Concentrations of fat-soluble vitamins in human milk, requirement in the diet, and effects in deficiency states

Because of limited transplacental transfer, mammalian newborns typically have low stores of vitamins A, D, E, and K (251,260,261) during parturition. Thus, the neonate is highly dependent on colostrum and milk consumption to establish normal tissue stores of these vitamins. Unfortunately, breast milk contains low levels (between 40 and 50 IU/L) of vitamin D as well as vitamin K, and hence AI of these vitamins cannot be met with human milk (251,252). The complicated processing of vitamins A, D, E, and K and their metabolites is beyond the scope of this review.

Digestion and Apical Uptake

The digestion of fat-soluble vitamins requires bile salts and pancreatic lipases for micelle formation and enzymatic hydrolysis, respectively, similar to the process described above for dietary lipids (262). Although diffusive intestinal absorption remains the only pathway described for vitamin D and K, the uptake of vitamins A and E may also be mediated by lipid transporters (Fig. 3). SR-BI is clearly involved in the uptake of vitamin E in Caco2 cells (263), whereas FAT/CD36 is not. NPC1L1 and ABCA1 may also play a role in the facilitated transport of the carotenoids, a form of vitamin A, in Caco2 cells (264). However, vitamin A does not appear to be transported by CD36 or SRB1 (264). These findings were made in cultured cells, however, and the role of the active transporters in the intestinal absorption of fat-soluble vitamins remains little known in neonates.

Intracellular and Basolateral Transport

After absorption into enterocytes, vitamin A or retinol is bound to cellular retinol-binding protein type II (CRBPII, Rbp2, 3q23) (265). The retinol/CRBPII complex serves as a substrate for reesterification of the retinol by the enzyme lecithin:retinol acyltransferase (266). The retinol esters are then incorporated into chylomicrons containing other dietary lipids (267). The molecular and cellular mechanisms involved in the intracellular trafficking, assembly, and/or efflux of dietary vitamin E in intestine-derived lipoproteins remain mostly unknown. It is possible that intestinal Apo E and Apo A-IV may contribute to intestinal vitamin E absorption because subjects bearing different single nucleotide polymorphisms in Apo A-IV and Apo E display lower plasma levels of vitamin E (268). In CaCo2 cells, the main subcellular destinations of vitamin E are microsomal membranes (269); then vitamin E is incorporated into chylomicrons and secreted with I-HDL, which require, respectively, MTP for chylomicron assembly and ABCA1 for HDL excretion (269).

There is little information available on the intestinal transcellular processing of vitamin K. The subcellular destination of vitamin K and the identity of its intracellular transporters in the enterocytes remain to be discovered. After absorption from the intestinal lumen, vitamin K becomes associated with chylomicrons in the blood (270,271).

Genetic Disorders of Intestinal Lipid Transporters

Several diseases are associated with mutations of various lipid transporters. However, because these transporters are involved in lipid transport in many tissues other than the intestinal mucosa, the disorders resulting from their mutation appear to be linked to a deficiency of lipid transport into those cells rather than that in the intestine. Thus, only a few mutations have consequences that are clearly linked to impairment of intestinal lipid uptake. For example, mutations of ABCG5 or ABCG8 lead to the development of sitosterolemia (272,273). This rare autosomal, recessively inherited disorder is characterized by hyperabsorption of cholesterol and phytosterols and reduced secretion of these sterols into bile. Hypobetalipoproteinemia, an autosomal-dominant disorder, is defined by low levels (<5th percentile) of total apoB and/or of low-density lipoprotein cholesterol in plasma. Hypobetalipoproteinemia is due to a variety of genetic defects in MTP (274), impairing the assembly of lipids with apoB in lipoprotein production. Interestingly, hypobetalipoproteinemia is also associated with vitamin E deficiency (275).


There is another extremely serious defect of nutrient malabsorption that is not specific to any nutrient but is induced by nutrients in general. Loss of function mutations in the transcription factor Neurog3 results in almost total absence of neuroendocrine cells in the small intestine (276). Patients had chronic unremitting diarrhea that was malabsorptive in nature. Feeding of carbohydrate-free, water plus tryglycerides, water plus amino acids, fructose-free, soy-based, and even oral rehydration solutions each resulted in diarrhea. Only water feeding did not cause diarrhea. This extremely rare disease highlights the selective pressures imposed by GI malabsorption on their carriers who survive only with modern medical care.


For water-soluble vitamins, it is not known why dietary supplementation of some vitamins does not translate into increase in blood levels of the vitamin being supplemented.

Because the recently identified lipid transporters are found in low abundance in the neonatal intestine, they may mainly facilitate the absorption of essential LC-PUFAs critical for neonatal development. Unfortunately, the ontogenetic development of these transporters as well as their interaction with lipids present in breast milk has not been studied. Specifically, there is little information available concerning the expression, functionality, and potential growth-limiting role of these lipid transporters in preterm versus full term.

There is also little known about the uptake, intracellular processing, and basolateral transport of vitamins E and K. Developmental regulation of transport systems participating in the transport of the vitamins A, E, and K needs further study.

The postnatal development of nutrient transporter has mostly been described in suckled animals. However, considering the difference in nutrient composition between formulas and breast milk on one hand and the extreme reactivity of transporter to dietary modulations on the other hand, studies on the exact postnatal evolution of transporters in bottle-fed animals would certainly help to improve formulas to closely match formulas' composition with bottle-fed neonate requirements and absorptive capacities.

Besides their classical transport function, a novel role in immunomodulation of the mucosa has been recently described for some transporters (SGLT1, PEPT1) (277,278). The neonatal period is a critical period in terms of dialogue between the GALT and the microbiota, which shapes the gut immune system for the rest of our life. The role of such transporters in immunomodulation of the gut during the neonatal period and later in health consequences warrants further studies.


The authors thank Ms Jackie Lee for valuable help in manuscript preparation.


