S330S330 DOI 10.1002/mnfr.200900099 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Review Phytate in foods and significance for humans: Food sources, intake, processing, bioavailability, protective role and analysis Ulrich Schlemmer1, Wenche Frψlich2, Rafel M. Prieto3, 4 and Felix Grases3, 4 1 Department of Physiology and Biochemistry of Nutrition, Max Rubner-Institut,Federal Research Institute of Nutrition andFood, Karlsruhe, Germany 2 University of Stavanger, Norwegian School of Hotel Management, Jar, Norway 3 Laboratory ofRenal Lithiasis Research, University Institute of Health Sciences Research (IUNICS), Universityof Balearic Islands Ctra,Palmade Mallorca 4 CIBERFisiopatologa ObesidadyNutricin (CB06/03), Instituto de Salud Carlos III, Spain The article gives an overview of phytic acid in food and of its signif icance for human nutrition. It summarises phytate sources in foods and discusses problems of phytic acid/phytate contents of food tables. Data onphytic acid intake areevaluatedand dailyphytic acid intake depending on food habits is assessed. Degradation of phytate during gastro-intestinal passage is summarised, the mechanism of phytate interacting with minerals and trace elements in the gastro-intestinal chyme described and the pathway of inositol phosphate hydrolysis in the gut presented. The present knowledge of phytate absorption is summarised and discussed. Effects of phytate on mineral and trace elementbioavailability are reportedandphytate degradation during processingand storageis described. Beneficial activitiesof dietaryphytate suchasitseffectson calcificationand kidney stoneformationandonlowering blood glucose and lipids are reported. The antioxidative property of phytic acid and its potentional anticancerogenic activities are briefly surveyed. Development of the analysis of phytic acid and other inositol phosphates is described, problems of inositol phosphate determination and detection discussed and the need for standardisation of phytic acid analysis in foods argued. Keywords: Absorption/Antioxidant/Degradation/Inositol phosphates/Phytic acid Received:March7,2009;revised:May25,2009; accepted:May31,2009 1 Introduction magnesium but neither contained proteins nor lipids [3]. The name , phytin’was createdbyvirtueof the fact that this The discovery of phytate dates from 1855 to 1856 when Hartigfirst reported small round particlesinvarious plant seeds similarinsizeto potato starchgrains[1,2].Usingthe iodine test he showedthat the particles were free of starch and concluded that they must contain reserve nutrients for the germination of seeds. Later it was discovered that the isolated particles were rich in phosphorous, calcium and Correspondence: Dr. U. Schlemmer, Department of Physiology and Biochemistry of Nutrition, Max Rubner-Institut,Federal Research Institute of Nutrition and Food, Haid-und-Neu-Straίe 9, 76131 Karlsruhe, Germany E-mail: ulrich.schlemmer@mri.bund.de; Schlemmer.van-Ruiten@t-online.de Fax: +49-721-6625-404 Abbreviations: HAP, hydroxyapatite; ICP, inductively coupled plasma; PAR, 4-(2-pyridylazo) resorcinol; TBARS, thiobarbituric acid reacting substances i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim substance is of plant origin, having been detected neither in meat nor in dairy products, and it originally described the classical calcium – magnesium phytate deposits of plant seeds [3].Winterstein [4], Schulze andWinterstein [5] and Posternak [6] showed that hydrolysing phytin by hydrochloric acid liberated phosphoric acid and inositol. To explain the high phosphorous, calcium and magnesium contents of phytin, paired phosphoric acids were discussed as possible structures [7, 8]. Other molecular structures, however, were also under controversial debate for many years [8, 9]. In 1914, Anderson [10] presented the molecular structure of myo-inositol-1,2,3,4,5,6-hexakis dihydrogen phosphate, also calledphytic acid (Fig.1), whichis still valid and was confirmed by various modern analytical methods [11 – 13]. Phytate, the salt of phytic acid, is widely distributed in the plant kingdom. It serves as a storage form ofphospho www.mnf-journal.com S331S331 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Figure1.myo-inositol-1,2,3,4,5,6-hexakisphosphateatpH6–7.Underphysiologicalconditionsthenegativechargesarecounterbalancedmostlikelybysodiumionsorbyothercations.Conformation:5axial/1equatorial(modifiedtoEmseyandNiazi[11]). rous and minerals and contains l75% of total phosphorous of the kernels [14]. Other parts of plants such as roots, tubersand turions,however,areverylowinphytate(l0.1% dw) [15]. Besides phytate, other inositol phosphates such as inositol pentaphosphates and inositol tertraphosphates are also present in seeds, however, to a much lower extent (a15%) [16]. During the germination of seeds, phytate is hydrolysed [17, 18] and phosphate along with minerals such as calcium and magnesium becomes susceptible for germination and development of the seedlings, explaining its significant rolein plant metabolism. Phytate is predominantlypresent in unprocessed food,but canbedegraded during processing,soabroad rangeof inositol phosphatesmaybe consumed.Ithasbeen estimatedthat the dailyintake of phytate and other inositol phosphates on the basisofWesternstyle dietsvaries from l0.3 to 2.6g[19] andina global range from 0.180to 4.569g [20], strongly dependingonthediet selected;lowin normalWestern diets and high in vegetarian diets. Under heat treatment up to l1008C (home cooking, roasting, pressure cooking, etc.) phytateisquite stable[21,22]whilefood processingwiththe aidofphytasesmayresultinstrongphytatehydrolysis[23]. For decades phytate has been regarded as an antinutrient, as, during gastro-intestinal passage, it may inhibit the absorption of some essential trace elements and minerals, which under certain dietarycircumstances maylead to calcium, iron and zinc deficiencies [24 – 29]. Thus, intensive research has been carried out to remove phytate from food byproper processing, to improve the bioavailability and to avoid deficiencies of essential trace elements and minerals. In the last20years,however, beneficial propertiesofphytate havebeen observed and antioxidant [30] and anticancer activities [31] were reported. Inhibition of calcium salt crystallisation and prevention of renal stone formation i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Table 1. Sources of dietary energy consumption according to the FAO (kcal/capita/day, 2001 – 2003)a) Sources Developed % Developing % countries countries Fruits and vegetables 308 9.3 295 11.1 Cereals 1020 30.7 1391 52.4 Pulses 286 8.6 198 7.5 Sugar 427 12.9 194 7.3 Animal products 712 21.5 311 11.7 Oils and fats 566 17.1 267 10.1 Total 3319 100.0 2656 100.0 a) Adapted from [43]. through dietaryphytatewere described [32]. Reductionof starch digestion along with slowing down of the glycemic index of foods [33, 34] have also been reported, as well as positive effects on blood glucose and blood cholesterol [35, 36]. Thesefindingshave revived discussions about the significanceofphytateand other inositol phosphatesin human nutrition and for human health. Of central interest thereforeisto understandhowphytate exertsits beneficialeffectsinorgansandcells,whatthefate ofphytateis duringthe digestioninthegutandhowphytate and its degradations products can be absorbed. Under physiological pH(l6–7)phytateis highlynega- tivelycharged[13](Fig.1)andasno adequate carriershave been detectedbynow,ithaslongbeen assumedthatphytate cannot cross the lipid bilayer of plasma membranes and in consequence, its absorption in the gut has been considered rather improbable. However, recent studies in humans and rats have shown increasing levels of phytate in plasma and enhanced urinary phytate excretion after application of sodium phytate [37, 38]. Studies with radioactive labelled phytate in rats also provided some evidence for the absorptionofphytateoratleastofpartsofits degradationproducts [39, 40]. Cellular uptake studies with MCF-7 cells also provide evidence of phytate absorption [41] and recent experimentswithHeLashowthat cellular uptakeofphytatemight occur via pinocytosis [42]. Moreover, thegreat number of studies showing anticancer activity of phytate in skin, lung, liver, mammary, prostate, soft tissue, etc. also suggest that phytate or phytate degradation products have to be absorbed to a certain extent, even though the absorption mechanism still remainstobe clarified [41]. This reviewgives anoverviewof the main dietary food sources of phytate and the estimated daily intake. It describes the degradation of phytate during gastro-intestinal digestion and the passage throughout the gut and discusses the present knowledge of cellular uptake, absorption and bioavailability of phytate and lower phosphorylated inositol phosphates. Moreover, it surveys the adverse and beneficial activitiesofphytate and reports thedevelopment and progress made in the analysis of phytate and other inositol phosphates in food. www.mnf-journal.com S332S332 U. Schlemmer et al. Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 2. Content of phytic acid/phytate in cereals Cereals Phytic acid/phytatea) References Common names Taxonomic names g/100 g (dw) Maize Zea mays 0.72–2.22 [47–57] Maize germ 6.39 [20] Wheat Triticum spp. (l25species) 0.39–1.35 [48,49,51,57 –60] Wheat bran 2.1 – 7.3 [20, 46, 51, 53, 59, 60, 61, 72] Wheatgerm 1.14–3.91 [20,46,74] Rice Oryza glaberrima/sativa 0.06–1.08 [47–50,52,55,62,63,114] Ricebran 2.56–8.7 [46,51,59,60,75] Barley Hordeum vulgare 0.38–1.16 [51,53,57,58,60,64-66,99] Sorghum Sorghum spp. (l30 species) 0.57–3.35 [50,51,55,67,73] Oat Avena sativa 0.42–1.16 [51,54,56 –58,60,61,64,67,68] Rye Secale cereale 0.54–1.46 [47,48,53,56,60,64,69] Millet Pennisetum sp., etc. 0.18–1.67 [50,55,70,75,114] Triticale Triticale secale 0.50–1.89 [70,71] Wild rice Zizania sp. 2.20 [53] a) Depending on the data published. Table 3. Content of phytic acid/phytate in legumes Legumes Phytic acid/phytatea) References Common names Taxonomic names g/100 g (dw) Kidney beans Phaseolus vulgaris 0.61 – 2.38 [47 – 50, 53, 75 – 83] Haricot beans Pinto beans Navy beans Blackeye beans Broad beans Vicia faba 0.51–1.77 [79,85 –87,100 –105] Peas Pisum sativum var. arvense 0.22–1.22 [48,65,84 –90] Dry cowpeas Vigna unguiculata 0.37 – 2.90 [48, 50, 55, 82, 91 – 98] Black-eyed peas Chickpeas Cicer arietinum 0.28 – 1.60 [48, 50, 65, 83, 85, 86, 92, 95, 99] (Garbanzo/Bengal gram) Lentils Lens culinaris 0.27 – 1.51 [50, 65, 82, 83, 86, 95, 106 – 111] a) Depending on the data published. 2 Sources of phytate in foods Themain sourcesofphytateinthedailydietare cerealsand legumes, including oil seeds and nuts. They are important for human nutrition and represent l40 and l60% of total caloric intake for humans in developed and in developing countries, respectively(Table1)[43]. In cereals, phytate is located in the aleurone layer and the germ whilethe endospermis almost freeofphytate[44 – 46]. Approximately80% of phytate is located in the aleurone layer of small grains (wheat, rice, etc.), which represents l20%ofthis tissue'sdryweightand demonstratesthe enormous phytate reservoir which can be stored in special tissues [14, 44]. Cereals are rich in phytate and contain l1% phytic acid on the dry matter basis, ranging from l0.06 to l2.2% (dw) (Table 2) [46 – 73]. Cereal food products, however, may show higher phytic acid contents [46]. For wheat germs, wheat bran and special wheat bran fractions the phytic acid concentrations of 1.1– 3.9, 2.0 – 5.3 i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim and 7.3%, respectively, were reported [46, 51, 72, 74] and for rice bran it ranged up to 8.7% [75]. Inlegume seedsphytate predominantlyoccursinthe protein bodies of the endosperm or the cotyledon, containing up to 90% of the total phytic acid. In the whole seed the phytic acid content varies from 0.2 to 2.9% (dw) (Table 3) andis higherin the cotyledons(a3.7%) [48 –50, 53, 55, 65, 75– 111]. In oilseeds such as sunflower kernels, soybeans, soybean products, sesame seeds, linseeds and rape seeds the phytic acid content ranges from l1 to 5.4% (dw) (Table 4) [46, 49–52,55,58,74,85,91,112 –117]and specialfoodproducts such as dehulled sesame seeds or rape seed protein concentrates showphytic acid contentsupto l5.4 and 5.3 – 7.5%, respectively [74, 113]. In soy concentrates a maximumphytic acid contentof 10.7%is reported [75]. In nuts, the forth group of phytate-rich food, such as hazelnuts, walnuts, almonds and cashew nuts, the phytic acid content varies of l0.1 – 9.4% (dw) (Table 5) [47, 49, www.mnf-journal.com Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 4. Content of phytic acid/phytate in oilseeds S333 Oilseeds Taxonomic names Phytic acid/phytatea) References Common names g/100 g (dw) Soybeans Glycine max 1.0–2.22 [50–52,55,58,91,106,112,114 –116] Soy concentrate 10.7 [75] Tofu 0.1–2.90 [20, 46, 49, 51, 112, 115] Linseed Linum usitatissimum 2.15–3.69 [20,46,85,117] Sesame seed Sesmun indicum 1.44–5.36 [20,46,47,49,51,74,99,113,114,117] Rapeseed Brassica napus 2.50 [117] Rapeseedproteinconcentrate 5.3–7.5 [113] Sunflower meal Helianthus annuus 3.9–4.3 [51] a) Depending on the data published. Table 5. Content of phytic acid/phytate in nuts Nuts Taxonomic names Phytic acid/phytatea) References Common names g/100 g (dw) Peanuts Arachis hypogaea 0.17 – 4.47 [47, 49, 76, 117, 118] Almonds Prunus dulcis 0.35 – 9.42 [47, 49, 76, 117, 118] Walnuts Juglans regia 0.20 – 6.69 [47, 49, 76, 103, 117, 118] Cashew nuts Anacardium occidentale 0.19 – 4.98 [47, 49, 76, 103, 118] Brazil nuts Bertholetia excelssa 0.29 – 6.34 [47, 117 – 119] Pistachios Pistachia vera 0.29 – 2.83 [49, 110, 117] Hazelnuts Corylus avellana 0.23 – 0.92 [110, 117] Macadamia nuts Macadamia integrifolia 0.15 – 2.62 [49, 76, 118] Pecans Carya illinoinensis 0.18 – 4.52 [47, 49, 76, 118] Pine nuts Pinus pinea 0.20 [118] a) Depending on the data published. 76, 103, 116, 118, 119]. Especial high phytate contents of 9.4, 6.7 and 6.3% (dw) are found in almonds, walnuts and Brazil nuts, respectively. However, these results require confirmationbyspecific methods forphytic acid analysis. Table2–5showhugevariationsofthephytic acidorphytate contentindifferentrawand unprocessed foods.Formillet, rice, almonds, walnuts, peanuts, etc. variations of about one orderof magnitude exist.Formost legumes half thisvariation is present. These enormous ranges of phytic acid or phytate concentrations published do not only reflect the great number of botanical varieties of seeds, various environmental or climatic conditions of growing but also the different stages of seed maturation. All these factors influence thephytic acid contents presented, along with differences resulting from various unspecific and specific methods forthe determinationofphyticacidorphytatein food. Confusion mayarise if both phytic acid and phytate contents are mixed in food tables. Analysing phytate in food, all analytical procedures determine phytic acid following acidic extraction from food. This, however, eliminates any information on the phytate cations and in consequence no exact molecular weight of phytates occurring naturally can be calculated. Therefore, results should be given correctly as phytic acid even though it is not present in plants. How phytate which also does not exist in nature but is widely used as calibration standard and should not be confounded withthe naturallyoccurringphytatesin foods. To avoid confusion and to compare results on a common basis, international standardisation of specific methods for the determination of phytic acid/phytate and other inositol phosphates in foods and of listing either the phytic acid or thephytate contentinfoodtablesis absolutelyessential. 3 Estimation of dietary phytate intake Systematic studies on the mean dailydietaryphytic acid or phytate intake in humans of various countries are very rare. Thus, the available intake data from different countries were collected and the dailydietaryphytic acid intakes estimated( Table6). 3.1 Europe In the UK earlier studies showed phytate intake varying from504to844mgfor adults[82,120,121],whilea recent study calculated the mean dailyphytate intakein men (aged 40 years) at 1436 l 755 mg [122]. ever,if food tables present , phytate’ contents of food, this Earlystudies from Italy, evaluating 13 Italian diets, also , phytate’ismostlycalculatedonthebasisof dodecasodium reportedabroadrangeof112 –1367mgphyticacid per day i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com S334S334 U. Schlemmer et al. Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 6. Daily intake of phytic acid/phytatea) in different countries Country Sex, age and conditions Daily intake of phytic acid/phytate References mean value, mean l SD or range (mg) Europe United Kingdom Male (>40 years) 1436 l 755 [122] Male–female 600–800 [82] Male–female 504–848 [120,121] Italy Male–female 219(112 –1367) [123] Male–female,averageItaliandiet 293(265 –320) [124] Male–female,north-west 288 [124] Male–female,north-east 320 [124] Male–female,south 265 [124] Sweden Male–female 180 [125] Male–female(35 –76years)Westerntypediets 369(230 –532) [126] Male–female(35 –76years)vegetariandiets 1146(500 –2927) [126] 6– 8 m, commercial milk-based cereals and porridge 124 l 82 (lmol phytate/day) [127] 6– 8 m, infant formula and porridge 26 l 18 (lmol phytate/day) [127] 9– 11 m, commercial milk-based cereals and porridge 189 l 78 (lmol phytate/day) [127] 9– 11 m, infant formula and porridge 62 l 47 (lmol phytate/day) [127] Finland Male–female 370 [128] North and Central America USA Infants (<1 years) 166 l 167 [129] Children (1 – 3 years) 390 l 231 [129] Children (4 – 5 years) 501 l 271 [129] African-Americanmale –female(medianintake) 538(253 –1352) [131] African-American females (median intake) 512 [131] African-American males (median intake) 608 [131] Male–female(19 –35years) 1293 l 666 [133] Female(18 –24years) 395 l 334 [134] Female omnivorous (self-selected) 631 (590 – 734) [135] Male omnivorous (self-selected) 746 (714 – 762) [135] Female vegetarians l1250 l 450 [135] Male vegetarians l1550 l 550 [135] Male lacto-ovo-vegetarian 5577 [136] Male lacto-ovo-vegetarian 972 [137] Average American (bw, 75 kg) 750 [136] Canada Female (4 – 5 years) 250 (132 – 318) [130] Male(4 –5years) 320(203 –463) [130] Lacto-ovo diets – Asian immigrants 1487 l 791 [132] Mexico Male–female(18 –30m) 1666 l 650 [138] Male–female(7 –9years) 3380 l 1070 [139] Guatemala Female(15 –37years) 2254 [140] Asia India Male–female 670 [141] Male–female(4 –9years) 720–1160 [142] Male–female(10 –19years) 1380–1780 [142] Male–female(20 –45years) 1560–2500 [142] Male–female(>45years) 1290–2080 [142] Females (16 – 20 years) l840 [143] Thailand Females nonurban 1139 l 481 [144] Males nonurban 1104 l 965 [144] Females urban 997 l 435 [144] Males urban 1304 l 956 [144] PeoplesRepublic Male–female 1186(823 –1603) [145] ofChina Male–femaleurban 781(443 –1205) [145] Male–femalenonurban 1342(970 –1757) [145] Republic of China Females 690 l 189 [146] Males 915 l 330 [146] i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 6. Continued S335 Country Sex, age and conditions Daily intake of phytic acid/phytatea) References mean value, mean l SD or range (mg) South Korea Male (21 – 70 years) 839 l 400 [114] Female (21 – 70 years) 752 l 407 [114] Female (20 – 24 years) 322 l 220 [147] Female (64 – 75 years) 496 l 252 [147] Africa Egypt 18 – 30 months 796 l 248 [138] 7 – 9 years 1270 l 280 [139] Kenya 18 – 30 months 1066 l 324 [138] 7 – 9 years 2390 l 480 [139] Malawi Preschool girls (4 – 6 years) 1621 l 660 to 1729 l 592 [148] Preschoolboys(4 –6years) 1857 l 530 to 2161 l 684 [148] Gambia 1 – 17 months 10 – 560 [149] Ghana children 578 l 161 [150] Nigeria Male–female 2200 [77] Ethiopia Female pregnant 1033 l 843 [99] a) Depending on the data published. with an estimated mean intake of 219 mg/day[123], while a later report stated the mean phytic acid intake of the national Italian diet at 293 mg, defined as the average of typical diets from the north-west (288 mg), from the northeast (320 mg) and from the south of Italy (265 mg) [124]. The study indicates, moreover, that cereals, contributing the highestportiontothephyticacid intake(52 –57%),arelowest in the south and highest in the north-east, resulting in the highestphyticacid intakeinthenorth-easternpartofItaly. In Sweden, the mean phytic acid intake of adults was reportedat180mg[125].However,a laterstudyfromSwedenin adults(35 –76years)showed aphytic acid intakeof 230–532mg (meanvalue 369mg) andof 500 –2927mg (mean value 1146 mg) for Western type diets and vegetarian diets, respectively[126], whereas thephytic acid intake was recalculated on the basis of the phytate phosphorus, reported in the paper. In Swedish infants (6 –8m), nourished with two different diets, one being an infant formula and porridge and the other one a commercial milk-based cereal diet, the mean dailyphytate intakewas26 l 18 and 124 l 82 mg, respectively. With increasing age (9–11m), enhanced dailyphytate intake of 62 l 47 and 189 l 87 mg, for both diets, respectively,wasreported [127]. For Finland the averageper capita intake of phytic acid from cereal products, the main dietaryphytate source,was estimated at 370 mg [128]. 