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Alpers disease

Alpers' disease, also called progressive infantile poliodystrophy, is a progressive degenerative disease of the central nervous system that occurs in infants and children. It is an autosomal recessive disorder that is sometimes seen in siblings. First signs of the disease, which include intractable seizures and failure to meet meaningful developmental milestones, usually occur in infancy, after the first year of life, but sometimes as late as the fifth year. more...

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Primary symptoms of the disease are developmental delay, progressive mental retardation, hypotonia (low muscle tone), spasticity (stiffness of the limbs) possibly leading to quadriplegia, and progressive dementia. Seizures may include epilepsia partialis continua, a type of seizure that consists of repeated myoclonic (muscle) jerks. Optic atrophy may also occur, often leading to blindness. Deafness may also occur. And, although physical signs of chronic liver dysfunction may not be present, many patients suffer liver impairment leading to liver failure. While some researchers believe that Alpers' disease is caused by an underlying metabolic defect, no consistent defect has been identified. Pathologically, there is status spongiosus of the cerebral grey matter.


There is no cure for Alpers' disease and, currently, no way to slow its progression. Treatment is symptomatic and supportive. Anticonvulsants may be used to treat the seizures. However, caution should be used when selecting valproate as therapy since it may increase the risk of liver failure. Physical therapy may help to relieve spasticity and maintain or increase muscle tone.


The prognosis for individuals with Alpers' disease is poor. Those with the disease usually die within their first decade of life. Liver failure is usually the cause of death, although cardiorespiratory failure may also occur.


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Advances in nutrition and gastroenterology: Summary of the 1997 A.S.P.E.N. Research Workshop
From JPEN: Journal of Parenteral and Enteral Nutrition, 1/1/98 by Klein, Samuel


ABSTRACT. Background: The 1997 A.S.PE.N. Research Workshop was held at the annual meeting in San Francisco, on January 26, 1997. The workshop focused on advances in clinical and basic research involving the interface between nutrient and luminal gastroenterology. Methods: Presentations on the genetic regulation of gastrointestinal development, the molecular biology of small intestinal adaptation, the effect of nutrition support on intestinal mucosal mass, the relationship between nutrition and gastrointestinal motility, nutrient absorption, and gastrointestinal tract substrate metabolism were made by the preeminent

leaders in the field. Results: The investigators presented an insightful analysis of each topic by reviewing data from their own laboratories and the published literature. Conclusions: This workshop underscored the important interactions between nutrition and luminal gastroenterology at the basic science, metabolic/ physiologic, and clinical levels. The integration of presentations from the different disciplines provided a unique interaction of information and ideas to advance our understanding of nutrition and gastrointestinal tract. (Journal of Parenteral and Enteral Nutrition 22:3-13, 1998)

The major function of the gastrointestinal tract is to digest and absorb nutrients. Therefore, an understanding of gastrointestinal tract development, metabolic requirements, and function are critical to our understanding of clinical nutrition. The 1997 American Society of Parenteral and Enteral Nutrition (A.S.PE.N.) Research Workshop, held at the annual meeting in San Francisco, brought together basic science, metabolic/physiologic, and clinical investigators to review the interactions between nutrition and luminal gastroenterology. The program consisted of eight presentations by leading authorities in the field followed by an open group discussion of each topic. This workshop demonstrated the importance of combining both basic science and clinical investigation to advance our understanding of this important area.


The development of the gastrointestinal tract is characterized by an unfolding of genetic programs leading to the formation of individual organs and specialized cell types.' The essential processes are "hard wired," similar among species, and time dependent. However, multiple extrinsic factors, such as peptides, hormones and growth factors in fetal blood or amniotic fluid, epithelial/mesenchymal interactions, and neural influences, also control differentiation.2 Indeed, newer models of development, such as chimeric and transgenic mice, suggest a basic commonality of mechanisms controlling intestinal differentiation in different species. This allows some extrapolation of data obtained in experimental animals to patterns identified in humans.

In the intestine, differentiation in the context of development occurs in three gradients: chronological, vertical (crypt-villus), and horizontal (proximal-distal). Such patterns have been established from the study of fetuses using a variety of analytical techniques: biochemical assays of differentiation markers, immunohistochemistry, in situ hybridization, measurement of transcriptional rates of specific genes, and transgenic mice expressing specific promoter-reporter constructs.37

A number of microvillus enzymes have been used as indicators of intestinal epithelial maturation. For example, lactase-phlorizin hydrolase appears at the time of cellular differentiation at fetal day 18 in the rat,34 whereas sucrase-isomaltase (SI) and ileal lipid-binding protein are low or absent at this time and increase as weaning begins.5' Although this pattern is somewhat different in the human (sucrase rises early and lactase later), it is now clear that these patterns are intrinsically regulated in each enterocyte and that each enzyme is independently controlled.28

A characteristic feature of the intestine is the pattern of renewal that occurs along the vertical, or crypt-villus, gradient. Cell proliferation occurs in cells toward the base of the crypt, producing cell lineages responsible for each of the four differentiated epithelial cell types (enterocytes, enteroendocrine cells, goblet cells, and Paneth cells).8 Cell migration from crypt to villus tip requires approximately 3 days in the rat and 5 days in the mature human; in the neonate, cell migration rate is slower. As the cells migrate, they acquire their characteristic phenotypes and site-specific expression of differentiation markers. For example, SI activity is highest toward the upper villus region.

The proximal-to-distal pattern of gene expression in the intestine has stimulated great interest. Indeed, a number of studies are devoted to the analysis of the molecular mechanisms that might be responsible for this horizontal gradient.3.7 Interestingly, individual enterocyte-specific proteins exhibit different patterns of expression in the same cells.4 For example, lactasephlorizin hydrolase and SI are expressed maximally in the midjejunum, whereas ileal lipid-binding protein, the apical sodium-dependent bile salt transporter, and cobalamin-intrinsic factor complex receptor are expressed maximally in the ileum. These patterns are part of the genetic program in each enterocyte, because they are recapitulated correctly even when fetal intestine is transplanted into an ectopic site in a mature recipient.2

Efforts to unravel the molecular control of this horizontal gradient have focused mainly on the analysis of promoter function, and the cis-acting elements and transacting factors that might be responsible for individual patterns of expression of fusion genes. The genes for fatty acid-binding protein, SI, and lactase-phlorizin hydrolase have been best studied.3- 7 By using transgenic mice, it has been possible to assess the function of individual promoter constructs linked to a human growth hormone reporter. Each construct yields specific expression patterns.r7 The overall interpretation of a large number of experiments is that multiple cis-acting elements and combinations of trans-acting factors are responsible for the correct expression of specific proteins along the horizontal axis.

In summary, intestinal differentiation is a complex process with multiple levels of control. Differentiation occurs in a tightly regulated chronological and spatial organization. Understanding of the integrated processes involved will require detailed analysis of multiple individual mechanisms and careful comparison of such events. Continued study of appropriate genes may identify regulatory pathways common to those proteins expressed in specific regions of the intestine during development.


The mammalian small intestine contains a rapidly proliferating and perpetually differentiating epithelium. Anchored stem cells in the crypts of Lieberkuhn give rise to four major differentiated cell types, including enterocytes, goblet, enteroendocrine, and Paneth cells. The gut epithelium has a remarkable capacity to adapt to a constantly changing environment. After the loss of functional small bowel surface area due to surgical resection, intestinal bypass, vascular injury, or inflammatory bowel disease, the remaining intestine undergoes an adaptive response. This response is characterized by increased crypt cell proliferation leading to villus cell hyperplasia, increased villus height and crypt depth, and enhanced fluid, electrolyte, and nutrient absorption. Insufficient gut adaptation in humans results in the important clinical problem of short bowel syndrome.

A rat 70% small intestinal resection model has been used commonly to define the regulatory mechanisms for intestinal cell epithelial proliferation/differentiation and mucosal regeneration. To define this regulation on a cellular and molecular level, subtractive hybridization techniques were used to isolate genes that are specifically induced or repressed during adaptation.9 A number of complementary DNAs (cDNAs) were isolated that encode likely regulators of adaptation, including cell cycle regulators, immediate early genes, and proteins that control protein synthesis and degradation (Table I). This cohort of ileal-remnant-induced genes provides a template for monitoring the intestinal response to therapeutic interventions designed to augment the adaptive response. Studies of the expression of these genes during adaptation and ontogeny, particularly at the time of crypt-villus morphogenesis, have demonstrated that the adaptive response is not limited to stimulation of crypt cell proliferation, but also includes specific changes in gene expression in enterocytes lining the villus.

