Dietary lipids modify the age-associated changes in intestinal uptake of fructose in rats

L. Drozdowski,1 T. Woudstra,1 G. Wild,2 M. T. Clandinin,1 and A. B. R. Thomson1

1Nutrition and Metabolism Group, Division of Gastroenterology, Department of Medicine, University of Alberta, Edmonton, Alberta; and 2Department of Anatomy and Cell Biology, McGill University, Montreal, Quebec, Canada

Submitted 21 July 2003 ; accepted in final form 16 September 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Because reduced nutrient absorption may contribute to malnourishment in the elderly, age and diet modulate fructose uptake in mice, and alterations in fructose uptake may be paralleled by changes in the abundance of fructose transporters, the objectives of this study were to determine 1) the effects of aging on fructose absorption in rats, 2) the effect of feeding diets enriched with saturated fatty acids (SFA) vs. polyunsaturated fatty acids (PUFA), and 3) the mechanisms of these age-and diet-associated changes. Male Fischer 344 rats aged 1, 9, and 24 mo received isocaloric diets enriched with SFA or PUFA. The uptake of 14C-labeled D-fructose was determined in vitro using the intestinal sheet method. Northern and Western blot analyses and immunohistochemistry were used to determine the abundance of sodium-independent glucose and fructose transporters (GLUT)2 and GLUT5. When expressed on the basis of mucosal surface area, jejunal fructose uptake was increased in 9 and 24 mo compared with 1-mo-old animals fed SFA. PUFA-fed animals demonstrated increased fructose uptake at 24 mo compared with younger animals. Ileal fructose uptake was increased with SFA vs. PUFA in 9-mo-old rats but was reduced with SFA in 1- and 24-mo-old rats. Variations in GLUT2 and GLUT5 abundance did not parallel changes in uptake. These results indicate that 1) age increases fructose uptake when expressed on the basis of mucosal surface area, 2) age influences the adaptive response to dietary lipid modifications, and 3) alterations in fructose uptake are not explained by variations in GLUT2 or GLUT5.

fatty acids; sugar; absorption; small intestine; aging


INCREASES IN THE NUMBER of older people has focused attention on the physiological processes associated with aging, as well as strategies to improve the quality of life for the elderly. The elderly are at a high risk for malnutrition. Although there are many physiological and social factors involved, a reduction in nutrient absorption may contribute to this malnourishment. A study using breath hydrogen analysis after a carbohydrate meal showed evidence of malabsorption in the elderly (7). Similarly, transport experiments using isolated brush-border membrane (BBM) vesicles demonstrated a reduction in Na+-dependent D-glucose uptake in older patients (34). In contrast, a study by Wallis et al. (36) did not find changes in Na+-dependent glucose transport in BBM vesicles isolated from human duodenal biopsies.

Results from experiments using rodent models of aging also demonstrate conflicting results. Several studies (6, 10, 20) show reductions in D-glucose absorption in aged rats. However, a normal or increased absorptive capacity along the length of the small intestine was found in a study using everted intestinal segments from old vs. young rats (5). Results from studies in mice do not offer conclusive results on the effect of aging on nutrient absorption. Ferraris et al. (8) showed a reduction in uptake and site density of SGLT1 in aged mice. This is in contrast to the findings of Thompson et al. (27), who showed an increase in intestinal glucose uptake in aged mice (27).

Discrepancies in the results from human, rat, and mouse studies may be due to the differences in the methodologies used. Whereas some investigators studied uptake using BBM vesicles (6, 10, 20, 34, 36), others used everted intestinal rings (5, 8, 27). The method of expressing results is also important, as well. Most studies have expressed uptake based on intestinal weight, and therefore failed to take into account any potential age- and diet-associated changes in mucosal weight or surface area. The strain of the animals used, the age of the animals, and the site of the intestine used may also differ between studies and may explain the variability in the results.

Uptake of fructose has been studied in aging mice. Ferraris et al. (8) showed that D-fructose uptake per milligram of tissue was higher in the jejunum of young compared with old animals. The effect of diet on D-fructose uptake in aging animals was also examined (9). D-fructose uptake was also found to be higher in younger animals fed a high-carbohydrate, low-protein diet. No differences in D-fructose uptake were seen between young and old animals fed a no-carbohydrate, high-protein diet.