1. Kennaugh JM, Hay WW Jr. Nutrition of the fetus and newborn. West J Med 1987; 147:435–448.
2. Hirayama BA, Loo DD, Diez-Sampedro A, et al. Sodium-dependent reorganization of the sugar-binding site of SGLT1. Biochemistry 2007; 46:13391–13406.
3. Manolescu AR, Augustin R, Moley K, et al. A highly conserved hydrophobic motif in the exofacial vestibule of fructose transporting SLC2A proteins acts as a critical determinant of their substrate selectivity. Mol Membr Biol 2007; 24:455–463.
4. Adibi SA. Regulation of expression of the intestinal oligopeptide transporter (Pept-1) in health and disease. Am J Physiol 2003; 285:G779–G788.
5. Broer A, Tietze N, Kowalczuk S, et al. The orphan transporter v7-3 (slc6a15) is a Na+-dependent neutral amino acid transporter (B0AT2). Biochem J 2006; 393:421–430.
6. Gilbert ER, Wong EA, Webb KE. Peptide absorption and utilization: implications for animal nutrition and health. J Anim Sci 2008; 86:2135–2155.
7. Perez AV, Picotto G, Carpentieri AR, et al. Minireview on regulation of intestinal calcium absorption. Emphasis on molecular mechanisms of transcellular pathway. Digestion 2008; 77:22–34.
8. Murer H, Forster I, Biber J. The sodium phosphate cotransporter family SLC34. Pflugers Arch 2004; 447:763–767.
9. Lonnerdal B. Trace element nutrition of infants—molecular approaches. J Trace Elem Med Biol 2005; 19:3–6.
10. Quamme GA. Molecular identification of ancient and modern mammalian magnesium transporters. Am J Physiol Cell Physiol 2010; 298:C407–C429.
11. Said HM. Recent advances in carrier-mediated intestinal absorption of water-soluble vitamins. Annu Rev Physiol 2004; 66:419–446.
12. Said HM, Mohammed ZM. Intestinal absorption of water-soluble vitamins: an update. Curr Opin Gastroenterol 2006; 22:140–146.
13. Hui DY, Howles PN. Molecular mechanisms of cholesterol absorption and transport in the intestine. Semin Cell Dev Biol 2005; 16:183–192.
14. Bonen A, Chabowski A, Luiken JJFP, et al. Mechanisms and regulation of protein-mediated cellular fatty acid uptake: molecular, biochemical, and physiological evidence. Physiology 2007; 22:15–28.
15. Groh-Wargo S, Thompson M, Cox JH. Nutritional Care for High-Risk Newborns. Chicago, IL: Precept Press; 2000.
16. Ferraris RP, Hsiao J, Hernandez R, et al. Site density of mouse intestinal glucose transporters declines with age. Am J Physiol 1993; 264:G285–G293.
17. Wright EM. I. Glucose galactose malabsorption. Am J Physiol 1998; 275:G879–G882.
18. Wright EM, Turk E, Martin MG. Molecular basis for glucose-galactose malabsorption. Cell Biochem Biophys 2002; 36:115–121.
19. David ES, Cingari DS, Ferraris RP. Dietary induction of intestinal fructose absorption in weaning rats. Pediatr Res 1995; 37:777–782.
20. Wales JK, Primhak RA, Rattenbury J, et al. Isolated fructose malabsorption. Arch Dis Child 1990; 65:227–229.
21. Wasserman D, Hoekstra JH, Tolia V, et al. Molecular analysis of the fructose transporter gene (GLUT5) in isolated fructose malabsorption. J Clin Invest 1996; 98:2398–2402.
22. Nobigrot T, Chasalow FI, Lifshitz F. Carbohydrate absorption from one serving of fruit juice in young children: age and carbohydrate composition effects. J Am Coll Nutr 1997; 16:152–158.
23. Corpe CP, Burant CF, Hoekstra JH. Intestinal fructose absorption: clinical and molecular aspects. J Pediatr Gastroenterol Nutr 1999; 28:364–374.
24. Douard V, Choi HI, Elshenawy S, et al. Developmental reprogramming of rat GLUT5 requires glucocorticoid receptor translocation to the nucleus. J Physiol 2008; 586:3657–3673.
25. Douard V, Cui XL, Soteropoulos P, et al. Dexamethasone sensitizes the neonatal intestine to fructose induction of intestinal fructose transporter (Slc2A5) function. Endocrinology 2008; 149:409–423.
26. Jiang L, Ferraris RP. Developmental reprogramming of rat GLUT-5 requires de novo mRNA and protein synthesis. Am J Physiol Gastrointest Liver Physiol 2001; 280:G113–G120.
27. Cui XL, Jiang L, Ferraris RP. Regulation of rat intestinal GLUT2 mRNA abundance by luminal and systemic factors. Biochim Biophys Acta 2003; 1612:178–185.
28. Lenzen S, Lortz S, Tiedge M. Effect of metformin on SGLT1, GLUT2, and GLUT5 hexose transporter gene expression in small intestine from rats. Biochem Pharmacol 1996; 51:893–896.
29. Kellett GL, Brot-Laroche E, Mace OJ, et al. Sugar absorption in the intestine: the role of GLUT2. Annu Rev Nutr 2008; 28:35–54.
30. Barone S, Fussell SL, Singh AK, et al. The Slc2a5 (Glut5) is essential for the absorption of fructose in the intestine and generation of fructose-induced hypertension. J Biol Chem 2009; 284:5056–5066.
31. Stumpel F, Burcelin R, Jungermann K, et al. Normal kinetics of intestinal glucose absorption in the absence of GLUT2: evidence for a transport pathway requiring glucose phosphorylation and transfer into the endoplasmic reticulum. Proc Natl Acad Sci U S A 2001; 98:11330–11335.
32. Peduto A, Spada M, Alluto A, et al. A novel mutation in the GLUT2 gene in a patient with Fanconi-Bickel syndrome detected by neonatal screening for galactosaemia. J Inherit Metab Dis 2004; 27:279–280.
33. Santer R, Steinmann B, Schaub J. Fanconi-Bickel syndrome—a congenital defect of facilitative glucose transport. Curr Mol Med 2002; 2:213–227.
34. World Health Organization. Proteins and Amino Acid Requirements in Human Nutrition: A Report of a Joint FAO/WHO/UNU Expert Consultation. Geneva: WHO; 2007.
35. Hay WW. Strategies for feeding the preterm infant. Neonatalogy 2008; 94:245–254.
36. Macé K, Steenhout P, Klassen P, et al. Protein quality and quantity in cow's milk-based formula for healthy term infants: past, present and future. In: Rigo J, Ziegker EE, editors. Protein and Energy Requirements in Infancy and Childhood. Vevey: Nestec Ltd; 2006. pp. 189–205.
37. Dupont C. Protein requirements during the first year of life. Am J Clin Nutr 2003; 77:1544–1549.
38. Hill DJ, Murch SH, Rafferty K, et al. The efficacy of amino acid-based formulas in relieving the symptoms of cow's milk allergy: a systematic review. Clin Exp Allergy 2007; 37:808–822.
39. Koo WW, Lasekan JB. Rice protein-based infant formula: current status and future development. Minerva Pediatr 2007; 59:35–41.
40. Lönnerdal B. Recombinant human milk proteins. In: Rigo J, Ziegker EE, editors. Protein and Energy Requirements in Infancy and Childhood. Vevey: Nestec Ltd; 2006. pp. 207–217.
41. Axelsson I. Effects of high protein intakes. In: Rigo J, Ziegler TR, editors. Protein and Energy Requirements in Infancy and Childhood. Vevey: Nestec Ltd; 2006. pp. 121–131.
42. Koletzko B. Long-term consequences of early feeding on later obesity risk. In: Rigo J, Ziegler TR, editors. Protein and Energy Requirements in Infancy and Childhood. Vevey: Nestec Ltd; 2006. pp. 1–18.