A group of enterocyte-specific genes involved in nutrient trafficking were cloned because they were induced in the remnant intestine within 48 hours after intestinal resection. This group includes liver fatty acid-binding protein, ileal lipid-binding protein, apolipoprotein AIVand cellular retinol-binding protein II (CRBP II). The early induction of genes involved in nutrient trafficking after resection may occur as an adaptive mechanism to facilitate assimilation of lipids and retinoids or to regulate the adaptive response or as a nonspecific response to changes in nutrient delivery to remnant intestine. The case for a regulatory role is best for the induction by 8 hours of CRBP II, a protein that is clearly important for the absorption and metabolism of vitamin A. CRBP II messenger RNA (mRNA) levels were elevated by 8 hours after resection and remained elevated for at least 2 weeks.ll This increase occurred before the reintroduction of the chow diet and the change was distinct from other genes involved in lipid/retinoid trafficking such as apolipoprotein AI. It is well established that CRBP II functions to facilitate intestinal vitamin A absorption and metabolism, that vitamin A assimilation can be regulated by CRBP II,12,13 and that CRBP II expression is subject to regulation by retinoic acid.14 Furthermore, vitamin A is essential for normal cell growth, differentiation, and maintenance of epithelial tissues.

On the basis of these considerations, one might expect changes in CRBP II levels after resection to reflect a regulatory role for vitamin A in adaptation. To directly address the effects of vitamin A in adaptation, Wang et all' administered retinoic acid or vehicle IV to rats immediately after 70% small bowel resection. Compared with vehicle, all-trans retinoic acid significantly stimulated crypt cell proliferation in the adapting remnant intestine by 6 hours after surgery. In addition, the intestinal adaptive response was attenuated markedly in vitamin A-deficient rats (D. A. Swartz-Basile, PhD, D. C. Rubin,MD, M. S. Levin, MD, unpublished data). These data suggest that retinoic acid acts to modulate intestinal proliferation in the adapting small intestine after loss of functional small bowel surface area.


Numerous animal studies have assessed the importance of luminal nutrients on intestinal mucosal mass and function. The clinical use of enteral supplements to maintain mucosal mass and its barrier function is based on studies performed in animals, mostly rats, demonstrating that total parenteral nutrition (TPN) or starvation induced mucosal atrophy. However, TPN is a major stress in rats and leads to alterations in mucosal permeability. Moreover, there are species differences in TPNinduced atrophy"5 and in increased mucosal permeability,'6 both phenomena being much more significant in rats than in humans or other animals. In humans, only prolonged starvation is associated with marked changes in intestinal mucosa.17-18In adult patients, the use of TPN without oral feedings for up to 2 weeks did not cause mucosal atrophy" and short-term TPN in normal volunteers caused only a mild decrease in mucosal thickness, which was not related to altered cell number.20 Similarly, prolonged TPN in children has been shown to cause only mild focal villous atrophy.21A comparison of the route of feeding on small intestinal mucosa was made in patients with chronic pancreatitis who were given preoperative enteral or parenteral nutrition." Both enteral and parenteral nutrition were associated with decreased jejunal mucosa thickness, but the decrease in villus height was greater in patients given enteral nutrition compared with those given parenteral nutrition. However, the diets were not isocalorically administered and preoperative nutritional status was not reported.

It has been hypothesized that translocation of intestinal bacteria into the bloodstream may be responsible for infectious complications in malnourished patients, critically ill patients, and patients with gastrointestinal diseases. Although the phenomenon of bacterial translocation into mesenteric lymph nodes has been demonstrated in humans, it was not correlated with nutritional status.23 Intestinal permeability has been used as a marker for bacterial translocation because of the difficulty in evaluating bacterial translocation directly in humans. However, no data exist in humans linking the presence of altered intestinal permeability with bacterial translocation.24 Moreover, assessment of permeability may depend on the marker used. Short-term starvation in normal volunteers led to a decrease in intestinal permeability of mannitol but no change in the permeability of EDTA or lactulose.25

Glutamine is an important nutrient for intestinal mucosa. Therefore, it has been hypothesized that TPN, which does not contain glutamine, is suboptimal for maintaining gastrointestinal tract mass and function. Unfortunately, the issue of whether glutamine supplementation improves mucosal mass in humans receiving TPN has not been adequately studied.zo There are also limited data evaluating the benefit of glutamine-supplemented TPN on mucosal function. However, Tremel et al27 found that critically ill patients given glutamine-dipeptide-supplemented parenteral nutrition had greater D-xylose absorption compared with patients given standard TPN, but no measurements of mucosal mass were performed. Thus, the reason for the functional change was not apparent. The clinical efficacy of glutamine, growth hormone, and a high-carbohydrate, low-fat diet in patients receiving home TPN was evaluated recently.28Forty percent of patients (19/47) were able to be weaned off TPN. However, it is not known if treatment success was associated with increased mucosal mass because no parameters of mucosal mass were evaluated.

A few studies in preterm infants and neonates have documented a response to enteral nutrients by following surrogate markers of bowel growth and development.29"31 Several growth-promoting gut hormones have been shown to increase with initial low-caloric feedings in very-low-birth-weight infants.293 These findings have confirmed a standard of care for premature infants in several centers, which promote small oral feedings (5% to 10% of projected needs) during the first weeks of life. 30,31

In summary, there is not enough information in humans to support the concept that TPN induces intestinal mucosal atrophy or altered function. Moreover, it is not clear whether the lower rate of septic complications observed in critically ill patients given enteral rather than parenteral nutrition32 is due to beneficial effects of enteral feeding on the gastrointestinal tract or to the increased risk of infection associated with TPN.


Digestion and absorption of nutrients are time-demanding events. To achieve optimal nutrition, the speed of movement of food must be tightly controlled. Gastric emptying is the first point of traffic control for the meal. Gastric emptying is regulated by the load of the meal, the length of intestinal exposure, the nutrient itself, and the region of exposure and the phase of the meal.33-35 Loaddependent inhibition of gastric emptying serves to prevent uncontrolled entry of nutrients so that the absorptive sites of the proximal small intestine are not overwhelmed. In an experimental animal model that allowed selected lengths of the small intestine to be exposed to glucose, the intensity of the inhibitory feedback on gastric emptying depended on the number of inhibitory sensors recruited for the response.3334 Thus, after a large meal a greater number of stimulated sensors are recruited along a longer length of the small intestine to participate in a more intense inhibitory response. In addition, the intensity of inhibition was nutrient-specific; inhibitory feedback of fat was approximately 37 times as potent as glucose.34 This difference can be explained by the slower rate of absorption of oleic acid. With less rapid removal of the nutrient from the lumen, a longer length of the small intestine must be exposed to participate in the inhibitory feedback. The control of transit was also region- and phase-specific. Although there was no difference in liquid emptying when glucose was delivered to the proximal or distal quarter of the small intestine, distal quarter glucose was approximately three times more potent than the proximal quarter in inhibiting solid emptying.35

Once food is emptied from the stomach, controlled intestinal transit becomes the next point of regulation. The idea that fat absorption is completed within the proximal small intestine was suggested by the intestinal intubation study of Borgstrom et al,36 in which fat absorption after an emulsion meal was reported to approach 100% after the first 100 cm of the small intestine. Recently, this question was reexamined, and it was found that the completion of fat absorption by the midpoint of the small intestine was dependent on the load of fat.3 After a 60 g fat meal, 17 g of fat was not absorbed by the proximal half of small intestine. Exposing the distal half of gut to fat was critical for optimal fat absorption, because regulated intestinal transit depended on the inhibitory feedback that was generated from the distal gut. Although intestinal transit is known to be inhibited by fat in the ileum,3839 fat confined to the proximal half of the small intestine also inhibited intestinal transit,4 but was less potent than when perfused into the ileum.41 Such a region-specific control system permits adjustment of intestinal transit according to fat load. A small load of fat is absorbed before reaching the distal small intestine and does not trigger maximal intestinal slowing, whereas a large fat load may reach the distal small intestine and cause more intense slowing of intestinal transit.

A variety of neural and humoral mediators are involved in the control of intestinal transit. Fat-induced jejunal brake is dependent on the accelerating effect of cholecystokinin and the slowing effect of a peripherally located opioid pathway.42 By using the technique of peptide immunoneutralization, peptide tyrosine tyrosine (PYY) was shown to mediate the fat-induced ileal brake response.43

Intestinal transit also is regulated by other dietary factors, such as the load of protein entering the small intestine.44 In addition, intact whey soy protein inhibited intestinal transit more than hydrolyzed protein.45 Furthermore, when either a low-residue or a high-fiber formula was perfused into the small intestine at 50 or 100 mL/h, it was found that increasing the rate of formula delivery had a very different effect on intestinal transit for the two formulas. Although there was a flow-dependent accelerating effect with the low-residue formula, this expected effect was not seen with the high-fiber formula.46 Instead, intestinal transit was slowed profoundly at either of the two rates of formula delivery. This effect of fiber supplementation was dependent on the displacement of nutrients into the distal small intestine.


