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Effect of diet containing phytate and phytase on the activity and messenger ribonucleic acid expression of carbohydrase and transporter in chickens

N. Liu^*, Y. J. Ru, F. D. Li^*,1 and A. J. Cowieson

^* Faculty of Animal Science and Technology, Gansu Agricultural University, Lanzhou, China 730070; and Danisco Animal Nutrition, Science Park III, Singapore, 117525; and Danisco Animal Nutrition, Marlborough, Wiltshire, SN8 1XN, United Kingdom

	Abstract

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The effect of dietary phytate and phytase on carbohydrase activity and hexose transport was investigated in broiler chickens. Diets containing phytate P (2.2 or 4.4 g/kg) with different phytase dose rates (0, 500, or 1,000 phytase units/kg) were fed to 504 female Cobb chicks for 3 wk. Diets containing high phytate concentrations depressed (P < 0.05) BW and G:F, whereas phytase supplementation improved (P < 0.05) the performance of birds. In the duodenum, phytate decreased (P < 0.05) the activities of disaccharidases, Na⁺K⁺-ATPase, and glucose concentrations by 5 to 11%, but phytase enhanced (P < 0.05) the concentrations of amylase, sucrase, maltase, Na⁺K⁺-ATPase, and glucose by 5 to 30%. In the jejunum, phytate decreased (P < 0.05) the concentrations of amylase, sucrase, Na⁺K⁺-ATPase, and glucose by 10 to 22%, and phytase alleviated the negative effect of phytate on the above variables. Ingestion of diets containing phytate also decreased (P < 0.05) serum amylase activity and glucose concentration, and phytase enhanced (P < 0.05) serum concentrations of amylase, sucrase, maltase, Na⁺K⁺-ATPase, and glucose. There were also interactions (P < 0.05) between phytate and phytase on the concentrations of serum amylase, duodenal amylase, sucrase, and jejunal glucose. Enzymatic analysis at a molecular level showed that neither phytate nor phytase influenced the mRNA expression of sucrase-isomaltase in the small intestine. Also, the investigation into the sodium glucose cotransporter gene may challenge the mechanism by which phytate interferes with glucose utilization, as partly indicated by bird performance, and transmembrane transport because diets containing increased phytate upregulated (P < 0.05) the mRNA expression of the sodium glucose cotransporter gene in duodenum and did not influence it in the jejunum. These results indicate that phytate can impair endogenous carbohydrase activity and digestive competence, and phytase can ameliorate these effects for chickens.

Key Words: adenosine triphosphatase • carbohydrase • chicken • phytase • phytate • sodium-glucose cotransporter-1

	INTRODUCTION

Top Abstract INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

 
The effects of dietary phytate and phytase on the performance and nutrient digestibility of intensively farmed livestock have received considerable attention in the scientific literature. Recent evidence demonstrated that the ingestion of phytate can have substantial adverse effects on endogenous secretion and energy utilization in chickens (Cowieson et al., 2004, 2006; Ravindran et al., 2006; Cowieson and Ravindran, 2007; Liu et al., 2008). Although increased endogenous AA losses with the ingestion of phytate will influence the energy value of poultry diets, it may be of greater direct significance that phytate can reduce amylase activity and starch digestion in vitro (Cawley and Mitchell, 1968; Knuckles and Betschart, 1987) and in vivo (Dilworth et al., 2004), and modify the activity of intestinal Na⁺K⁺-ATPase (Dilworth et al., 2005). It is reported that elevated dietary sugar concentrations induced increases in the expression of sucrase-isomaltase (Miyamoto et al., 1993) and sodium glucose transporter-1 (SGLT-1; Kishi et al., 1999) in the rat jejunum. The evidence that dietary phytate interferes with sugar digestion and blood glucose in rats (Dilworth et al., 2004) and mice (Lee et al., 2006) leads to the hypothesis that antinutritional factors capable of altering nutrient utilization might regulate associated gene expression. Further, it has been recently established that phytate and phytase may influence the partitioning of Na, perhaps mediated via hypersecretion of NaHCO₃ in response to increased concentrations of dietary phytate, which are partially ameliorated by phytase (Cowieson et al., 2004; Ravindran et al., 2008). Given that these extra-phosphoric effects are not well understood, the current study was designed to assess the effect of dietary phytate and phytase on the activity of amylase, sucrase, maltase and Na⁺K⁺-ATPase, glucose, and the mRNA expression of sucrase-isomaltase and SGLT-1 in the intestine and serum of chickens.