3.2 North and Central America Preschool childrenin theUS showed a mean dailyphytate intake of 166 l 167 mg for up to 1 year of age, 390 l 231 mg for 1 –3 years, and 501 l 271mg for4 –5 years [129] while Canadian girls and boys of the same age (4–5 years) showed lower phytate intakes of 250 and 320 mg, respectively[130]. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim In African-American adults in the US, the median of the dailyphytate intakewas538 mg, with strong differences for females (512 mg) and males (608 mg) [131]. In adult Asian immigrants to Canada, consuming predominatelylacto- ovovegetariandiets,high meandailyphytate intake of 1487 l 791mgwere also found [132].Astudy in American students and university faculty staffmembers (19–35 years) showed high mean daily phytate intake of 1293 l 666 mg, ranging from 198 to 3098 mg [133] and in a study withyoung Americanwomen (18–24years)alow mean daily phytate intake of 395 l 334 mg was estimated [134]. In self selected diets of omnivorous females and males the phytate intake was found to be 631 mg (590– 734 mg) and 746 mg (714 – 762 mg) and in female and male vegetarians l1250 l 450 and l1550 l 550 mg, respectively [135]. The maximum phytate intake ever reported in humans was 5.770 mg for lacto-ovo vegetarians (Trappist monks) in 1978 [136] which, however, was much lower with 972 mg when the study was repeated 10 years laterin the samegroupofTrappist monks [137].Foraverage Americans witha bodyweightof75kga studyof 1976 calculatedthedailyphytate intakeat750mg[136]. Infants in Mexico (18 –30 m) showed the daily phytate intake of 1666 l 650 mg [138] and pupils (7 –9 years) of 3380 l 1070 mg [139]. Studies from Guatemala also indicated a high daily phytate consumption of 2254 mg for females (15 –37years) [140]. 3.3 Asia In India the mean dailyphytate intake ranges from 670 to 2500 mg [141, 142]. For children (4 –9 years) it is 720 – 1160 mg, for adolescents (10–19 years) 1350 – 1780 mg, for adults (20 –45 years) 1560 – 2500 mg and for elderly people(A45 years) 1290 – 2080 mg [142]. Another study, www.mnf-journal.com S336S336 U. Schlemmer et al. however, reported comparablelower dailyphytate intakeof l840 mg for females (16 –20years) [143]. In Thailand the mean dailyphytate intake forwomenwas 1139 l 481 mg and for men 1104 l 965 mg, reported from Ubon Ratchathaniinthe northernpart ofThailand.Forthe city of Bangkok, however, it was 997 l 435 mg (M) and 1304 l 956 mg(m)[144]. In thePeoples Republic of China the median dailyphytate intakewas1186mg,rangingfrom823to1603mg,and strong differences in the median daily phytate intake for urban population (781 mg, ranging from 443 to 1205 mg) and rural population (1342 mg, ranging from 970 to 1757 mg)were described [145].For students and university members in the Republic China (Taiwan), the mean phytate intake per dayfor females was 690 l 189 mg and for males 915 l 330mg [146],fittingwellwiththedataofthephytate intakeof urban populationinthePeoples Republicof China [145]. In the Republic of Korea the daily phytate intake for males (21 –70 years) was 839 l 400 mg and for females 752 l 407 mg [114]. Differences for young (mean 23 years) and aged females (mean 70 years) were 322 l 220 and 496 l 52 mg, respectively[147]. 3.4 Africa Infantsin Egypt(18 –30 m)showeddailyphytate intakeof 796 l 248 mg [138] and pupils (7–9 years) of 1270 l 280mg [139].For children fromKenyaofthe same age a daily phytate intake of 1066 l 324 mg (18 –30 m) [138] and of 2390 l 480 mg (7 –9 years) was described [139]. The mean dailyphytate intake for girls andboys(4 – 6 years) from Malawi ranged from 1621 to 1729 mg and from 1857 to 2161 mg, respectively[148]. Infants in Gambia( 1–17 m)showed a dailyphytate intakeof10–560mg [149] and children in Ghana a phytic acid intake of 578 l 161 mg [150]. A mean daily intake of phytate for pregnant women in Ethiopia was 1033 l 843 mg [99] and for Nigerian people an average phytate intake of 2200 mg per dayand personwasestimated [77]. 3.5 Conclusions The data collectedgive anoverviewofthe mean dailyphytate intake in humans of countries from all over the world, stronglyvaryingin sexand ages. For infants(a1years) in Sweden the mean dailyphytate intake reaches 26 –189mg [127]. Whether this infant formula is representative and comparable to other industrialised countries remainstobe clarifiedbutitconveysanidea ofthelevel ofphytate intakein infantsin Europe. Children in the US and in Canada (1–5 years) show a mean daily phytate intake of 166– 501 mg [129, 130] which is much lowercomparedto Egypt,KenyaandMexico,where infants (l1–2years)showa meandailyphytate intakeof796, i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 1066 and 1666 mg and pupils (7 –9 years) show one of 1270, 2390 and 3380 mg, respectively[138, 139].For children from Malawi (4 –6years) a very high mean dailydietary phytate intake of 1622 – 1729 mg (M) and 1857 – 2161 mg(m)was also reported [148]. For adults, studies from Sweden, Finland and Italyreport alow mean dailyphytate intakeof 180–370 mg,ifWestern style diets are consumed [123 – 126, 128], and for vegetarians a much higher one of 1146 mg (Sweden) [126]. In the UKtwolevelsofphytate intake are also present, oneinthe range of 504– 848 mg [120, 121] and another one of 1436 mg [122]. Similar data exist for the US, indicating for women a low range of the mean daily phytate intake of 395 mg (18 –24 years) [134] and a high one of 1250 mg (19–35 years) [135]. In 1971 the mean phytate intake of average Americans was estimated at 750 mg [136]. In Nigeria and Guatemala much higher dailyphytate intakes of 2200 and 2254 mg, respectively, were described [77, 140]andthe maximumdailyphytate intakeever reportedin humans was 5770 mg for lacto-ovo vegetarians (Trappist monks) [136]. The data suggest that for adults different levels of the dailydietaryphytate intake exist: (i) atalowlevelof l200 to l350 mg, probably due to Westernstyle dietslowinphytate rich plant foods. (ii)Ata higherlevelof l500 – l800 mg, probablydue to primarily Western style diets with enhanced portions of cereals,wholegrain productsandotherphytaterich foods. (iii) At a high level of A1000 mg, probably due to diets rich in plant and phytate containing foods such as vegetarian diets. In developing countries, due to the high content of cereals and legumes in the traditional diet, obviouslyquite high levelsofphytate intakeofupto 2000mgand more canbe assumed, althoughonlyafew studiesareavailable[77,139, 140, 150]. Strong differences between the mean phytate intake in rural population (1342 mg) and urban population (781 mg) are evident in the Republic of China, probably reflecting the changing of dietaryhabits from traditional to moreWesterntypedietsin citiesand metropolises[145]. Evaluating the results of the daily phytate intake in humans, strong differences are apparent between developing and industrialised countries, between urban and rural areas, between females and males, between young and old and between omnivores and vegetarians (Table 6). These differences certainlyderive from differences in the phytate content of cultivated plant foods, different contents in plant foods in the dailydiet, different amounts of foods consumed and different processing and preparation of vegetables, legumes, pulses, cereals and whole grain products, etc. Moreover,different methodsforthe determinationofphytic acid may also play a role in the high variation of reported phytate intake.To obtaina reliableoverviewof thephytic acid intake in different countries, further studies with detailed documentation of the social and nutritional back www.mnf-journal.com Mol. Nutr. Food Res. 2009, 53, S330 – S375 S337 Figure 2. Hydrolysis of phytate from a diet rich in intrinsic feed phytases during the passage throughout the stomach, small intestine and large intestine as well as in the faeces of pigs 5 h after feeding [159]. Inositol phosphates are listed from the front to the back in the following row: InsP2, Ins(1,2,3)P3/Ins(1,2,6)P3, Ins(1,5,6)P3, Ins(1,2,3,4)P4/Ins(1,3,4,6)P4, Ins(1,2,5,6)P4, Ins(1,2,3,4,6)P5, Ins(1,2,3,4,5)P5, Ins(1,2,4,5,6)P5, Ins(1,3,4,5,6)P5 and InsP6. Figure 3. Hydrolysis of phytate from an extruded diet with inactivated phytases, during the gastro-intestinal passage throughout the stomach, small intestine and large intestine as well as in the faeces of pigs 5 h after feeding [159]. *InsP6 concentrations were different (p a 0.05). Inositol phosphates are listed from the front to the back in the following row: InsP2, Ins(1,2,3)P3/Ins(1,2,6)P3, Ins(1,5,6)P3, Ins(1,2,3,4)P4/Ins(1,3,4,6)P4, Ins(1,2,5,6)P4, Ins(1,2,3,4,6)P5, Ins(1,2,3,4,5)P5, Ins(1,2,4,5,6)P5, Ins(1,3,4,5,6)P5 and InsP6. ground and specific, evaluated methods for the analysis of degraded in the human gut. In 1935 McCane and Widphytic acid and lower phosphorylated inositol phosphates dowson already observed36– 63% of phytate in the stool of in food are required (see discussion in Section 2). adults after applying diets, rich in alimentary phytate, and calculated the mean phytate degradation of 54 l 8% [151]. This was confirmed by a later experiment, showing phytate 4 Mechanism of phytate hydrolysis in the gut phosphorus absorption of 50% [24]. Similar results were described by Hoff-Jψrgensen et al. [152] for infants (1 – Studies on the phytate degradation in the human gut are 11 m) and children (10 years), indicating a gastro-intestinal scarce. Early experiments showed that some phytate is phytate hydrolysis of l30– 47%. However, in 1945 Cruicki 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com S338S338 U. Schlemmer et al. shank et al. [153] reported an almost complete phytate phosphorus digestibility in adults, implying an almost total phytatehydrolysis, andin 1947Walker et al. [154] pointed out that the dietary calcium level effects phytate degradation in the gut (43 – 90%). Although it was not clear at that timeto whichextentphytate mightbehydrolysed exactlyin the human gut – which was also due to the lack of an adequate and specific method for determining phytate – these earlystudies already showedthat phytate is degraded stronglyduring the gastro-intestinal digestion in humans. 4.1 Stomach In ileostomy patients Sandberg et al.in 1986 [155–158] foundphytate degradationof56 –66% for the total passage through the stomach and small intestine when the diet contained active food phytases. If the dietary phytases were inactivated eitherby extrusion or heat treatmentof the diet, the total phytate degradation in the upper part of the gut decreasedto0 –28% [155 –158]. As studies in ileostomypatients allowonlylimited informationonthephytate degradationinthegutandasfor ethical reasons detailed studies in humans on the phytate hydrolysis in the different parts of the digestive tract are nonfeasible, Schlemmer et al. [159] studied the phytate hydrolysis in detail in the stomach, the small intestine, the large intestine, the faeces and the enzymes involved in pigs which is an excellent model for simulating digestion processes of humans [160]. Figures2 and3show the distribution of inositol phosphates in the chyme of the different parts of the gut and of the faeces of pigs fed a diet rich in intrinsic feed phytases (control diet) (Fig. 2) and the same but extruded diet with inactivated feed phytases (extruded diet) (Fig. 3). The two diets were applied to differentiate betweenphytases from dietaryand from endogenous origin, possiblyinvolvedin the gastro-intestinalhydrolysisofphytate. Moreover, the inositol phosphates in the liquid and solid phase of the chyme were separated to differentiate between soluble, degradable and possibly physiologically active inositol phosphates and insoluble, nondegradable and probablyphysiologicallyinactive ones.Figure3shows that no phytate degradation occurs in the stomach if no active feedphytases are present. This confirms earlierfindings in ileostomy patients [157, 158]. In the presence of active foodphytases,however, stronghydrolysisof soluble phytate occurs and lower phosphorylated inositol phosphates are formedbystepwise inositol phosphate degradation (Fig. 2). From the inositol phosphate isomers determined in the gastric chyme and the respective cleaving specificity of the enzymes, it can be concluded that intrinsic feed phytases of plant origin (6-phytases) degrade phytate in the stomach, as the main inositol pentaphosphate formed is DL-Ins(1,2,3,4,5)P5 which is specific for 6-phytases (EC 3.1.3.26) (see Section 7.2). The samefindings are reported from ileostomy patients, also showing DL-Ins(1,2,3,4,5)P5 i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 as the predominant inositol pentaphosphate in the gastrointestinal content [161]. As no Ins(1,2,4,5,6)P5, specific for 3-phytases (EC 3.1.3.8), is formed during gastric digestion, it can be excluded that endogenous phytases, e.g. described as 3-phytases in human and rat small intestinal mucosa cells [162 – 165], are involved in the phytate degradation in the pigs’stomach. In order to confirm thesefindings an additional ex vivo experimentwascarriedoutandNa –phytatewashydrolysed byenzymes purified from the pigs feed and from the stomach chyme of pigs fed the same diet. The kinetic of the phytatehydrolysis for both enzymeswas the same and the inositol phosphate isomers formed were identical [159]. This shows clearlythat the enzymes, activeinhydrolysingphytate in the gastric chyme, are phytases of feed origin and no evidence is given that endogenous phytases (3-phytases) takepartinthephytatehydrolysistoaphysiologicallyrelevantdegree. Basedonthe quantitativeevaluationoftheinositol phosphate isomers,a major pathwayofphytatehydrolysisby 6-phytasesof plant originin the stomachwas proposed [159]: InsP6 fi DL-Ins(1,2,3,4,5)P5 fi DL-Ins(2,3,4,5)P4 +DL-Ins(1,2,3,4)P4 fi DL-Ins(2,3,4)P3 +Ins(1,2,3)P3 fi InsP2 (seeFig. 5). This is similar to the pathway reported for cereal phytases from wheat, barley,rye and oat [161]. Due to the fact that only l2/3 of total phytate is soluble and present in the liquid phase of the gastric chyme of the pigs (Fig. 2), this means that l1/3 of the phytate is still bound to the feed matrix and should be hardly available for enzymatichydrolysis. Thus, incomplete phytate liberation from feed matrix during digestion in the stomach consequently results in incomplete gastric phytate hydrolysis. This explains whyneither in ileostomypatients [155 – 158] nor in pig studies [159, 166, 167] complete phytate degradation during passage throughout the stomach and small intestine hasever been detected. Moreover,itconfirms earlier assumptionsofKemme et al. [168] that the gastro-intestinalphytatehydrolysis mightbegovernedbythevelocity ofphytate liberation from feed. In orderto studythe maximumphytatehydrolysisinthe upper part of the gut, phytase activity in the diets of pigs was enhanced by adding microbial phytases to the pig's diets(a1800 FTU Aspergillus niger phytase/kg feed) and thephytate phosphorousavailabilitywas determined.With increasingphytase activityphytate phosphorousavailability in pigs increased to a plateau at 60– 66% [169, 170]. This implies that there is not only a maximum of phytate phosphorousavailability but alsoa maximumofphytatehydrolysis of l60– 66% during digestion in the stomach and small intestine which indicates that total phytate degradation in www.mnf-journal.com S339S339 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 7. Solubility of inositol phosphates in the intestinal chyme of pigsa) Small intestine pH 6.6 l 0.2 [%] Large intestine pH 6.2 l 0.2 [%] InsP2 InsP3 InsP4 InsP5 InsP6 75 31 8 7 2 24 6 0 3 2 a) Chyme was removed from the small and large intestine of pigs (jejunum and colon, respectively) fed a normal pigs diet with active feed phytases (control diet) [159]. Chyme was centrifuged and the inositol phosphates of the liquid and solid phase separated [173]. Inositol phosphates were determined by HPLC on Mono-Q (HR 5/5) and quantified by Fe3+-detection [159]. Inositol phosphate isomers with the same numbers of phosphate groups were summed. Results are mean values on the basis of double determinations. Solubility of inositol phosphates was defined as the ratio of soluble to total inositol phosphates (%) determined in the chyme. The pH given is the mean pH of the respective homogenised chyme sample (n = 3). the upper part ofthe gut of pigs is unavailable even at very high dietaryphytase activity.It confirms the assumption mentioned above that most probably incomplete liberation of phytate from the feed matrix is responsible for the incompletephytatehydrolysisinthe upperpartofthegut. Gastric degradation of dietaryphytatebysplitting phosphategroupsfromphytateisnotonlyrelevantfor monogastric animals, such as pigs and broilers, to enhance their phosphoroussupply,butisalsosignificantfor humans.By hydrolysing phytate to lower phosphorylated inositol phosphates (InsP5, InsP4)inositol phosphates with higher solubility are formed [47], improving on the one hand the susceptibility for further enzymatic hydrolysis and accelerating on the other hand the stepwise degradation to low phosphorylated inositol phosphates, mainly to InsP3 but also to InsP2.Aslower inositol phosphatesin comparisontophytate are assumed to showlower affinity to minerals [171, 172] and higher solubility of the complexes with metal ions [47],phytatehydrolysisin the stomachbythis reduces the inhibition of the intestinal absorption of essential trace elements and minerals in humans and animals. 