Dietary lipids are a major energy source and provide essential fatty acids, fat soluble vitamins, and cholesterol to the body. The absorption of fat is efficient and 95% of dietary loads up to 500 g/d are absorbed. Fat absorption is made complex because of its insolubility in water and the mechanisms required to package absorbed fats for distribution to depot sites throughout the body.47

Triacylglcyerol Absorption

Triacylglycerol (TG), the major dietary fat, is insoluble in water and therefore forms globules in water with a small surface area exposed to the aqueous phase. The first step in fat absorption is to increase its surface area by dispersing it as an emulsion. Cooking, grinding, chewing, and intragastric mixing of fat help form emulsions. Dietary phospholipids, denatured proteins, and cholesterol are poorly soluble in water but are attracted to oil/ water interfaces and thereby decrease the energy required to form emulsions. Another emulsification mechanism is provided in the stomach by gastric lipase, which helps to create and stabilize emulsions by generating free fatty acids (FFAs) from TG.

When the emulsified lipid enters the small intestine, its surface area is much greater than the original fat globules. The surface of the emulsion is covered by a monolayer of phospholipids, fatty acid, free cholesterol, and denatured protein. The large oil/water surface makes an ideal substrate for pancreatic colipase-dependent lipase, which is only active at oil/water interfaces. In the presence of physiologically relevant concentrations of bile salts in the duodenum 5 to 10 mmol/L), which is above their critical micellar concentration (CMC) of 1.8 mmol/ L, the lipase desorbs from the interface and becomes inactive.48 However, a colipase, also secreted by the pancreas, anchors the lipase to the interface even in the presence of bile salts in concentrations above their CMC, and thus lipolysis proceeds quickly under physiologic conditions.

Hydrolysis of TG by lipase generates two FFAs and one sn-2 monoacylglycerol (MG). As these products appear, they are able to interact with water to a much greater extent then their parent TG. First lipolysis continues to reduce the size of the emulsion particles, but soon the hydrolytic products are present in excess. These now bud off the emulsions as liquid crystals and multilayered liposomes. In the presence of bile salts, these quickly form small unilamellar vesicles that further interact with bile salt micelles to form mixed bile salt micelles. These lipid-saturated micelles deliver dietary lipid to the apical membrane of the enterocytes. The lipid products of the micelle are absorbed in the proximal small intestine, whereas the bile salt component is absorbed in the terminal ileum.

The many steps involved in the absorption process take place quickly. Within 60 cm from the pylorus, 80% of the lipid from a meal is absorbed by the small intestinal mucosa.

Intracellular Events in Lipid Absorption

Exactly how the FFA and MGs penetrate the apical membrane of the enterocyte is unclear. It is likely that a specific transport protein is involved. Alternatively, FFAs could be reionized on the cytosolic face of the membrane and thus become trapped inside the cell. Once inside the cell, the FFA and MG likely use two fatty acid-binding proteins (I-FABP and L-FABP) expressed in intestinal cytosol to ferry them to the cytosolic face of the endoplasmic reticulum (ER). There are significant differences between the two FABPs: L-FABP binds two FFAs and I-FABP one; L-FABP delivers its FFA by diffusion whereas I-FABP's delivery mechanism is by collision; and MG binds to I-FABP but not I-FABP

Once on the ER membrane, FFA and MG are reesterified rapidly to TG by enzymes located on the cytosolic face of the ER. Resynthesis to TG occurs by two different routes. The primary route utilizes MG to which two FFAs are added progressively via specific acyl transferases. This route predominates, provided that adequate MG is available from the lumen. The second TG synthetic route utilizes 3-glycerophosphate as its glyceride-glycerol precursor. The diacylglycerol (DG) formed by the first route is not metabolically equivalent to the DG formed by the second pathway. The DG synthesized from MG goes only to TG, whereas DG synthesized via the second pathway can be used for TG or phospholipid synthesis. The FFAs utilized for TG synthesis via the MG pathway come predominantly from the diet.

The TG produced is exported from the intestine in the unique intestinal TG transport vehicle, the chylomicron. Therefore, fatty acids within chylomicron TGs reflect dietary intake. Prechylomicrons form in the ER by microsomal triglyceride transport protein, translocating apolipoprotein B48 across the ER membrane into the ER lumen, likely in concert with phospholipids (predominantly phosphatidylcholine) and a small amount of TG. The major portion of TG is added in a second step in the smooth ER. Other constituents of the chylomicron surface are also added such as free cholesterol and apolipoproteins A-I, A-IV, and the C lipoproteins. The prechylomicrons then go to the Golgi, where the proteins are modified by glycosylation.49 From the Golgi, the chylomicrons traverse the cytosol in another vesicle to the basolateral membrane. The vesicle fuses with the membrane and the enclosed chylomicrons are exocytosed into the lamina propria.

The composition of TGs in the intestinal mucosa do not reflect dietary TG fatty acids. However, chylomicron TG fatty acids are very similar to the composition of fatty acids in dietary TG. This potential paradox can be resolved by the realization that there are at least two pools of TG in the mucosa. One of these, termed pool A, supplies TG for chylomicron formation. This pool turns over rapidly, 0.6/h in the rat, and fatty acids can be shown to go from the intestinal lumen to the lymph in only 12 minutes. Pool A is comprised of dietary lipids. The second TG pool, pool B, turns over more slowly and exits the intestine via the portal vein, either as TG or as FA bound to albumin. Pool B TGs are composed of FFAs from plasma and TG from chylomicron remnants that have returned to the intestine.50,51

Regulation of Chylomicron Lymph Output and Its Physiologic Consequences

The small intestine regulates the proportion of absorbed TG that is released into the lymphatic system.52 In rats, only approximately 54% of the input rate exits the intestine via the lymph under large, but fully absorbed, TG loads. When phosphatidylcholine is added to the enteral infusion, 85% of intraduodenally infused TG appears in the lymph, whereas only 37% of the duodenal input rate exits in lymph in rats who have had bile duct diversion. It has been shown that the ability to increase or decrease lipid output into the lymph is due to an expansion or contraction, respectively, of pool A. Pool B has been documented to behave in the contrary manner, contracting when TG throughput is mostly into the lymph and expanding when TG is mainly transported via the portal route. When TG goes into the lymph, the TG in chylomicrons gets quickly metabolized in the periphery, mostly adipose tissue and muscle, generating chylomicron remnants and ultimately high-density lipoprotein. When a large quantity of the absorbed lipid goes to the liver, much of it may be exported as very-low-density lipoprotein.


The uptake of sugars by the small intestine is accomplished by the integrated action of digestive enzymes and transport proteins. Three major dietary monosaccharides, glucose, galactose, and fructose, represent the vast majority of ingested sugars. The et 1-4 glucosidic bonds of starches are cleaved by salivary and intestinal amylase. The resulting maltose, maltotriose, and alphalimit dextrins as well as natural disaccharides, sucrase and lactose, are degraded further to monosaccharides by the action of the large glycoproteins, maltase, SI, and lactase, which are attached to the brush border membranes of the small intestine.

In mammals, the transit of glucose and other carbohydrates across the plasma membrane is carried out by two classes of transport proteins, both of which are represented in the small intestine.53 The first is the sodium/ glucose cotransporter (SGLT) system, which is an energy-dependent secondary active transport system that transports glucose against its concentration gradient. The other class of glucose carriers is the facilitative glucose transporter gene family, which mediates the bidirectional, energy-independent, stereo-specific transfer of glucose and other sugars across cell membranes. cDNAs for six functional members of this gene family (designated GLUT 1/erythrocyte, GLUT2/hepatocyte, GLUT3/brain, GLUT4/muscle-fat, GLUT5/small intestine, and GLUT7/microsomal) have been isolated and characterized to date.53 Direct evidence that SGLT1 is the primary glucose transporter of the small intestine comes from the observed mutations in SGLT1 resulting in malabsorption of glucose and galactose.4 However, mutations in SGLT1 do not result in diminished fructose uptake. Subsequently, GLUT5 was identified as the transporter that mediates the uptake of fructose in the small intestine as well as in other tissues.55

SGLT1 and GLUT5 reside on the apical or brush border membrane of the enterocyte whereas GLUT2 is targeted to the basolateral membrane. The present model suggests that glucose and galactose is actively transported from the intestinal lumen by SGLT1 and concentrated in the cell where it exits via passive diffusion through GLUT2 into the bloodstream. Because GLUT2 also has fructose transport activity, it is likely that enterocytic fructose uptake is mediated by GLUT5 and the exit via GLUT2. Although evidence has been presented for a substantial amount of carbohydrate uptake under certain conditions via a pericellular route,56 recent studies suggest that this route may be overstated significantly in the awake, intact animal.57