	MATERIALS AND METHODS

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All procedures were approved by Gansu Agricultural University Institutional Animal Care and Use Committee.

Treatments

The experiment used a 2 x 3 factorial arrangement of treatments based on corn-soybean meal diets with 2 concentrations of phytate P (2.2 or 4.4 g/kg of diet, as-fed basis), and the addition of Escherichia coli-derived phytase (Phyzyme XP, Danisco Animal Nutrition, Wilt-shire, UK) at 0, 500, or 1,000 phytase units (FTU)/kg diet (as-fed basis). The low and high phytate diets contained the same nutrient specification, differing only in the concentration of dietary phytate P (Table 1). Rice bran and corn germ meal were used to regulate phytate P concentrations.

A total of 504 female Cobb chicks (1 d of age) were randomly allocated into 6 treatments, each of which had 6 replicates of 14 chicks per replicate. All chicks were raised in 3-layered cages and given ad libitum access to diets and water, continuous lighting, and controlled ventilation. Temperature was maintained at 32°C for the first 5 d and then gradually reduced according to normal management practices until a temperature of 25°C was achieved at d 21. Body weight and diet consumed per cage basis were recorded weekly to monitor the performance. At 7 d of age, the chickens were inoculated with Newcastle disease virus and infectious bronchitis vaccine (Yikang Biological Products Corp., Liaoyang, China) by intranasal and intraocular administration.

Sampling

At 21 d of age, 3 chickens with average BW from each replicate were selected. Before sampling, the birds were offered diets and water as usual without fasting, considering that phytate with diet ingestion may influence the activity of digestive enzymes. Blood was obtained from each chick by cardiac puncture, and serum was prepared as described previously (Liu et al., 2008) and stored at –20°C. Birds were then killed by cervical dislocation. The duodenum and jejunum were excised and flushed with 0 to 4°C NaCl solution (0.15 M), and fragments (1 cm) at the middle of duodenum or jejunum were collected, combined by replicate, frozen in liquid nitrogen, and kept at –70°C for mRNA assay. The lumen of the duodenum or jejunum was cut longitudinally to expose brush border cells. Mucosa was gently scraped off with a glass microscope slide and pooled by replicate. Each mucosal sample was homogenized (1:4, wt/vol) with 0 to 4°C NaCl solution (0.15 M) and then centrifuged at 8,000 x g for 3 min at 4°C; the supernatant was immediately frozen in liquid N and kept at –70°C until analysis. The pooling of samples (except for serum samples) by replicate is a means of normalizing the individual differences in enzyme activity and gene expression analysis.

Protein Concentration

After thawing, an aliquot mucosal supernatant was taken for protein determination using the method of Folin and Ciocalteu’s phenol (No. F9252, Sigma-Aldrich, Bellefonte, PA; Lowry et al., 1951). Working standards were prepared using dilute solutions of BSA (No. P0914, Sigma-Aldrich). The absorbance was read at 545 nm on a semi-auto biochemical analyzer GF-D600 (Nanjing T-Bota Scietech Instruments and Equipment, Nanjing, China). The protein concentration was expressed as milligrams per milliliter of mucosa supernatant.

Amylase Activity Assay

The activity of amylase (EC 3.2.1.1) was determined by the iodine-starch method (Gitlitz and Frings, 1976). After 0.5 mL of 0.4 mg/mL of soluble starch substrate (No. S9745, Sigma-Aldrich) was preheated for 5 min at 37°C; 10 mL of mucosal supernatant was added. The mixture was incubated for 30 min at 37°C, and 0.5 mL of 10 mM iodine solution and 3.0 mL distilled water were added and mixed for reading the absorbance value at 660 nm. One amylase unit is defined as the amount of enzyme that was required to hydrolyze 10 mg of starch in 30 min at 37°C. Amylase activity in serum or mucosa was expressed as micromoles per milliliter of serum or per milligram of mucosal protein.

Disaccharidase Activity Assay

The activities of sucrase (EC 3.2.1.48) and maltase (EC 3.2.1.20) in the serum and intestine were detected based on the method by Dahlqvist (1984). Briefly, 0.1 mL of serum or mucosal supernatant incubated with 0.1 mL of 56 mM disaccharide substrate (sucrose or maltose) in 25 mM malate buffer at pH 6.4 for 30 min at 37°C. The disaccharidase activity is then stopped by the addition of Tris. The glucose liberated is measured with a glucose oxidase reagent by a Glucose Detection Kit (Shanghai Rongsheng Biotech, Shanghai, China). Enzyme activity was expressed as units per milliliter of serum or milligrams of protein. Glucose concentration in serum or mucosa was expressed as micromoles per milliliter of serum or per milligram of mucosal protein.

