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Phytic Acid Chemistry: Influence on Phytin-Phosphorus Availability and Phytase Efficacy1
2002 Poultry Science Association, Inc.
Phytic Acid Chemistry: Influence on Phytin-Phosphorus Availability and Phytase Efficacy1 R. Angel,*,2 N. M. Tamim,* T. J. Applegate,† A. S. Dhandu,* and L. E. Ellestad*
Primary Audience: Feed Formulators, Researchers SUMMARY Poultry diets are primarily composed of seed-based ingredients and contain a high proportion of their P in the phytic acid molecule, making this P poorly available. Phytic acid (IP6) is a highly reactive acidic compound that readily binds mineral cations, and in this complexed form is called phytin. The chemical characteristics of IP6 influence exogenous (diet) and intestinal phytase efficacy and the availability of both phytin-P (PP) and any minerals bound to IP6. Research has shown that minimizing IP6-mineral complexes in the digesta of poultry can lead to increased PP availability in the absence of dietary phytase, as well as increased efficacy of dietary phytases. Understanding these binding interactions and how they influence the efficacy of different phytases can provide a valuable tool in choosing when to use phytase and what phytase to use under different situations. This understanding can provide the foundation for developing new methods, as well as minimizing the cost and maximizing efficacy of current methods, to reduce the amount of P excreted by poultry. Key words: phytin-phosphorus, phytase, availability, phosphorus excretion 2002 J. Appl. Poult. Res. 11:471–480
DESCRIPTION OF PROBLEM The accumulation of P in soils and the threat to surface water quality that may result from P losses to waterways, due to runoff or leaching, are major challenges facing the poultry industry today. Worldwide concerns regarding P content in poultry litter have risen to the forefront among legislators, the public, poultry growers, and integrators. In specific regions, recent legislation has been implemented that limits the use of litter 1
application to soil (based partly on soil and litter P content). Legislation in the state of Maryland requires that all poultry feed contain phytase or other feed additives that decrease excreted P. Given limited land for litter application in areas where the greatest concentration of poultry production exist, the poultry industry must find strategies that reduce P in litter with minimal impact on the economics of producing sellable product. Why is P high in relation to other nutrients in poultry excreta? The primary constituents of
Presented as part of the Informal Nutrition Symposium “Making Sense of Scientific Research and Applying It Properly” at the 91st Annual Meeting of the Poultry Science Association, Newark, Delaware, August 11–14, 2002. 2 To whom correspondence should be addressed: [email protected].
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*4131 Animal and Avian Sciences, Department of Animal and Avian Sciences, University of Maryland, College Park, Maryland 20742; and †1151 Lilly Hall, Room 2-114, Department of Animal Science, Purdue University, West Lafayette, Indiana 47907
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DEFINITION OF TERMS Before delving into the chemistry of IP6, it is important to define terms. Myo-inositol 1,2,3,4,5,6-hexakis dihydrogen phosphate (IP6), an organic phosphate, is a phosphorylated cyclic sugar alcohol. In plants, IP6 exists in its anionic form, phytate. In mature seeds, IP6 is found as a complex salt of Ca, Mg, and K, and in some cases it is bound to proteins and starches. This complexed or chelated molecule of IP6 is known as phytin [1]. Confusion exists related to the terms used for the different forms of P. Total P is generally referred to as P and encompasses any and all forms of P in the diet. Available P (aP) refers to the P that is absorbed from the diet by the animal (i.e., feed P minus P in the distal ileum). Retained P refers to the P that stays in the body (i.e., feed P minus excreta P). As mentioned earlier, most of the P in plant ingredients of seed origin is present in the phytin molecule and is referred to as PP. Any P that is
not bound to the phytin molecule is referred to as nonphytin P (nPP). This nPP can be chemically determined by subtracting analyzed PP from analyzed P. A key difference between aP and nPP is that the term aP includes all absorbed forms of P and will include inorganic P (Pi) and organic P (including PP), whereas nPP excludes any PP available to the animal. The 1994 NRC [2] nPP recommendations for broilers are based mostly on research done using book values of aP for different feed ingredients. NRC states that the nPP requirement for chicks from hatch to 3 wk of age remain unchanged from those published in the previous NRC [3], even though the previous (1984) recommendations were given as aP. Thus, it appears that the most recent NRC [2] used aP and nPP interchangeably, yet aP and nPP are not equivalent as per previous definitions.