43. Adibi SA. Intestinal transport of dipeptides in man: relative importance of hydrolysis and intact absorption. J Clin Invest 1971; 50:2266–2275.
44. Himukai M, Konno T, Hoshi T. Age-dependant change in intestinal absorption of dipeptides and their constituent amino acids in the guinea-pig. Pediatr Res 1980; 14:1272–1275.
45. Ganapathy V, Ganapathy ME, Leibach FH. Intestinal transport of peptides and amino acids. In: Barrett KE, Donowitz M, eds. Gastrointestinal Transport. Molecular Physiology. San Diego: Academic Press. 2001:379–12.
46. Ogihara H, Saito H, Shin BC, et al. Immuno-localization of H+/peptide cotransporter in rat digestive tract. Biochem Biophys Res Commun 1996; 220:848–852.
47. Chen H, Wong EA, Webb KE. Tissue distribution of a peptide transporter mRNA in sheep, dairy, cows, pigs and chickens. J Anim Sci 1999; 77:1277–1283.
48. Freeman TC, Bentsen BS, Thwaites DT, et al. H+/di-tripeptide transporter (Pept1) expression in the rabbit intestine. Pflugers Arch 1995; 430:394–400.
49. Terada T, Shimada Y, Pan X, et al. Expression profiles of various transporters for oligopeptides, amino acids and organic ions along the human digestive tract. Biochem Pharmacol 2005; 70:1756–1763.
50. Erickson RH, Guam JR Jr, Lindstrom MM, McKean D, Kim YS. Regional expression and dietary regulation of rat small intestinal peptide and amino acid transporter mRNAs. Biochem Biophys Res Commun 1995; 216:249–257.
51. Howard A, Goodlad RA, Walters JRF, et al. Increased expression of specific intestinal amino acid and peptide transporter mRNA in rats fed by TPN is reversed by GLP-2. J Nutr 2004; 134:2957–2964.
52. Rome S, Barbot I, Windsor E, et al. Regionalization of Pept1, NBAT and EAACS transporters in the small intestine of rats are unchanged from birth to adulthood. J Nutr 2002; 132:1009–1011.
53. Tanaka H, Miyamoto K, Morita K, et al. Regulation of the PepT1 peptide transporter in the rat small intestine in response to 5-fluorouracil-induced injury. Gastroenterology 1998; 114:714–723.
54. Shiraga T, Miyamoto K, Tanaka H, et al. Cellular and molecular mechanisms of dietary regulation on rat small intestinal H+/peptide transporter PepT1. Gastroenterology 1999; 116:354–362.
55. Ihara T, Tsujikawa T, Fujiyama Y, et al. Regulation of Pept1 peptide transporter expression in the rat small intestine under malnourished conditions. Digestion 2000; 61:59–67.
56. Lis MT, Crampton RF, Mattews DM. Effect of a dietary changes on intestinal absorption of L-methionine and L-methionyl-L-methionine in the rat. Br J Nutr 1972; 27:159–167.
57. Ferraris RP, Diamond J, Kwan WW. Dietary regulation of intestinal transport of the dipeptide carnosine. Am J Physiol 1988; 255:G143–G150.
58. Ferraris RP, Diamond JM. Specific regulation of intestinal nutrient transporters by their dietary substrates. Annu Rev Physiol 1989; 51:125–141.
59. Shen H, Smith DE, Brosius FC. Developmental expression of PEPT1 and PEPT2 in rat small intestine, colon and kidney. Pediatr Res 2001; 49:789–795.
60. Broër S. Amino acid transport across mammalian intestinal and renal epithelia. Physiol Rev 2008; 88:249–286.
61. Dave MH, Schulz N, Zecevic M, et al. Expression of heteromeric amino acid transporters along the murine intestine. J Physiol 2004; 558:597–610.
62. Anderson CM, Howard A, Walters JR, et al. Taurine uptake across the human intestinal brush-border membrane is via two transporters: H+-coupled PAT1 (SLC36A1) and Na+- and Cl(-)-dependent TauT (SLC6A6). J Physiol 2009; 587:731–744.
63. Casirola DM, Vinnakota RR, Ferraris RP. Intestinal amino acid transport in mice is modulated by diabetes and diet. J Nutr 1994; 124:842–852.
64. Karasov WH, Solberg DH, Diamond JM. Dependence of intestinal amino acid uptake on dietary protein or amino acid levels. Am J Physiol 1987; 252:G614–G625.
65. Stein ED, Chang SD, Diamond JM. Comparison of different dietary amino acids as inducers of intestinal amino acid transport. Am J Physiol 1987; 252:G626–G635.
66. Mourad FH, Barada KA, Khoury C, et al. Amino acids in the rat intestinal lumen regulate their own absorption from a distant intestinal site. Am J Physiol Gastrointest Liver Physiol 2009; 297:G292–G298.
67. Wenzel U, Meissner B, Doring F, et al. PEPT1-mediated uptake of dipeptides enhances the intestinal absorption of amino acids via transport system b(0,+). J Cell Physiol 2001; 186:251–259.
68. Levin RJ, Kodolvsky O, Hoskova J, et al. Electrical activity across human foetal small intestine associated with absorption processes. Gut 1968; 9:206–213.
69. Malo C. Multiple pathways for amino acid transport in brush border membrane vesicles isolated from the human fetal small intestine. Gastroenterology 1991; 100:1644–1652.
70. Ohno C, Nakanishi Y, Honma T, et al. Significance of system L amino acid transporter 1 (LAT-1) and 4F2 heavy chain (4F2hc) expression in human developing intestines. Acta Histochem Cytochem 2009; 42:73–81.
71. Bartsocas CS. Developmental aspects of amino acid transport. Suggestion of switching-on and switching-off mechanisms. Biol Neonate 1977; 31:60–64.
72. Weiss MD, Donnelly WH, Rossignol C, et al. Ontogeny of the neutral amino acid transporter SNT1 in the developing rat. J Mol Hist 2005; 36:301–309.
73. Buddington RK, Malo C. Intestinal brush-border membrane enzyme activities and transport functions during development of pigs. J Pediatr Gastroenterol Nutr 1996; 23:51–64.
74. Zhang H, Malo C, Buddington RK. Suckling induced rapid intestinal growth and changes in brush border digestive functions of newborn pigs. J Nutr 1997; 127:418–426.
75. al-Mahroos FT, Abumrad N, Ghislan FK. Developmental changes in glutamine transport by rat basolateral membrane vesicles. Proc Soc Exp Biol Med 1990; 194:186–192.
76. al-Mahroos FT, Bulus N, Abumrad N, et al. Maturational changes in glutamine transport by rat jejunal brush border membrane vesicles. Pediatr Res 1990; 27:519–524.
77. Moyer MS, Goodrich AL, Rolfes MM, et al. Ontogenesis of intestinal taurine transport: evidence for a beta-carrier in developing rat jejunum. Am J Physiol 1988; 254:G870–G877.
78. Murphy S, Daniel VG. Postnatal amino acid uptake by the rat small intestine. Changes in membrane transport systems for amino acids associated with maturation of jejunal morphology. J Dev Physiol 1979; 1:111–126.
79. Toloza EM, Diamond J. Ontogenic development of nutrient transporters in rat intestine. Am J Physiol 1992; 263:G593–G604.
80. Yoneshige A, Sasaki A, Miyazaki M, et al. Developmental changes in glycolipids and synchronized expression of nutrient transporters in the mouse small intestine. J Nutr Biochem 2010; 21:214–226.