Na⁺K⁺-ATPase Activity Assay

The method of measuring Na⁺K⁺-ATPase (EC 3.6.1.3) was identical to the description by Whealty and Hentry (1987). Briefly, phosphate liberated from ATP-Na₂ (No. A7699, Sigma-Aldrich) by each sample was measured in 2 media: medium I (all ATPases system) had optimum concentrations of all ions, containing 0.1 mL of enzyme sample, 0.2 mL of buffer solution (200 mM NaCl, 40 mM KCl; 12 mM MgCl₂, and 50 mM Tris-HCl, pH 7.8); medium II (Na⁺K⁺-ATPase restrained system) lacked K and contained 0.1 mL of enzyme sample and 0.2 mL of buffer solution (240 mM NaCl, 12 mM MgCl₂, 50 mM Tris-HCl pH 7.8, and 10 mM ouabain, which was added to prevent stimulation of Na⁺K⁺-ATPase by K⁺ present in the samples).

After incubation at 30°C for 5 min, the 2 media 0.1-mL solutions (60 mM ATP-Na₂; 50 mM Tris-HCl pH 7.8) were added and then were incubated again in the 30°C water bath for 30 min. The reaction was stopped by adding 0.4 mL of 30% cold trichloroacetic acid. Phosphorus concentrations of the 2 media were measured using the method of phosphomolybdic blue (Fiske and Subbarow, 1925). The activity of Na⁺K⁺-ATPase was calculated as the difference between phosphates liberated by each homogenate in the 2 media and was expressed as micromoles of phosphates per milligram homogenate protein or per milliliter of serum per hour.

Messenger RNA Quantification

The mRNA expressions of sucrase-isomaltase (EC 3.2.1.48) and SGLT-1 in duodenum and jejunum were analyzed by real-time PCR. In brief, total RNA from pools of 3 chickens per replicate was prepared using RNAiso reagent (TaKaRa, Dalian, China). After removal of potential contaminating DNA using DNase I (Ambion, Austin, TX), the extracted RNA pellets were reverse transcribed using M-MLV RTase cDNA Synthesis Kit (TaKaRa). Primers were designed according to the reported sequences (Table 2). Primer specificity was checked by systematic sequencing of PCR products. Quantification of different mRNA was performed on iCycler iQ Real Time PCR Detection System (Bio-Rad, Hercules, CA) using SYBR Premix Ex Taq (Ta-KaRa). The quantification of mRNA expression was normalized to β-actin.

Statistical Analysis

The experiment was a 2 x 3 factorial arrangement (n = 6 for mucosa and gene expression samples, n = 18 for serum samples) with dietary phytate P and phytase dose rate being the main factors. A general linear model was used to assess the effect of dietary phytate and phytase and their interaction (SAS Inst. Inc., Cary, NC). Differences of initial copies of target genes normalized to β-actin copies were separated using Duncan’s multiple-range test at P < 0.05 level of significance. Values in the tables were means and pooled SEM.

	RESULTS

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The chickens were healthy throughout the experiment with a mortality of <2% that was unrelated to dietary treatment. Supplementation with phytase improved (P < 0.05) feed intake, BW, and G:F of birds (Table 3), but no differences were found between phytase at 500 and 1,000 FTU/kg of feed. Birds fed the diet with a high phytate concentration had lighter BW and poorer G:F than those fed on the low phytate diets (P < 0.05). No interaction between phytate and phytase was detected on the performance variables of birds.

	DISCUSSION

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It has been reported that phytate is capable of depressing enzymatic digestion in vitro (Singh and Krikorian, 1982; Deshpande and Ceryan, 1984; Knuckles and Betschart, 1987; Knuckles, 1988) and in vivo (Dilworth et al., 2004). The underlying mechanisms by which phytate inhibits the activity of digestive enzyme in the gastrointestinal tract of animals include chelation with co-factors required for optimum enzyme activity, binding the digestion products, as well as forming phytate-protein complexes at pH below the isoelectric point of proteins (Cawley and Mitchell, 1968; Katayama, 1997). In this study, birds fed the diets containing high phytate concentrations had a reduced activity of endogenous carbohydrase, indicating that a decrease in intestinal enzyme activity by phytate may result in more nutrients passing through the digestive system unabsorbed.