Regulation of Glucose and Fructose Transporter Expression in the Small Intestine

Glucose uptake in the small intestine is regulated by a number of factors. During postnatal development, increases in glucose and fructose uptake and decreases in galactose uptake occur as a "hardwired" program, which is independent of carbohydrate intake.68 This suggests that a genetic program is responsible for the increase in transport. During adulthood, however, the levels of glucose transport in the brush border and the basolateral membrane is regulated rapidly and reversibly by the amount of glucose in the diet. When rats are switched from a low- to a high-carbohydrate diet, glucose transport is rapidly doubled because of an induction of phloridzin-inhibitable binding (phloridzin binding being equivalent to the Na+-dependent glucose transporter, SGLT1) at the brush border membrane. In other studies, an equal or increased effect on the up-regulation of the basolateral membrane facilitative glucose transport was seen in animals fed diets enriched for glucose, which corresponds to an increase in GLUT2 levels.59 The increase in basolateral glucose transport also can be seen with IV infusion of glucose and may be due to both increases in number and functional activity of GLUT2.

The regulation of fructose uptake in the small intestine is similar to glucose in that the largest increase is seen with its cognate substrate. For example, when rats were fed 65% fructose in their diet, there was a 2.5-fold stimulation in fructose uptake after 3 days and no significant change in glucose uptake compared with a diet consisting of 30% glucose." Dr Burant's group61and others,62,63 recently found that an increase in small intestinal GLUT5 protein and mRNA occurs in response to a fructose diet. The rise in GLUT5 level is rapid, as is the decrease when fructose is withdrawn.

Besides up-regulation by diet, the absorption of fructose in the small intestine can be augmented by the concomitant administration of glucose in both humans and in rats. Holdsworth and Dawson63' first described the stimulatory effect of glucose on fructose absorption, which has been subsequently confirmed in other studies. In fact, in both humans and in rats, the effect of glucose is best if it is in a 1:1 molar ratio with fructose. This potentiation can be inhibited by acarbose, an inhibitor of SI, which suggests a role for SI in the cotransport of glucose and fructose.64 However, recent studies have found that amino acids, in particular those associated with water absorption (eg, L-alanine) resulted in significantly better facilitated fructose absorption when compared with glucose. These data suggest that the movement of osmotically active solutes (eg, Na+ and amino acids) result in enhanced absorption of other solutes (ie, fructose).

Regulation of Gene Expression in Intestinal Epithelia

The rate of migration of the polarized epithelial cells is such that the average life span is approximately 3 days. The absorptive epithelial cells constitute 90% to 95% of the cells of the villus and are derived from a crypt precursor pool. The identity of the precursor pool and the mechanism for renewal of the villus cells are unknown. The highest levels of SI, lactase, GLUT2, GLUT5, and SGLT1 mRNA and protein occur in the midvillus region in the small intestine under normal conditions.5 In general, little is known about the mechanism of differentiation of the crypt enterocytes and activation of gene transcription. Transgenic mice have provided some information about the regulation of SI during the differentiation profile of the villus cells. The proximal 324 bp of the SI gene is able to direct intestine-specific expression. Four different domains upstream form the TATA box of the SI gene have been identified by deoxyribonuclease 1 footprint analysis. One of these cis-acting sequences was found to be related to a homeobox family of genes, Cdx2.65 Two of the other sites were found to bind HNF-1 proteins. Additional potential transcription factor binding sites have been identified without formal functional properties identified in the SI gene promoter.

Recently, the 5' upstream region of the GLUT5 gene from mouse has been cloned and found to have high homology in the first 600 bp of the human GLUT5 gene (C. Corpe, MD, and C. Burant, MD, PhD, unpublished results). At least three binding sites for regulatory proteins have been found in this region. One of the binding sites has homology to HNF-4 but is clearly different as determined by competition studies (D. Belt, MD, and C. Burant, MD, PhD, unpublished data).

Adaptation of Sucrase and Hexose Transporter Expression

Adaptation of the small intestine can occur with concomitant changes in carbohydrate digestive and absorptive functions. In pregnancy, the small intestine hypertrophies with and increase in the absorptive capacity for glucose.66 In hyperthyroidism, an increase in glucose transporter expression and activity is described that may be in response to the hyperphagia of this condition." The small intestine also has the remarkable ability to adapt to changes in nutrient load, which varies with the order of mammals studied. With starvation there is an atrophy of the small intestinal villi with a decrease in glucose transport capacity.66 TPN in rats"8 and humans" can attenuate the decrease in glucose transport. In contrast, small bowel resection causes a marked hypertrophy of the remaining intestinal segments with a marked increase in the levels of sodium-dependent glucose transporter mRNA expression.

Insulinopenic diabetes is another condition that alters intestinal glucose transport, as well as other digestive and nutritive processes. Besides hypertrophy of specific mucosal regions, there are alterations in hydrolases and water, electrolyte, and nutrient absorption. Glucose absorption is increased by diabetes in both the brush border membrane and in the basolateral membranes of the enterocyte. The levels of both protein and mRNA corresponding to SGLT1, GLUT2, and GLUT5 increase markedly after induction of insulinopenic diabetes, which is reversed by insulin treatment.70 In contrast, in models of noninsulin-dependent diabetes in the mouse (ob/ob)71 and rats (Zucker-Diabetic Fatty) (C. Corpe, MD and C. Burant, MD, PhD, unpublished results) there is hypertrophy of the gut without a significant change in transporter expression.


Although the gastrointestinal tract accounts for approximately 25% of whole body energy expenditure, there is relatively little quantitative in vivo information regarding intestinal oxidative substrate metabolism. In this context, there are two critical questions: (1) Do these tissues metabolize specific, or preferred, substrates? and (2) Given the dual substrate supply, do the mucosal cells have a preferred source of substrates?

To answer these questions, in vivo experiments using conscious animals receiving nutritionally adequate feedings are needed, so that the physiologic rates of luminal and systemic delivery of an appropriate mixture of potential substrates are maintained. Furthermore, techniques that allow the independent quantification of the uptake and metabolism of luminal and systemic substrates should be used.

Tracer Methodology

The major problem of studying intermediary metabolism with carbon tracers is its cyclic nature, and conclusions that rely only on measurements of the movement of isotopically labeled carbon atoms through intermediary metabolism can be quite misleading. Recent developments in mass spectrometry and nuclear magnetic resonance spectroscopy have allowed the practical exploitation of a technique"273 termed mass isotopomer analysis. The technique measures labeled molecules (ie, molecules bearing 1, 2,. . . x 13C-atoms) rather than labeled atoms. By using a tracer in which all the carbon atoms are labeled (so-called uniformly labeled, [U-13C], tracers), mass isotopomer analysis allows the separate quantification of the following: (1) administered tracer molecules, a key measurement with regard to nutrient absorption74; (2) tracer molecules that have been recycled through metabolism, which is critical to the quantification of first-pass metabolism; and (3) the rates of specific pathways of tracer metabolism.73 The combination of the isotopic technique with enteral and systemic infusions and with portal and arterial sampling is a particularly powerful approach to studies of gut metabolism.

An illustration of the technique is provided by recent studies of dietary nucleotide utilization.75'6 The consumption of nucleotide-free diets has been associated with mucosal atrophy and it has been proposed that nucleosides are essential nutrients for the mucosa.77,78 Earlier studies with 14C-labeled nucleosides or amino acids (summarized in Grimble?7 and LeLeiko and Walsh's) have led to the general conclusion that this nutritional essentiality reflects a specific requirement for preformed (dietary) nucleosides as precursors for mucosal nucleic acid synthesis. To examine this proposition, mice were fed diets that contained nutritionally significant amounts of [U-"C]nucleosides and the incorporation into mucosal RNA of [U-13C]- and [13C5]purines was quantified." The [U-13C]-isotopomer arises only from the direct incorporation of the tracer molecules, whereas the [13C5]isotopomer can only reflect incorporation via the salvage pathway. Although 13C incorporation into RNA was readily demonstrable, 80% of the labeled purines contained only single 13C-labeled atoms. This indicated that the U-13C-tracer nucleosides had been degraded almost entirely and that substantial quantities of 13C had been reincorporated via de novo purine synthesis. This supports the conclusion that the mucosal nucleic acid nucleotides had arisen predominantly from de novo synthesis and that measurements of the incorporation of 13C (ie, the analysis that would be typical of a 14C experiment) would have overestimated the direct incorporation of intact dietary nucleosides by at least 20-fold. The result does not deny the trophic action of dietary nucleosides, but questions whether the effect bears any direct relationship to nucleic acid synthesis.