81. Younoszai MK, Smith C, Finch MH. Comparison of in vitro jejunal uptake of L-valine and L-lysine in the rat during maturation. J Pediatr Gastroenterol Nutr 1985; 4:992–997.
82. Gilbert ER, Emmerson DA, Webb KE, et al. Developmental regulation of nutrient transporter and enzyme mRNA abundance in the small intestine of broilers. Poultry Sci 2007; 86:1739–1753.
83. Ferraris RP, Diamond J. Regulation of intestinal sugar transport. Physiol Rev 1997; 77:257–302.
84. Subcommittee on the Tenth Edition of the RDA FaNB, Commission on Life Sciences, National Research Council. Recommended Dietary Allowances. Washington DC: National Academy Press; 1989.
85. Zhuang L, Peng JB, Tou L, et al. Calcium-selective ion channel, CaT1, is apically localized in gastrointestinal tract epithelia and is aberrantly expressed in human malignancies. Lab Invest 2002; 82:1755–1764.
86. Peng JB, Chen XZ, Berger UV, et al. Molecular cloning and characterization of a channel-like transporter mediating intestinal calcium absorption. J Biol Chem 1999; 274:22739–22746.
87. Peng JB, Chen XZ, Berger UV, et al. Human calcium transport protein CaT1. Biochem Biophys Res Commun 2000; 278:326–332.
88. Meyer MB, Watanuki M, Kim S, et al. The human transient receptor potential vanilloid type 6 distal promoter contains multiple vitamin D receptor binding sites that mediate activation by 1,25-dihydroxyvitamin D3 in intestinal cells. Mol Endocrinol 2006; 20:1447–1461.
89. Song Y, Peng X, Porta A, et al. Calcium transporter 1 and epithelial calcium channel messenger ribonucleic acid are differentially regulated by 1,25 dihydroxyvitamin D3 in the intestine and kidney of mice. Endocrinology 2003; 144:3885–3894.
90. Benn BS, Ajibade D, Porta A, et al. Active intestinal calcium transport in the absence of transient receptor potential vanilloid type 6 and calbindin-D9k. Endocrinology 2008; 149:3196–3205.
91. Van Cromphaut SJ, Rummens K, Stockmans I, et al. Intestinal calcium transporter genes are upregulated by estrogens and the reproductive cycle through vitamin D receptor-independent mechanisms. J Bone Miner Res 2003; 18:1725–1736.
92. Jantarajit W, Thongon N, Pandaranandaka J, et al. Prolactin-stimulated transepithelial calcium transport in duodenum and Caco-2 monolayer are mediated by the phosphoinositide 3-kinase pathway. Am J Physiol Endocrinol Metab 2007; 293:E372–E384.
93. Huybers S, Naber TH, Bindels RJ, et al. Prednisolone-induced Ca2+ malabsorption is caused by diminished expression of the epithelial Ca2+ channel TRPV6. Am J Physiol Gastrointest Liver Physiol 2007; 292:G92–G97.
94. Wasserman RH, Chandler JS, Meyer SA, et al. Intestinal calcium transport and calcium extrusion processes at the basolateral membrane. J Nutr 1992; 122:662–671.
95. Larsson D, Nemere I. Vectorial transcellular calcium transport in intestine: integration of current models. J Biomed Biotechnol 2002; 2:117–119.
96. Lee GS, Lee KY, Choi KC, et al. Phenotype of a calbindin-D9k gene knockout is compensated for by the induction of other calcium transporter genes in a mouse model. J Bone Miner Res 2007; 22:1968–1978.
97. Liang CT, Barnes J, Imanaka S, et al. Alterations in mRNA expression of duodenal 1,25-dihydroxyvitamin D3 receptor and vitamin D-dependent calcium binding protein in aged Wistar rats. Exp Gerontol 1994; 29:179–186.
98. Yamagishi N, Miyazaki M, Naito Y. The expression of genes for transepithelial calcium-transporting proteins in the bovine duodenum. Vet J 2006; 171:363–366.
99. Eto N, Tomita M, Hayashi M. NaPi-mediated transcellular permeation is the dominant route in intestinal inorganic phosphate absorption in rats. Drug Metab Pharmacokinet 2006; 21:217–221.
100. Xu H, Bai L, Collins JF, et al. Molecular cloning, functional characterization, tissue distribution, and chromosomal localization of a human, small intestinal sodium-phosphate (Na+-Pi) transporter (SLC34A2). Genomics 1999; 62:281–284.
101. Huber K, Roesler U, Muscher A, et al. Ontogenesis of epithelial phosphate transport systems in goats. Am J Physiol Regul Integr Comp Physiol 2003; 284:R413–R421.
102. Kirchner S, Muduli A, Casirola D, et al. Luminal fructose inhibits rat intestinal sodium-phosphate cotransporter gene expression and phosphate uptake. Am J Clin Nutr 2008; 87:1028–1038.
103. Xu H, Bai L, Collins JF, et al. Age-dependent regulation of rat intestinal type IIb sodium-phosphate cotransporter by 1,25-(OH)2 vitamin D3. Am J Physiol Cell Physiol 2002; 282:C487–C493.
104. Douard V, Asgerally A, Sabbagh Y, et al. Dietary fructose inhibits intestinal calcium absorption and induces vitamin D insufficiency in CKD. J Am Soc Nephrol 2010; 21:261–271.
105. Radanovic T, Wagner CA, Murer H, et al. Regulation of intestinal phosphate transport. I. Segmental expression and adaptation to low-P(i) diet of the type IIb Na(+)-P(i) cotransporter in mouse small intestine. Am J Physiol Gastrointest Liver Physiol 2005; 288:G496–G500.
106. Giral H, Caldas Y, Sutherland E, et al. Regulation of rat intestinal Na-dependent phosphate transporters by dietary phosphate. Am J Physiol Renal Physiol 2009; 297:F1466–F1475.
107. Stauber A, Radanovic T, Stange G, et al. Regulation of intestinal phosphate transport. II. Metabolic acidosis stimulates Na(+)-dependent phosphate absorption and expression of the Na(+)-P(i) cotransporter NaPi-IIb in small intestine. Am J Physiol Gastrointest Liver Physiol 2005; 288:G501–G506.
108. Sugiura SH, Ferraris RP. Contributions of different NaPi cotransporter isoforms to dietary regulation of P transport in the pyloric caeca and intestine of rainbow trout. J Exp Biol 2004; 207:2055–2064.
109. Elin RJ. Magnesium: the fifth but forgotten electrolyte. Am J Clin Pathol 1994; 102:616–622.
110. Schweigel M, Martens H. Magnesium transport in the gastrointestinal tract. Front Biosci 2000; 5:D666–D677.
111. Meneely R, Leeper L, Ghishan FK. Intestinal maturation: in vivo magnesium transport. Pediatr Res 1982; 16:295–298.
112. Ghishan FK, Meneely RL. Intestinal maturation: the effect of glucocorticoids on in vivo net magnesium and calcium transport in the rat. Life Sci 1982; 31:133–138.
113. Voets T, Nilius B, Hoefs S, et al. TRPM6 forms the Mg2+ influx channel involved in intestinal and renal Mg2+ absorption. J Biol Chem 2004; 279:19–25.
114. Schlingmann KP, Weber S, Peters M, et al. Hypomagnesemia with secondary hypocalcemia is caused by mutations in TRPM6, a new member of the TRPM gene family. Nat Genet 2002; 31:166–170.
115. Schlingmann KP, Sassen MC, Weber S, et al. Novel TRPM6 mutations in 21 families with primary hypomagnesemia and secondary hypocalcemia. J Am Soc Nephrol 2005; 16:3061–3069.
116. Abboud S, Haile DJ. A novel mammalian iron-regulated protein involved in intracellular iron metabolism. J Biol Chem 2000; 275:19906–19912.