Alpha-amylase from exocrine pancreas or enterocytes enters the intestinal lumen in an active form to catalyze hydrolysis of 1 to 4 glycosidic bonds in starch and dextrin. The end products of starch digestion by amylase are maltose, maltotriose, and -limit dextrin. The decrease in the jejunal activity of amylase caused by dietary phytate in this study indicated that the digestibility of starch may be depressed. Similar results have been reported that phytate significantly decreased the activity of amylase in the lower intestine in rats, but not for the proximal segment (Dilworth et al., 2004). Furthermore, amylase activity was significantly compromised by phytate in model systems (Deshpande and Ceryan, 1984; Knuckles and Betschart, 1987), which may be caused by the chelation of Ca, a necessary cofactor for the stability and optimal functioning of amylase in the intestine. Because phytic acid can decrease Ca balance in chickens (Simons et al., 1990), phytate may either impair amylase stability or activation, or regulate the process of digestion by binding to digestion products. Additionally, it is evident that amylase secretion and its concentration in the intestine and serum can be influenced by the dietary amount of carbohydrate and protein (Messiha and Watson, 1989), so in this study, dietary phytate and phytase modified nutritional status in the intestine, which may be in part responsible for the alteration in secretion function of acinar cells and subsequently enzymatic level of intestine and serum.

After the action of amylase, completion of starch digestion is catalyzed by saccharidases attached to the brush border of the small intestine. Maltase breaks down maltose and maltotriose to glucose, and isomaltose is hydrolyzed to glucose by isomaltase. In this study, the decreased maltase activity in the duodenum of birds fed the high phytate diets may indicate that liberation of glucose from dextrin is compromised by phytate ingestion. Further, the concentration of glucose in the intestine and serum was also decreased by dietary phytate, which may be a consequence of the inhibition of brush border enzyme activity in this study. Dilworth et al. (2004) reported that blood glucose concentrations in rats fed phytic acid extract from sweet potato or commercial phytic acid were numerically but not statistically reduced. Beneficially, in this study, phytase enhanced the blood glucose concentrations by 32%, for which there is a similar report that phytase significantly increased the blood glucose concentration by 21% in commercial pigs (Kies et al., 2005).

The inhibition of sucrase activity by phytate in both the duodenum and jejunum in this study was of interest. Sucrose is the most important dietary disaccharide and extensively exists in vegetable foods (Somogyi and Trautner, 1974). Brush border sucrase is involved in the hydrolysis of sucrose to fructose and glucose. Katayama (1997) reported that dietary phytate can protect sucrose-fed rats against an accumulation of hepatic lipids due to the depression in the conversion of sugar to lipid in the body. Also, Onomi et al. (2004) reported that rats fed a high-sucrose diet with 0.02 to 10% Na phytate had reduced formulation of hepatic and serum lipid. However, Dilworth et al. (2005) reported that there was no significant change in the activities of intestinal disaccharidases in Wistar rats fed diets enriched with phytic acid, though dietary phytate significantly reduced blood glucose concentrations.

Phytate contains a total of 12 dissociable protons with pK_a values from 1.5 to around 10 (Angel et al., 2002). The least pK_a values correspond to the dissociation of the first proton from each of the 6 phosphate groups with pK_a value increasing as each subsequent proton dissociates. The low pK_a of phytate, especially in a fully protonated form, as well as the release of phosphate anions by phytase, may be contributory factors in the increased secretion of Na into the gastrointestinal tract as a buffer. Cowieson et al. (2004) speculated that phytate induced the movement of Na into the gut lumen to buffer this polyanionic molecule. Additionally, the entry of acidic chyme into the duodenum stimulates Ca, Na, and K secretion from the gallbladder, pancreas, and mucosa (Ruckebusch et al., 1991). It seems feasible that, if phytate can increase endogenous Na secretion into the lumen, Na may become limiting for nutrient transport functions in instances of marginal Na provision. It is possible, therefore, that phytate may decrease acid-base homeostasis and compromise Na-dependent transport mechanisms involved in the intestinal uptake of glucose as well as AA (Gal-Garber et al., 2003; Ravindran et al., 2008). Indeed, in this study the effect of phytate on the activity of Na-dependent transporters was particularly marked. Absorption processes of nutrients in the intestine are mainly driven by Na⁺K⁺-ATPase that generates a Na⁺ and K⁺ concentration gradient and electric potential difference, allowing the absorption of various molecules (Schultz et al., 1974; Mandel and Balaban, 1981; McBride and Kelly, 1990). Increased Na⁺K⁺-ATPase activity has been shown to improve glucose translocation across the cell membrane by creating an inward Na⁺ gradient (Kies et al., 2005). It is reported that the activity of Na⁺K⁺-ATPase in lower intestine was significantly reduced in Wistar rats fed diets with added phytic acid compared with controls (Dilworth et al., 2005). Apart from the effect of Na, the lack of other electrolytes including P and K resulted from phytate mineral chelation or diet-deficient P necessary to promote energy synthesis may result in a decrease in the bioactivity of Na⁺K⁺-ATPase.