Uptake of fructose across the BBM is mediated by sodium-independent glucose and fructose transporters (GLUT)5 (32). The transport of these sugars out of the enterocyte across the basolateral membrane (BLM) occurs via the facilitative GLUT2 (32). In addition to its role as a BLM transporter, GLUT2 has recently been localized in the BBM where it has been suggested as contribing to the uptake of sugars into the enterocyte (12, 13, 18, 19).

Intestinal adaptation is a process by which the intestine changes both morphologically and/or functionally in response to alterations in environmental stimuli. Dietary manipulations have been shown to modify intestinal adaptation and nutrient absorption. Young animals fed a diet enriched in saturated fatty acids (SFA) have greater glucose uptake than do animals fed an isocaloric diet high in polyunsaturated fatty acids (PUFA) (29–31). Similarly, adult rats fed SFA show increases in fructose uptake compared with those fed PUFA (26).

The intestinal adaptive process may be compromised during the process of aging and may, thereby, contribute to malabsorption and malnutrition. Ferraris and Vinnekota (9) showed in aging rats that adaptive increases in uptake in response to a high carbohydrate diet were reduced in old compared with young animals, and the changes were limited to the more proximal regions of the intestine (9). In other studies (15, 16), after a 3-day starvation period, animals were refed for 1 day and enzyme-specific activities were measured. On refeeding, the specific activities of lactase, sucrase, and maltase demonstrated an exaggerated enzyme response in aged animals. It is not known how the aging rat responds to diets high in PUFA or SFA or whether modifications in dietary lipids could be used to alter the expected age-associated changes in nutrient absorption.

The objectives of this study were to determine 1) the effects of aging on the in vitro uptake of fructose in rats, 2) the effect of feeding SFA vs. PUFA, and 3) the molecular mechanisms of these age- and diet-associated changes.


    MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Animals

Principles for the care and use of laboratory animals, approved by the Canadian Council on Animal Care and the Council of the American Physiological Society, were observed in the conduct of this study. Thirty-six male Fischer 344 rats, ages 1, 9, and 24 mo were obtained from the National Institute of Aging colony and Harlan Laboratories. Pairs of rats were housed at a temperature of 21°C in a 12:12-h light-dark cycle. Water and food were supplied ad libitum.

Animals were fed standard Purina rat chow for 1 wk, and then fed one of two diets for a further 2 wk: 1) a semipurified diet containing 20% (wt/wt) fat and enriched with either SFA or 2) the same diet enriched with PUFA (Tables 1 and 2). There was a total of six animals in each age and diet combination. The isocaloric semipurified diets were nutritionally adequate, providing for all known essential nutrient requirements. Animal weights were recorded at weekly intervals.


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Table 1. Fatty acid composition of the semisynthetic diets

 

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Table 2. Macronutrient composition of the semisynthetic diets

 
Uptake Studies

Probe and marker compounds. The 14C-labeled D-fructose was supplied by Amersham Biosciences (Piscataway, NJ), and the unlabeled fructose by Sigma (St Louis, MO). Concentrations used were 8, 16, 32, and 64 mM. [3H]inulin was used as a nonabsorbable marker to correct for the adherent mucosal fluid volume.

Tissue preparation. The animals were killed by an intraperitoneal injection of Euthanyl (pentobarbital sodium, 240 mg/100 g body wt). The small intestine was removed from the ligament of Treitz to the ileocecal valve and rinsed with cold saline. This segment was then divided into thirds, with the proximal third considered to be the "proximal" small intestine, and the distal third considered to be the "distal" small intestine. The intestine was opened along its mesenteric border, and pieces of the proximal segment (jejunum) and the distal segment (ileum) were cut and mounted as flat sheets in the transport chambers. A 5-cm piece of each jejunal and ileal segment was gently scraped with a glass slide to determine the percentage of the intestinal wall comprised of mucosa. The chambers were placed in preincubation beakers containing oxygenated Krebs-bicarbonate buffer (pH 7.2) at 37°C, and tissue disks were preincubated for 15 min to allow the tissue to equilibrate at this temperature. The rate of fructose uptake was determined from the timed transfer of the transport chambers to the incubation beakers containing [3H]inulin and 14C-labeled D-fructose in oxygenated Krebs-bicarbonate (pH 7.2, 37°C). Preincubation and incubation chambers were mixed with circular magnetic bars at identical stirring rates precisely adjusted by using a strobe light. Stirring rates were reported as revolutions per minute. A stirring rate of 600 rpm was selected to achieve low effective resistance of the intestinal unstirred water layer (21, 37, 38).