117. Gunshin H, Mackenzie B, Berger UV, et al. Cloning and characterization of a mammalian proton-coupled metal-ion transporter. Nature 1997; 388:482–488.
118. Vulpe CD, Kuo YM, Murphy TL, et al. Hephaestin, a ceruloplasmin homologue implicated in intestinal iron transport, is defective in the SLA mouse. Nat Genet 1999; 21:195–199.
119. Leong WI, Bowlus CL, Tallkvist J, et al. DMT1 and FPN1 expression during infancy: developmental regulation of iron absorption. Am J Physiol Gastrointest Liver Physiol 2003; 285:G1153–G1161.
120. Frazer DM, Wilkins SJ, Anderson GJ. Elevated iron absorption in the neonatal rat reflects high expression of iron transport genes in the distal alimentary tract. Am J Physiol Gastrointest Liver Physiol 2007; 293:G525–G531.
121. Nemeth E, Tuttle MS, Powelson J, et al. Hepcidin regulates cellular iron efflux by binding to ferroportin and inducing its internalization. Science 2004; 306:2090–2093.
122. De Domenico I, Lo E, Ward DM, et al. Hepcidin-induced internalization of ferroportin requires binding and cooperative interaction with Jak2. Proc Natl Acad Sci U S A 2009; 106:3800–3805.
123. Nemeth E, Roetto A, Garozzo G, et al. Hepcidin is decreased in TFR2 hemochromatosis. Blood 2005; 105:1803–1806.
124. Domellof M, Lonnerdal B, Abrams SA, et al. Iron absorption in breast-fed infants: effects of age, iron status, iron supplements, and complementary foods. Am J Clin Nutr 2002; 76:198–204.
125. Liuzzi JP, Cousins RJ. Mammalian zinc transporters. Annu Rev Nutr 2004; 24:151–172.
126. Dufner-Beattie J, Wang F, Kuo YM, et al. The acrodermatitis enteropathica gene ZIP4 encodes a tissue-specific, zinc-regulated zinc transporter in mice. J Biol Chem 2003; 278:33474–33481.
127. Huang ZL, Dufner-Beattie J, Andrews GK. Expression and regulation of SLC39A family zinc transporters in the developing mouse intestine. Dev Biol 2006; 295:571–579.
128. Wang K, Zhou B, Kuo YM, et al. A novel member of a zinc transporter family is defective in acrodermatitis enteropathica. Am J Hum Genet 2002; 71:66–73.
129. Liuzzi JP, Bobo JA, Cui L, et al. Zinc transporters 1, 2 and 4 are differentially expressed and localized in rats during pregnancy and lactation. J Nutr 2003; 133:342–351.
130. Murgia C, Vespignani I, Cerase J, et al. Cloning, expression, and vesicular localization of zinc transporter Dri 27/ZnT4 in intestinal tissue and cells. Am J Physiol 1999; 277:G1231–G1239.
131. Tacnet F, Lauthier F, Ripoche P. Mechanisms of zinc transport into pig small intestine brush-border membrane vesicles. J Physiol 1993; 465:57–72.
132. Kelly EJ, Quaife CJ, Froelick GJ, et al. Metallothionein I and II protect against zinc deficiency and zinc toxicity in mice. J Nutr 1996; 126:1782–1790.
133. IOM. Dietary Reference Intakes: Thiamin, Riboflavin, Niacin, Vitamin B6, Folate, Vitamin B12, Pantothenic Acid, Biotin, and Choline. Food and Nutrition Board, ed, Washington, DC: National Academy Press; 1998.
134. Montalto MB, Benson JD, Martinez GA. Nutrient intakes of formula-fed infants and infants fed cow's milk. Pediatrics 1985; 75:343–351.
135. Specker BL, Black A, Allen L, et al. Vitamin B-12: low milk concentrations are related to low serum concentrations in vegetarian women and to methylmalonic aciduria in their infants. Am J Clin Nutr 1990; 52:1073–1076.
136. Said HM, Ortiz A, Subramanian VS, et al. Mechanism of thiamine uptake by human colonocytes: studies with cultured colonic epithelial cell line NCM460. Am J Physiol Gastrointest Liver Physiol 2001; 281:G144–G150.
137. Bonjour JP. Biotin. In: Machlin LJ, editor. Handbook of Vitamins. New York, NY: Marcel Dekker; 1991. pp. 393–427.
138. Rindi G, Laforenza U. Thiamine intestinal transport and related issues: recent aspects. Proc Soc Exp Biol Med 2000; 224:246–255.
139. Dutta B, Huang W, Molero M, et al. Cloning of the human thiamine transporter, a member of the folate transporter family. J Biol Chem 1999; 274:31925–31929.
140. Eudy JD, Spiegelstein O, Barber RC, et al. Identification and characterization of the human and mouse SLC19A3 gene: a novel member of the reduced folate family of micronutrient transporter genes. Mol Genet Metab 2000; 71:581–590.
141. Ganapathy V, Smith SB, Prasad PD. SLC19: the folate/thiamine transporter family. Pflugers Arch 2004; 447:641–646.
142. Nabokina SM, Reidling JC, Said HM. Differentiation-dependent up-regulation of intestinal thiamin uptake: cellular and molecular mechanisms. J Biol Chem 2005; 280:32676–32682.
143. Reidling JC, Said HM. Adaptive regulation of intestinal thiamin uptake: molecular mechanism using wild-type and transgenic mice carrying hTHTR-1 and -2 promoters. Am J Physiol Gastrointest Liver Physiol 2005; 288:G1127–G1134.
144. Reidling JC, Subramanian VS, Dudeja PK, et al. Expression and promoter analysis of SLC19A2 in the human intestine. Biochim Biophys Acta 2002; 1561:180–187.
145. Said HM, Balamurugan K, Subramanian VS, et al. Expression and functional contribution of hTHTR-2 in thiamin absorption in human intestine. Am J Physiol Gastrointest Liver Physiol 2004; 286:G491–G498.
146. Boulware MJ, Subramanian VS, Said HM, et al. Polarized expression of members of the solute carrier SLC19A gene family of water-soluble multivitamin transporters: implications for physiological function. Biochem J 2003; 376:43–48.
147. Merrill AH, Froehlich JA, McCormick DB. Isolation and identification of alternative riboflavin-binding proteins from human plasma. Biochem Med 1981; 25:198–206.
148. McCormick DBaG, HL, Vitamins. In: Burtis CA, Edward R. Ashwood MD, Tiez NW, eds. Tietz Textbook of Clinical Chemistry. Philadelphia: WB Saunders; 1994:1275–1316.
149. Said HM, Ma TY, Grant K. Regulation of riboflavin intestinal uptake by protein kinase A: studies with Caco-2 cells. Am J Physiol 1994; 267:G955–G959.
150. Jusko WJ, Levy G. Absorption, metabolism, and excretion of riboflavin-5′-phosphate in man. J Pharm Sci 1967; 56:58–62.
151. Mayersohn M, Feldman S, Gibaldi M. Bile salt enhancement of riboflavin and flavin mononucleotide absorption in man. J Nutr 1969; 98:288–296.
152. Said HM, Mohammadkhani R. Uptake of riboflavin across the brush border membrane of rat intestine: regulation by dietary vitamin levels. Gastroenterology 1993; 105:1294–1298.
153. Said HM, Ortiz A, Moyer MP, et al. Riboflavin uptake by human-derived colonic epithelial NCM460 cells. Am J Physiol Cell Physiol 2000; 278:C270–C276.
154. Sundaram U. Regulation of intestinal vitamin B(2) absorption. Focus on “Riboflavin uptake by human-derived colonic epithelial NCM460 cells”. Am J Physiol Cell Physiol 2000; 278:C268–C269.