Glucose, as the major monosaccharide available for absorption from most practical diets of farm animals, is subjected to carrier-mediated transfer through the brush border membrane by SGLT-1 (Wright, 1993). It is reported that SGLT-1 could be regulated by dietary saccharides (Miyamoto et al., 1993) and proteins (Gilbert et al., 2008), but little is known about the regulation from dietary antinutritive factors. In this study, diets containing high phytate upregulated the mRNA concentrations of SGLT-1 in the duodenum, but this cannot explain the depression of phytate on blood glucose and final performance of birds, and the exact reason for this effect on the transcellular transport of glucose molecules should be investigated further. Theoretically, as with Na⁺K⁺-ATPase, in the scheme of SGLT-1, Na⁺ is the primary solute transported and glucose is a cosolute. In the absence of Na⁺, glucose cannot be transported. The disequilibrium of intracellular Na homeostasis caused by phytate in the intestine (Cowieson et al., 2004) may interfere with the Na⁺:glucose stoichiometric ratio, Na pump activity, and glucose transport efficiency via repartitioning of Na for NaHCO₃ production. Indeed, Ravindran et al. (2008) demonstrated that much of the beneficial effect of phytase could be removed by the addition of greater concentrations of Na to the diet (in the form of NaHCO₃). This, along with data presented herein, indicates that a Na matrix value for phytase may be justified as repartitioning of Na from pH balance functions to nutrient recovery should be captured in commercial feed formulation packages.

The investigation into the enzymatic activity at a molecular level showed both sucrase activity and sucrase-isomaltase mRNA peaked in the jejunum of chickens in this study, which was consistent with the report of Leeper and Henning (1990). However, the changes in the activity of membrane enzymes generated by dietary factors were not parallel to the mRNA expression of associated genes, indicating that dietary phytate may decrease the enzymatic activity through mechanisms of complex formation or inactivation rather than a quantitative change in secretion.

In summary, this study confirmed that phytate is capable of adversely modifying secretory and absorptive physiology in broiler chickens. The preponderance of data observed here indicates that dietary phytate is not only antinutritional but antiphysiological in the procurement of carbon and other nutrients for intensively farmed chickens. Importantly, the inclusion of phytase showed a pronounced effect on improving the activity of endogenous enzymes; thus, phytase may play a compensatory role in luminal and membrane digestion and recovery of nutrients.

¹ Corresponding author: lifd{at}gsau.edu.cn

Received for publication June 14, 2008. Accepted for publication August 4, 2008.

	LITERATURE CITED

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Angel, R., N. M. Tamim, T. J. Applegate, A. S. Dhandu, and L. E. Llestad. 2002. Phytic acid chemistry: Influence on phytin-phosphorus availability and phytase efficacy. J. Appl. Poult. Res. 11:471–480.[Abstract/Free Full Text]

Cawley, R. W., and T. A. Mitchell. 1968. Inhibition of wheat alpha-amylase by bran phytic acid. J. Sci. Food Agric. 19:106–108.