4.2 Small intestine In the small intestine most inositol phosphates are precipitated and present in the solid phase of the chyme (Fig. 2). Only InsP2 shows higher contents in the liquid than in the solid phase and the solubility of InsP3 in the chyme is higher than that of the other inositol phosphates such as InsP4, InsP5 and InsP6. Due to interactions with multivalent cations such as iron, zinc, calcium, etc. and increasing pH during the passage from the stomach (pylorus, pH l 2) to the i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim small intestine (pH l 5–7), inositol phosphates with increasing phosphorylation degrees precipitate, especially phytate (Fig. 2). By precipitation, inositol phosphates reduce the availability of these metal ions in the chyme and thus interfering in the intestinal absorption of these essential minerals and trace elements. By means of an additional ex vivo experiment, the distribution of soluble and insoluble inositol phosphates in the intestinal chyme was studied due to the physiological significance for the understanding of the mechanism of inositol phosphates interfering in the intestinal absorption of metal ions. The results show that the solubility of the inositol phosphatesinthechyme,definedasthe ratioof soluble to total inositol phosphates (%), varies with the degree of phosphorylation. The higher phosphorylated the inositol phosphates are, the loweris their solubility in the intestinal chyme and vice versa (Table 7) [173]. Earlier discussions on the role of phytate in the small intestine, assuming that the more phytate is degraded the more inositol phosphates with lower metal affinity are formed [171], seems to be insufficientto fully explain the inhibitory effectofphytic acid on mineral intestinal absorption. It seems more probable, thatthe inhibitionofthe metal intestinal absorptionby phytate is the direct consequence of reduced solubility of minerals and trace elements, interacting with phytates in the small intestinal chyme. The results presentedinFigs.2and3 on the soluble inositol phosphatesin the gastro-intestinalchyme show thatby phytate degradation in the stomach the concentration of solublephytateinthe gastricchyme decreases, reducingthe interactions with metal cations in the gastric chyme. In parallel the concentration of lower phosphorylated inositol phosphates in the gastric chyme increases (pH 4.6) and even at pH 6.2 in the small intestine the concentration of soluble inositol-di-and inositol-triphosphates remains considerably high. High concentration of soluble inositol-diand inositol-triphosphates in the small intestinal chyme seemstobe relevanttoavoidprecipitationof metal-inositol phosphates and keeping trace elements and minerals in solution in the small intestine, the predominant location of the metal absorptionin the gut.As iron uptakebyCaco-2 cells isnotshowntobe impairedbylower phosphorylated inositol phosphates such as InsP3 and InsP4 at a molar ratio of 2:1 (inositol phosphates/iron) [174], it can be assumed that the higher the concentration of the soluble InsP2 and InsP3 is in the small intestinal chyme, the less affected is the intestinal absorption of minerals and trace elements. This might best explain why the degradation of phytate improves the intestinal absorption of minerals and trace elements in humans and animals. During the passage from the stomach to the small intestine, constant digestion and absorption of absorbable nutrients lead to increased concentration of non-and lowabsorbable compounds such as inositol phosphates in the chyme. Consequently, the concentration of inositol phos www.mnf-journal.com S340S340 U. Schlemmer et al. Mol. Nutr. Food Res. 2009, 53, S330 – S375 Figure 4. Hydrolysis of phytate from extruded feed with addition of defined Aspergillus niger phytases activities (diet A: a50 FTU/kg, diet B: 150 FTU/kg and diet C: 900 FTU/kg) during the passage through the stomach, duodenum and the ileum of pigs [177]. Inositol phosphates are listed from the front to the back in the following row: InsP2, Ins(1,2,3)P3/Ins(1,2,6)P3, Ins(1,5,6)P3, Ins(1,2,3,4)P4/Ins(1,3,4,6)P4, Ins(1,2,5,6)P4, Ins(1,2,3,4,6)P5, Ins(1,2,3,4,5)P5, Ins(1,2,4,5,6)P5, Ins(1,3,4,5,6)P5 and InsP6. phates increases during the passage from the stomach to the small intestine and is higher in the small intestinal than in the stomachchyme(Figs.2and3).Inpigs,fedadietfreeof activephytases,thephytate concentrationinthe small intestine is higher than in pigs fed the diet with active feed phytases, due to the lack of anyrelevant gastric phytate degradation( Figs.2and3). Moreover,the absenceofanyphytate hydrolysis products in the intestinal chyme clearlyindicates that no endogenous phytases are involved in the intestinal phytate hydrolysis (Fig. 3). This also confirms that only phytases of food or feed origin are relevant for the phytate degradation in the small intestine of humans and pigs. Inpigsfedadietwithactivephytasesno changeoftheinositol phosphates pattern(the relation of the different inositol phosphate isomers to each other) shows that obviously no further degradation of phytate and the other inositol phosphates occurs (Fig. 2). This is certainlythe consequence of strong inactivationofthe intrinsicplantfeedphytasesbythe peptic digestioninthepylorus[175]andbyanunfavourable pHin the duodenal chyme (pH l 6.5–7)atwhichplantphytasesshowonlyverylowactivity [176].If,however,thepigs’ diet is enriched with microbial phytases such as Aspergillus niger phytases,toimprovephytatehydrolysis,the degradation of inositol phosphates continues in the small intestine (Fig.4)[177]. Similar resultswere also describedbyRapp et al. [178]. As DL-Ins(1,2,4,5,6)P5is the major inositol pentaphosphateformedinthe duodenumand ileumand typicalfor Aspergillus Niger phytase (3-phytase, EC 3.1.3.8), this clearly shows that when feeding a diet supplemented with Aspergillus Niger phytases, these microbial phytases are obviously responsible for continuingphytatehydrolysisin the small intestine (Fig. 4). These results show, moreover, that microbial phytases are more stable than intrinsic plant feed phytases towards peptic digestion and inactivation in thepylorusand duodenuminpigs. 4.3 Large intestine In the large intestinal chyme the inositol phosphates are almost completelypresent in the solid phase with high concentrations of phytate and very low ones of the lower phosphorylated inositol phosphates (InsP2– InsP5). This is true for both diets (Figs.2and 3). The most prominentphytate degradation products are DL-Ins(1,2,3,4,5)P5 and DLIns( 1,2,4,5,6)P5, indicating that 6-and 3-phytases are involvedinthephytatehydrolysisinthelarge intestine.As feed phytases of plant origin (6-phytases) are no longer active in the intestine (see Section 4.2), the DLIns( 1,2,3,4,5)P5 detectedmusthavebeenformedby Escherichia coli phytases, which are also 6-phytases [179] and prominent in the micro flora, while the DL-Ins(1,2,4,5,6)P5 is generatedby3-phytasesof other microbial origin. Evaluating the inositol phosphate isomers formed during thephytatehydrolysisinthelarge intestine,twomajorpathwaysof thephytatehydrolysisby bacterial 6-phytases and microbial 3-phytaseswere established [159]: i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 8. Activity of phytases and alkaline phosphatases in feed and gastro-intestinal chyme of pigs fed the control and the extruded diet S341 Feed Stomach Small intestine Colon [mU/mg protein] Control diet Phytases 43.1 l 2.1a) 3.7 l 0.6b) 0.9 l 0.3 2.2 l 0.1 Alkaline Phosphatases – – 146 l 37d) 29.9 l 6.9e) Extruded diet Phytases 0.2 l 0.1c) 0.3 l 0.2 0.7 l 0.2 1.8 l 0.1 Alkaline Phosphatases – – 153 l 43f) 39.3 l 5.8g) means l SD, n =3 a vs. b, d vs. e and f vs. g are significantly different p a 0.125. a vs. c is significantly different p a 0.05. Adapted from [159]. (i) InsP6 fi DL-Ins(1,2,4,5,6)P5 fi DL-Ins(1,2,5,6)P4 fi DL-Ins(1,2,6)P3 fi InsP2. (ii) InsP6 fi DL-Ins(1,2,3,4,5)P5 +DL-Ins(1,2,4,5,6)P5 fi DL-Ins(1,2,3,4)P4 +DL-Ins(1,2,5,6)P4 fi DL-Ins(1,2,3)P3/ DL-Ins(1,2,6)P3 fi InsP2 (seeFig. 5). As in pigs fed the diet with active phytases (control diet) the concentration of InsP6in the large intestinalchymewas as high as in the small intestine, it demonstrated that no phytate degradation could have been taken place during the passage from the small to the large intestine (Fig. 2). This is in contrast to the phytase inactivated diet (extruded diet), showing onlyhalf of the phytate concentration in the large intestine in comparison to the small intestine, indicating strongphytate degradationinthelarge intestineforthephytase free diet (Fig. 3). Applying the concentration of free inorganic phosphate and inositol monophosphates as a marker for intensivephytatehydrolysis, the high concentration of free inorganic phosphate and inositol monophosphates confirmed that phytate was degraded more intensively in pigs fed the diet free of phytase than in the diet withactivephytases[159].Evenifthe reasonforthis stronger InsP6hydrolysis remains to be clarified, the results allow the assumption, that the higher the concentration of InsP6 is in the small intestinal chyme, the stronger is the InsP6hydrolysis in the large intestine. Experimentsin rats also focussingonphytatehydrolysis inthe large intestine [180] reported56%ofphytatehydrolysed in conventional rats while in germfree rats almost no phytatehydrolysiswas detected. Thisfinding also emphasisesthe significanceof microbialphytasesforphytatedegradation during the digestion in the gut and mayexplain the strongphytatehydrolysisinthelarge intestine. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim 4.4 Faeces In the faeces very low phytate concentrations are present, which demonstrate continuing strong phytate hydrolysis (Figs.2and3)[159].Thisistrueforboth diets.Asthemain phytatehydrolysis products are DL-Ins(1,2,3,4,5)P5and DLIns( 1,2,4,5,6)P5,it indicates that the same bacterial 6-phytases and microbial 3-phytases as in the large intestine are responsibleforthe continuationofthephytatehydrolysisin faeces. Phytate degradation in the gut depends on the calcium level in the diet. This was already shown in humans by Walkeret al.in 1947 [154]. Sandberg et al. [181] found in pigs that the total phytate degradation throughout the gut decreased significantly 97, 77 and 45%(p a 0.001) when the dietary calcium intake increased by 4.5, 9.9 and 15 g/ day, respectively.Asthephytate degradationinthe stomach and small intestinewas almost unaffected, different supplementation of calcium carbonate predominantly affected the hydrolysis of phytate in the large intestine. Similar results were reported for ratsbyPileggi et al. [182] andWise and Gilbert [180]. Most probably, high dietarycalcium content affects the phytate solubility in the gastro-intestinal chyme and thereby reduces the accessibility of phytate for enzymatic hydrolysis. A similar effect on phytate degradation was also assumed for magnesium [46]. Thephytatehydrolysis in the gut and the role of calcium in the intestinal phytate degradation were previously discussedbyWise [183] andbySandberg[161]. 4.5 Enzymes To assess the contribution of enzymes involved in the gastro- intestinal degradation of phytate, enzyme activities of phytases and alkaline phosphatases were determined in the feed and the chyme during the passage throughout the gut. Inthedietwithactivephytases (controldiet),thephytase activity was high (43.1 mU/mg protein), decreased to l1/ 10 in the stomach chyme (3.7 l 0.6 mU/mg protein) and www.mnf-journal.com S342S342 U. Schlemmer et al. Mol. Nutr. Food Res. 2009, 53, S330 – S375 was only l1/40 in the small intestine (0.9 l 0.3 mU/mg protein) (Table 8). This means that strong inactivation of feed phytases occurs during digestion in the stomach and the small intestine. Lower phytase activity in the small intestine thanin the stomachwas also observedbyJongbloed et al. [167] and Rapp et al. [178] and reduced phytase activity as a consequence of proteolytic digestion in the proximal small intestinewas already describedbyScheuermann et al. [176].In rats Miyazawa andYoshida [184] also observedlowerphytase activityin the colorectal thanin the upper small intestinal chyme. In the diet free of phytases (extruded diet) the phytase activitywasverylow(0.2 l 0.1 mU/mg protein) and did not change significantlyin the stomach chyme (0.3 l 0.2 mU/ mg protein).Aslight increaseofthephytaseactivityinthe small intestine was probably due to phytases of microbial origin and of detached mucosal cells (0.7 l 0.2 mU/mg protein) and was the same in both diets (Table 8). This phytase activity in the small intestine, however, is too low to contributetoanyphysiologicallyrelevant degradationof inositol phosphates in the small intestine (Figs. 2 and 3). If microbial phytases are added to the diet, phytase activity is higherandphytatehydrolysisalso occursinthe small intestine( Fig.4, dietsBandC) [167, 177].Lowphytate degradationinthe small intestineisalsoin accordancewithPointillart et al. [185] Hu et al. [186], Iqbal et al. [165] and Davies and Flett [162]who reportedlowphytase activityin mucosa cell homogenates of pigs, humans and rats, respectively. The lack of detectable inositol phosphates degradation in the small intestine, however, is not only due to the low activity of endogenous phytases but also to the low solubility of inositol phosphates at intestinal pH 6.6 l 0.2 (Table 7) [159] which may restrict their susceptibility to enzymatichydrolysis. Activity of alkaline phosphatases in the diet and also in the stomach is nondetectable, but high in the small and large intestineandis independentofthedietfed(Table8).It indicates that alkaline phosphatases are of endogenous and not of dietary origin. Compared to phytases, the activity of i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Figure 5. Phytate hydrolysis during the gastro- intestinal digestion in pigs – pathways of the inositol phosphates. Plant phytases —; microbial phytases ---; major pathways fi; minor pathways fi: *, **, and ***: enantiomers of inositol phosphates are not distinguishable by HPLC, *: Ins(1,2,5,6)P4 is equivalent to L-Ins(2,3,4,5)P4, **: Ins(1,2,6)P3 equivalent to L-Ins(2,3,4)P3, and DL-Ins(3,4,5)P3 equivalent to DLIns( 1,5,6)P3 (with permission from Taylor & Francis, London, UK, http://www. informaworld. com [159]). alkaline phosphatases in the small intestine is about two orders of magnitude higher(l150 mU/mg protein) (Table 8). This is in accordance with Iqbal et al. [165] who found in small intestinal mucosal homogenates of humans 1000 times higher activity of alkaline phosphatase than that of phytases. In the colon, the alkaline phosphatase activity declines(l35 mU/mg protein) (Table 8), probably due to subsequent inactivation in the large intestine and the lower secretion of alkaline phosphatases in the colon rather than in the small intestine. Similar resultswere also reported for rats [184]. As the activity of alkaline phosphatases is much higher thanthatofphytasesinthe intestine,the questionwasraised whether or not the high alkaline phosphatase activity explains the stronghydrolysis of the inositol phosphates in the large intestine. Thus, in an ex vivo experiment [173] aliquots of small intestinal chyme of the pigs fed a diet with activephytases[159]were either incubatedatpH6.2or8.7 to differentiate betweenphytases and alkaline phosphatases which might be active in hydrolysing inositol phosphates under the intestinal conditions [173]. The pH 6.2 was selected as it is almost optimal for phytases but nonoptimal for alkaline phosphatases, while pH 8.7 is nonoptimal for phytases but almost optimal for alkaline phosphatases [187]. The results show that hydrolysis of phytate and the lower phosphorylated inositol phosphates was high at pH6.2,whileatpH8.7nosignificant changesofthe inositol phosphate concentrations(p a 0.05) occurred but the concentration of InsP6onlydecreasedbytrend [173]. These findings indicate that onlymicrobial phytases are involved inthehydrolysisof inositol phosphateswhile alkalinephosphatasesobviouslydo not contributetoa relevant degreeto the degradation of inositol phosphates in the large intestinal chyme.Thisistrueevenifthephytase activityislow(Table 8), but the residual time of the chyme in the large intestine isobviouslysufficientlongforthe strongphytatedegradation observed in the large intestine and faeces. Bitar and Reinhold [163] studied the pH dependence of phytases and alkaline phosphatases from human mucosal homogenates www.mnf-journal.com S343S343 Mol. Nutr. Food Res. 2009, 53, S330 – S375 and found pH optima for phytases and alkaline phosphatases of pH l 7.5 and l9.5, respectively. Since at pH 8.7, selected for the incubation of small intestinal chyme, the degradation of inositol phosphates in the pigs’ chyme was very low, it indicates that phytases and alkaline phosphatases from human mucosa cells also might not be involved in relevant degree in the intraluminalhydrolysis of inositol phosphates in the small intestinal chyme in pigs [173]. 4.6 Pathway of the stepwise phytate degradation in the gut Based on the analysis of the inositol phosphates isomers in the gastro-intestinal chyme, Schlemmer et al. [159] elucidated the pathway of the stepwise phytate degradation in the different parts of the gut (Fig. 5). The inositol phosphates analysis does not only show that phytate is hydrolysedby6- and 3-phytases from feed and microbial origin, respectively, but that further inositol phosphates, such as DL-Ins(1,4,5)P3 and DL-Ins(1,3,4,5)P4, which are not formed within the known pathwayof the 3-and 6-phytases, are also present in the gastric chyme. As Ins(1,4,5)P3 and Ins(1,3,4,5)P4 possess second messenger activity and are highly important for the intracellular metabolism, it has been assumed earlier that most probably these inositol phosphates will not exist extracellular to prevent disordered cell metabolism after their absorption. Thus, it is highly remarkable that their existence could be proven extracellular in the gastric chyme of pigs [159]. Even though their concentration is low, their physiological role in the gut requirestobe clarified. 4.7 Phytate balance during the passage throughout the gut Phytate degradation for the passage through the digestive tract was nearlycomplete for both diets with high (control diet: 97.4 l 2.3%) and low intrinsic feed phytases (extruded diet: 97.7 l 2.2%) [159]. This shows that during digestion inthegutphytatewillbe almost completelydegraded, independentofthe intrinsicfeedphytaseactivity.Ifactivephytases are presentin the diet, strongphytatehydrolysis then occursin the stomach.Ifphytases are inactivatedby processing, phytate degradation then mainly occurs in the large intestinebymicrobialphytases. This is comparable with humans, as most food containing phytate is heat treated in one way or the other, thus inactivating phytases. If diets contain high amounts of wheat or rye bran, strongphytatehydrolysis can thenbe assumedin the stomach and the rest of phytate will be degraded almost completelyinthe large intestine. Similar phytate degradation in the whole gut(l92– 100%) has also been reported by other groups [167, 181, 188]. It should be pointed out, however, that phytate degradation in the lower part ofthe intestine depends on the diet i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim arycalcium level. The redistribution of inositol phosphates from solidto liquid phaseis probablyaffectedbyincreasing calcium content in the feed and as a result enzymatic hydrolysis is subsequentlyreduced. Brune et al. [189] studied the effect of high phytate intake over a long period of time on the intestinal absorption of nonheme iron. In vegetarians with a high phytate intake(l1100 mg phytic acid/day) and in control subjects with lowphytate intake(l370 mg phytic acid/day) over several years, they found an almost identical inhibition of iron absorption from wheat rolls with iron labelled bran (55Fe, 59Fe). This indicates that in the course of high phytate consumption, no adaptation to reduced iron absorption occurs over time and allows the assumption that high phytate intake does not induce endogenous or microbial phytase activity in the intestine. 4.8 Conclusions Studiesin humans showedthat37 –66%of dietaryphytate is degraded during digestion in the stomach and small intestine when the diet is rich in plant food phytases [151, 156, 158]. As most plant food such as whole grain products, cereals and legumes – the main sources of dietaryphytate intake – are processed or heat treated either during food productionor preparationin oneway orthe other,phytases in prepared food should probablybe inactivated toa large extent. This means that in humans, consuming Western style diets withlowphytase activity,phytatedegradationin the stomachandthesmall intestinebyfoodphytasesisvery limited.From studiesinpigs, sheddinglightonphytatedegradation in different parts of the gut, can be concluded that in humans the mainphytatehydrolysis occurs in the large intestine by means of microbial phytases [159]. Studying the stepwise phytate degradation in the gut, clarity could be gained on the pathwayof the gastro-intestinal inositol phosphate degradation in different parts of the gut as well as the enzymes involved [159]. Even though the activity of alkaline phosphatases is much higher than that of phytases in the large intestine, the ex vivo studies let assume that alkaline phosphatases do not contribute to the inositol phosphate degradation to a relevant degree and microbial phytases are responsibleforthe strongphytatehydrolysisinthe large intestine [173]. During the stepwise degradationof dietaryphytatein the gut, large numbers of inositol phosphates are formed. Among the inositol phosphates typicallyformedby 3-and 6-phytases, other inositol phosphates such as DLIns( 1,3,4,5)P4 and DL-Ins(1,4,5)P3, showing intracellular signal transduction function, are detected in the gastric chyme [159]. Even though their concentration is low, their extracellularexistenceis remarkable.As theirphysiological propertiesinthegut arenotyetunderstood, theirphysiological role in the gastric chyme needs to be clarified. This is also true for the major inositol phosphates formed during www.mnf-journal.com S344S344 U. Schlemmer et al. phytate digestion in the gut. The higher phosphorylated inositol phosphates are, the lower the solubility of their metal complexes in the gastro-intestinal chyme. Consequently, high phosphorylated inositol phosphates, and especially phytic acid, bind strongly to minerals and trace elements under the acidic conditions of the gastric chyme and form soluble complexes. During the passage from the stomach to the small intestine and with increasing pH they precipitate. Thus, the availability of bound trace elements and minerals is reduced, the intestinal absorption of these elements affected and under certain dietary conditions and imbalanced nutrition this may lead to serious deficiencies of these elements in humans and animals. The changing of the inositol phosphate solubility in the gastro-intestinal chyme during the passage throughout the gut confirms the long assumed mechanism for the phytate inhibition of the bioavailability of trace elements and minerals in the gut [159]. For optimum physiological benefit of phytate, the phytate content in the upper part of the gut has to be low to avoid adverse mineral interactions but high to use its antioxidative and anticancer activity as well as its contribution of preventing kidney stone formation in humans. Thus, the fateofphytate during digestioninthegutisof principlesignificanceto assessitsroleinhuman nutrition. 