Determination of uptake rates. After incubating the disks in labeled solutions for 6 min, the experiment was terminated by removing the chamber and rinsing the tissue in cold saline for ~5 s. Exposed mucosal tissue was then cut out of the chamber with a circular steel punch, placed on a glass slide, and dried overnight in an oven at 55°C. The dry weight of the tissue was determined, and the tissue was transferred to scintillation counting vials. The samples were saponified with 0.75 M NaOH, scintillation fluid was added, and radioactivity was determined by means of an external standardization technique to correct for variable quenching of the two isotopes (21).

Rates of uptake of fructose were determined as nanomoles per 100 milligrams tissue per minute, nanomoles per 100 milligrams mucosal tissue per minute, nanomoles per centimeter square serosal surface area per minute, and nanomoles per centimeter square mucosal surface area per minute. Because the relationship between uptake and fructose concentration was linear, to determine statistical differences, the slopes of the lines were calculated and compared.

Morphology, mRNA, and Protein Analysis

Tissue preparation. An additional 24 animals (4 in each of the 6 age/diet groups) were raised and killed similarly as for the uptake studies. A 5-cm portion from each of the proximal jejunum and distal ileum was rinsed, quickly harvested, snap frozen in liquid nitrogen, and stored at –80°C for later mRNA isolation. Mucosal scrapings were harvested from the remaining proximal and distal small intestine, snap frozen in liquid nitrogen, and stored at –80°C for subsequent isolation of BBM and BLM. For morphology and immunohistochemistry analysis, two 1-cm pieces of proximal and distal small intestine were fixed in 10% formalin.

Morphological measurements. Morphometric data were obtained from hematoxylin- and eosin-stained paraffin sections. Vertical cross sections of the intestinal tissue were used to obtain measurements of villous height, villous width at one-half villous height, villous bottom width, and crypt depth. The tissue block was then reoriented 90° laterally, and sections were taken to determine the villous depth at one-half villous height. Magnification was calibrated by using a micrometer. Mucosal surface area was calculated as described previously (31). The number of villi per millimeter of serosal length was measured in longitudinal and horizontal cross sections and then multiplied together to obtain the number of villi per square millimeter serosa. When this villous density was multiplied by villous surface area, the result was the mucosal surface area, expressed as square millimeters per square millimeter of serosa. At least 10 villi were assessed per section. The two following formulas were used (31). Villous surface area (µm2/villus)= (2 x M x H) + (2 x M – A) x D + (2 x D) x [(A – M)2 + (H)2]0.5 x 1,000, where H is villous height, M is villous width at one-half height, A is villous bottom width, and D is villous thickness. Mucosal surface area (mm2/mm2 serosa) = number of villi/mm2 serosa x villous surface area (µm2/villus)/1,000.

Messenger RNA abundance. Intestinal pieces were homogenized in a denaturing solution containing guanidinium thiocyanate, using the Fast Prep cell disruptor (Savant Instruments, Holbrook, NY) After the addition of 2 M sodium acetate, a phenol chloroform extraction was performed. The upper aqueous phase containing the RNA was collected. RNA was precipitated with isopropanol overnight at –80°C, with a final wash with 70% ethanol. Concentration and purity of RNA was determined by spectrophotometry at 260 and 280 nm. Samples were stored at –80°C until use for Northern blot analysis.

Fifteen micrograms of total RNA was fractionated by agarose gel electrophoresis and transferred to nylon membranes by capillary diffusion. RNA was fixed to the membranes by baking at 80°C for 2 h. Northern blot analysis was performed used the DIG Easy Hyb method, according to the manufacturer’s protocol (Roche Diagnostics).

GLUT2 and GLUT5 plasmids were kindly donated by Dr. G. I. Bell, of the Howard Hughes Medical Institute, University of Chicago (Chicago, IL).

Density of the mRNA bands was determined by transmittance densitometry (imaging densitometer model GS-670; Bio-Rad, Hercules, CA). Quantification of the 28S ribosomal RNA units from the ethidium bromide-stained membranes was used to account for loading discrepancies.