Glucose, Glutamine, and Amino Acid Metabolism

The pioneering work of Windmueller and Spaeth79-81 suggested that amino acid (especially nonessential amino acid) metabolism plays a critical role in mucosal energetics. Although this early work involved a preparation (the in situ vascularly perfused isolated loop) in which there would have been changes in the relative rates of delivery of luminal and systemic substrates, subsequent in vivo studies generally support the idea that amino acids may be the preferred substrates for mucosal energy metabolism.

The fact that the uptake of arterial glucose by the portal drained viscera in the fasted state82 and the first-pass dietary glucose oxidation in the fed state"8 are both minimal largely excludes glucose as a major energy source for the portal-drained viscera. In contrast, it is clear that considerable quantities of dietary amino acids are utilized by the tissues of the splanchnic bed,' probably by the intestinal mucosa.s1 Furthermore, the ingestion of protein is associated with a considerable portal output of ammonia and alanine.86 Recent experiments in piglets with either enterally administered [U-13C]amino acids5 or with dual IV-intragastric infusions of 2H- and '3C-phenylalanine suggest that >50% of the measured first-pass intestinal utilization of amino acids is directed to catabolism and that mucosal protein synthesis may preferentially utilize arterial amino acids.

Perhaps the most significant observation of Windmueller and Spaeth's was the substantial ability of the mucosa to use glutamine as a metabolic substrate. Despite the caution of the authors' conclusions in this regard, this seminal observation has been taken to signify that glutamine is the major oxidative substrate. Although it seems likely that uptake of arterial glutamine is important for mucosal function,87 this might not reflect the role of glutamine as a major oxidative fuel. With some notable exceptions (eg, Jungas et al86), a critical observation of Windmueller and Spaeths has often gone unrecognized, ie, when glutamate or glutamine were presented to the gut from the lumen, the ability of the intestine to oxidize glutamate was at least as great as its ability to oxidize glutamine. This observation has been confirmed recently in vivo with stable isotopic experiments in A recent study"9 also has shown that the atrophic effects of a glutamine plus glutamate-free diet on the mucosa can be reversed by the provision of enteral glutamate. Furthermore, studies of the metabolism of enteral [U-13C]glutamate in young pigs have shown the following: (1) 95% the dietary glutamate is metabolized in first pass, presumably in the mucosa; (2) at least 65% of this enters the mucosal Krebs cycle; (3) there is little or no mucosal glutamine synthesis from the absorbed glutamate; and (4) the contribution of enteral glutamate to the mucosal glutamate/glutamine pool is at least fourfold higher than the contribution of arterial glutamine uptake. Therefore, the burden of evidence favors the idea that in the fed state enteral glutamate is a particularly important source of energy to the mucosa.

These results, of course, leave unanswered the related questions of why intestinal uptake of arterial glutamine continues in the fed state and what is the basis of its functionally beneficial effects in the mucosa? It is possible that continuing glutamine utilization reflects a basal need for glutamate that is not satisfied by a dietary source, although this seems unlikely. It is also possible that glutamine plays a regulatory role that does not require its metabolism. Finally, in view of the results from the nucleotide labeling experiments discussed above, it seems likely that glutamine may well function as a key precursor for the nitrogen utilized for mucosal nucleic acid synthesis.


Short-chain fatty acids (SCFAs) are fermentation end products of colonic bacterial polysaccharidase degradation of dietary fiber and undigested starch. These SCFAs are either utilized as energy for bacterial maintenance and proliferation or absorbed by the colonic epithelium. The physiologic effects of SCFAs include the following: (1) enhanced sodium absorption; (2) increased colonocyte proliferation; (3) metabolic energy production; (4) enhanced colonic blood flow; (5) stimulation of the autonomic nervous system; and (6) increased gastrointestinal hormone production. These effects all maintain the colonic milieu. Acetate, propionate, and n-butyrate account for approximately 85% of all SCFAs produced in the human colon.91

SCFAs and Sodium Absorption

Rapid absorption of SCFAs from the colonic lumen is principally a nonsaturable, transcellular process. Protonated SCFAs cross into colonocytes apically by nonionic diffusion in a concentration-dependent manner.92-94 For nonionized diffusion to occur, SCFA anions are protonated at the apical colonocyte membrane by Na--H+ exchange.94 SCFA absorption is coupled to sodium absorption by this method of H+ recycling. After entering the colonic mucosal cells, protonated SCFAs dissociate and release hydrogen ions, which in turn, are transported back into the colonic lumen in exchange for sodium. This overall process leads to increased sodium, and consequently water, absorption by colonocytes. This is the rationale for the hypothesis that SCFAs derived from dietary fiber may have an antidiarrheal effect.

SCFAs and Colonocyte Proliferation

Studies with radiolabeled thymidine show increased colonic crypt cell turnover and migration in animals fed fiber-supplemented diets." In rats given separate guar gum- and pectin-supplemented diets, colonocyte proliferation increases more than in those fed cellulose- or fiber-free diets. Low-fiber diets produce colonic atrophy characterized by mucosal hypoplasia and decreased colonocyte proliferation.oo Dietary components such as cellulose may sustain the colonic mucosa, but the trophic effects in the colon are generally believed to be mediated by SCFAs.97 Although the precise mechanism is unknown, SCFAs may exert their trophic effect by providing energy to colonocytes or by stimulating release of enterotrophic gastrointestinal hormones.97,99-100

Of the primary SCFAs, butyrate plays a major role in colonocyte proliferation and consequent colonic mucosal growth. SCFAs have a dose-dependent stimulatory effect on crypt cell proliferation in the following order of effectiveness: butyrate > propionate > acetate.9%101 When delivery of dietary fiber to the colonic lumen is decreased, butyrate production diminishes, resulting in mucosal atrophy.0o Accordingly, butyrate infusions into the colonic lumen promote mucosal growth characterized by increased mucosal mass, DNA content, and mitotic indices.97-100,12-104 In rats, intraluminal perfusion of butyrate concentrations of 20, 40, and 150 mmol/L into cecectomized colon increase segmental DNA by 100%, 100%, and 50%, respectively. Butyrate infusions of 20 mol/ L stimulate colonic mucosal growth similar to physiologic concentrations of butyrate, propionate, and acetate combined. However, butyrate concentrations 7.5 times the normal colonic concentration (20 mol/L) do not significantly increase colonocyte proliferation and mucosal growth.105

SCFAs and Metabolic Energy Production

As the preferred metabolic fuel for the colonic mucosa, SCFA provides approximately 70% of the energy supply to the colonic mucosa. Of the three principal SCFAs, butyrate is the preferred metabolic fuel of normal rat colonocytes.101,106 When compared with other common cellular fuels, butyrate is the principal respiratory fuel for rat colonocytes with acetoacetate, with L-glutamine and D-glucose following in the sequential order of importance.101,107 Regional differences demonstrate that butyrate oxidation predominates in the distal colon, whereas glucose and glutamine oxidation are more pronounced in the proximal colon. In humans, colonocyte preference for metabolic fuels is similar to that found in rats.101,106

Because SCFAs are not endogenously synthesized, the colonic mucosa can only obtain these metabolic fuels from bacterial fermentation. Thus, in states of SCFA deprivation, such as decreased dietary fiber ingestion or colonic microflora reduction, diminished SCFA oxidation leading to reduced adenosine triphosphate production may severely impair colonocyte function and lead to mucosal breakdown.

SCFAs and Colonic Blood Flow

The trophic effects of SCFAs on the colonic mucosa are probably mediated partially by enhanced mesenteric blood flow.'08'19 Intraluminal infusion of SCFAs produces a 24% increase of colonic blood flow, which suggests that SCFAs directly dilate the colonic vasculature. Of the main SCFAs, acetate produces the greatest increase of blood flow.'os SCFAs individually and in combination produce a significant concentration-dependent dilation of resistant arteries in resected human colon." This vasodilatory effect suggests that SCFAs may improve colonic microcirculation in vivo, thereby having a trophic effect on colonic mucosa. These results may explain why modest doses of SCFAs have trophic effects on intestinal mucosa even after parenteral administration. 110-112

SCFAs and the Autonomic Nervous System

Accumulating evidence suggests that the autonomic nervous system has a role in mediating the enterotrophic effects of SCFAs. In rats, SCFAs rapidly perfused into the colon increase the mitotic and labeling indices of colonocytes.ll; This trophic effect is abolished by preceding a surgical vagotomy or chemical sympathectomy with guanethidine sulfate.104,113,114 Furthermore, acute SCFA infusions significantly increase crypt cell production rate in both normally innervated and extrinsically denervated jejunal segments."4 These results indicate that SCFAs can stimulate jejunal enterocyte proliferation systemically without the prerequisite for efferent autonomic nerve connections.