5 Absorption and bioavailability of phytate and other inositol phosphates Under physiological pH (l6–7), phytic acid is highly chargedandeightoftwelvehydroxylgroups carrynegative charges (Fig. 1). Due to the small size of the inositol molecule, phytic acid shows an extremelyhigh negative charge density.For these reasons,it has been assumed thatphytic acid or phytate in all probability cannot cross the lipid bilayer of plasma membranes. As adequate carriers have not yet been detected in the gut, the gastro-intestinal absorptionofphytatewasconsidered rather improbable. Recent studies in cell lines, rats and humans, however, give some evidence to the gastro-intestinal absorption of phytate.Duetovariousdifficultiesinthe sensitive determination of phytate and other inositol phosphates in body fluids, such as blood plasma or urine, only a few studies have been carried out so far. 5.1 Cellular uptake of phytate and phytate absorption in rats NahapetianandYoung[39] observedin1980thatthe radioactivity of 14C labelled phytate was almost quantitatively absorbed in rats and recovered in blood, organs, bones, urine and expiredCO2. Thiswas one of thefirst reports indicating absorption of phytate or of its degradation products in rats. Sakamoto et al. [40] also administered radiolabelled phytate to rats and found broad distribution of radio- i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 Table 9. Phytate concentration in organs and fluids of rats fed either a diet free of phytate (AIN 76 A) or the same diet with added calcium –magnesium phytate (AIN 76 A + 1% Ca – Mg – phytate) for 12 wka) Organs/ Phytate Diets liquids content Phytate free Phytate free diet (AIN 76 A) diet + 1% Ca – Mg – Phytate Brain lg/g (dw) 2.55 l 0.30 24.89 l 1.14 Kidney lg/g (dw) 0.048 l 0.005 1.71 l 0.06 Bone lg/g (dw) n.d. 1.79 l 0.06 Urine mg/L 0.06 l 0.04 3.20 l 0.06 Plasma mg/L 0.021 l 0.004 0.22 l 0.01 Values in each line were significantly different (p a 0.05). a) Adapted from [191]. activityinorganssuchastheliver, kidneysandgut.Inblood and urine, however, onlytraces of radioactivity were measured. Quitealotof radioactivitywasdetectedinthe GI-tract and the chromatographic analysis revealed that InsP3 was possiblythe main inositol phosphate in the gastric epithelial cells along with traces of other low phosphorylated inositol phosphates while high phosphorylated inositol phosphates like InsP5and InsP6were absent. Whether or not InsP3really was present in this tissue as the predominant inositol phosphate remains to be clarified as also other organic phosphates such as nucleotides, being not removed during the sample preparation, may coelute during chromatographic separation from the column under the same conditions as some of the inositol triphosphate isomers (see Section 9.4 andFig. 9). The same experimental uncertainties also exist for a cellular uptake study using 3H-InsP6 in different cells (YAC-1, K562 and HT-29 cells), showing major radioactivity in the chromatic fractions of InsP1, InsP2 and InsP3 [190] which, however, could also derive from other compounds such as nucleotides. These findings do not prove beyond anydoubt the cellular uptake of lowphosphorylated inositol phosphates formed by phytate hydrolysis. Recent studies on cellular uptake of radiolabelled InsP6byMCF-7 breast cancer cells, however, showed 86% of the absorbed radioactivity in the cytosol, 7.4% in the nuclear or membrane pellet and 58% localised chromatographicallyin the InsP6 fraction. Besides InsP6, also InsP4was detected,giving evidence to cellular uptake of InsP6 and its metabolic degradation product InsP4[41]. Ferry et al. [42] also found some evidence for the internalisation of InsP6into HeLa cells via pinocytosis as colchicine, a probable pinocytosis inhibitor, blocked the cellular uptake of phytate completelyand as the chromatographical analysis of the cells after incubation with 3H-InsP6 showed most radioactivity in the InsP6 and other high phosphorylated inositol phosphate fractions. Even if the procedure of washing the cells to separate intracellular from intracellular inositol phosphateswas not describedin detail, thesefind www.mnf-journal.com Mol. Nutr. Food Res. 2009, 53, S330 – S375 S345 Figure6.Phytateplasmaconcentrationafteringestionofasingledoseof1400mgdodecasodiumphytate(1515lmolofInsP6)inhumans;(a)significantlydifferentfromphytatecon- centrationattime0h(pa 0.05)(withpermissionfromIOSPressBV,Amsterdam,NL[195]). Figure7.Urinaryphytateexcretioninhumans2–8hafterapplicationofthreedifferentphytatesupplements:400mgLit- Stop(Ca-Mg-Phytate,467lmolInsP6),3200mgCell-Forte' (Ca-Mg-Phytate,3734lmolInsP6)and1400mgNa–phytate(1515lmolInsP6)(a)vs.2h(pa 0.05);(b)vs.4h(pa 0.05). (withpermissionfromIOSPressBV,Amsterdam,NL[195]). ingsmaygivefurtherevidenceto cellular uptakeofInsP6 bycultivated HeLa cells. Grases et al. [191] feda purified control diet freeofphytate and the same diet with addition of 1% sodium phytate to rats for7days. The plasma concentrationofphytic acid was significantly higher(p a 0.05) for rats receiving the sodium phytate containing diet (0.022 l 0.01 mg/L) than for the rats fed thephytate free control diet (0.21 l 0.04 mg/ L). The same was observed in the urine. After feeding the phytate diet, significant(p a 0.05) higher phytic acid concentration in the urine was determined (3.20 l 0.06 mg/L) compared to the control diet (0.06 l 0.04 mg/L). In the organs of rats, fed the phytate containing diet, significant higher phytic acid concentrations(p a 0.05) compared to the control dietwere also observed(Table9), with ten times higher phytic acid content in the brain (24.89 l 1.14 lg/g) than in the kidney (1.71 l 0.06 lg/g). The study indicates that thelevelofphytic acidin body fluids and organsvaries stronglywiththephytate contentofthediet applied. Another study on rats followedthephytic acid concentrationintheurineafter applicationofaliquiddietwithdifferent contents of phytic acid (61, 182 and 425 mg/L) for 100 days [192]. With increasing dietary phytic acid content, enhanced phytic acid excretion in the urine was detected. Maximum urinary excretion of phytic acid was found for the applied phytic acid concentration of 182 mg InsP6/L, correspondingtoadailyintakeof6.9mgInsP6 per rat or to 20.9 mg InsP6/kg body weight. When phytate was omitted from the diet, the InsP6 concentration in the urine declined immediately and became undetectable(a0.1 mg/L InsP6) after22days [192]. Thesefindings showvarying urinary excretions of phytate in rats depending on different dietary phytate levels and also indicate absorption of phytate in the i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim rat's gut. Other studies in rats also show strong effects of high dietaryphytate intakeonphytic acid concentrationin plasma and brain [193, 194]. 5.2 Phytate absorption in humans Grases et al. [195] showed in three women and four men that the plasma concentration of phytic acid was low (0.106 l 0.015 lmol InsP6/L)whenthe dietwaslowinphytate( a100 mg phytate intake per day) and reached a maximum of 0.181 l 0.030 lmol InsP6/L4haftera bolus application of 1.4g sodium phytate (1.5 mmol InsP6)(Fig. 6). When a diet with an average phytate content (l700 mg phytate per day)wasconsumedfor16days,the plasma concentration of phytic acid reached a level of 0.393 l 0.045 lmol/L. Applying 0.4g calcium– magnesium phytate (0.46 mmol InsP6) or 1.4g sodium phytate (1.5 mmol InsP6) after a 15-day period on a low phytate diet, the urinary excretion of phytic acid was 0.008 mg (12.1 nmol InsP6) and 0.013 mg (19.7 nmol InsP6) after 2h, respectively, and reaching after8h a significant(p a 0.05) higher urinary phytic acid excretion of 0.052 mg (79 nmol InsP6)and 0.058 mg (88 nmol InsP6), respectively (Fig.7)[195].As higher applicationsof calcium–magnesiumphytate such as 3.2g(3.7 mmol InsP6)did not result in significantly higher urinaryphytic acid excretion compared to 0.4g calcium– magnesium phytate (0.46 mmol InsP6)or 1.4g sodiumphytate (1.5 mmol InsP6), this lets assume that obviously limited capacity of phytic acid absorption and/or urinaryphytic acid excretion in humans exists [195]. This study shows varying phytate concentrations in plasma and different phytate excretion in urine due www.mnf-journal.com S346S346 U. Schlemmer et al. to changing dietaryphytate consumption,giving someevidence to phytate absorption in humans depending on the dietary phytate applied. Phytic acid was measured in this studyby a nonspecific but highly sensitive method, determining phytic acid on the basis of the inositol content after anion exchange separation onAG1X8 which mayinclude also other high phosphorylated inositol phosphates and hence may result in some higher phytic acid content but with the same error for all plasma and urine samples [195]. Other studies also indicateastrong relation between dietaryphytate intake andphytate absorptionin humans. One study showed significantlylower phytate excretion (p a 0.05) when phytate intake in healthy subjects and in calciumoxalate kidney stone formerswas changed from freely chosen phytate containing diets to an experimental diet free ofphytate [196]. Another studyshoweda strong increaseof urinaryphytate excretion up to 440 lgphytate(a667 nmol InsP6)within8hafter applicationof 400mgof calcium– magnesiumphytate (0.46 mmol InsP6)[197]. 5.3 Conclusions Fromthedifferent studiesitcanbeconcludedthatincultured cells,in ratsand humans,phytate absorptionmayoccureven thoughthe mechanismof absorption remainstobe clarified. Support for phytate absorption in humans comes from the tight correlationofphytate consumedand increasingphytate concentration in plasma and enhanced urinary excretion [195–197]. Similar resultswere foundin rats[191 –194].As the absorptionofdietaryphytateis independentofthefilling stageofthe stomach [197],itis assumed thatphytate absorption probablytakes place in the small intestine. The studies, moreover, suggest that the phytate absorption in the gut is verylow and does not exceedafew percent(f2%) [192], whichwasalso concludedfrompig studies, determiningthe disappearance ratesofphytateand other inositol phosphates from the gastro-intestinal tract [177]. The great number of in vitro experiments and animal studies, showing anticancer or anticalcif ication activities after application of sodium phytate or calcium-magnesium phytate, moreover, stronglysupports the absorption of phytate or of its degradation products in the gut, as without gastro- intestinal absorption the observedeffects are unexplainable[ 32,41].The assumptionsthatphytatemightbehydrolysed completelyin the gut and be rephosphorylated intracellularily after absorption of myo-inositol and phosphate requires to be proven. The studies in rats and humans, applying phytate and detecting phytic acid in plasma and urine after comparablyshort time [191 – 197], do not seem to supportthesehypotheses. Whetherphytate or other inositol phosphates might be absorbed via active transports, pinocytosisor otherwaysof absorptionorbya combination of these, also remainstobe clarified. A recent study of Irvine and coworkers [198], however, found no detectable InsP6 either in human serum and plate- Mol. Nutr. Food Res. 2009, 53, S330 – S375 let-free plasma(a1nM InsP6)orin human urine(a5nM), using an enzymatic method, which determines InsP6 specifically and with high sensitivity after 32P-labelling and HPLC separation. As the statistical basis of thesefindings is vague and detailed information on the dietary phytate level of the persons is missing, it is difficult to assess whether or not these results reallymaycontradict the earlier reports in humans and rats, discussed above. 6 Influence of phytate on intestinal mineral absorption Health authorities from all over the world universally recommend increasing consumption of whole grains and legumes for health promoting diets. Wholegrain foods are valuable sources of carbohydrates, dietaryfibre, numerous bioactive compounds, vitamins, minerals and trace elements which arein shortsupplyin manycountries. Mineral malnutrition is a global problem affecting industrialised and developing countries as well. Children, infants and women at childbearing age are primarily affected [199, 200]. Under nonvaried and nonbalanceddietaryconditions, phytate may affect the bioavailability and in consequence the statusof iron,zincand calcium[24 –29,201].It should be stressed that in many countries wholegrain cereals and legumes are among the most important food sources for minerals and trace elements but also contain high amounts of phytate and polyphenols. Thus, when advice is given for good dietary sources of minerals and trace elements, various interactions between the minerals and trace elements andphytic acidhavetobe taken into considerationto ensure high bioavailability and adequate supply. 6.1 Phytic acid: Chelating properties, binding capacity for minerals and inhibition of mineral absorption Phytate occurs in all edible plant seeds such as grains, legumes, oilseeds and nuts but also in lower content in roots, tubers and vegetables(Tables2 –5).Asphytic acidis stronglynegativelycharged under physiological conditions (Fig.1)[13], it showsgreat potential for complexing positivelycharged multivalent cations, especially of iron, zinc, magnesium and calcium [202]. These complexes are soluble under the acidic conditions in the stomach and precipitate at neutral pH in the intestine, resulting in poor absorption of minerals and trace elements (see Section 4). They also may affect peptides and proteins, leading to reduced protein bioavailability [155, 156, 203 – 206] and impaired enzymatic activity [205, 207]. Even if phytate seems to be most effective for impairing the mineral absorption other components like inorganic phosphate, polyphenols and nondigestible dietaryfibre reducethe absorptionof mineral and trace elements aswell [208, 209]. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com S347S347 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Studies, both in animals and humans, have shown that foodsor diets richin dietaryfibremayalter mineral metabolisminthe presenceofphytate[210,211]whileotherstudies indicate noeffectof dietaryfibre on mineral absorption [212, 213]. Numerous studies have described the negative effect of phytate on the bioavailability of minerals and trace elements [171, 205, 214 – 221] which recently was extensivelyreviewed[ 222–226]. There are, however, a number of dietary components which counteract the inhibitory effects of phytate on mineral absorption. Some evidence exists from human studies for improving calcium absorption after application of fermentable carbohydrates [227 – 229]. Other studies, which first showeddecreased calcium absorption in the presence of phytate, reported enhanced calcium availability after degradationofphytate[230,231].Thisisduetothe reduced formation of the low soluble Ca –phytate which not only affects the mineral availability of calcium in the intestine but is also poorlysusceptible to enzymatic dephosphorylation to lower phosphorylated inositol phosphates [23, 181]. Organic acids obtainedbyfood fermentation also counteractthe inhibitoryeffectsofphytateand enhancezincabsorption in the presence of phytate [232]. The same effect was reported for dietaryprotein, wherebythe content and type of protein along with the content of zinc are important for the improvementofzinc absorption[233,234]. Dietslowinanimal protein result in low zinc absorption in the presence of phytate [235] and high calcium content increases the inhibitory effect on zinc bioavailability by forming calcium – zinc–phytates. Insufficientzinc intake,however,isthemain causeforzincdeficienciesinhumans[236]. Irondeficiencyis oneofthe mostprevalentdeficiencies in theworld and mainly causedby insufficient iron intake [237]. Moreover, the sources of iron, hem or nonhem iron andthetotal compositionofthedietareofgreatimportance for iron bioavailability. The content of phytate in food has beencloselyrelatedtoironabsorptionandhighphytatecontent results in lower iron absorption [126]. Phytic acid decreasesthe solubilityofironbyforminglowsolubleiron– phytate [238] and for this reason iron availability from the chyme is affected and the intestinal iron absorption inhibited. This inhibition can be counteracted by complexing agents like proteins, peptides, beta-carotene, organic acids and ascorbic acid [239 – 246]. Ascorbic acid,moreover,stops the oxidation of ferrous to ferric iron and thus preventing the formationoftheverylowsolubleFe3+–phytates. 6.2 Phytic acid interactions with toxic trace elements (Cd, Pb) Duetothehigh bindingaffinityofphyticacidtometalions the question was raised whether or not phytate could be applied to affect the bioavailability of toxic trace elements such as cadmium or lead. In infant cynomolgus monkeys (Macaca fascicularis) strong reduction of the blood lead i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim concentration following the addition of phytate to the diet was observed [247]. In adult human volunteers, who ingested 203Pb as lead acetate, the absorption of lead was affectedby a mealhighin calciumandinphytate [248].In ratstheeffectof calciumandphytatewas studiedregarding theabsorptionofleadand cadmium.The additionofcalcium or phytate significantly reduced the lead accumulation in bones(pa0.001) and in blood and liver(pa0.05) [249] and applying phytate and calcium together, the strongest inhibitionofthetissuelead retentionwasreported.Forcadmiuma significant increaseintheliverand kidney accumulationby calcium(pa 0.05) was observed which after supplementationbyphytate, was reduced againandnofurthereffectof phytate on the cadmium tissue levels was detected [249]. Applying a fish-meal diet in rats either with or without sodiumphytate,Yannaiand Sachs foundnoeffectonthecadmium, leadand mercuryabsorptioninthisdietafter addition ofsodiumphytate [250].Inastudy with albino rats, Rimbach et al. [251]fed three diets basedoneggwhiteand cornstarch and supplementedbyzinc(15mg/kg)and cadmium(5mg/ kg). The control diet (diet 1) was free of phytate and active phytases while the both experimental diets contained either 0.5% sodium phytate (diet 2) or 0.5% sodium phytate plus microbial phytases (2000 U/kg) (diet 3). Liver cadmium concentrationin ratsfeddiet2was significantlyhigherthan in rats fed diet 1 (control diet) or diet 3. By trend similar resultswere foundfor kidney cadmium accumulation. These results show that the high cadmium accumulation in liver and kidneys were due to the sodium phytate added as after phytate degradationbyphytases the high cadmium accumulationwasreduced [251]. The resultswere confirmedbythe same authors in another rat study,showing also significantly higher cadmium liver and kidneyaccumulation after applicationofasodiumphytate containingdiets(0.5%)andalsoa reductionof the accumulated cadmiumbysupplementation of microbialphytases (2000 U/kg) [252].Alater rat studyby Rimbach and Pallauf [253] showed, however, only a slight increase in the cadmium accumulation in the kidneys and no significant alteration of the liver cadmium content with increasing dietaryphytateupto1.4%(3.5 –14.0gNa –phytate/ kg). As the zinc content of these diets was more than twice as much (225 mg Zn) as that of the earlier ones (100 mgZn) [251, 252], the authors concluded that under the conditionsofhigh dietaryzinc contents,phytatemighthave only littleeffect on the carry overof cadmiumingrowing rats [253].In mice Lind et al. [254] studied the accumulation ofcadmiumfromfibrerichdietsbasedonwheatbran,sugarbeetfibreand carrotsin comparisontoa semisynthetic controldiet supplementedwithCdCl2.Alldiets containedacadmium content of 0.05 mg Cd/kg. The wheat-bran diet showed significantlylower fractional cadmium accumulationintheliverand kidneys(%oftotalCd intake), indicating alower absorption of cadmium. The authors discussed that most probably the higher dietary content of inositol hexaand inositol pentaphosphates, forming with cadmium low www.mnf-journal.com S348S348 U. Schlemmer et al. soluble Cd–phytate complexes, contributed more to the lower cadmium absorption than the elevated Zn level [254]. Similar resultswere reportedbyWing [255]whodetermined the fractional cadmium accumulation(109Cd) from different diets based on either whole wheat, bran or endosperm with different levels of phytate (4.2, 7.6 and 0.3 mmol IP6/kg, respectively). The fractional accumulation of 109Cd in the liver and the kidneyswas significantlylower(pa 0.001) in rats fed the phytate rich whole wheat and bran diets comparedtothelowphytate endospermdiet (controldiet). The bioavailability of cadmium from cow's milk formula, soyformula, wheat/oat/milk formula, whole meal/milk formula and water were compared in rat pups [256]. The pups received a single oral dose of one diet labelled with 109Cd, 0.1or0.3mgCd/kgbodyweight.Thelowest cadmiumbioavailability was found in the cereal-based formulas, explainedbythe cadmium binding to dietaryfibre andphytate [256]. 