Protein analysis. BBMs, BLMs, and enteroctye cytosol were isolated from rat intestinal mucosal scrapings by differential centrifugation and Ca2+ precipitation (22, 24, 25). Aliquots were stored at –80°C. For Western blot analysis, GLUT5 was measured in the BBM and GLUT2 was measured in the BLM. Each group was represented in each probed blot. Results were expressed as a percentage of the total density on each blot to normalize for differences in development. We previously determined that 15 µg of protein was sufficient for detecting a signal without saturating the detection system.

The protein concentration of the samples was determined by using the Bio-Rad Protein Assay (Life Science Group, Richmond, CA). Proteins were separated by SDS-PAGE and were transferred to nitrocellulose membranes. Transfer efficiency was verified by 3-hydroxy-4-[2-sulfo-4-(4-sulfophenylazo)phenylazo]-2,7-naphthalenedisulfonic acid staining of membranes and by Coomassie blue staining of gels. Membranes were blocked by incubation overnight in bovine lacto transfer technique optimizer containing 5% wt/vol dry milk in TTBS (0.5% Tween 20, 30 mM Tris, 150 mM NaCl).

Membranes were washed in TTBS (3 x 10 min each) and subsequently probed with specific rabbit anti-rat antibodies to GLUT5 (Chemicon International, Temecula, CA) and GLUT2 (Biogenesis, Poole, England) for 2 h at room temperature. The antibodies were prepared in 5% dry milk in TTBS at a dilution of 1:500. Membranes were subsequently washed with TTBS to remove the residual unbound primary antibody and were then incubated for 1 h with goat anti-rabbit antibody (1:20,000 in 2% dry milk in TTBS) conjugated with horseradish peroxidase (HRP) (Pierce Biotechnology, Rockford, IL).

Membranes were washed again in TTBS to remove residual secondary antibody, and were briefly incubated with Supersignal chemiluminescent-HRP substrate (Pierce Biotechnology). Membranes were exposed to X-OMAT AR films (Medtec, Burnaby, BC, Canada) and the relative band densities were determined by transmittance densitometry using Bio-Rad model GS-670 imaging densitometer.

Immunohistochemistry. Jejunal and ileal tissue was embedded in paraffin, and 4- to 5-µm sections were mounted on glass slides, dewaxed in xylene, and hydrated after incubation in a series of ethanol incubations. Slides were then incubated in solution of 20–50% H2O2/80% methanol for 6 min, rinsed in tap water, and counterstained with Harris hematoxylin (10 s). Slides were then air-dried and the tissue was encircled with hydrophobic slide marker (PAP pen; BioGenex, San Ramon, CA). After rehydration in PBS, the slides were incubated for 15 min in blocking reagent (20% normal goat serum) followed by primary antibody to GLUT5 for 30 min. Slides were then washed in PBS and incubated in LINK and LABEL according to the manufacturer’s protocol. Solutions were subsequently incubated for 5 min in diaminobenzidine, rinsed in water, dehydrated in absolute ethanol, and cleared in xylene. Negative controls were processed on the same slide in an identical manner, excluding the incubation with the primary antibody. A Leitz Orthoplan Universal Largefield microscope and a Leitz Vario Orthomat 2 automatic microscope camera were used to photograph the slides. Chromagen staining was quantified by using a Pharmacia LKB-Imagemaster DTS densitometer and Pharmacia Imagemaster 1D (version 1.0) software (Amersham Biosciences). Four villi per animal were quantified, and the results were normalized to the negative control values.

Expression of Results

Results were expressed as means ± SE. Sigmastat (version 1.0) was used to perform two-way ANOVA (P < 0.05) to determine the effect of age and diet and any age-diet interactions. Individual differences between ages were determined by using a Student-Newman-Keuls multiple-range test.


    RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
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 DISCUSSION
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Animal Characteristics

The rate of body weight change (g/day) fell among 1, 9, and 24 mo in rats fed SFA or PUFA (Fig. 1). At each age, the body weight decline was greater with SFA than with PUFA. Food intake was not influenced by the age of the rats, regardless of whether they were fed SFA or PUFA (data not shown).