155. Horwitt MK, Harper AE, Henderson LM. Niacin-tryptophan relationships for evaluating niacin equivalents. Am J Clin Nutr 1981; 34:423–427.
156. Nabokina SM, Kashyap ML, Said HM. Mechanism and regulation of human intestinal niacin uptake. Am J Physiol Cell Physiol 2005; 289:C97–C103.
157. Bechgaard H, Jespersen S. GI absorption of niacin in humans. J Pharm Sci 1977; 66:871–872.
158. Shibata K, Gross CJ, Henderson LM. Hydrolysis and absorption of pantothenate and its coenzymes in the rat small intestine. J Nutr 1983; 113:2107–2115.
159. Fenstermacher DK, Rose RC. Absorption of pantothenic acid in rat and chick intestine. Am J Physiol 1986; 250:G155–G160.
160. Johnson LR. Physiology of the Gastrointestinal Tract. New York: Academic Press; 2006.
161. Stein ED, Diamond JM. Do dietary levels of pantothenic acid regulate its intestinal uptake in mice? J Nutr 1989; 119:1973–1983.
162. Said HM, Ortiz A, McCloud E, et al. Biotin uptake by human colonic epithelial NCM460 cells: a carrier-mediated process shared with pantothenic acid. Am J Physiol 1998; 275:C1365–C1371.
163. Baker EM, Canham JE, Nunes WT, et al. Vitamin B6 requirement for adult men. Am J Clin Nutr 1964; 15:59–66.
164. Hansen CM, Leklem JE, Miller LT. Vitamin B-6 status indicators decrease in women consuming a diet high in pyridoxine glucoside. J Nutr 1996; 126:2512–2518.
165. Miller LT, Leklem JE, Shultz TD. The effect of dietary protein on the metabolism of vitamin B-6 in humans. J Nutr 1985; 115:1663–1672.
166. Nakano H, McMahon LG, Gregory JF 3rd. Pyridoxine-5′-beta-glucoside exhibits incomplete bioavailability as a source of vitamin B-6 and partially inhibits the utilization of co-ingested pyridoxine in humans. J Nutr 1997; 127:1508–1513.
167. Said HM, Ortiz A, Ma TY. A carrier-mediated mechanism for pyridoxine uptake by human intestinal epithelial Caco-2 cells: regulation by a PKA-mediated pathway. Am J Physiol Cell Physiol 2003; 285:C1219–C1225.
168. Said ZM, Subramanian VS, Vaziri ND, et al. Pyridoxine uptake by colonocytes: a specific and regulated carrier-mediated process. Am J Physiol Cell Physiol 2008; 294:C1192–C1197.
169. Bhagavan HN, Brin M. Drug–vitamin B6 interaction. Curr Concepts Nutr 1983; 12:1–12.
170. Alpers DH. What is new in vitamin B(12)? Curr Opin Gastroenterol 2005; 21:183–186.
171. Quadros EV, Nakayama Y, Sequeira JM. The binding properties of the human receptor for the cellular uptake of vitamin B12. Biochem Biophys Res Commun 2005; 327:1006–1010.
172. Seetharam B. Receptor-mediated endocytosis of cobalamin (vitamin B12). Annu Rev Nutr 1999; 19:173–195.
173. Mock DM. Biotin. In: Ziegler EK, Filer LJ, eds. Present Knowledge in Nutrition. Washington, DC: International Life Sciences Institute Press; 1996.
174. Said HM. Biotin bioavailability and estimated average requirement: why bother? Am J Clin Nutr 1999; 69:352–353.
175. Ma TY, Dyer DL, Said HM. Human intestinal cell line Caco-2: a useful model for studying cellular and molecular regulation of biotin uptake. Biochim Biophys Acta 1994; 1189:81–88.
176. Reidling JC, Nabokina SM, Said HM. Molecular mechanisms involved in the adaptive regulation of human intestinal biotin uptake: a study of the hSMVT system. Am J Physiol Gastrointest Liver Physiol 2007; 292:G275–281.
177. Rodriguez MS. A conspectus of research on folacin requirements of man. J Nutr 1978; 108:1983–2075.
178. Said HM, Redha R. A carrier-mediated transport for folate in basolateral membrane vesicles of rat small intestine. Biochem J 1987; 247:141–146.
179. Hamid A, Kiran M, Rana S, et al. Low folate transport across intestinal basolateral surface is associated with down-regulation of reduced folate carrier in in vivo model of folate malabsorption. IUBMB Life 2009; 61:236–243.
180. Inoue K, Nakai Y, Ueda S, et al. Functional characterization of PCFT/HCP1 as the molecular entity of the carrier-mediated intestinal folate transport system in the rat model. Am J Physiol Gastrointest Liver Physiol 2008; 294:G660–G668.
181. Balamurugan K, Said HM. Ontogenic regulation of folate transport across rat jejunal brush-border membrane. Am J Physiol Gastrointest Liver Physiol 2003; 285:G1068–G1073.
182. Dudeja PK, Torania SA, Said HM. Evidence for the existence of a carrier-mediated folate uptake mechanism in human colonic luminal membranes. Am J Physiol 1997; 272:G1408–G1415.
183. Selhub JaRIH, Folic acid. In: Ziegler EK, Filer LJ, eds. Present Knowledge in Nutrition. Washington, DC: International Life Sciences Institute Press; 1996:206–219.
184. Wilson JX. Regulation of vitamin C transport. Annu Rev Nutr 2005; 25:105–125.
185. Subramanian VS, Marchant JS, Boulware MJ, et al. A C-terminal region dictates the apical plasma membrane targeting of the human sodium-dependent vitamin C transporter-1 in polarized epithelia. J Biol Chem 2004; 279:27719–27728.
186. Boyer JC, Campbell CE, Sigurdson WJ, et al. Polarized localization of vitamin C transporters, SVCT1 and SVCT2, in epithelial cells. Biochem Biophys Res Commun 2005; 334:150–156.
187. Liang WJ, Johnson D, Ma LS, et al. Regulation of the human vitamin C transporters expressed in COS-1 cells by protein kinase C. Am J Physiol Cell Physiol 2002; 283:C1696–C1704.
188. Uauy R, Castillo C. Lipid requirements of infants: implications for nutrient composition of fortified complementary foods. J Nutr 2003; 133:2962S–2972S.
189. Klein CJ. Nutrient requirements for preterm infant formulas. J Nutr 2002; 132:1395S–1577.
190. Ailhaud G, Massiera F, Weill P, et al. Temporal changes in dietary fats: role of n-6 polyunsaturated fatty acids in excessive adipose tissue development and relationship to obesity. Prog Lipid Res 2006; 45:203–236.
191. Demmelmair H, von Schenck U, Behrendt E, et al. Estimation of arachidonic acid synthesis in full term neonates using natural variation of 13C content. J Pediatr Gastroenterol Nutr 1995; 21:31–36.
192. Smithers LG, Gibson RA, McPhee A, et al. Effect of long-chain polyunsaturated fatty acid supplementation of preterm infants on disease risk and neurodevelopment: a systematic review of randomized controlled trials. Am J Clin Nutr 2008; 87:912–920.
193. Calder PC, Krauss-Etschmann S, de Jong EC, et al. Early nutrition and immunity—progress and perspectives. Br J Nutr 2006; 96:774–790.
194. Foody JM, Milberg JA, Robinson K, et al. Homocysteine and lipoprotein(a) interact to increase CAD risk in young men and women. Arterioscler Thromb Vasc Biol 2000; 20:493–499.
195. Innis SM, Gilley J, Werker J. Are human milk long-chain polyunsaturated fatty acids related to visual and neural development in breast-fed term infants? J Pediatr 2001; 139:532–538.