Findings from studies of chronic infusions, however, differ from the aforementioned acute study. Administration of SCFAs for 10 days into normally innervated rat cecum out of continuity produce trophic changes in jejunum.ll" When extrinsic denervation of rat ceca is performed, the jejunotrophic effects of SCFAs infused via the ceca are abolished. This chronic study suggests that afferent innervation is essential for the jejunotrophic effects of cecally infused SCFA, whereas the previous acute study suggests transection of efferent innervation does not alter the trophic effects of SCFA.

In a recent study to determine which component of the autonomic nervous system (ie, parasympathetic or sympathetic) is responsible for jejunotrophism, rats received cecal infusions of either SCFAs or saline for 10 days after surgical vagotomy, chemical sympathectomy, or sham operation.llE In such rats, both parasympathetic and sympathetic systems mediate the trophic effects of cecal SCFAs on jejunum. However, only disrupted parasympathetic connections fully block these effects on jejunal function, such as glucose absorption.l'6 When considered together, these findings suggest that autonomic receptors play an important role in SCFA-induced regulation of enterocyte proliferation.

SCFAs and Gastrointestinal Hormones

SCFAs may produce their enterotrophic effects by enhanced production of gastrointestinal hormones. Gastrin, enteroglucagon, and PYY are most often implicated in the mediation of intestinal proliferation and mucosal growth. Increased colonocyte proliferation stimulated by fermentable fiber, the substrate precursor for SCFAs, is closely associated with increased plasma enteroglucagon levels throughout the intestinal tract and elevated PYY levels in the colon. There is no significant correlation between increased colonocyte proliferation and levels of plasma gastrin.117In rats with normally innervated ceca, however, SCFA infusions have a systemic jejunotrophic effect associated with increased levels of jejunal tissue gastrin.115 In denervated rat ceca, SCFAs do not elevate tissue gastrin levels or promote a jejunotrophic effect Jejunal PYY levels do not increase significantly with innervated or denervated cecal infusions of SCFAs. Clearly, more studies are needed to clarify the relationship between SCFAs and gastrointestinal hormones.

SCFAs: Antineoplastic Effects

In contrast to their stimulus for proliferation of normal colonocytes, SCFAs (especially butyrate) inhibit growth of colorectal cancer cells in vitro. A recent study demonstrated that butyrate inhibited growth of hepatic colorectal metastases in vivo.l's In another study butyrate decreased in vivo crypt surface hyperproliferation in the normal rat colon.lls This effect was associated with an increased colonic expression of c-Jun. Finally, recent studies have shown that continuous infusing of butyrate into the hepatic artery significantly decreases the number of hepatic metastases in an in vivo murine model of metastatic colorectal cancer.120


The authors appreciate the efforts of the A.S.P.E.N. Research and Data Committee, Edward Bernstein, and other A.S.PE.N. staff in organizing this workshop. This work was supported in part by National Institutes of Health Grant DK-52290.


1. Grand RJ, Watkins JB, Torti F1: Development of the human gastrointestinal tract (Invited Review). Gastroenterology 70:790(810, 1976 2. Winter HS, Hendren RB, Fox CH, et al: Human intestine matures as nude mouse xenograft. Gastroenterology 100:89-98,1991 3. Krasinski SD, Estrada G, Yeh K-Y, et al: TranScriptional regulation of intestinal hydrolase biosynthesis during postnatal development in rats. Am J Physiol 267(Gastrointest Liver Physiol 30):G584594, 1994 4. Rings EHHM, Krasinski SD, Van Beers EH, et al: Restriction of lactase gene expression along the proximal-to distal axis of rat small intestine occurs during postnatal development. Gastroenterology 106:122S1232, 1994

5. Sacchettini JC, Hauff. SM, Van Camp SL, et al: Development and structural studies of an intracellular lipid binding protein expressed in the ileal epithelium. J Biol Chem 265:1919919201, 1990

6. Suh E, Traber PG: An intestine-specific homeobox gene regulates proliferation and differentiation. Mol Cell Biol 16:619625,1996 7. Simon TC, Roberts LJ, Gordon JI: A20-nucleotide element in the intestinal fatty acid binding protein gene modulates its cell lineage-specific, differention-dependent, and cephalocaudal patterns of expression in transgenic mice. Proc Natl Acad Sci USA 92:8&5W689, 1995 8. Henning SJ: Postal development: Coordination of feeding, digestion, and metabolism. Am J Physiol 241(Gastrointest Liver Physiol 4):G 199-214, 1981

9. Dodson BD, Wang JL, Swietlicki E, et al: Subtractive hybridization cloning and characterization of cDNAs differentially expressed in the adapting remnant small intestine following massive small bowel resection. Am J Physiol 271:G347356, 1996

10. Rubin DC, Swietlicki EA, Wang JL, et al: Enterocytic gene expression in intestinal adaptation: Evidence for a specific cellular response. Am J Physiol 270:G143152, 1996

11. Wang JL, Swartz-Basile D, Rubin DC, et al: Retinoic acid stimulates early cellular proliferation in the adapting remnant rat small intestine after partial resection. J Nutr 127:1297-1303,1997 12. Levin MS: Cellular retinol binding proteins are determinants of retinol uptake and metabolism in stably transfected Caco-2 cells. J Biol Chem 268:8267-8276, 1993

13. Lissoos T, Davis AE, Levin MS: Retinyl ester synthesis and secretion in Caco-2 cells transfected with cellular retinol binding proteins. Am J Physiol 268:G224-231, 1995

14. Levin MS, Davis AE: Retinoic acid increases cellular retinol binding protein II mRNA and retinol uptake in the human intestinal Caco-2 cell line. J Nutr 127:13-17, 1997

15. Sitrin HS, Bryant M, Ellis LM: Species differences in TPN-induced intestinal villus atrophy (abstract). JPEN 16 (Suppl):30S, 1992 16. Pappenheimer JR: On the coupling of membrane digestion with intestinal absorption of sugars and amino acids. Am J Physiol 265:G409-417, 1993

17. Brunser O, Reid A, Monckeberg F, et al: Jejunal mucosa in infant malnutrition. Am J Clin Nutr 21:976-981, 1968

18. Winick M (ed): Hunger Diseases: Studies by the Jewish Physicians in the Warsaw Ghetto, Vol 7. John Wiley & Sons, New York, 1979 19. Guedon C, Schmitz J, Lerebours E, et al: Decreased brush border hydrolase activities without gross morphologic changes in human intestinal mucosa after prolonged total parenteral nutrition of adults. Gastroenterology 90:373-378, 1986

20. Buchman AL, Moukarzei AA, Bhuta S, et al: Parenteral nutrition is associated with intestinal morphologic and functional changes in humans. JPEN 19:453-460, 1995

21. Rossi TM, Lee PC, Young C, et al: Small intestinal mucosa changes, including epithelial cell proliferative activity, of children receiving total parenteral nutrition (TPN). Dig Dis Sci 38:16091613, 1993 22. Groos S, Hunefeld G, Luciano L: Parenteral versus enteral nutrition: Morphological changes in human adult intestinal mucosa J Submicrosc Cytol Pathol 28:61-74, 1996

23. Sedman PC, Macfie J, Sagar P, et al: The prevalence of gut translocation in humans. Gastroenterology 107:643649, 1994

24. Lipman TO: Bacterial translocation and enteral nutrition in humans: An outsider looks in. JPEN 19:156-165,1995

25. Elia M, Goren A, Behrens R, et al: Effect of total starvation and very low calorie diets on intestinal permeability in man. Clin Sci 73:205-210, 1987

26. Payne-lames JJ, Grimble GK: The present state of glutamine. Curr Opin Gastroenterol 11:161-167, 1995

27. Tremel H, Kienle B, Weilemann L, et al: Glutamine-dipeptide supplemented parenteral nutrition maintains intestinal function in the critically ill. Gastroenterology 107:15951601, 1994 28. Byrne TA, Persinger RL, Young LS, et al: A new treatment for patients with short-bowel syndrome: Growth hormone, glutamine, and a modified diet. Ann Surg 222:243255, 1995

29. Aynsley-Green A, Lucas A, Lawson GR: Gut hormones and regulatory peptides in relation to enteral feeding, gastroenteritis and necrotizing enterocolitis in infancy. J Pediatr 117:524-532, 1990 30. Berseth CL: Effect of early feeding on maturation of the preterm infant's small intestine. J Pediatr 120:947-953, 1992