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Fig. 1. Effect of age and dietary lipids on body weight change. Values are means ± SE. Different letters indicate a significant effect of age. *Significant diet effect; statistically significant differences were determined by using a 2-way ANOVA (P < 0.05). SFA, saturated fatty acids; PUFA, polyunsaturated fatty acids.

 
Age and diet had no effect on jejunal or ileal tissue weight, mucosal weight, or percentage of the intestinal wall comprised of mucosa (Table 3).


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Table 3. Effect of age and diet on intestinal weight

 
There were no differences in the mean values of the heights of the villi of the jejunum or ileum of rats aged 1, 9, or 24 mo (data not shown). In animals fed SFA, the jejunal and ileal mucosal surface areas were lower at 9 and 24 mo, compared with 1 mo (Fig. 2). In those fed PUFA, both the jejunal and ileal mucosal surface area were lower at 24 compared with 1 and 9 mo. In the jejunum, diet had no effect on mucosal surface area at 1, 9, or 24 mo (Fig. 2). In the ileum, the mucosal surface area was increased in 1-mo-old animals fed SFA compared with PUFA. Diet did not affect ileal mucosal surface area at 9 or 24 mo of age.



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Fig. 2. Effect of age and dietary lipids on the mucosal surface area of the small intestine. Values are means ± SE. Different letters indicate a significant effect of age. *Significant age-diet interaction; statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 
The age-associated decline in jejunal mucosal surface area occurred between 9 and 24 mo in rats fed PUFA, and between 1 and 9 mo in those fed SFA. Thus feeding PUFA slowed the age-associated decline in the mucosal surface area of the jejunum and ileum. Thus the surface area of the mucosa had to be taken into account when expressing the rate of uptake of fructose.

Fructose Absorption

When fructose uptake was expressed on the basis of the weight of the wall of the intestine (nmol·100 mg–1·min–1), neither age nor diet significantly affected fructose uptake in the jejunum or ileum (Fig. 3).



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Fig. 3. Effect of age and dietary lipids on D-fructose uptake expressed on the basis of intestinal weight. Values are means ± SE. No statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 
Similarly, when fructose uptake was expressed on the basis of the weight of the mucosa (nmol·100 mg mucosa–1·min–1), neither age nor diet significantly affected fructose uptake in the jejunum or ileum (Fig. 4).



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Fig. 4. Effect of age and dietary lipids on the uptake of D-fructose expressed on the basis of mucosal weight. Values are means ± SE. No statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 
Similarly, when fructose uptake was expressed on the basis of the serosal surface area (nmol·cm–2·min–1), neither age nor diet significantly affected fructose uptake in the jejunum or ileum (Fig. 5).



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Fig. 5. Effect of age and dietary lipids on the uptake of D-fructose expressed on the basis of serosal surface area. Values are means ± SE. No statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 
When uptake was expressed on the basis of mucosal surface area in animals fed SFA, both the jejunal and ileal uptake of fructose was greater at 9 or 24 mo compared with 1 mo (Fig. 6). In animals fed PUFA, both the jejunal and ileal uptake of fructose was higher at 24 than at 9 or at 1 mo. Diet influenced ileal fructose uptake. At 9 mo, uptake was higher in animals fed SFA compared with animals fed PUFA. In contrast, at 24 mo, uptake was higher in animals fed a PUFA diet compared with animals fed SFA.



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Fig. 6. Effect of age and dietary lipids on the uptake of D-fructose expressed on the basis of mucosal surface area. Values are means ± SE. Different letters indicate a significant effect of age. *Significant age-diet interaction; statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 
Transporter Protein Abundance and Immunohistochemistry

In animals fed SFA or PUFA, the jejunal and ileal abundance of GLUT5 determined by Western blot analysis was similar at 1, 9, and 24 mo (Fig. 7). Diet did not significantly affect jejunal or ileal GLUT5 protein abundance.