196. Innis SM, Dyer RA. Brain astrocyte synthesis of docosahexaenoic acid from n-3 fatty acids is limited at the elongation of docosapentaenoic acid. J Lipid Res 2002; 43:1529–1536.
197. Innis SM. Polyunsaturated fatty acids in human milk: an essential role in infant development. Adv Exp Med Biol 2004; 554:27–43.
198. Lapillonne A, Carlson SE. Polyunsaturated fatty acids and infant growth. Lipids 2001; 36:901–911.
199. Innis SM. Dietary lipids in early development: relevance to obesity, immune and inflammatory disorders. Curr Opin Endocrinol Diabetes Obes 2007; 14:359–364.
200. Gaillard D, Negrel R, Lagarde M, et al. Requirement and role of arachidonic acid in the differentiation of pre-adipose cells. Biochem J 1989; 257:389–397.
201. Massiera F, Saint-Marc P, Seydoux J, et al. Arachidonic acid and prostacyclin signaling promote adipose tissue development: a human health concern? J Lipid Res 2003; 44:271–279.
202. Scopesi F, Zunin P, Mazzella M, et al. 7-ketocholesterol in human and adapted milk formulas. Clin Nutr 2002; 21:379–384.
203. Boleman SL, Graf TL, Mersmann HJ, et al. Pigs fed cholesterol neonatally have increased cerebrum cholesterol as young adults. J Nutr 1998; 128:2498–2504.
204. Pond WG, Mersmann HJ, Su D, et al. Neonatal dietary cholesterol and alleles of cholesterol 7-alpha hydroxylase affect piglet cerebrum weight, cholesterol concentration, and behavior. J Nutr 2008; 138:282–286.
205. Reiser R, Sidelman Z. Control of serum cholesterol homeostasis by cholesterol in the milk of the suckling rat. J Nutr 1972; 102:1009–1016.
206. Li JR, Bale LK, Kottke BA. Effect of neonatal modulation of cholesterol homeostasis on subsequent response to cholesterol challenge in adult guinea pig. J Clin Invest 1980; 65:1060–1068.
207. Harris WS, Connor WE, Lindsey S. Will dietary omega-3 fatty acids change the composition of human milk? Am J Clin Nutr 1984; 40:780–785.
208. Henderson RA, Jensen RG, Lammi-Keefe CJ, et al. Effect of fish oil on the fatty acid composition of human milk and maternal and infant erythrocytes. Lipids 1992; 27:863–869.
209. Jensen RG, Ferris AM, Lammi-Keefe CJ. Lipids in human milk and infant formulas. Annu Rev Nutr 1992; 12:417–441.
210. Jensen RG. Lipids in human milk—composition and fat-soluble vitamins. In: E. Lebenthal, ed. Textbook of Gastroenterology and Nutrition in Infancy. New York: Raven Press; 1989: 157–208.
211. Kennedy K, Fewtrell MS, Morley R, et al. Double-blind, randomized trial of a synthetic triacylglycerol in formula-fed term infants: effects on stool biochemistry, stool characteristics, and bone mineralization. Am J Clin Nutr 1999; 70:920–927.
212. Zoppi G, Andreotti G, Pajno-Ferrara F, et al. Exocrine pancreas function in premature and full term neonates. Pediatr Res 1972; 6:880–886.
213. Lowe ME, Kaplan MH, Jackson-Grusby L, et al. Decreased neonatal dietary fat absorption and T cell cytotoxicity in pancreatic lipase-related protein 2-deficient mice. J Biol Chem 1998; 273:31215–31221.
214. Yang Y, Sanchez D, Figarella C, et al. Discoordinate expression of pancreatic lipase and two related proteins in the human fetal pancreas. Pediatr Res 2000; 47:184–188.
215. Hamosh M. Lipid metabolism in pediatric nutrition. Pediatr Clin North Am 1995; 42:839–859.
216. Besnard P, Niot I. Role of lipid-binding proteins in intestinal absorption of long-chain fatty acid. In: Christophe AB, De Vriese S, eds. Fat Digestion and Absorption. Champaign, IL: AOCS Press; 2000:96–118.
217. Kamp F, Zakim D, Zhang F, et al. Fatty acid flip-flop in phospholipid bilayers is extremely fast. Biochemistry 1995; 34:11928–11937.
218. Proulx P, Aubry H, Brglez I, et al. Studies on the uptake of fatty acids by brush border membranes of the rabbit intestine. Can J Biochem Cell Biol 1985; 63:249–256.
219. Schoeller C, Keelan M, Mulvey G, et al. Role of a brush border membrane fatty acid binding protein in oleic acid uptake into rat and rabbit jejunal brush border membrane. Clin Invest Med 1995; 18:380–388.
220. Schwieterman W, Sorrentino D, Potter BJ, et al. Uptake of oleate by isolated rat adipocytes is mediated by a 40-kDa plasma membrane fatty acid binding protein closely related to that in liver and gut. Proc Natl Acad Sci U S A 1988; 85:359–363.
221. Stremmel W, Lotz G, Strohmeyer G, et al. Identification, isolation, and partial characterization of a fatty acid binding protein from rat jejunal microvillous membranes. J Clin Invest 1985; 75:1068–1076.
222. Trotter PJ, Ho SY, Storch J. Fatty acid uptake by Caco-2 human intestinal cells. J Lipid Res 1996; 37:336–346.
223. Milger K, Herrmann T, Becker C, et al. Cellular uptake of fatty acids driven by the ER-localized acyl-CoA synthetase FATP4. J Cell Sci 2006; 119:4678–4688.
224. Hui DY, Labonte ED, Howles PN. Development and physiological regulation of intestinal lipid absorption. III. Intestinal transporters and cholesterol absorption. Am J Physiol Gastrointest Liver Physiol 2008; 294:G839–G843.
225. van Bennekum A, Werder M, Thuahnai ST, et al. Class B scavenger receptor-mediated intestinal absorption of dietary beta-carotene and cholesterol. Biochemistry 2005; 44:4517–4525.
226. Schulthess G, Compassi S, Werder M, et al. Intestinal sterol absorption mediated by scavenger receptors is competitively inhibited by amphipathic peptides and proteins. Biochemistry 2000; 39:12623–12631.
227. Werder M, Han CH, Wehrli E, et al. Role of scavenger receptors SR-BI and CD36 in selective sterol uptake in the small intestine. Biochemistry 2001; 40:11643–11650.
228. Altmann SW, Davis HR Jr, Zhu LJ, et al. Niemann-Pick C1 Like 1 protein is critical for intestinal cholesterol absorption. Science 2004; 303:1201–1204.
229. Yu L, Bharadwaj S, Brown JM, et al. Cholesterol-regulated translocation of NPC1L1 to the cell surface facilitates free cholesterol uptake. J Biol Chem 2006; 281:6616–6624.
230. Berge KE, Tian H, Graf GA, et al. Accumulation of dietary cholesterol in sitosterolemia caused by mutations in adjacent ABC transporters. Science 2000; 290:1771–1775.
231. Baier LJ, Bogardus C, Sacchettini JC. A polymorphism in the human intestinal fatty acid binding protein alters fatty acid transport across Caco-2 cells. J Biol Chem 1996; 271:10892–10896.
232. Besnard P, Niot I, Poirier H, et al. New insights into the fatty acid-binding protein (FABP) family in the small intestine. Mol Cell Biochem 2002; 239:139–147.
233. Hanhoff T, Lucke C, Spener F. Insights into binding of fatty acids by fatty acid binding proteins. Mol Cell Biochem 2002; 239:45–54.
234. Lehner R, Kuksis A. Biosynthesis of triacylglycerols. Prog Lipid Res 1996; 35:169–201.
235. Black DD. Intestinal lipoprotein metabolism. J Pediatr Gastroenterol 1995; 20:125–147.