31. Troche B, Harvey-Wilkes K, Engle WD, et al: Early minimal feedings promote growth in critically ill premature infants. Biol Neonat 67:172181, 1995

32. Heyland DK, Cook DJ, Guyatt GH: Enteral nutrition in the critically ill patient: A critical review of the evidence. Intensive Care Med 19:435 442, 1993

33. Lin HC, Doty JE, Reedy TJ, et al: Inhibition of gastric emptying by glucose depends on the length of the intestine exposed to the nutrient. Am J Physiol 256:G204411, 1989

34. Lin HC, Doty JE, Reedy TJ, et al: Inhibition of gastric emptying by sodium oleate depends on the length of intestine exposed to the nutrient. Am J Physiol 259:G1030-1036, 1990

35. Lin HC, Kim BH, Elashoff JD, et al: Gastric emptying of solid food is most potently inhibited by carbohydrates in the canine distal ileum. Gastroenterology 102:793-801, 1992

36. Borgstrom B, Dahlqvist A, Lundh G: Studies of intestinal digestion and

absorption in the human. J Clin Invest 36:1521-1536, 1957 37. Lin HC, Zhao X-T, Wang IJ: Fat absorption is not complete by midgut but dependent on load of fat. Am J Physiol 34:G62-67, 1996 38. Spiller RC, Trotman IF, Higgins BE: The ileal brake-inhibition of jejunal motility after ileal fat perfusion in man. Gut 25:365-374, 1984 39. Read N, McFarlane A, Kinsman R, et al: Effect of infusion of nutrient solutions into the ileum on gastrointestinal transit and plasma levels of neurotensin and enteroglucagon. Gastroenterology 86:274-290, 1984 40. Lin HC, Zhao X-T, Wang L-J: Jejunal brake: Inhibition of intestinal transit by fat. Dig Dis Sci 41:326-329, 1996

41. Lin HC, Zhao X-T, Wang LI: Intestinal transit is more potently inhibited by fat-induced ileal than jejunal brake. Dig Dis Sci 42:19-25, 1997 42. Lin HC, Zhao X-T, Wang I-J: Intestinal transit of fat in proximal gut depends on accelerating effect of CCK and slowing effect of opioid pathway (abstract). Dig Dis Sci 41:1884, 1996

43. Lin HC, Zhao X-T, Wang L-J, et al: Fat-induced ileal brake depends on peptide YY. Gastroenterology 110:1491-1495, 1996 44. Zhao X-T, Miller RH, McCamish MA, et al: Protein absorption depends on load-dependent inhibition of intestinal transit in dogs. Am J Clin Nutr 64:319-332, 1996

45. Zhao X-T, McCamish MA, Miller RH, et al: Intestinal transit and protein absorption depend on load and degree of hydrolysis of soy protein (Abstr). Gastroenterology lO:A849, 1996

46. Lin HC, Zhao X-T, Chu AW, et al: Fiber-supplemented enteral formula slows intestinal transit by intensifying feedback from the distal gut. Am J Clin Nutr 65:1840-1844, 1997

47. Carey MC, Hernell O: Digestion and absorption of fat. Semin Gastrointest Dis 3:189-208, 1992

48. Hofmann AF: Bile acids. IN The Liver Biology and Pathobiology, 3rd ed, Arias IM, Boyer JL, Fausto N, et al (eds). Raven Press, New York, 1994

49. Kumar NS, Mansbach CM II: Determinants of triacylglycerol transport from the endoplasmic reticulum to the Golgi in intestine. Am J Physiol 273:G1&G30, 1997

50. Mansbach CM II, Dowell RF: Uptake and metabolism of circulating fatty acids by rat intestine. Am J Physiol 263:G927-933, 1992 51. Mansbach CM II, Dowell RF: The role of the intestine in chylomicron remnant clearance. Am J Physiol 269:144-152, 1995 52. Mansbach CM II, Arnold A: Steady state kinetic analysis of triacylglycerol delivery into mesenteric lymph. Am J Physiol 251:G2&3269, 1986

53. Bell GI, Burant CF, Takeda J, et al: Molecular biology of glucose transporters. J Biol Chem 268:19161-19164, 1993

54. Wright EM, Turk E, Zabel B, et al: Molecular genetics of intestinal glucose transport. J Clin Invest 88: 1451440, 1991 55. Burant CF, Takeda J, Brot-Laroche E, et al: GLI sT5 is the fructose transporter of the small intestine and sperm. J Biol Chem 265:13276-13282, 1992

56. Pappenheimer JR, Reiss KZ: Contribution of solvent drag through intracellular junctions to absorption of nutrients by the small intestine of the rat. J Membr Biol 100:123136, 1987

57. Uhing MR, Kimura RE: The effect of surgical bowel manipulation and anesthesia on intestinal glucose absorption in rats. J Clin Invest 95:27902798,1995

58. Ferraris RP, Diamond JM: Specific regulation of intestinal nutrient transporters by their dietary substrates. Annu Rev Physiol 51:125141, 1989 59. Cheeseman CI, Harley B: Adaptation of glucose transport across rat enterocyte basolateral membrane in response to altered dietary carbohydrate intake. J Physiol 437:563575, 1991

60. Crouzoulon G, Korieh A: Fructose transport by rat intestinal brush border membrane vesicles. Effect of high fructose diet followed by return to standard diet. Comp Biochem Physiol 100:175-182, 1991 61. Burant CF, Saxena MD: Rapid, reversible substrate regulation of fructose transporter (GLUT5) expression in rat small intestine and kidney Am J Physiol 267:G71-79, 1994

62. Miyamoto K, Hase K Takagi T, et al: Differential responses of intestinal glucose transporter mRNA transcripts to levels of dietary sugars. Biochem J 295:211-215, 1993

63. Holdsworth CD, Dawson AM: Absorption of fructose in man. Proc Soc Exp Biol Med 118:142-145, 1965

64. Fujisawa T, Riby J, Kretchmer N: Intestinal absorption of fructose in the rat. Gastroenterology 101:360-367, 1991

65. Traber PG, Silberg DG: Intestine-specific gene transcription. Annu Rev Physiol 58:275-297, 1996

66. Philpott DJ, Butzner JD, Meddings JB: Regulation of intestinal glucose transport. Can J Physiol Pharmacol 70:1201-1207, 1992 67. Khoja SM, Kellett GL: Effect of hypothyroidism on glucose transport

and metabolism in rat small intestine. Biochim Biophys Acta 1179:76 80, 1993

68. Miura S, Tanaka S, Yoshioka M, et al: Changes in intestinal absorption of nutrients and brush border glycoproteins after total parenteral nutrition in rats. Gut 33:484-489, 1992

69. Inoue Y, Espat NJ, Frohnapple DJ, et al: Effect of total parenteral nutrition on amino acid and glucose transport by the human small intestine. Ann Surg 217:604-612, 1993

70. Burant CF, Flink S, DePaoli AMI, et al: Small intestine hexose transport in experimental diabetes: Increased transporter mRNA and protein expression in enterocytes. J Clin Invest 93:578-585, 1994 71. Ferraris RP, Vinnakota RR: Intestinal nutrient transport in genetically obese rice. Am J Clin Nutr 62:54046, 1995 72. Tserng KY, Kalhan SC: Estimation of glucose carbon recycling and glucose turnover with [U;C]glucose. Am J Physiol 245:E476-482,1983 73. Kalderon B, Gopher A, Lapidot A: Metabolic pathways leading to liver glycogen repletion in vivo, studied by GC-MS and NMR. FEBS Lett 204:2932, 1986

74. Berthold HK, lachey DL, Reeds PJ, et al: Uniformly labelled algal protein used to determine amino acid essentiality in vivo. Proc Natl Acad Sci USA 88:8091-8095, 1991

75. Berthold HK, Crain P, Reeds PJ, et al: Dietary pyrimidines but not dietary purines make a significant contribution to nucleic acid synthesis. Proc Natl Acad Sci USA 92:10123-10127, 1995

76. Boza JJ, Jahoor F, Reeds PJ: Ribonucleic acid nucleotides in maternal and fetal tissues derive almost exclusively from synthesis de novo in pregnant mice. J Nutr 126:1749-1758, 1996 77. Grimble GK: Dietary nucleotides and gut mucosal defense. Gut 1 (Suppl):S46-51, 1994

78. LeLeiko NS, Walsh MJ: The role of glutamine, short-chain fatty acids, and nucleotides in intestinal adaptation to gastrointestinal disease. Pediatr Clin North Am 43:451-469, 1996

79. Windmueller HG, Spaeth AE: Uptake and metabolism of plasma glutamine by the small intestine. J Biol Chem 249: 5070-5079, 1974 80. Windmueller HG, Spaeth AE: Intestinal metabolism of glutamine and glutamate from the lumen as compared to glutamine from blood. Arch Biochem Biophys 171:662672, 1975