Cowieson, A. J., T. Acamovic, and M. R. Bedford. 2004. The effects of phytase and phytic acid on the loss of endogenous amino acids and minerals from chickens. Br. Poult. Sci. 45:101–108.[CrossRef][Medline]

Cowieson, A. J., T. Acamovic, and M. R. Bedford. 2006. Phytic acid and phytase: Implications for protein utilization by poultry. Poult. Sci. 85:878–885.[Abstract/Free Full Text]

Cowieson, A. J., and O. Adeola. 2005. Carbohydrases, protease, and phytase have an additive beneficial effect in nutritionally marginal diets for broiler chicks. Poult. Sci. 84:1860–1867.[Abstract/Free Full Text]

Cowieson, A. J., and V. Ravindran. 2007. Effect of phytic acid and phytase on the flow and amino acid composition of endogenous protein at the terminal ileum of growing broiler chickens. Br. J. Nutr. 98:745–752.[Medline]

Dahlqvist, A. 1984. Assay of intestinal disaccharidases. Scand. J. Clin. Lab. Invest. 44:169–172.[Medline]

Deshpande, S. S., and M. Ceryan. 1984. Effects of phytic acid, divalent cations, and their interactions on -amylase activity. J. Food Sci. 49:516–519.[CrossRef]

Dilworth, L. L., F. O. Omoruyi, and H. N. Asemota. 2005. Digestive and absorptive enzymes in rats fed phytic acid extract from sweet potato (Ipomoea batatas). Diabetol. Croat. 34:59–65.

Dilworth, L. L., F. O. Omoruyi, O. Simon, E. Y. Morrison, and H. N. Asemota. 2004. Hypoglycemia and faecal minerals in rats fed phytate. Nutr. Food Sci. 34:60–64.

Fiske, C. H., and Y. Subbarow. 1925. The colorimetric determination of phosphorus. J. Biol. Chem. 66:375–400.[Free Full Text]

Gal-Garber, O., S. J. Mabjeesh, D. Sklan, and Z. Uni. 2003. Nutrient transport in the small intestine: Na⁺,K⁺-ATPase expression and activity in the small intestine of the chicken as influenced by dietary sodium. Poult. Sci. 82:1127–1133.[Abstract/Free Full Text]

Gilbert, E. R., H. Li, D. A. Emmerson, K. E. Webb Jr., and E. A. Wong. 2008. Dietary protein quality and feed restriction influence abundance of nutrient transporter mRNA in the small intestine of broiler chicks. J. Nutr. 138:262–271.[Abstract/Free Full Text]

Gitlitz, P. H., and C. S. Frings. 1976. Interferences with the starch-iodine assay for serum amylase activity, and effects of hyperlipemia. Clin. Chem. 22:2006–2009.[Abstract/Free Full Text]

Katayama, T. 1997. Effects of dietary myo-inositol or phytic acid on hepatic concentrations of lipids and hepatic activities of lipogenic enzymes in rats fed on corn starch or sucrose. Nutr. Res. 17:721–728.

Kies, A. K., W. J. J. Gerrits, J. W. Schrama, M. J. W. Heetkamp, K. L. van der Linden, T. Zandstra, and M. W. A. Verstegen. 2005. Mineral absorption and excretion as affected by microbial phytase, and their effect on energy metabolism in young piglets. J. Nutr. 135:1131–1138.[Abstract/Free Full Text]

Kishi, K., T. Tanaka, and M. Lgawa. 1999. Sucrase-isomaltase and hexose transporter gene expression are coordinately enhanced by dietary fructose in rat jejunum. J. Nutr. 129:953–956.[Abstract/Free Full Text]

Knuckles, B. E. 1988. Effect of phytate and other myo-inositol phosphate esters on lipase activity. J. Food Sci. 53:250–252.[CrossRef]

Knuckles, B. E., and A. A. Betschart. 1987. Effect of phytate and other myo-inositol phosphate esters on -amylase digestion of starch. J. Food Sci. 52:719–721.[CrossRef]

Lee, S. H., H. J. Park, H. K. Chun, S. Y. Cho, S. M. Cho, and H. S. Lillehoj. 2006. Dietary phytic acid lowers the blood glucose level in diabetic KK mice. Nutr. Res. 26:474–479.

Leeper, L. L., and S. J. Henning. 1990. Development and tissue distribution of sucrase-isomaltase mRNA in rats. Am. J. Physiol. Gastrointest. Liver Physiol. 258:52–58.