6.3 Conclusion Dietary phytate probably affects the bioavailability and retention of lead in rats, monkeys and humans [247 – 249]. Inhibition of the cadmium bioavailability and retentionby phytate in mice was reported [254, 255], while under certain dietary conditions, such as low dietary zinc content, the cadmium absorption and retention in kidneys was even improved by addition of sodium phytate [251, 252], and normalised after phytate degradation or at high dietary zinc content [253]. Thus, the concentration and relation of various minerals, trace elements and phytate in the diet seems to be important for the effects of phytate on the cadmium bioavailability and retention. From the different results it can be concluded that in respect to the bioavailability and retention of cadmium, phytate as an endogenous part of the food matrix probably reacts differently than sodium phytate supplemented to the diet. Moreover, as the study with a fish-meal based diet shows no inhibitory effect of phytate on the intestinal absorption of cadmium, lead and mercury [250], it can be assumed that the kind of proteins consumed also might play an important role in phytate effecting the bioavailability and retention of toxic trace elements. 6.4 Determinants of the phytate – mineral interaction Several factors are present which govern the inhibitory effect of phytate on mineral bioavailability: pH, content of minerals and phytate, solubility of phytates and concentrationof enhancers or inhibitors. Moreover,greatvariations exist for the affinity of phytate complexes with different valent cations showing increasing binding strength from mono-to multivalent cations(e.g.Na+,Ca2+,Fe3+)[257]. Furthermore, different phytate – mineral ratios in foods and i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 diets seem to be also important for phytate inhibition of the intestinal absorption of minerals and trace elements and the phytate concentration in the diet has to exceed a certain level to have a substantial effect on the bioavailability of minerals and trace elements [258].To predict the bioavailability of minerals from diets just based on the phytate content in foods is not reliable as all other factors involved in the phytic acid – mineral interaction have to be taken into consideration [223]. The same is true for the quotient of Zn/ Ca/phytic acid, widely applied for assessing the mineral bioavailability [132, 135, 137], which also disregards the complexing agents in the diet, competing with phytic acid for the binding of metal ions. 6.5 Conclusions Growing interestinwholegrainsandwholegrain products in developing and industrialised raised the questions on these foodsonthe mineral status.High contentofphytatein these products has been considered a major factor for limited mineral bioavailability, resulting in iron, zinc and calcium deficiencies. The inhibition of the intestinal metal absorption, however, can be counteracted by many food compounds such as organic acids and complexing agents, ascorbic acid, food fermentation products, etc. competing with phytic acid in the binding of minerals and trace elements. Thus, it will be assumed that in well balanced diets the inhibitory effects of phytic acid is low and little evidence exist from nutritional surveys that in well nourished populationgroups dietaryphytate may seriously effect the status of iron, zinc and calcium. Under malnutrition and nonbalance dietslowin minerals and essential trace elements but high in phytate, however, the situation is completely different.Vulnerablegroups in developing and developed countries with inadequate intake or deficiencies of minerals and trace elements need to increase total intake of these elements via the dailydiet or to improve the bioavailability of these elements under consideration of all factors inhibiting or enhancing the bioavailability of the minerals and trace elements in the diet. Adequate strategies to prevent deficiencies of these essential elements adjusted to the specific situation are required and different approaches are possible eitherbysupplementationof the respective elements,byincreasing the contents of competing and complexing agents orby removingphytate from food. 7 Effects of preparation, processing and storage on phytate and other inositol phosphates Preparation and processing can make food healthier, tastier and safer and increase its storage stability. Preparation and processing,however, can alsoaffect the nutritionalvalueby www.mnf-journal.com S349S349 Mol. Nutr. Food Res. 2009, 53, S330 – S375 destroying labile nutrients like vitamins or by removing phytate to improve the bioavailability of mineral and trace elements. As phytate is quite heat stable up to l1008C[22], it cannot be easily removedby conventional heat treatment like cooking. Enzymatic degradationofphytate,however, either by phytases occurring naturally during food processing or by adding phytases, effectively degrades phytate in food [23]. 7.1 Thermal hydrolysis of phytate During boiling and heat treatment, phytate in food shows a high stability up to l1008C for a boiling time of 1h [22]. Only9% ofphytateisdegraded under these conditionsin soybeans(Soja hispida Max.).Table10shows the stepwise phytatehydrolysis in brown beans(Phaseolus vulgaris L.) at various temperatures and boiling times, applied for legume processing and household preparation (U. Schlemmer, 1996 unpublished results). To cock brown beanswell,a temperatureof 1108Cfor 1.5his needed after soaking the beans over night [22]. Under these conditions InsP6is degraded from 86 to 66% and lower phosphorylated inositol phosphates (InsP5– InsP3) are formed. The total sum of inositol phosphates (InsP3– InsP6), however, does not decline but increases slightly, probably due to a better release of phytate from the bean matrix. This shows that preparing beans for consumption under normal household conditions l1/4 of phytate in raw beans is degraded and quantitatively transformed to InsP5– InsP3.If the temperature is raised veryhigh, e.g.to 1408C, which is beyond any practical conditions of household preparation but relevant for certain industrial processing at a quite extended boiling time of 45 min, phytate is only reduced from 86 to 36% along with a total loss of inositol phosphates (InsP6– InsP3) of only28%. These results demonstrate the high stability of phytate during thermal preparation of brown beans. Table 10givesanoverviewonhow boiling temperature and time governthethermalhydrolysisofphytateinlegumes during processing and preparation and confirms earlier reports [17, 18, 125, 222, 259, 260]. 7.2 Phytases Phytases (myo-inositol hexaphosphate phosphohydrolase, EC 3.1.3.8 and EC 3.1.3.26) are found in plants, microorganisms and in animal tissues as well [163, 261– 263]. Theyhydrolysephytate aswell as other organic phosphates like lower inositol phosphates, nucleotides, etc., however, withdifferentaffinityandefficacy[187,264].The stepwise cleavingof phosphategroups leadstolower phosphorylated inositol phosphates and inorganic phosphate andfinally stops at myo-inositol. Different phytases show different cleaving specificities for hydrolysing phosphate groups formphytate, resulting in different inositol pentaphosphate i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Table 10. Stepwise hydrolysis of phytate (%) in brown beans by hydrothermal treatmenta) P InsPs InsP6 InsP5 InsP4 InsP3 [%] Raw brown 100 86 14 – – beans 1108C 30 min 113 85 23 2 3 60 min 107 72 24 5 3 90 min 104 66 26 10 6 1208C 30 min 111 68 28 10 5 60 min 102 58 27 11 6 90 min 91 47 23 11 10 1308C 15 min 113 68 29 11 5 45 min 96 50 25 13 8 90 min 79 38 21 10 10 1408C 15 min 90 54 24 9 3 45 min 72 36 21 10 5 90 min 58 21 21 10 6 a) Brown beans (Phaseolus vulgaris L., 20 g dw/50 mL H2O) were heated under pressure in a pressure cocker and the thermal degradation of inositol phosphates determined [22]. After heating, the brown beans were freeze dried, ground (a1 mm, particle size) and extracted with HCl (2 g/ 40 mL 2.4% HCl, 3 h, 228C). Samples were prepared and inositol phosphates with different numbers of phosphate groups were determined by RP HPLC in according to Sandberg and Ahderinne [398]. Total inositol phosphates P (InsPs) of the respective samples were summed to 100% and the total loss of InsP6 – InsP3 during the heat treatment was calculated. Results are mean values of double determination. isomers [187, 265 – 267]. There are twointernationallyclassifiedphytases: 3-phytase (EC 3.1.3.8) and 6-phytase (EC 3.1.3.26), named afterthe positionofthefirstphytatephosphorester bond hydrolysed [209, 268]. The 3-phytases are primarily of microbial origin [261– 263] while the 6-phytases are mainlyofplant sources [269]. There are,however, exceptions: Escherichia coli phytases are 6-phytases and soybean phytases 3-phytases. One predominant inositol phosphate seems to be formed at anystage of the stepwise inositol phosphate degradation and the rate of hydrolysis decreases with the degree of phosphorylation [187, 202, 265 – 267]. The optimal temperature for phytase in cereals is 458C, with an approximate stability limit at 558C andapHoptimum at 5.0– 5.6 [269]. Microbial phytases produced by fungi often show two different pH optima, one at 2.5 and oneat5.0[187,270 –272]witha temperature optimumof 588C, and with nearlyno activity A688C. Bacterial phytases such as Bacillus subtilis showa neutralto alkalinepH opti www.mnf-journal.com S350S350 U. Schlemmer et al. mum (pH l7.0– 7.5) [270] and most of them have optimum activity at l508C. Phytases have been studied in a great number of plants and have been isolated and characterised from different plant sources. High phytase activity is described in seeds [273] where they seem to be mainly associated with the aleurone tissue but to a minor extent also with the endospermand scutellum. Cerealphytases are present withvarying activity in all unprocessed cerealgrains, with the highest activity found in rye and with lower activity in wheat, oat, spelt and corn [274]. Phytase activity also varies with the harvestyear and cultivars [275]. Unprocessed oat shows similar phytase activity as wheat [276] but due to the conventional heat treatment of oat to avoid lipid rancidity, oat phytases are largely inactivated [277, 278]. During food processing and preparation, predominantlyduring soaking, malting, germination, fermentation and bread making, phytases of plant or microbial origin are widely applied to reduce phytate content in foods to improve the bioavailability of minerals and trace elements. 7.3 Soaking, malting and germination of cereals Soakingisacommonlyused methodas pretreatmentofgermination, malting, etc. Soaking may last for short periods (15–20min) or long periods (12–16h). The soaking medium used depends upon the type of seed. In the household, cereals and legumes are most often soaked in water overnight. At optimal conditions for phytases (558C, pH4.5–5.0)phytate mightbe reducedeffectivelybysoaking [188].Asphytateiswater soluble, somephytate removal canbeobtainedbydiscardingthesoakwater[172]. Phytase activity increases considerably during germination [279], but there aregreat differences between different cereals. Barley has shown an increase of phytase activity up to 11 times of the original one [280], while phytases in wheat,rye and oats increased 4.5, 2.5 and9times from the original activity [277]. Thephytate contentwas reducedby 16% in barley [280], and 30% in wheat and rye and 17% in oats [277]. In malted wheat, rye and barley, ground and soaked for 2h, almost total hydrolysis of phytate was obtained. Oats needed a longer time (up to 17 h) to reach complete reduction [281]. Optimal conditions for hydrothermal processes of whole kernels of barley tohydrolyse phytate up 95– 96% have been developed, using two wet stepsandtwodry steps, followedbydrying [282]. Degradation seems to be highest in the scutellum cells and less in the aleurone layer due to changes in the microstructure of the phytate globoids in the barley during hydrothermal processing [283]. 7.4 Soaking and germination of legumes Supplementation of cereals with legumes rich in protein is consideredtobeeffectivetofight protein-calorie malnutri i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 tion in many developing countries. A common method of processinglegumesto reducephytateis soakingandgermination. As these foods are the main sources for protein in many developing countries, it is important that phytate is reduced to a minimum to obtain bioavailability as high as possible for both protein and minerals. Before pulses and legumes are consumedthey are dehulled and prepared with subsequent soaking, germination, fermentation, roasting and autoclaving [284]. The decrease in phytate content due to germination has shown to be highest in pigeon peas (65.8%), followed by chickpeas (64.1%), bean curd (40.6%), soybeans (38.9%) and mung beans (37.2%) [285]. Other studies on beans [17, 286] have shown considerably lower reduction than reported in this study. Even if fermentation has been shown to result in a decrease of phytate content, this method was less effective than germination. It should however be pointed out that these two methods are the most effective waystolowerthephytate contentinlegumesand pulses. Soaking of peas for up to 12h decreased the content of phytate no more than 9% [287] while other studies showed no effect of soaking (16 h, 228C) on the phytate content in peas, lentils or beans [22]. A small reduction could be caused by leaching phytate into the soaking water [288]. The most effective way of reducing phytate degradation was at pH7.0 and 458C [289]. Germination results in the reduction of phytate in peas – the longer the germination period,thegreaterthephytate degradation. Whilealossof 6–8%ofphytic acidwas observed after12h,a lossof67 – 83% occurred after48h[288].Itis recommendedto dehull andsoaklegumes before consumptionto reducethephytate content and therebyincrease the nutritional quality of proteinsin these foods.With quinoa seedsitwas also demonstrated that soaking, germination and lactic acid fermentation resultedin reducedphytate contentupto98% [290]. 7.5 Fermentation and bread making Fermentation has been used for processing and preservation offoodsforalongtimeinhistory.Dueto productionoflactic acidand otherorganicacidsinthedoughpHislowered,phytases activated and phytate degraded. Part of the phytate reduction is due to the action of endogenous phytases, but exogenous microbial phytases mayalso be active in phytate degradation during fermentation [265]. It seems as if phytases normallypresent in cereals are ofgreater importance for phytate reduction, than yeast phytases added to cereals [291].Differentyeast specieshavebeen identifiedas possible sourcesofphytase[292–294],andcertain bacteria[295, 296] mayalso provide viable sources of exogenous phytase. Fermentation of maize, soybeans and sorghum has been shown to reduce the phytate content in foods [296, 297]. It has been demonstrated that combined germination and lactic acid fermentation of white sorghum and maize cruels can resultinalmost completedegradationofphytate [298]. www.mnf-journal.com S351S351 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Phytate hydrolysis occurs during the different stages of bread making, depending of the flour used for the bread, both type and extraction rate being of importance. The acidity of the dough is of utmost importance for the degradation ofphytate during both scalding and sourdough fermentation of the bread [299, 300]. Sourdough fermentation has been shown to reduce the phytate content more effectively than yeastfermented bread [301, 302].Aslight acidificationof pHto5.5inthe bread dough, obtained eitherbysourdough or by lactic acid, resulted in increased phytate breakdown [303, 304]. It is remarkable that this minor acidif ication is sufficientto degradethephytateeffectively.Evenifalower pH due to higher addition of sourdough seems to be more effective for endogenous phytases in the grain, together with the added microbial phytases, pH at 5.5 seems to be more acceptedbyconsumerswho dislikethe acidic taste.In rye bread, phytic acid is almost totally degraded to lower inositol phosphates and free inorganic phosphate during bread making when long fermentation times are used, but the degradationislowerwhenwholegrainis includedinthe recipe [305]. 7.6 Addition of exogenous phytases Phytases can be added to food during processing to reduce the phytate content. Commercial sources of the enzyme are available from wheat bran, yeast or microbial sources. Phytases seem to havea broad specificity and when isolated, e.g. from wheat, theyalso degrade phytate in oats and other cereals [276, 306]. Phytases from fungal origin mayachieve complete phytate degradation, while endogenous phytases just reducephytateby73–80% [307]. 7.7 Extrusion cooking Extrusion cookingisahigh-temperature, shorttime process using high shear forces at elevated pressure and temperature. As this method gives desirable texture to the foods, it has become a widely used technique in food manufacture, especially for cereals and legumes, like breakfast cereals, weaning foods, crispbread, snacks, sweets and vegetable proteins. Extrusion is extremely versatile with respect to ingredients and production capacity,in addition to supplying variety to shapes and textures. An increase in dietary fibre has been reported after extrusion cooking [308], but little information has been given regarding the fate of phytate. One might expect hydrolysis of phytate during this process, however, the time of exposure is too short for significant reduction. Forcereals therewereonlyslight changesin total inositol phosphates via extrusion cooking [154, 303, 309] and only a small decrease in phytate accompanied by a small increase in inositol pentaphosphates were observed [310]. Similar observations were made in oilseeds, and soybean meal was least affected by extrusion cooking [311]. The i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim observed phytate degradation during extrusion cooking is due to the high temperature and pressure occurring during the process. Theeffectof extrusion onphytatehydrolysis, however, seems to be strongly dependent on the extrusion conditions applied, as studies extruding total animal feed didnotshowanychangeinthe inositol phosphatepatternof the extruded diet [159]. 7.8 Removal of phytate by mechanical processes Mechanical removal ofphytateis dependentonthetypeof seed processed, but also on the morphological distribution ofphytateinseeds.Foralargenumberofoilseedsand cereals, themainpartofthephyticacid appearstobe locatedin the aleurone layer, but also to a minor degree in the germ [44– 46]. This is, however, not the case for, e.g. soybeans, where it seems more evenly distributed in the whole seed. In millet and oat, phytate seems to be evenly distributed both in the bran, kernel and germ [303, 312]. Milling of cereals, in which phytate is located in the outer layer ofthe seed, can cause up to 90% reduction of phytate. In corn where phytate is mainly located in the germ, removing of thispart ofthegrain willeffectivelyresultin strong reductionofphytate. Inlegumes,where the majorityofphytateis located in the protein bodies in the endosperm [215, 303], dehulling will therefore remove phytate only to a minor degree. Mechanical separation of the phytate containing compartmentsofthe seeds,however,willalsoleadtoaloss of nutrients andvaluable bioactive compounds. 7.9 Storage Several studies have indicated a decrease in phytate content both for legumes and cereals [66, 116, 313 – 315] during long storage. This reduction depends on the storage conditions (especiallyhumidity and temperature), the type of seeds and the age of the seeds [316, 317]. When stored under dryand cool conditionsno decreaseinphytateis assumed.Inastudy with cowpea flour, added to macaroni to increase phytate content, no change in phytate was observed during the six months storage under optimal conditions [318]. 