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Fig. 7. Effect of age and dietary lipids on sodium-independent glucose and fructose transporter (GLUT)5 protein abundance. Values are means ± SE. No statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 
Jejunal and ileal villi were divided into five equal sections starting from the tip of the villi down to the crypt region. The abundance of GLUT5 protein was found to be evenly distributed along the crypt-villus axis (data not shown). The jejunal abundance of GLUT5 as determined by immunohistochemistry was not influenced by age or diet (Fig. 8). In contrast, in SFA-fed animals, ileal GLUT5 was reduced at 9 and 24 mo compared with 1 mo (Fig. 9). In PUFA-fed animals, the reduction in GLUT5 was evident only at 24 mo compared with 1- or 9-mo-old animals. At both 1 and 24 mo, SFA-fed animals had increased GLUT5 abundance compared with PUFA-fed animals. There was a significant interaction between age and diet (P = 0.000956). The abundance of GLUT2 in the BLM was similar at 1, 9, and 24 mo in animals fed SFA or PUFA (data not shown).



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Fig. 8. Effect of age and dietary lipids on GLUT5 abundance as determined by immunohistochemistry. Values are means ± SE. Different letters indicate a significant effect of age. *Significant age-diet interaction; statistically significant differences were determined by using a 2-way ANOVA (P < 0.05).

 


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Fig. 9. GLUT5 immunohistochemistry on ileal sections from 1-mo-old rats fed PUFA (A); 9-mo-old rats fed PUFA (B); 24-mo-old rats fed PUFA (C); 1-mo-old rats fed SFA (D); 9-mo-old rats fed SFA (E); and 24-mo-old rats fed SFA (F).

 
Transporter mRNA Expression

The expression of GLUT5 as well as GLUT2 in the jejunum was similar at 1, 9, and 24 mo in animals fed SFA or PUFA (data not shown).


    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
Variations in dietary lipids influence the changes in intestinal surface area seen with aging. Animals fed SFA exhibited a reduction in surface area at both 9 and 24 mo (Fig. 2). This decrease in jejunal surface area seen with aging is delayed by feeding PUFA, because animals fed this diet showed reductions in surface area only at 24 mo of age. In addition, feeding PUFA prevents the age-associated decline in ileal mucosal surface area seen with SFA. This signifies that the intestine of the old rats remains capable of adapting its morphology in response to dietary lipid manipulations. This observation also stresses the point that failure to take into account the animal’s diet may lead to errors in the interpretation of the effects of aging on the morphology of the intestine. Finally, the rate of decline in jejunal and ileal mucosal surface area seen in rats fed SFA can be slowed by feeding PUFA.

The simplest way of expressing the rate of in vitro uptake of nutrients is on the basis of the weight of the full thickness of the wall of the intestine. However, if an experimental manipulation alters the weight of the intestine or the mucosal surface area, then there may be variations in the rate of nutrient uptake that are understandable in light of there simply being more mucosal tissue or a greater surface area. For this reason, where there are treatment-associated variations in mucosal mass or the surface area of the villous membrane, as was the case in this study (Table 1 and Fig. 2), then it is more appropriate to express uptake on the basis of mass of the transporting mucosal tissue or the mucosal surface area. Clearly, no significant differences in fructose uptake are seen when expressed on the basis of intestinal (Fig. 3), mucosal (Fig. 4), or serosal weight (Fig. 5). However, significant increases in jejunal and ileal fructose uptake with both the SFA and PUFA diet are seen when expressed on the basis of mucosal surface area (Fig. 6). Thus the ability to show an effect of age or diet on fructose uptake depends on the way in which the rate of uptake is expressed.

Previous studies (26) have shown that fructose uptake is increased in young rats fed SFA compared with PUFA. Our results confirm these findings in 9-mo-old animals. However, in the older 24-mo-old animals, the diet effect appears to be reversed, because fructose uptake is increased with PUFA compared with SFA. This suggests that dietary lipids affect the absorption of nutrients differently depending on the age of the animal. This finding is in agreement with other work (9, 15, 16), which shows a reduction in adaptive capabilities in response to diet with aging. Indeed, the results of this study demonstrate that the ability of SFA to increase fructose uptake is impaired with aging. Ferraris and Vinnekota (9) noted that the effect of dietary changes in aged mice was limited to the proximal small intestine, but the effect of dietary lipids on glucose uptake in aged rats occurs in both the jejunum and ileum (Figs. 36; Ref. 9). Thus aging modifies the adaptability of the intestine in response to dietary manipulation. Clearly, the influence of a manipulation that results in intestinal adaptation in young rats does not necessarily apply in older animals.