Liu, N., G. H. Liu, F. D. Li, J. S. Sands, S. Zhang, A. J. Zheng, and Y. J. Ru. 2007. Efficacy of phytases on egg production and nutrient digestibility in laying hens fed reduced phosphorus diets. Poult. Sci. 86:2337–2342.[Abstract/Free Full Text]

Liu, N., Y. J. Ru, A. J. Cowieson, F. D. Li, and X. CH. Cheng. 2008. Effects of phytate and phytase on the performance and immune function of broilers fed nutritionally marginal diets. Poult. Sci. 87:1105–1111.[Abstract/Free Full Text]

Lowry, O. H., N. J. Rosebrough, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the folin-phenol reagent. J. Biol. Chem. 193:265–275.[Free Full Text]

Mandel, L. J., and R. S. Balaban. 1981. Stoichiometry and coupling of active transport to oxidative metabolism in epithelial tissues. Am. J. Physiol. 240:357–371.

McBride, B. W., and J. M. Kelly. 1990. Energy cost of absorption and metabolism in the ruminant gastrointestinal tract and liver: A review. J. Anim. Sci. 68:2997–3010.[Abstract]

Messiha, N., and R. R. Watson. 1989. The long term effects of varying dietary protein, lipid and vitamin E on intestinal and serum amylase. Nutr. Res. 9:1237–1247.

Miyamoto, K., K. Hase, T. Takagi, T. Fujii, Y. Taketani, H. Minami, T. Oka, and Y. Nakabou. 1993. Differential responses of intestinal glucose transporter mRNA transcripts to levels of dietary sugars. Biochem. J. 295:211–215.[Medline]

NRC. 1994. Nutrient Requirements of Poultry. 9th rev. ed. Natl. Acad. Press, Washington, DC.

Onomi, S., Y. Okazaki, and T. Katayama. 2004. Effect of dietary level of phytic acid on hepatic and serum lipid status in rats fed a high-sucrose diet. Biosci. Biotechnol. Biochem. 68:1379–1381.[Medline]

Ravindran, V., A. J. Cowieson, and P. H. Selle. 2008. Influence of dietary electrolyte balance and microbial phytase on growth performance, nutrient utilization, and excreta quality of broiler chickens. Poult. Sci. 87:677–688.[Abstract/Free Full Text]

Ravindran, V., P. C. Morel, G. G. Partridge, M. Hruby, and J. S. Sands. 2006. Influence of an Escherichia coli-derived phytase on nutrient utilization in broiler starters fed diets containing varying concentrations of phytic acid. Poult. Sci. 85:82–89.[Abstract/Free Full Text]

Ruckebusch, Y., L.-P. Phaneuf, and R. Dunlop. 1991. Physiology of Small and Large Animals. BC Decker Inc., Philadelphia, PA.

Schultz, S. G., R. A. Frizzell, and H. N. Nellans. 1974. Ion transport by mammalian small intestine. Annu. Rev. Physiol. 36:51–91.[CrossRef]

Selle, P. H., and V. Ravindran. 2007. Microbial phytase in poultry nutrition. Anim. Feed Sci. Technol. 135:1–41.[CrossRef]

Shelton, J. L., L. L. Southern, L. A. Gaston, and A. Foster. 2004. Evaluation of nutrient matrix values for phytase in broilers. J. Appl. Poult. Res. 13:213–221.[Abstract/Free Full Text]

Simons, P. C. M., H. A. J. Versteegh, A. W. Jongbloed, P. A. Kemme, P. Slump, K. D. Bos, M. G. E. Wolters, R. F. Beudeker, and G. J. Verschoor. 1990. Improvement of phosphorus availability by microbial phytase in broilers and pigs. Br. J. Nutr. 64:525–540.[CrossRef][Medline]

Singh, M., and A. D. Krikorian. 1982. Inhibition of trypsin activity in vitro by phytate. J. Agric. Food Chem. 30:799–800.[CrossRef]

Somogyi, J. C., and K. Trautner. 1974. Glucose, fructose and sucrose content in various vegetables. Schweiz. Med. Wochenschr. 104:177–182.[Medline]

Whealty, M. G., and R. P. Hentry. 1987. Branchial and antennal Na⁺K⁺-dependent ATPase and carbonie anhydrase activity during salinity acclimation of the euryhaline crayfish Pacifastacus leniusculus. J. Exp. Biol. 133:73–86.[Abstract/Free Full Text]

Wright, E. M. 1993. The intestinal Na⁺/glucose cotransporter. Annu. Rev. Physiol. 55:575–589.[CrossRef][Medline]

This Article Abstract Full Text (PDF) All Versions of this Article: jas.2008-1234v1 86/12/3432    most recent Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Liu, N. Articles by Cowieson, A. J. Search for Related Content PubMed PubMed Citation Articles by Liu, N. Articles by Cowieson, A. J.