8 Beneficial properties 8.1 Preventing pathological calcification Kidney stones areprevalentin humans.Itis assumed that5 – 10%ofhumanssufferfromtheformationofkidney stonesin the urinary collecting system. The formation of kidney stones is dependant on the solubility of calcium salts in the urine. The solubility, the maximum content of calcium, remaining soluble in an aqueous solution at given temperatures, is an equilibrium parameter which is independent of time and thermodynamics. It mainlydepends on the stability of the crystal lattice and the stability of the formed aqueous www.mnf-journal.com S352S352 U. Schlemmer et al. solvates (soluble species). Solubility can alsobeaffectedby the composition of the media (mainlyionic strength) due to its influence on the reactivity (related to chemical potential) of the solvates. When a system contains higher content of a solute than that corresponds to the solubility (saturated system), the system remains in an unstable state (supersaturated) and sooneror later mustevolvetothe stable conditions (thermodynamic equilibrium) through the crystallisation of the excess of solute. Precisely, the driving force that pushes the crystallisation processes is the difference between the equilibrium conditions (solubility) and the actual ones. These time-dependent processes, studiedbythe kinetics, can last for seconds or years. The general mechanism of the formation of single crystals can be explained as a result of the combination of two independent steps: nucleation and crystalgrowth. The time necessaryto generateacrystal mainly depends on the nature of the crystal, the supersaturation of the solution, the presence of performed solid particles (the so-called heterogeneous nucleants)and the concentration of crystallisation inhibitors. These latter ones, due to their structure, may interact with the nucleus or interfere in the crystallisation processes [319]. Most of the human fluids are supersaturated with regard to some substances. Thus, blood, interstitial liquid and intracellular liquid, due to their pH value (pH A 7.0), free calcium ion concentration and phosphate concentration, are supersaturated with respect to calcium phosphate (hydroxyapatite, HAP). Urine is always supersaturated with respect to calcium oxalate and depending on its pH value, is also supersaturated with respect to uric acid (pH a 5.5) or calcium phosphates (pH A6.0). In spite of this, normal crystallisation processes onlytake place in biologicallycontrolled situations, like the formation of bone and teeth. Nevertheless, uncontrolled pathological crystallisation is also frequent: e.g.in tissue calcif ication associated to cancer, calcification in cardiovascular system and calculi formation (renal, biliar, sublingual). The questionis,whycrystallisationdoesnot occur indiscriminatelyin all human fluids and only appears in pathological situations? The answer is clear: there are four main aspects which must be considered in explaining pathological crystallisation: (i) supersaturation higher than usual of the crystallising substance, and/or (ii) the presence of heterogeneous nucleants (crystallisation inducers), and/or (iii)deficitofcrystallisation inhibitors, and/or (iv)afailureofthe immune system. The crystallisation inhibitors actbydelaying the crystallisationof supersaturated substances,by avoidingthecrystallisation before the renovation of the corresponding fluid orbypermitting that the immune system eliminates cellular debris(crystallisation inducers)oreven incipient calcifications [320]. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 It has been known since the 1930s that the presence of trace amounts of molecules, such as polyphosphates, can act aswater softeners through their inhibitionofthecrystallisation of calcium salts, such as calcium carbonate. However, the useof such compoundsas natural regulatorsof calcification under physiological conditions was not explored until the 1960s. During that decade, Fleisch et al. [321, 322] showedthatpyrophosphate,anaturallyoccurringpolyphosphate, is present in serum and urine and can prevent calcificationbybindingtoHAP. However,studiesinanimal models found thatpyrophosphate can inhibit ectopic calcificationin blood vessels and kidneys only when injected rather than ingested. Oral administration causeshydrolysis and hence inactivationofpyrophosphate, resultinginasearchfor more stable analogues. Bisphosphonates, a group of polyphosphates, showhighaffinityforHAPandprevent calcification both in vitro and in vivo,even when administered orallyto animals [323]. Crystallisation inhibitors bind to crystal nuclei or crystal faces and disturb crystal development. The adsorption of such compounds to crystal faces can also inhibit crystal dissolution. It was shown that bisphosphonates may inhibit HAP crystal dissolution [324, 325] and bone resorption [324, 326, 327]. Manystudies, both in vitro experimentsand clinical trials,havealsoshownthatvarious bisphosphonates inhibit the osteoclast-mediated bone resorption [327 – 329]. It is well known that proteins are active in modulating calcif ication in mammalian tissues. These proteins can either enhance or inhibit the ability of macrophages to destroy HAP deposits (i.e. osteoclastic activity)[330 –332].Acommon characteristicof these proteins involved in the calcif ication is that they show high affinity to calcium ions. These proteins include osteopontin [333 – 336], osteoprotegerin [337 – 339], matrix Gla protein [340 – 343] and osteocalcin [332, 343]. They have shown some crystallisation inhibitor activity, however, under in vitro conditions with nonphysiological high protein, calcium and phosphate concentrations [344– 346]. Moreover, these proteinshavealsocalcification promoteractivityduetotheir heterogeneous nucleant capacity [347 – 349]. It appears that the major calcif ication modulator role of these proteins is to regulate osteoclast/osteoblast cell activity [330, 331, 350]. Recently,ithasbeen demonstrated invitrothatphytate exerts potent action as crystallisation inhibitor of calcium salts (oxalate and phosphate) [351– 354]. 8.1.1 Phytate and renal lithiasis Beyond proteins, phytate has been shown to possess strong activity of inhibiting the crystallisation of calcium oxalates or calcium – phosphates [351 – 354]. Phytate is naturally present in urine at similar concentrations to those used in the in vitro studies [355] and urinaryphytate concentration also depends on dietaryintake [190, 191]. This demonstrates the potential therapeutic effects of phytate in the treatment of calcium renal lithiasisinpreventing calculusdevelopment. www.mnf-journal.com S353S353 Mol. Nutr. Food Res. 2009, 53, S330 – S375 The effects of phytate on urolith development in a nephrolithiasis animal model using ethyleneglycolwere studied [356].Inthegroupof rats treated withphytate,the number of calcif ications on the papillarytips and the total calcium amount of the papillary tissue was significantly reduced when compared with the controlgroup treated exclusively with ethyleneglycol. A purified rodent diet (AIN-76 A), free ofphytate, has shown to possess some activity to cause kidney calcifications in female rats which was absent with nonpurified rodent diets. This suggestsa nutritional factor whichavoids these kidney calcifications. One possible candidate was phytate.Theeffectsofphytate,addedtothe AIN-76Adiet, was therefore studied in the calcif ication of kidneytissue in female Wistar rats. Rats were randomly distributed into three groups and fed the AIN-76 A diet, the AIN-76 A diet+1%phytate and the standard nonpurified diet. No phytatewas detectableinthe urineofthe ratsfedthe AIN76Adiet, freeofphytate. Urinaryphytatelevelsoftherats fed the AIN-76A+1%Na –phytate diet and the standard dietwere significantlyhigher than thoseofthe ratsfedthe phytate free AIN-76Adiet. The concentrations of calcium and phosphorus in kidneys were greater in the AIN-76 A group thanin AIN-76A+1%phytate and standardgroups. Onlyratsofthe AIN-76Agroup displayed mineral deposits at the corticomedullary junction. Thesefindings demonstrate clearlythat renal calcificationin female ratswaslow inthe presenceofphytateandhighinthe absenceofphytate [357]. Hypertension, induced by nicotine, combined with hypercalcemia, induced by vitamin D was used to induce calcificationin renal tissuein maleWistar rats whichwere feed a purified phytate free diet. These rats developed significant calcium depositsin kidneysand papillae,aswellas in kidney tubules and vessels, whereas calcium deposits were absent when phytate was added to this phytate free diet. Thesefindings showthatphytate acts as crystallisation inhibitor both in the intrapapillarytissue and in urine [358]. Furthermore, the effect of phytate on dystrophic calcification, chemically inducedby subcutaneous injectionofa 0.1% KMnO4 solution,was studied. MaleWistar ratswere randomlydivided into fourgroups treated for 31 days. A: Animals were given a purified diet free of natural phytate but with added1%Na –Phytate.In thisgroup, thephytate plasma levels (0.393 l 0.013 lM) were similar to those observed in rats consuming a standard diet. B: Animals consuming only the purified diet free of natural occurring phytate. In this case the phytate plasma levels decreased (0.026 l 0.006 lM); C: Animals consuming the same purified diet asgroupBbut received dailysubcutaneous injections of 50 lg/kg etidronate(a crystallisation inhibitor) during the last 14 days. In this case the phytate plasma levels were also very low (0.025 l 0.007 lM); D: Animals consumingthe samedietasgroupBbut6%of carobgerm,rich in phytate, was added. The phytate plasma levels (0.363 l 0.035 lM) were also similar to those observed in i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim rats consuminga standard diet. After21days plaqueformation was induced. Calcification plaques were allowed to proceed for 10 days, after which the plaque material present was excised, dried and weighed. The results show that the presence of phytate in plasma at normal concentrations (0.3 – 0.4 lM) clearly inhibits the development of dystrophic calcifications [359]. Recent studies demonstrated that phytate is naturally present in human urine and normal levels oscillated between 0.5 and3mg/L [355, 358]; the urinaryconcentrations foundinagroupof calciumoxalate active stone-formerswere significantlylower than those foundinagroup of healthy people [196] (see also Section 5). Ingestion of dietaryphytatesignificantlyreducedtherisktodevelopcalcium stones in humans [360, 361]. Thus, a clinical study in 36 calcium oxalate active stone-formers with positive urinary risk to develop calcium stones was performed. In a subgroup of 19 stone-formers the urinary risk to develop calcium stones was re-evaluated after 15 days. The other groupof17 stoneformerswas treated withphytate(120mg of phytate/day as calcium – magnesium salt phytin)for 15 days and then the urinaryrisk was re-evaluated. Other urinary lithogen parameters were also determined. The obtained results show that whereas the ordinary urinary lithogen parameterswere not modifiedbyingestionofphytate, the urinaryrisk to develop calcium stones was significantly reduced, demonstrating an interesting therapeutic efficacyin usingphytate asacrystallisation inhibitor. The urinary risk factor was evaluated using a test specially developed andvalidated for this purpose [360]. During an 8-year period a prospective study examined the association between dietaryfactors and the risk of incident symptomatic kidney tones in 96 245 female participants. The study lets assume that a high intake of dietary phytate decreases the risk of kidney stone formation and dietaryphytate mightbeanewand safewaytoprevent kidney stone formation [362]. Itisimportantto remarkthatphytatewasearlyusedinthe treatment of renal lithiasis at an earlystage. Thus, in 1958, Henneman et al. [363]usedhighdosesofphytateas sodium salt (8.8 g/day)to treat stone-former patients with idiopathic hypercalciuria. Nevertheless, the objective and basis of such treatment was clearlydifferent to that presented here. Thus, high doses of phytate were supplied to hypercalciuric patients with the aim to form nonsoluble complexes in the intestinal tract in order to prevent the absorption of dietary calcium and subsequentlytodecrease urinarycalcium excretion. However, the low dose supplied in the newlyproposed treatment aims at raising the urinary excretion of phytate, increasing the inhibitorycapacity of urine towards calcium saltscrystallisation(oxalateand phosphate). Consequently, all in vitro and in vivo results clearlyindicate thatphytateplaysa significant roleascrystallisation inhibitor of calcium salts in biological fluids and is an alternativeinthe treatmentof calciumoxalate renal lithiasis. www.mnf-journal.com S354 hydrogen peroxide(H2O2)atleast one free coordination site occurs in the heartvessels. In general, the formation of cal cific vascular lesions involves complex physicochemical ofironisrequiredandthus,inthe presenceofphytate,thefor mationofhydroxyl radicals(OH9)fromH2O2via theFenton and molecular events. Calcif ication (HAP) is caused by injury and progresses through promoter factors and/or the reaction under assistance of the Haber-Weiss reaction in a deficitof calcification modulators. The capacityofphytate twostep reaction – as a potential inhibitor of cardiovascular calcification was 9O2+Fe3+=O2+Fe2+ (reductionof ironbysuperoxide assessedin rats subjectedto calcinosis inductionbyvitamin radical) D plus nicotine [358] or by a macro dose of vitamin D [365]. In both cases phytate demonstrated significant Fe2++H2O2=Fe3++OH–+OH9 (Fenton reaction) i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com U. Schlemmer et al. Figure 8. Molecular structure of mono ferric-phytate in which iron (Fe3+-ion) is bound to phytic acid so that all six coordination sites of iron are occupied. Iron bound in this configuration (phosphate groups in position 1, 2 and 3 and in the axial, equatorial and axial position) does not take part in the formation of hydroxyl radicals from H2O2 via the Fenton reaction under assistance of the Haber – Weiss reaction in a two step reaction [368–370]. Under physiological conditions the negative charges are counterbalanced either by protons, sodium ions or other cations depending on pH and affinity. 8.1.2 Phytate and sialolithiasis Sialolithiasis is a common disease of salivaryglands. Little is known about the aetiology of these calculi and their exact mechanismofformationisunknown.Inastudy,the composition and structure of 21 sialoliths were studied and the compositionofthesalivaofeachcorresponding patientwas determined (pH, calcium, magnesium, phosphorus, citrate and phytate). Eighteen sialoliths exhibited similar macro and microstructures, consisting of HAP and organic matter. The salivaryCa concentration ofpatients with HAP calculi was significantlyhigherandthe salivaryphytate concentrationwas significantlylower than thoseof the healthy sub- jects.Itwas concludedthatadeficitofcrystallisation inhibitorssuchasphytateisalsoanimportant aetiologic factorof sialolith development [364]. 8.1.3 Phytate and cardiovascular calcification Calcification is an undesirable disorder, which frequently Mol. Nutr. Food Res. 2009, 53, S330 – S375 effects on decreasing calcif ications of the cardiovascular system. From these results it can be concluded that phytate is important in preventing cardiovascular calcification and further human studies are needed to fully understand its role in this process. 8.2 Blood glucose and lipid lowering effects Under physiological conditions phytic acid interacts with proteinsbyforming low soluble complexes. Thus, enzyme activity mightbe inhibitedbyphytate. Thompson et al. [33] showedthat in vitro formation of glucose from white bean flourwasreducedinthe presenceof intrinsic dietaryphytate and after addition of 1% sodium phytate. Administering the same flour to humans, the effect of phytate on the glycemic index was studied. Intrinsic dietary phytate as well as the addition of 1% sodium phytate to the flour significantly reduced(pa0.05)blood glucoselevelsinsix volunteers[33]. Inarecentlypublishedstudyin diabetic KK-mice,Lee et al. [34] found reduced blood glucose levels depending on the dietaryphytate contents(0 –1.0%).In another studyin diabetic KK-mice Lee et al. [366] reported decreased serum total cholesterol and LDL cholesterolbyphytate(0 –1.5%). Noeffectsofphytateonthetriglyceride concentrationcould be detected. The mechanism of these phytic acid effects remainstobeclarifiedbutitwasassumedthatthe cholesterol loweringeffectsin blood causedbyphytate mightbein part related to an increase in faecal bile acid and lipid excretion [367]andareductioninhepatic cholesterol synthesis. These results let assume that dietary phytate may also benef icially affect the blood glucose and blood cholesterol levels. 8.3 Antioxidative property The antioxidativepropertyisoneofthemostimpressivecharacteristicsofphytate. Itis mainlybasedon complexingiron between three phosphategroupsin positions1,2and3andin the axial, equatorial and axial position, respectively[368 – 370]. These phosphategroups are flexible and bind the iron ion in such a waythat all six co-ordination sites of iron are occupiedby –OHgroups (Fig.8)and the labile boundwater in thefirst coordination sphere of iron is removed. As the stabilityofthisiron–phytate complexishigh(Fe3+–phytate complex Kf1017 [257]), other ligands withloweraffinity cannot interact with ironin this cage’.For the interaction with , S355S355 Mol. Nutr. Food Res. 2009, 53, S330 – S375 is inhibited. It can be assumed that the molecular structure of mono ferrous-phytate (Fe2+-ion) is similar to that shown for mono ferric-phytate [368–370] as thehydroxyl radical formation fromH2O2 via theFenton reaction is also inhibited when ferrous iron (Fe2+-ion) is complexed by phytic acid [371]. For this reason phytic acid is an antioxidant unlike other antioxidants, such as ascorbic acid or b-carotene, whichprimarilyactas radicalscavengers.Evenifphytate inhibitsthe oxidationoftheFe2+-ioninthephytate complexbyH2O2, due to the lack of anyfree coordination sites and of any available aquo coordination sites of iron, this does not mean that any oxido-reduction process of the phytate bound ironis prevented.Forexample, the iron oxida- tionintheFe2+-phytate complexbymolecularoxygen(O2) is accelerated by a concentration dependent manner, whereas the reduction of the Fe3+-ion in the mono ferricphytate complex in the presence of ascorbic acid is inhibitedbyphytate [30]. However, for the inhibition of the hydroxyl radical formation certain ratios of phytic acid to iron are important. While Graf et al. [368] found a large range of 1:4 to 20:1 (InsP6/Fe), Rimbach and Pallauf [371] pointed out that the inhibition of hydroxyl radicals via the Fenton reaction requires a ratio A5:1, confirming earlier results of Hawkins et al. [370].For stoichiometric reasonsa molecular ratio A 1seems to be more reasonable as phytic acid onlyinteracts one iron ion in this specific configuration. Beyond its antioxidative property, phytic acid also reduces the iron mediated lipid peroxidation and inhibits the formation of thiobarbituric acid reacting substances (TBARS) [30, 372]. This effect has been applied in meat processing to prevent the oxidation of myoglobin to metmyoglobin, which changes the colour in homogenised meat from fresh red to a brownish shade during preparation and storage [373 – 375]. Porres et al. [376] studied in pigs the effect of different dietaryphytateonthe protectionofoxidative stress induced by moderately high iron intake in colon and liver. They found that colonic lipid peroxidation, measured as TBARS, wasloweredbyphytateto someextentinthe colon mucosa, while it was unaffected in the liver. Highest lipid peroxidation was observed with high dietary iron content (FeSO4) and in the absence of phytate. The authors concluded that phytate is protective against lipid peroxidation in the colon induced by a moderately high iron intake. However, no effectwasobserved withanormal iron intake. In Sprague –Dawley rats, Rao et al. [377] studied the effectofphytate on the damage causedby experimentally induced ischemia-reperfusion injury. The results show that phytate, injected intravenously (7.5 –15mg/100g BW) prior to cardiac excision, protects the myocardium from damage. Thiswas measuredbymarkersof muscle damage, heartfunction and lipid peroxidation. Rimbach and Pallauf [371] studied the effect of iron, phytate and vitamin E on the antioxidative status in male albino rats for 28 days. At marginal dietary iron supply, phytate reduced the iron bioavailability, but no effect was observed on oxidative stress measured by determining reduced glutathione, TBARS and protein carbonyls. In a follow upstudy, also in male albino rats, Rimbach and Pallauf [378] fed diets with different levels of magnesium and sodiumphytate (300mgMg and0, 7.5 and15gNa –phy- tate/kg).With increasing dietaryphytate, magnesium bioavailability and magnesium content in plasma and femur decreased. Thiswas accompaniedby an increasein hepatic TBARS and protein carbonyls along with a moderate decline of reduced glutathione levels in the liver. The results show a decreased antioxidative effect of phytate mediated by magnesium. A reversed effect of dietary phytate was observedbyShanandDavis[379]in chickens.Herephytate increased the selenium concentration in tissues andbythis the activity of glutathione peroxidase (GSHPx) in blood and heart was increased, along with decreased GSHPx activity in the kidneys. Comparing diets high andlowin intrinsic dietaryphytate in postmenopausal women, Engelman et al. [380] found only small and nonsignificant effects on oxidative stress measured by proteincarbonyls, 8-iso-prostaglandin-F-2 alpha and oxidised LDL. The assumption that oxidative stress and reactiveoxygen species mightbe significant for thedevelopmentof cancer, arteriosclerosis, neurodegenerative diseases, cirrhosis,irradiation damages and other civilisation diseases [381] raised the question on the role of phytate as antioxidant in human nutrition and disease prevention. In vivo studies reported, however,showanequivocalpicture.Ontheonehand antioxidative activity of phytic acid is obvious while on the other hand no antioxidative activity can be observed. This might be due to the different modes of action by which phytate takes part in antioxidative processes in the metabolism, mainlymediatedbybindingvarious bi-and trivalent cations such as iron and copper. Although under clearlydefined in vitro conditions, the antioxidative property of phytate, such as the inhibition of thehydroxyl radical formation, can be determined exactly,itisdifficulttodothis invivo incomplex body fluids, tissues and organs. Thus, future studies will haveto elucidatethe significanceofthephytatefor disease preventionbyinhibitinghydroxyl radical formation.As the concentration of phytate in mammalian cells is comparable high, reachingalevelof l15– 100 lM[382, 383], much has been speculated on its real role in the cellular metabolism. – AsH2O2and reactive oxygen species such as 9O2 are ubiquitous produced and iron is widespread in the metabolism, the intracellular significanceofphytic acid couldbetogovern intracellularformationofhydroxyl radicals whichisof vital interest to anyliving cell and organism. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com S356S356 U. Schlemmer et al. 8.4 Anticancer activity The anticancer activity of phytic acid is one of the most important beneficial activitiesofphytic acid.Itwas demonstratedinvariouskindsof cancer,suchascolon,liver,lung, mammary, prostate, skin and soft tissue cancer of mice and/ or rats [384 – 387]. Moreover, different mechanistic steps of the anticancer activity of phytic acid in various, mostly human cell lines, were observed. Mainly sodium phytate was added either to the diet, to the drinking water or to the cell medium [384]. Intrinsic dietaryphytate, however, has also shown to effect aberrant crypt foci (ACF) and silomu- cin-producingACF, both earlymarkersof colon cancer formation [388]. Thus,it canbe concluded that dietaryphytate also shows anticancer effects whereas applying high content of sodium phytate might show more pronounced effects. As most conclusions of the anticancer activity of phytic acid have been drawn from carcinogen-induced cancer models and cell line experiments, there is a strong need for evaluating the anticancer activity of phytic acid in humans. Given that excellent and extensive reviews on the anticancer activity of phytic acid have been published very recently [384 – 388], a further review here of this topic is not yetrequired. 9 Determination of phytic acid/phytate and other inositol phosphates in foods – development of analytical methods 9.1 Nonspecific methods The analysis of phytic acid dates back to the method of Heubner and Stadler in 1914 [389]. Essentiallythis method is based on the extractionofphytic acid fromground cereal powderbyhydrochloric acid and subsequent precipitation asFe3+–phytate from thefiltrate after stepwise additionof FeCl3.Thephyticacid contentis deducedonthebasisofthe iron content of theFe3+–phytate precipitate. Due to problems detecting the exact endpoint of this titration, McCance and Widdowson [390] changed this procedure and determined the phosphorous contentof theFe3+–phytate precipitate. Phytic acid content was calculated on the assumption that the total phosphorous originated from phytate. Due to inconsistent stoichiometric ratios ofFe3+-ions or phosphoroustophyticacidintheFe3+– phytate precipitate, Harland and Obeleas [391] omitted the Fe3+ –phytate precipitation and purified the phytate-containing HCl extractby anionexchange chromatography onAG1resin. Separating inorganic phosphate from organic phosphatebystepwise NaCl elution they gained an elution fraction containing mainly phytate. However, it included other organic phosphates present in the sample. The phosphate content of this fraction was used to calculate the phytic acid content or more Mol. Nutr. Food Res. 2009, 53, S330 – S375 phosphates, the calculated phytic acid content is misleadinglyenhanced when phytate and other inositol phosphates or nucleotides are present. As this method was mainly appliedtorawand unprocessed food withalow contentof lower phosphorylated inositol phosphates(la15%, InsP1– InsP5) [16], this error is of minor practical relevance. In 1986amodifiedversionofthis method becametheofficial AOAC-methodin determiningphytatein food [392]. However, for processed food with a high content of phytate hydrolysis products or for biological samples rich in lower phosphorylated inositol phosphates (InsP1– InsP5), theAOAC method is inadequate and specific methods for the precise determination of phytic acid and other inositol phosphates are required. For extensive discussion of the nonspecific analysis the reader is referred to the excellent reviews of Oberleas and Harland [393] and Xu et al. [394]. 9.2 Specific methods In 1952 Smith and Clark [395]werefirst able to separate phytic acid and other inositol phosphates from soil by means of anion-exchange chromatography on a weak-base exchange resin and stepwise elution with increasing HCl concentration. Inositol phosphateswere determinedbyanalysing the phosphorous content of the different peaks. In 1963 Cosgrove [396] applied AG 1 as a strong anion exchange resin to separate various inositol phosphates obtained after acidichydrolysisofphytic acidby meansof HClgradients(0 –1.5NHCl). Quantificationofthe inositol phosphatesin the different peaks occurredbyanalysing the phosphorous-inositol ratio. In 1980Tangendjaja et al. [397] triedto determinephytic acidof ricebyRP chromatography on lBondapakC18 column and using sodium acetate (5 mmol/L) as mobile phase. Grafand Dintzis[74]improvedthisanalysisbyaddingapreceding purification and concentration step onAG1resin.For phytic acid detection they favoured the refractive index detection rather than the UV absorption. This purification and concentration proved to be very helpful for a good separation and detection of inositol phosphates by different HPLC methods, especially in complex matrix, and was widelyused.Duetothelowretentionofphyticacid(1.4min) on lBondapakC18 columns, other inositol phosphates could hardlybe discriminated [74].By applying terabutylammoniumhydroxideasan ion-pair reagent, Sandbergand Ahderinne [398] improved the retention of inositol phosphates on the stationaryphase and separated inositol phosphates with different numbers of phosphategroups (InsP3– InsP6).However, separation of the stereoisomers of the different inositol phosphates also remained impossible with this method. In 1984 in thefield of cell biology, Berridge and Irvine [399] observed that Ins(1,4,5)P3, cleavedfrom membranebound phosphatidylinositol 4,5-diphosphateby phospholicorrectly the content of , phytic acid equivalents’. As this pase C, shows a second messenger function in mobilising method does not discriminate between various organic intracellular calcium.This intensifiedthe searchto separate i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com S357S357 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Figure 9. (A) Separation of inositol phosphates on a Mono-Q (250 mm) column by using a HCl gradient (0 – 0.4 M). Inositol phosphates were detected by using the Ytrium –PAR complex and measured at 546 nm [409]. 1, Pi + InsP1;2, Ins(1,2)P2 + Ins(1,6)P2; 3, PPi; 4, unidentified; 5, Ins(1,3,5)P3 + Ins(1,4,6)P3; 6, Ins(1,3,4)P3; 7, Ins(1,2,6)P3 + Ins(1,2,3)P , +Ins(1,4,5)P3; 8, Ins(1,5,6)P3;9, Ins(4,5,6)P3; 10, Ins(1,2,3,5)P4/Ins(1,2,4,6)P4; 11, Ins(1,2,3,4)P4/Ins(1,3,4,6)P4; 12, Ins(1,3,4, 5)P4; 13, Ins(1,2,5,6)P4; 14, Ins(2,4,5,6)P4; 15, Ins(1,4,5,6)P4; 16, Ins(1,2,3,4,6)P5; 17, Ins(1,2,3,4,5)P5; 18, Ins(1,2,4,5,6)P5; 19, Ins(1,3,4,5,6)P5; 20, Ins(1,2,3,4,5,6)P6. (B) Nucleotides are also determined by this detection at 254 nm: a, AMP + CMP + NAD; b, cyclic AMP; c, GMP; d, NADH; e, UMP; f, NADP; g, ADP + ADP-ribose; h, ATP + CTP; i, GDP; k, IDP; m, UDP; n, ITP; o, UTP. They coelute with some InsP1 –InsP3, interfering their determination. Nucleotides are measured at 254 nm with optimum sensitivity (B), but also can be detected at 546 nm. The figure was adapted from [409]. the different inositol phosphatesbyadequate HPLC anionexchange chromatography methods. AG 1, silica based SAX, Partisil SAX 10 columns, etc. with aqueous mobile phasesof ammoniumformate, ammonium acetateor ammonium phosphate[400 –402]were successfullyusedtodifferentiate radiolabelled inositol phosphates, especially lower phosphorylated inositol phosphates (InsP1– InsP3). Using a modificationofthe ion-pairRPHPLCandvaryingtheACN concentration of the mobile phase, Sulpice et al. [403]were abletodiscriminate inositol phosphates such as Ins(1,4,5)P3 from other organic phosphates such asATPand 2,3-DPG. Irth et al. [404] later obtained distinct differentiation of Ins(1,2,6)P3 from other low phosphorylated inositol phosphates. Interestingly, the HPLC methods used in cell biology research have not been widely applied in food science or nutrition research. One reason for this might have been the i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim problems of adequate inline detection due to missing characteristic absorption spectra of and specific colorimetric reagents for inositol phosphates. As the detection of radiolabelled inositol phosphates in the mobile phase of highly concentrated aqueous ammonium formate or ammonium phosphate (maximum l1mol/L)was unproblematic, sensitive inline detection of nonradioactive inositol phosphates, such as from biological samples in the nano-or micromolar range, remaineda challenge. In 1985 Phillippyand Johnston [405] and later Phillippy et al. [15] showed good separation of a great number of different inositol phosphates on a strong anion exchange AS3 column (Dionex) by using an HNO3 gradient (0– 0.155M HNO3). This eluent offered reliable and sensitive inline detection of inositol phosphates by complexing the phosphate groups by means of Fe3+-ions. This detection showed a linear calibration curve up to l100 nmol phytic acid anda detection limitof1 –2nmolphytic acid, measuredat290nm[ 15].Itwas basedon earlier observationsby Imanari et al. [406], who foundFe3+-ions highly appropriated to form soluble complexes with inositol phosphates under acidic conditions, allowing for quantitative in-line determinationbymeansof postcolumn derivatisation. In 1986 Cilliers and Niekerk [407] and later Rounds and Nielsen [408] presented another method for inline detection ofphyticacidbymodifyingtheformerphyticacid determination of Latta and Eskins [106]. This determination based on an exchange reactionof the ligand(Fe3+)from theFe3+– sulfosalicylate complex(Wade reagent) to theFe3+–phytate, resulting in a bleaching of the intense purple colour of the ferric sulfosalicylate complex, measurable at 500 nm. In 1988 Mayr [409] proposed the separation of inositol phosphates on a Mono-Q column (L: 250 mm; 10 lm beads; strong anion exchanger; General Electric)by using anHClgradient(0 –0.4M)witha comparablelongrunning time of 60 –90min. This HPLC method provided verygood separation of most inositol phosphates including their stereoisomers (Fig. 9). Good separation of the higher phosphorylated inositol phosphates (InsP5– InsP6)as well as of the major lower phosphorylated inositol phosphates (InsP3– InsP4)can alsobe achievedona shortMono-Q column HR 5/5 (5650 mm) in less than half an hour (Fig. 10, below), which is adequate for most food analysis and many biological samplesaswell[159].Mayr[409] appliedforthe detection of inositol phosphates the coloured complex of Ytrium and 4-(2-pyridylazo)resorcinol (PAR). The Ytrium–PAR complexisbleachedin the presenceof inositol phosphates, due to the higher affinity of theY3+-ions to the phosphate groups of the inositol phosphates and the detection is called the metal-dye detection (MDD). The absorption is measured at 546 nm as a negative peak with a linear calibration curve up to 1000 pmol. In a modification of this method usinga Mini-QPC (3.2/3) column (the same anionexchangeras Mono-Qbuta bead sizeof3 lm) Guse et al. [410] describedthe detectionlimitof1 –3pmolInsP6. www.mnf-journal.com S358S358 U. Schlemmer et al. Mol. Nutr. Food Res. 2009, 53, S330 – S375 Figure 10. Separation of inositol phosphates on CarboPac PA-100 (above) and on Mono-Q HR 5/5 (0.5/5 cm) (below) by using HCl gradients (0 – 0.5 M). Inositol phosphates were detected with Fe3+-ions at 290 nm in accordance to [159]. 1, Pi + InsP1; 2, Ins(1,2)P2 + Ins(1,6)P2; 3, Ins(2,4)P2; 4, Ins(1,3,4)P3; 5, Ins(1,2,6)P3 + Ins(1,2,3)P3;6, Ins(1,4,5)P3; 7, Ins(1,5,6)P3; 8, Ins(4,5,6)P3; 9, Ins(1,2,3,5)P4/ Ins(1,2,4,6)P4; 10, Ins(1,2,3,4)P4/Ins(1,3,4,6)P4; 11, Ins(1,2,4, 5)P4; 12, Ins(1,3,4,5)P4; 13, Ins(1,2,5,6)P4; 14, Ins(2,4,5,6)P4; 15, Ins(1,4,5,6)P4; 16, Ins(1,2,3,4,6)P5; 17, Ins(1,2,3,4,5)P5; 18, Ins(1,2,4,5,6)P5; 19, Ins(1,3,4,5,6)P5; 20, Ins(1,2,3,4,5,6)P6. This method still is the most sensitive inline detection of nonradiolabelled inositol phosphate isomers after HPLC separation. However, other multivalent cations and espe- ciallyFe3+-ions interfere stronglyin the detectionbyaffectingtheY3+– PAR complex.Thusfor reliabledetermination, all iron containing materials in contact with the eluent have to be eliminated and all chemicals and standards used need highest possible purity. The longrunning timeof more than an hour for the long Mono-Q columns (250 mm), however, is disadvantageous for routine analysis andgivespreference for inositol phosphate separation on the shorter Mono-Q column (HR 5/5) or the Mini-Q column. In 1997 Skoglund et al. [411, 412] reported excellent separation of inositol phosphates including most of their stereoisomers by anion-exchange chromatography on Omni Pac PAX-100 (Dionex) and especially on CarboPac PA-100 (Dionex)byusingHClgradients(0 –0.5and0 –1.0 mol/L, respectively) along with the ferric ion detection [15]. This determination was proposed for mainly InsP3– InsP6.For lower phosphorylated inositol phosphates, such as InsP1– InsP2 and InsP3, good separation on Omni Pac PAX-100 (Dionex)byusing alkalinegradients with NaOH and 2-propanol in conjunction with anion micromembrane suppressor and conductivity detection in the picomole range of InsP1– InsP3was achieved[411, 413]. In 1988 and 1989 Smith and MacQuarrie [414, 415] already reported on the usefulness of the chemicallysuppressed conductivity detection, not only for lower phosphorylated (InsP1– InsP3)but also for higher i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim phosphorylated inositol phosphates (InsP4, InsP6)andTalamond et al. [416, 417] described it as an adequate detection alsoforthe determinationofphyticacidinfood. Very recently Letcher et al. [418] reported on an enzymatic method in which phytate can be determined after 32Plabelling and HPLC separation as InsP7.However,even if the method is highly sensitive and might allow phytate detection below the nanomolar range(a1nM InsP6), it is unfortunatelyrestricted to the determination of InsP6 only. Other inositol phosphates, also significant in cells and tissues, are not detectable. 9.3 Problems of detection Due to missing characteristic absorption spectra and of specific colorimetric reagents for inositol phosphates, further properties of the inositol phosphates had to be applied for analysis. In contrast to food samples, the concentration of phytate and other inositol phosphates in physiological samples such as tissues and cells is extremelylow. Therefore, highly sensitive methods and adequate procedures for the sample preparations from different matriceswere required. 9.4 Absorption Thehighaffinityofthe phosphategroupsof inositol phosphates to polyvalent cations, such asFe3+-,Y3+-and Cu2+ions, is the basis most commonlyused for the detection of www.mnf-journal.com S359S359 Mol. Nutr. Food Res. 2009, 53, S330 – S375 phytate and inositol phosphate in foods. Absorption can be measured eitherbydirect interaction between cations, such as Fe3+-ions and inositol phosphates [15], or by means of ligand exchange reactions, e.g. by removing Y3+ from the intensively colouredY 3+–PAR complexby inositol phosphates, resultingina bleachingof this complex [409]. Both methods, especiallythe latter one, provide sensitive detection of inositol phosphates. Nevertheless, other organic phosphates, such as nucleotides, interfere with this inositol phosphate determination as they coelute during chromatographic separation from the column with certain InsP1– InsP3 isomers and can also be determinedbythis detection method(Fig.9)[409].Duetothelow contentof nucleotides in food, their effect on the determination of inositol phosphates in food is also low. When cells or physiological samples such as blood plasma, urine, tissues and organs are analysed, nucleotides and phosphorylated proteins need to be separated by adequate sample preparation, such as additional anion-exchange chromatography, charcoal treatment, etc. [409, 419]. Meek and Nicoletti [420] proposed the determination of InsP2 and InsP3 after HPLC separation by cleaving the phosphategroups using immobilised alkaline phosphatase (calf intestine) in an on-line bioreactor, determining phosphatebymeansof ammoniummolybdateat340 nm. 9.5 Fluorescence detection A similar ligand exchange reaction as described for the Y3+ –PAR complex [409] was also applied for the fluorescence detection of inositol phosphates. Irth et al. [404] removedFe3+-ions from theFe3+–methylcalcein blue complexby inositol phosphates, showing higher affinity than the methylcalcein blue complextoFe3+-ions.By this ligand exchange reaction the quenching of the methylcalcein blue bymeansof theFe3+-ionswas attenuated and depending on the inositol phosphate concentration the fluorescence intensity increased. March et al. [421] used the activation of phytic acid on the oxidation of 2,29-dipyridyl keton hydrazone catalysed byCu2+-ions resulting in highly fluorescent reaction products to determine phytate in urine. The calibration curve for this determinationis linearover the rangeof76 –909 nmol/ Lwithadetection limitof45 nmol/L. Chen et al. [422] used the replacement of the Cu2+-ions from the Cu – gelatine complex by phytic acid. Thus the quenching of the Cu – gelatine complex was eliminated and the fluorescence intensity increased. The calibration curve for this determination is linear from 606 to 3.600 nmol/L with a detection limit of 348 nmol/L. In urine phytate was detected in the range of 0.49 – 0.75 mg/L with a recovery of 96.2 – 108.8%. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim 9.6 Light-scattering detection Light-scattering detection as in-line determination of phytate after HPLC separation ona strong anion exchange AS7 column (Dionex) was tested by Phillippy et al. [15] and comparedtotheFe3+-ion detection.The authors reporteda detection limit of 1.5 nmol InsP6 for the light-scattering detection which,however,ishigherthanthatoftheFe3+-ion detection with 758 pmol InsP6and so far it does not offer a real advantageover theFe3+-ion detection. 