Despite the decreases in the surface area of the intestine seen with aging, fructose uptake, when expressed on the basis of mucosal surface area, is increased with age in animals fed PUFA or SFA (Fig. 6). The aged intestine is therefore able to maintain a high absorptive capability, despite a reduction in absorptive surface area. However, changes in uptake seen with aging or with diet are not solely explained by alterations in mRNA and protein abundance of the fructose transporters. PUFA or SFA, however, show increases in fructose uptake without concomitant changes in GLUT5 abundance. In fact, in the ileum, reductions in GLUT5 abundance as determined by immunohistochemistry are in contrast to the increases in uptake seen with aging.

In models of diabetes, a "recruitment" of transporters in the lower part of the villi results in active transport occurring in this area and a resultant increase in glucose transport (4). Because most intestinal glucose uptake occurs in the upper third of the villi (28), a redistribution of GLUT5 to this area could explain altered uptake. In fact, GLUT5 was evenly distributed along the crypt-villous axis (data not shown), and therefore a change in the distribution of GLUT5 does not explain altered uptake.

One could speculate that changes in the fluidity of the BBM as a result of aging could also explain the apparent uncoupling of transport to both RNA abundance and protein abundance. There are reductions in the membrane fluidity of the BBM isolated from 117-wk-old Fischer 344 rats compared with younger animals (3). Similarly, the fluorescence polarization technique used by Wahnon et al. (35) showed reductions in membrane fluidity in 19-mo-old rats compared with 1- and 9-mo-old rats (35). Indeed, a study done using chickens demonstrated that reductions in membrane fluidity, as a result of changes in BBM lipid content, may be involved in the decrease in D-glucose uptake observed during posthatching development (33). With aging, declines in membrane fluidity are associated with increases in uptake (23). Alterations in the fluidity of the BBM as a result of dietary lipid manipulations might explain the uncoupling of transport to RNA abundance and protein abundance. Altering the fatty acid composition of the diet results in changes in the phospholipid content of the BBM of enterocytes (17). It is reasonable to speculate that changing the dietary lipids may have altered BBM fluidity and therefore GLUT5 function in the older animals. We may further speculate that transporters such as GLUT2 or GLUT5 may reside in specialized lipid rafts, and local changes in membrane fatty acids may be responsible for alterations in function. Further research is needed to fully characterize the effects of changes in raft-associated membrane lipids on intestinal nutrient transport.

It has been proposed that GLUT2 is present in the BBM, as well as the BLM, and transports glucose and fructose into the cell via facilitated diffusion (1, 2, 1114, 18, 19). The changes in uptake seen in this study may therefore be paralleled by increases in BBM GLUT2 protein. However, GLUT2 appears to only play a significant role in sugar uptake in the presence of high luminal sugar concentrations, such as those found after a high sugar meal or after a sugar bolus (11–14, 18, 19). In this study, we did not detect any GLUT2 protein in the BBM (data not shown). This is not surprising, because these animals were not fed diets high in sugar nor were they given a sugar bolus. Therefore, we do not believe that GLUT2 in the BBM played a significant role in intestinal sugar uptake in this model and is not likely to explain the changes in uptake that were observed.

In conclusion, despite age-related reductions in mucosal surface area, intestinal fructose uptake per unit mucosal surface area increases with age. Changes in fructose uptake were not paralleled by changes in BBM GLUT5 or BLM GLUT2 protein abundance. The intestinal adaptive response to PUFA- or SFA-enriched diets is also influenced by aging, suggesting that the results obtained from studies of intestinal adaptation in young populations cannot necessarily be generalized to older populations. Further studies are required to determine the potential signals involved in the age-related changes in intestinal fructose uptake.


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Dr. Gary Wild is a senior clinician scientist supported by the Fonds de la Recherche en Sante du Quebec.


    ACKNOWLEDGMENTS
 
We acknowledge Elizabeth Wierzbicki and Terri Canuel for technical assistance and Dr. Monika Keelan for invaluable advice regarding uptake calculations. We also acknowledge the hard work of Arnaud Brulaire, Stephanie Gabet, and Marion Garel.


    FOOTNOTES
 

Address for reprint requests and other correspondence: A. Thomson, Division of Gastroenterology, Univ. of Alberta, 205 College Plaza, 8215 112 St. Edmonton, AB T6G 2C8, Canada (E-mail: alan.thomson{at}ualberta.ca)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


    REFERENCES
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 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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