236. Besnard P, Niot I, Bernard A, et al. Cellular and molecular aspects of fat metabolism in the small intestine. Proc Nutr Soc 1996; 55:19–37.
237. Brunham LR, Kruit JK, Iqbal J, et al. Intestinal ABCA1 directly contributes to HDL biogenesis in vivo. J Clin Invest 2006; 116:1052–1062.
238. Iqbal J, Anwar K, Hussain MM. Multiple, independently regulated pathways of cholesterol transport across the intestinal epithelial cells. J Biol Chem 2003; 278:31610–31620.
239. Dutta-Roy AK. Transport mechanisms for long-chain polyunsaturated fatty acids in the human placenta. Am J Clin Nutr 2000; 71:315S–322S.
240. Levy E, Menard D, Suc I, et al. Ontogeny, immunolocalisation, distribution and function of SR-BI in the human intestine. J Cell Sci 2004; 117:327–337.
241. Sacchettini JC, Hauft SM, Van Camp SL, et al. Developmental and structural studies of an intracellular lipid binding protein expressed in the ileal epithelium. J Biol Chem 1990; 265:19199–19207.
242. Lu S, Huffman M, Yao Y, et al. Regulation of MTP expression in developing swine. J Lipid Res 2002; 43:1303–1311.
243. Wu JH, Semenkovich CF, Chen SH, et al. Apolipoprotein B mRNA editing. Validation of a sensitive assay and developmental biology of RNA editing in the rat. J Biol Chem 1990; 265:12312–12316.
244. Black DD, Rohwer-Nutter PL, Davidson NO. Intestinal apolipoprotein A-IV gene expression in the piglet. J Lipid Res 1990; 31:497–505.
245. Flores CA, Hing SA, Wells MA, et al. Rates of triolein absorption in suckling and adult rats. Am J Physiol 1989; 257:G823–G829.
246. Hubner C, Lindner SG, Stern M, et al. Membrane fluidity and lipid composition of rat small intestinal brush-border membranes during postnatal maturation. Biochim Biophys Acta 1988; 939:145–150.
247. Schwarz SM, Hostetler B, Ling S, et al. Intestinal membrane lipid composition and fluidity during development in the rat. Am J Physiol 1985; 248:G200–G207.
248. Black DD, Davidson NO. Intestinal apolipoprotein synthesis and secretion in the suckling pig. J Lipid Res 1989; 30:207–218.
249. Black DD, Hay RV, Rohwer-Nutter PL, et al. Intestinal and hepatic apolipoprotein B gene expression in abetalipoproteinemia. Gastroenterology 1991; 101:520–528.
250. Gonzalez-Vallina R, Wang H, Zhan R, et al. Lipoprotein and apolipoprotein secretion by a newborn piglet intestinal cell line (IPEC-1). Am J Physiol 1996; 271:G249–G259.
251. Van Winckel M, De Bruyne R, Van De Velde S, et al. Vitamin K, an update for the paediatrician. Eur J Pediatr 2009; 168:127–134.
252. Reeve LE, Chesney RW, DeLuca HF. Vitamin D of human milk: identification of biologically active forms. Am J Clin Nutr 1982; 36:122–126.
253. Tijerina-Saenz A, Innis SM, Kitts DD. Antioxidant capacity of human milk and its association with vitamins A and E and fatty acid composition. Acta Paediatr 2009; 98:1793–1798.
254. Sethuraman U. Vitamins. Pediatr Rev 2006; 27:44–55.
255. Biesalski HK. Vitamin E requirements in parenteral nutrition. Gastroenterology 2009; 137:S92–S104.
256. Greer FR. Vitamin K the basics—what's new? Early Hum Dev 2010 [Epub ahead of print].
257. Roche M, Dufour C, Loonis M, et al. Olive phenols efficiently inhibit the oxidation of serum albumin-bound linoleic acid and butyrylcholine esterase. Biochim Biophys Acta 2009; 1790:240–248.
258. Blomhoff R, Blomhoff HK. Overview of retinoid metabolism and function. J Neurobiol 2006; 66:606–630.
259. Mimouni FBAC, Shamir RBC. Vitamin D requirements in the first year of life. Curr Opin Clin Nutr Metab Care 2009; 12:287–292.
260. Debier C. Vitamin E during pre- and postnatal periods. Vitam Horm 2007; 76:357–373.
261. Ismadi SD, Olson JA. Dynamics of the fetal distribution and transfer of vitamin A between rat fetuses and their mother. Int J Vitam Nutr Res 1982; 52:112–119.
262. Leonard SW, Good CK, Gugger ET, et al. Vitamin E bioavailability from fortified breakfast cereal is greater than that from encapsulated supplements. Am J Clin Nutr 2004; 79:86–92.
263. Reboul E, Klein A, Bietrix F, et al. Scavenger receptor class B type I (SR-BI) is involved in vitamin E transport across the enterocyte. J Biol Chem 2006; 281:4739–4745.
264. During A, Dawson HD, Harrison EH. Carotenoid transport is decreased and expression of the lipid transporters SR-BI, NPC1L1, and ABCA1 is downregulated in Caco-2 cells treated with ezetimibe. J Nutr 2005; 135:2305–2312.
265. Crow JA, Ong DE. Cell-specific immunohistochemical localization of a cellular retinol-binding protein (type two) in the small intestine of rat. Proc Natl Acad Sci U S A 1985; 82:4707–4711.
266. Herr FM, Ong DE. Differential interaction of lecithin-retinol acyltransferase with cellular retinol binding proteins. Biochemistry 1992; 31:6748–6755.
267. Blomhoff R, Green MH, Norum KR. Vitamin A: physiological and biochemical processing. Annu Rev Nutr 1992; 12:37–57.
268. Borel P, Moussa M, Reboul E, et al. Human plasma levels of vitamin E and carotenoids are associated with genetic polymorphisms in genes involved in lipid metabolism. J Nutr 2007; 137:2653–2659.
269. Anwar K, Kayden HJ, Hussain MM. Transport of vitamin E by differentiated Caco-2 cells. J Lipid Res 2006; 47:1261–1273.
270. Ichihashi T, Kinoshita H, Takagishi Y, et al. Effect of bile on absorption of mepitiostane by the lymphatic system in rats. J Pharm Pharmacol 1992; 44:565–569.
271. Shearer MJ, McBurney A, Barkhan P. Studies on the absorption and metabolism of phylloquinone (vitamin K1) in man. Vitam Horm 1974; 32:513–542.
272. Berger J, Korosec T, Unterrainer G, et al. A de novo adrenoleukodystrophy gene (ABCD1) mutation S636I without detectable ABCD1 protein and a R104C mutation with normal amounts of protein from an Austrian patient collective. Hum Mutat 2000; 16:534.
273. Lee MH, Lu K, Hazard S, et al. Identification of a gene, ABCG5, important in the regulation of dietary cholesterol absorption. Nat Genet 2001; 27:79–83.
274. Berriot-Varoqueaux N, Aggerbeck LP, Samson-Bouma M, et al. The role of the microsomal triglygeride transfer protein in abetalipoproteinemia. Annu Rev Nutr 2000; 20:663–697.
275. Kayden HJ. The genetic basis of vitamin e deficiency in humans. Nutrition 2001; 17:797–798.
276. Wang J, Cortina G, Wu SV, et al. Mutant neurogenin-3 in congenital malabsorptive diarrhea. N Engl J Med 2006; 355:270–280.
277. Palazzo M, Gariboldi S, Zanobbio L, et al. Sodium-dependent glucose transporter-1 as a novel immunological player in the intestinal mucosa. J Immunol 2008; 181:3126–3136.
278. Merlin D, Si-Tahar M, Sitaraman SV, et al. Colonic epithelial hPepT1 expression occurs in inflammatory bowel disease: transport of bacterial peptides influences expression of MHC class A molecules. Gastroenterology 2001; 120:1666–1679.

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