9.7 Conductivity Smith and MacQuarrie [414, 415] showed that more than 20 different biologicallyimportant anions can be separated by anion-exchange chromatography on AS 4A columns (Dionex) and canbe detected with high sensitivitybyusing chemicallysuppressed conductivity. They reported a range of l20 pmol to 400 nmol InsP1 for the calibration line with a good separation not only between different phosphorylated organic compounds, such as nucleotides (ATP, ADP, GTP, etc.), glucose 6-phosphate, fructose 6-phosphate and various low phosphorylated inositol phosphates (InsP1– InsP3), but also of InsP4 and InsP6. Skoglund et al. [411, 413] and Talamond et al. [416, 417] later used this anion micromembrane suppressed conductivity detection either for low phosphorylated inositol phosphates or for phytic acid,inphysiological samplesandinfood, respectively. 9.8 NMR spectroscopy NMR spectroscopyisan adequate technique for analysing InsP6 and its various hydrolysis products (InsP1– InsP5), including the different stereoisomers. It has been successfully applied in both food and biomedical research [302, 423, 424]. The 31P-NMR spectroscopy has potential application as an analytical technique to determine phytate contentin plantsandaswellas human tissueandis appropriate to detect its form and binding to other components. Moreover, it is also adequate to study phytate metabolism and phytate degradation during food processing, offering the advantageofhigh accuracyand specificity.High resolution 31P-NMR has been used as a noninvasive method for the study of P-containing compounds in intact tissues and cell suspensions. In comparison with other techniques, such as HPLC, NMR allows direct detection and qualification of all phosphate compounds in the same experiment. Use of inositol phosphate standards for NMR analysis is not required, unlike with other analytical methods such as HPLC [425]. The low sensitivity of NMR methods might cause difficulties in detecting low inositol phosphate contents in biological and physiological samples, such as cells or tissues., However, the inositol phosphate concentration is high www.mnf-journal.com S360S360 U. Schlemmer et al. enough for a reliable 31P-NMR determination in foods, chyme and faeces. Comparing to other analytical methods, good agreement of phytate determination in foods by 31P NMR have been reported [426, 427]. NMR has also been successfullyapplied in routine analysis to differentiate inorganic phosphate from different inositol phosphates in plant extracts and in whole diet samples [428]. When analysing InsP6by31PNMR, four resonance peaks are observed since the molecule has plane symmetrythrough C-2 and C-5. Accordingly,thePsignals from C-1 and C-3, as wellasC-4andC-6 are identical.Inthe adopted system, proton decouplingisusedto eliminate interactionsbetweenprotons and 31P[425, 426]. Whenphytateishydrolysed,e.g.by phytases,newsignals appearduetotheformationofphytate hydrolysis products (InsP1–InsP5).Inositol pentaphosphate, thefirsthydrolysis productofthe stepwisephytate degradation, exhibitsfive equallystrong peaks and shows an asymmetrical molecule with phosphorylated carbon at positions2 and5[423,428].The assignmentofpeaks arisingfrom tetraand tris-phosphates becomes increasingly difficult when different isomers are generated. Inositol mono-and diphosphateseventually formed can alsobe identified. Phillippy applied 2-D 1H–1H–NMR to differentiate the two inositol trisphosphate isomers, Ins(1,2,3)P3 and D-Ins(1,2,6)P3, formedbywheatphytasehydrolysisofInsP6. Both inositol phosphate isomers cannotbe distinguishedbyHPLC [429], as they coelute during chromatographic separation from the column.InthisrespectMNRoffersadistinctadvantageover HPLC analysis. Anotheradvantage is that NMR is an excellent noninvasive analytical method withloweffortfor sample preparation. It should be stressed, however, that NMR analyses are expensive and require skilled expertise to correctly interpret the complex spectra from heterogeneous biological systems such as foods. 9.9 Mass spectrometric detection Studying inositol phosphates in cells, tissues and body fluids, concentrations a200 pmol inositol phosphates/mL org may occur.For this purpose, the mass spectrometric determination of inositol phosphates offers excellent sensitivity. March et al. reported a GC-MS method was reported for the determination of phytate [430]. It is based on the purification by anion-exchange chromatography, enzymatic hydrolysis of phytate tomyo-inositol and consecutive derivatisation to trimethysilyl derivative. Scyllo-inositol was applied as internal standard [431]. The method offers a linear calibration line of 15 –500 lgphytate/L withaCVof 1.9%andadetectionlimitof10nmolphytate/L.Ithasbeen successfullyappliedtoavarietyof biological samples, such asvarious rat organs (kidney,liver, brain and bone), human plasma, urine and kidneystones [430]. This method is tedious and requires highly active phytases with sufficient stability. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Mol. Nutr. Food Res. 2009, 53, S330 – S375 HPLC-MS was described as another method for the determination of phytate in human urine by [432]. The method is based on hydrolysing phytate and determining the myo-inositol [433]. Urine was purified and InsP6 separatedbyanion- exchange chromatographyand acidhydrolysis of phytate performed at 1208C for 11 h. Chromatographic separation was performed on an Aminex HPX-87C column with ultra pure, deionised H2O (18MX; Mili-Q system) as mobile phase and5 mMammonium acetatewas added consecutively (postcolumn). The detector counted positive ionsby monitoring m/z = 198, which corresponds to the adduct of myo-inositol with the ammonium cation. The RSDs obtained for standards containing 0.5, 1 and 1.5 mg phytate/L were 4.1, 3.0 and 2.7% respectively (n = 5). The LODwas60 lg/L ofphytate. Different urine samples were analysed both by this method andbythe GC-MS method, described above [430]. The resultsof both methodswere comparable,however, the HPLC-MS method is more suitable than the GC-MS method because derivation is avoided. Hydrolysis of phytate to inositol can be accomplishedbyextended acid heatingbutthis processhastobe carefullygovernedas myo-inositol easilydegradesin acidic media. Hsu et al. [434] applied a thermosprayliquid chromatographic/ mass spectrometric combination for the determination of inositol phosphates. They separated inositol phosphates (InsP1, InsP3, InsP6)byanion-exchange chromatography on Mono-Q with ammonium formate, heated at 2608C (pH 4) and determined mass spectrometrically the dephosphorylated inositol moiety as m/z ion 198 [NH4 N inositol]+. The method is sensitive with a detection limit of 100 pmol/lLfor Ins(1,2,6)P3. 31P An inductively coupled plasma-MS (ICP-MS) by method [435] for phytate determination in human urine based on total phosphorus determination of separated phytate was also described. Separation of accompanying inorganic phosphates,pyrophosphates or anyother phosphorus compounds from phytate is required by using anionexchange SPE. Separation of phytate and recovery were verifiedin artificial urine. The linear rangeof thephytate determination is 20– 600 lg phytate/L with a detection limit of5 lg/L. The lack of selectivity of phosphorous compounds when determining phosphorous via ICP-MS can be satisfactorily overcome by selective anion-exchange separationandpurificationofphytatein urine. Alternatively, an ICP atomic emission spectrometry (ICP-AES) for routine phytate analysis, also based on the determination of phosphorous, was developed [435]. This procedure also requires chromatographic separationofphytate from other phosphorous containing compounds and showsa linearworking rangeof0 –7 mgphytate/L witha detection limit of 64 lgphytate/L anda limitof quantification of 213 lgphytate/L. This method is less sensitive than the ICP-MS[436] describedabovebut shows sufficient sensitivity and is more suitable for routine phytate analysis in www.mnf-journal.com S361S361 Mol. Nutr. Food Res. 2009, 53, S330 – S375 urine. Comparison studies of both methods show consistent results(pa0.05) [436]. Forthe discussionofthe recentdevelopmentinmass spectrometric determination of phytic acid the reader is referred tothe interesting contributionofCooperet al. [438]. 9.10 Discussion and conclusions Surprisingly, some principles of the earlyphytate determinationby Heubner and Stadler [389] survived the last 100 years and are still being applied successfully. This is the acidic extractionofphytic acidbyhydrochloric acid from dryandgroundfoodpowderto liberatephyticacidfromthe food matrix on the one hand, and the use of Fe3+-ions for phytate detection on the other. While Heubner and Stadler applied ferric ions to precipitate phytate and by doing so determined the phytate content, Phillippy et al. usedFe3+ions to form soluble Fe3+ – inositol phosphate complexes, which could be applied for quantitative determination of inositol phosphates [15]. Graf and Dintzis [74] introduced the concentration and purification of the sample HCl extract on AG 1X8 resin. This is not onlyessential for the separation of inositol phosphatesby RP chromatography but is also helpful in maintaining good HPLC separation of the different stereoisomers of inositol phosphates on strong anion exchangers such as Mono-Q-or CarboPacPA-100 columns. Carlsson et al. [438], however, showed good separation of inositol phosphates of mainly food samples on CarboPacPA-100 without purificationonAG1X8 resinbutbydirect injectionof the ultrafiltrated sample HCl-extracts. Whether or not, this sample preparation is sufficient to prevent poisoning and blockage of HPLC columns, such as CarboPac PA-100, with high backpressure (>300 bar at ~1.3 ml/min), when long series of samples of complex biological matrices are analysed, remains to be clarified. Phillippy et al. [15] used purificationby meansofRP chromatography which might be an alternative for purification onAG1resins, andMayr [409] applied charcoal to the purification. This charcoal treatment does not only separate cations such as iron but also phosphopeptides and nucleotides, which all strongly interfere with the determination of inositol phosphates by using theY-PARreaction. Depending on the matrix, reliable determination of inositol phosphatesinthelongrunandbyusing automatedanalysisby means of autosamplers make it desirable to purify the sample HCl extract before HPLC analysis. This is relevant for complex biological matrices, such as gastro-intestinal content, blood plasma, urine and tissues. Whether this is also true for the analysis of raw and unprocessed food, containing mainly phytate and inositol pentaphosphates, remains to be clarified. Oberleas and Harland [407, 408] recently conducted an interlaboratory comparison trial for the specific determinationofphytic acidin foodbyseparating phytic acid on a short PL-SAX column (5064.6 mm) i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim and detectingitby meansoftheFe3+– sulfosalicylate com- plex(Wade reagent). Theyfound good correlation between the results of the samples (predominantly cereal samples) among the participating laboratories [439]. This is remarkable, as afterfiltration (0.45 lm) the phytic acid containing HCl sample extracts after filtration were directly applied for HPLC analysis on the unguarded anion exchange column withoutanyfurther purification. Unfortunately,itwas not reported how many injections can be successfully appliedbythis methodinarow,as poisoningandclogging of the column seems to be inevitable. Excellent separation of inositol phosphates can be achievedbyHPLC on strong anion exchangers such as CarboPacPA- 100 (25064mm) (Fig. 10, above) and Omni Pac PAX-100 (25064mm) [411, 412] and Mono-Q (Fig. 9) [409] and Mini-Q [410]. Mono-Q with a 250 mm column (25065mm; 10lm beads) shows optimum separation of the inositol phosphates at a long run of l60–90min, while onlyhalf of that time is required for excellent separation of inositol phosphates on CarboPac PA-100 (25064mm; 10 lm beads) and Mini-Q (5064.6 mm;3 lm beads). Currently separation on CarboPac PA-100 by anion-exchange chromatographyoffers the best discriminationof the different inositol phosphate isomers. However, running analysis on the CarboPacPA-100 column requires adequate equipment due to the high back-pressure of l350 bar at the requested elution velocity of 1.5 mL/min. Good separation of inositol phosphates alsowas achievedby Schlemmer et al. [159]onashortMono-Q columnHR5/5(5065mm) in less than half an hour, showing good separation of InsP6 and the four InsP5-isomers and sufficient differentiation of the main InsP4-and InsP3-isomers (Fig. 10, below). If more information on the inositol tetrakis-and inositol-trisphosphate isomers is needed, further gradients have to be applied and detailed analysis of the inositol mono-and inositol diphosphates requires extra separation anyway(e.g.by alkaline gradients). As the back-pressure on the Mono-Q column HR 5/5 does not exceed 30 bar during the run at a flow of 1.3 mL/min, many practical problems connected with high pressure analysis at Al300 bar as frequent leaking and blockage of the column, typical for long HPLC columns (250 mm) are absent. The analysis of inositol phosphates on Mono-Q column HR 5/5 (5065mm) is suitable for food samples [72] as well as for most studies in human and animal nutrition and has been successfully applied in various pig studies [159, 177]. The most sensitive detection of the different inositol phosphate isomers can be achievedbytheY-PARdetermination, with a detection limit of l1–3pmol InsP6 [409, 410]. Although mass spectrometric detection of inositol phosphates shows comparably high sensitivity [430, 432, 436],the separationof inositol phosphateonAG1X8 resin, with consecutive mass spectrometric detection of either the phosphorous or the inositol proportion, does not reach the excellent separation of the different inositol phosphate iso www.mnf-journal.com S362S362 U. Schlemmer et al. mers that can be achieved with Mono-Q or CarboPac 100 and subsequentY–PARdetection. The method of detecting inositol phosphates by using Fe3+-ions [15] offers good sensitivity, with a detection limit of l0.68 –2nmol InsP6, along with high stability and reproducibility [159]. It is very suitable and recommendablefor anyfood analysis [15] and manyphysiological samples with complex matrices such as the gastro-intestinal chyme [159] and is more sensitive than the ferric sulfosalicylate- complex detection [440]. If low phosphorylated inositol phosphates such as InsP1– InsP2arethe focusoftheanalysis, alkalinegradients along with conductivity detection in conjunction with anion micromembrane suppression seem to be the suitable detection. Alkaline gradients offer good separation of lower phosphorylated inositol phosphates in combination with sensitive detection in the picomole range [411, 413]. In food and nutrition research, HPLC methods for the determination of phytate and other inositol phosphates are most commonly used. Other methods such as CZE and capillary isotachophoresis [441 – 443] are also used but have not been widely applied. This is probably due to the lower sensitivity and discrimination of the inositol phosphate isomers compared to HPLC methods. In order to get more information on the role of phytate in nutrition and for health protection, simple and evaluated methods for the determination of phytic acid and other inositol phosphates present in food and diets are required (see the respective discussion in Sections2and 3). Most probably, the HPLC separation of phytic acid and other inositol phosphates from accompanying compounds after acidic extraction ofground samplesby applying anion-exchange chromatography along with a simple and evaluated detection seems to be the method of choice. Using long anion exchange columns (25064mm), such as Omni PacPAX100 or CarboPacPA-100 [411, 412, 438], excellent separation of the inositol phosphates can be obtained (see Fig. 10, above). Using short columns (L. 50 mm), such as the PLSAX column [439], a specific determination of phytic acid is also achievable. Although Oberleas and Harland [439] did not describe how other inositol phosphates also present in food can be discriminated from phytic acid by this method,it can be assumed that this should also be possible on this column. Using the Mono-Q column HR 5/5 (5065mm) [159], another short anion exchange column, sufficient discriminationof the relevant inositol phosphates present in foods and biological samples was shown by Schlemmer et al. [159] (Fig. 10, below). It should be pointed out, however, that the columns need to be guarded by precolumns to protect them and to guarantee reliable analysis in the long run, independent of the fact that the samples will be just filtered or further purified by additional sample preparations prior to the HPLC separation. For quantitative detection different methods are available, preferablyUVdetection, as these detectors are wide spread Mol. Nutr. Food Res. 2009, 53, S330 – S375 in laboratories. One method could be the detection by Rounds and Nielsen [408] using the Fe3+ – sulfosalicylate complex which was successfully applied by Oberleas and Harland in their interlaboratory comparison trial [439] or thehighlysensitivedeterminationofMayrby meansofthe Ytrium –PAR complex detection [409]. Another opportunity for in-line detection of inositol phosphates is the iron detection(Fe3+)ofPhillippyet al. [405] which has shown to be sensitive along with high stability and good reproducibility [159, 405]. There might be other methods of specific, simple, fast and reliable determination of phytic acid and other inositol phosphates in foods and the future will show which method of determination and detection will be most suitabletodeterminephyticacidinfoodsand diets. 10 Final conclusions Phytic acid is one of the most fascinating bioactive food compounds and is widely distributed in plant foods. It has different propertieswithvaryingeffectsfor humansandanimals. Due to its molecular structure, phytic acid shows a highaffinitytopolyvalent cations,suchas mineralsandtrace elements, and interferes in their intestinal absorption.With unbalanced nutrition or undernourishment this maylead to seriousdeficienciesandisofparticularlygreat significance for developing countries. However, with a well balanced nutrition this seemstobealess significant problem.Inindustrialised countries where various civilisation diseases are prevalent,thebeneficial propertiesofphyticacid,suchasits anticancer, antioxidative and anticalcif ication activities, are ofgreatimportance.Duetothe enormousproblemsofcivilisation diseases, anycontribution to prevent these diseases is highlysignificant.Ifphytate reallydoes show these beneficial propertiesin humansthenphytatewillbenolonger considered an antinutrient.Fordeveloping countries, however, where iron and zinc deficiencies are widelyspread,adequate strategies for preventing deficiencies of minerals and trace elements inducedbyphytateareofutmostsignificance.This canbedone eitherbysupplementationorbydegradingfood phytatesorbyimprovingthedailydietto obtainabetterbalanced supplyofessential nutrients. For optimum use of the beneficial phytate activities in thegut,phytateontheonehandhastobe degradedtoavoid inhibitory effects on the intestinal mineral absorption. On the other hand,if anticancer, antioxidative and anticalcification activities of phytate are to be used, any phytate hydrolysiswouldbe counterproductive.Thisisthe dilemma ofphytatein human nutrition! Thus, the actual demandofa population to either improve mineral and trace element bioavailability or to help prevent cancer, kidney stone formation or other civilisation diseases, will decide whether or not phytate will be welcome in our daily diet. Terms for phytate such as ,antinutrient’ or ,bad food compound’ should belong to the past. i 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.mnf-journal.com S363S363 Mol. Nutr. Food Res. 2009, 53, S330 – S375 Evaluating the literature, some tasks seem to be in the focus of future research: (i) the application of standardised and specific methods forthedeterminationofphyticacidandother inositolphosphates in foods and other biological samples. (ii) Additional information regardingphytic acid intake. (iii)Abetter understandingof the mechanismof the gastro- intestinal absorption and cellular uptake of inositol phosphates and of the role of phytate in the metabolism and of its significance for human health. This review originates from the COST-Concerted Action [14] Raboy, V., myo-Inositol-1,2,3,4,5,6-hexakisphosphates, Phytochemistry 2003, 64,1033 – 1043. [15] Phillippy,B.Q., Bland,J. M., Evens,T.J., Ion chromatographyofphytatein rootsand tubers, J. Agric.Food Chem.2003, 51, 350– 353. [16] Dorsch,J.A., Cook,A.,Young,K.A., Anderson,J.M., et al., Seed phosphorus and inositol phosphate phenotype of barley low phytic acid genotypes, Phytochemistry 2003, 62, 691– 706. 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