81. Windmueller HG, Spaeth AE: Respiratory fuels and nitrogen metabolism in vivo in small intestine of fed rats. Quantitative importance of glutamine, glutamate and spartate. J Biol Chem 255:107-112, 1980

82. Ebner S, Schoknecht P, Reeds PJ, et al: Growth and metabolism of gastrointestinal and skeletal muscle tissues in protein-malnourished neonatal pigs. Am J Physiol 266:R1736-1743, 1994

83. Moore MC, Pagliassotti MJ, Swift LL, et al: Disposition of a mixed meal by the conscious dog. Am J Physiol 266:E666675, 1994 84. Hoerr RA, Matthews DE, Bier DM, et al: Effects of protein restriction and acute refeeding on leucine and lysine kinetics in young men. Am J Physiol 264:E567-575, 1993

85. Stoll B, Burrin DG, Jahoor F, et al: Absorption of dietary amino acids studied with U13C-protein tracer in piglets (Abstr). J Pediatr Gastroenterol Nutr 22:438, 1996

86. Jungas RL, Halperin ML, Brosnan JT: Quantitative analysis of amino acid oxidation and related gluconeogenesis in humans. Physiol Rev 72:419-448, 1992

87. Souba WW, Smith RJ, Wilmore DW: Glutamine metabolism by the intestinal tract. JPEN 9:608-617, 1985

88. Matthews DE, Marano MA, Campbell RG: Splanchnic bed utilization of glutamine and glutamic acid in humans. Am J Physiol 264:E848454, 1993

89. Horvath K, Jami M, Hill ID, et al: Isocaloric glutamine-free diet and the morphology and function of rat small intestine. JPEN 20:128-134,1996 90. Cummings JH: Colonic absorption: The importance of short-chain fatty acids in man. Scand J Gastroenterol 19 (Suppl):89, 1984 91. Cummings JH, Branch WJ: Fermentation and the production of short chain fatty acids in the large intestine. IN Basic and Medical Aspects of Dietary Fiber, lst ed, Vahouny GV, Kritchevsky D (eds). Plenum Press, New York,1986, p 131

92. Ruppin H, Bar-Meir S, Sorgel KH: Absorption of short-chain fatty acids by the colon. Gastroenterology 78:1500, 1980 93. Rechkemmer G, Engelhardt W: Concentration and pH-dependence of short-chain fatty acid absorption in the proximal and distal colon of guinea pig (Cavia porcellus). Comp Biochem Physiol A 91:659, 1988 94. Engelhardt W: Absorption of short-chain fatty acids from the large intestine. IN Physiological and Clinical Aspects of Short-Chain Fatty Acids, Cummings JH, Rombeau JL, Sakata T (eds). Cambridge University Press, Cambridge, 1995, p 149

95. Vahouny GV, Cassidy MM: Dietary fiber and intestinal adaptation. IN Basic and Medical Aspects of Dietary Fiber, 1st ed, Vahouny GV, Kritchevsky D (eds). Plenum Press, New York, 1986, p 181 96. Bristol JB, Williamson CN: Large bowel growth. Scand J Gastroenterol 19 (Suppl):25, 1984

97. Sakata T: Stimulatory effect of short-chain fatty acids on epithelial cell proliferation in the rat intestine: A possible explanation for trophic effects of fermentable fibre, gut microbes and luminal trophic factors. Br J Nutr 58:95, 1987

98. Cameron IL, Ord VA, Hunter KE, et al: Quantitative contribution factors regulating rat colonic crypt epithelium: Role of parenteral and enteral feeding, caloric intake, dietary cellulose level and the colon carcinogen DMH. Cell Tissue Kinet 23:227, 1990

99. Clarke RM: "Luminal nutrition" versus "functional work-load" as controllers of mucosal morphology and epithelial replacement in the rat small intestine. Digestion 15:411, 1974

100. Chinery R, Goodlad RA, Wright NA: Soy polysaccharide in an enteral diet: Effects on rat intestinal cell proliferation, morphology and metabolic function. Clin Nutr 11:277, 1992

101. Roediger WEW: Utilization of nutrients by isolated epithelial cells of the rat colon. Gastroenterology 83:424, 1982

102. Goodlad RA, Wright NA Effects of addition of kaolin or cellulose to an elemental diet on intestinal cell proliferation in the rat. Br J Nutr 50:91, 1.n

103. Sakata T: Depression of intestinal epithelial cell production rate by hindgut bypass in rats. Scand J Gastroenterol 23:1200, 1988 104. Sakata T, Yajima T Influence of short chain fatty acids on the epithelial cell division of the digestive tract. Quant J Exp Physiol 69:639, 1984 105. Kripke SA, Fox AD, Berman JM, et al: Stimulation of intestinal mucosal growth with intracolonic infusion of short-chain fatty acids. JPEN 13:109, 1989

106. Roediger WEW: Role of anaerobic bacteria in the metabolic welfare of the colonic mucosa in man. Gut 21:793, 1980

107. Ardawi MSM, Newsholme EA: Fuel utilization on colonocytes of the rat. Biochem J 231:713, 1985

108. Kvietys PR, Granger DN: Effect of volatile fatty acids on blood flow and oxygen uptake by the dog colon. Gastroenterology 80:962, 1981 109. Demigne C, Remesy C: Stimulation of absorption of volatile fatty acids and minerals in the cecum of rats adapted to a very high fiber diet. J Nutr 115:53-60, 1985

110. Mortensen FV, Nielsen H, Mulvany MJ, et al: Short-chain fatty acids dilate isolated human colonic resistance arteries. Gut 31:1391, 1990 111. Koruda MJ, Rolandelli RH, Zimmaro-Bliss D, et al: Parenteral nutrition supplemented with short-chain fatty acids: Effect on the small bowel mucosa in normal rats. Am J Clin Nutr 51:685, 1990 112. Karlstad MD, Kileffer JA, Bailey JW, et al: Parenteral nutrition with short- and long-chain triglycerides: Triacetin reduces atrophy of small and large bowel mucosa and improves protein metabolism in burned rats. Am J Clin Nutr 44:1005, 1992

113. Sakata T, Engelhardt W: Stimulatory effect of short-chain fatty acids on the epithelial cell proliferation in the rat large intestine. Comp Biochem Physiol A 74:459, 1983

114. Sakata T: Stimulatory effect of short-chain fatty acids on the epithelial cell proliferation of isolated and denervated jejunal segment of the rat. Scand J Gastroenterol 24:886, 1989

115. Frankel WL, Zhang W, Singh A, et al: Stimulation of the autonomic nervous system mediates short-chain fatty acid-induced jejunotrophism. Surg Forum 43:24, 1992

116. Reilly K, Frankel W, Klurfeld D, et al: The parasympathetic (PSNS) and sympathetic (SNS) nervous systems mediate the systemic effects of short-chain fatty acids (SCFA) on jenunal structure and function. Surg Forum 44:230, 1993

117. Goodlad RA, Lerton W, Ghatei MA, et al: Effects of an elemental diet, inert bulk, and different types of dietary fiber on the response of the intestinal epithelium to refeeding in the rat and relationship to plasma gastrin, enteroglucagon, and PYY concentrations. Gut 28:171, 1987 118. Velazquez O, Jabbar A, DeMatteo R, et al: Butyrate inhibits seeding and growth of colorectal metastases to the liver in mice. Surgery 120:440448 1996

119. Velazquez O, Zhou, D, et al: In vivo crypt surface hyperproliferation is decreased by butyrate and increased by deoxycholate in normal rat colon: Associated in vivo effects on C-Fos and c)Jun expression. JPEN 20:243-250, 1996

120. Park JO, Zhou ED, Velazquez OC, et al: Continuous butyrate infusion into the hepatic artery inhibits growth of colorectal hepatic metastases in a murine model. Surg Forum 6:48:26-29, 1997

*Washington University School of Medicine, St Louis; ^Tufts University of Medicine, Boston; ^^Cedars-Sinai Medical Center, Los Angeles; (sec)University of Tennessee Medical Group, Menphis;//University of Chicago, Chicago; (para)US Department of Agriculture/Children's Nutrition Research Center, Houston; and #Hospital of the University of Pennsylvania, Philadelphia

Received for publication, August 29, 1997. Accepted for publication, September 8, 1997. Correspondence and reprint requests: Samuel Klein, MD, Washington University School of Medicine, 660 South Euclid Avenue, Box 8127, St Louis, MO 63110-1093.

Copyright American Society for Parenteral and Enteral Nutrition Jan/Feb 1998
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