PHYTIN Phytin is found primarily in seeds [4], with its location within the seed differing among plants. Ninety percent of the phytin in corn is found in the germ portion of the kernel, whereas in wheat and rice most of the phytin is in the aleurone layers of the kernel and the outer bran [5]. In most oilseeds and grain legumes, phytin is associated with protein and concentrated within subcellular inclusions called globoids that are distributed throughout the kernel; however, in soybean seeds, there appears to be no specific location for phytin [4]. Location of phytin within the seed and its chemical associations with other nutrients influence its availability. Phytin constitutes between 1 and 3%, by weight, of many of the cereals and oilseeds used in animal feeds [6]. In seeds, the role of phytin is as follows: 1) a P reserve, 2) an energy store, 3) a competitor for adenosine triphosphate during the rapid biosynthesis of phytin near seed maturity when seed metabolism is inhibited and dormancy is induced, 4) an immobilizer of divalent cations needed for the control of cellular processes and that are released during germination upon the action of intrinsic plant phytases, and 5) a regulator of readily available seed Pi level [7]. The IP6 molecule contains 28.2% P and is the main source of P in seed-based poultry diets. Phytin accounts for approximately 50 to 80% of the P in seed feedstuffs [4]. The importance of
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poultry diets are seed-based ingredients. Most of the P stored in seeds is present as phytin-P (PP) a form poorly available to poultry. The availability of PP varies within and among ingredients. Therefore, P from inorganic sources, such as deflourinated or dicalcium phosphate, is added to poultry diets to meet requirements and results in diets with a much higher total P than that needed by the bird. This excess P, mainly from poorly available PP, is excreted. To solve the challenge of reducing P in poultry litter in the most economical way, it is important to understand the role the P form most prevalent in poultry diet, PP, has in determining phytase efficacy, PP availability to the bird and the amount of P in excreta. Understanding the interactions that occur in the gastrointestinal tract (GIT) between phytic acid (IP6) with other seed, diet, and digesta nutrients and how these interactions affect efficacy of the different phytases can provide a valuable tool in the applied decision-making process of which dietary phytase to use, when to use it, and how much to use. This understanding may explain why an enzyme capable of hydrolyzing P from phytin, present in the intestinal mucosa of poultry appears to have very limited efficacy and how this efficacy could be improved.
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Chemistry of Phytic Acid Phytic acid has 12 replaceable protons or reactive sites. Six are strongly acidic, with pK of 1.5 to 2.0; two are weakly acidic with a pK of approximately 6.0, and four are very weakly acidic with pK of 9.0 to 11.0 [10, 11]. This means that at pH normally encountered in feeds and in the digestive tract, phytin will carry a strong negative charge and is capable of binding di- and trivalent cations such as Zn2+, Cu2+, Ni2+, Co2+, Mn2+, Fe2+, and Ca2+ in very stable complexes [12, 13, 14, 15, 16, 17, 18], thus reducing the availability of these complexed minerals as well as that of the PP to the animal [9]. The order of potency of selected minerals to inhibit PP hydrolysis by phytase at pH 7 was Zn2+ >> Fe2+ > Mn2+ > Fe3+ > Ca2+ > Mg2+ [14]. The ability of the different metal ions to inhibit PP hydrolysis was related to the stability of the complex, the pH of the solution, and the phytate to mineral molar ratio. Despite the fact that the IP6-Ca complex has a lower stability than other IP6-metal ion complexes, it may be dietary Ca that plays the most critical role in minimizing the effectiveness of both exogenous and endogenous phytases. The level of Ca added to poultry diets is eightto 40-fold higher than that of Zn, and the impact of the much higher concentrations of Ca may be more powerful than its lower affinity due to simple mass action. The impact of Ca may be exacerbated by the presence of other cations, as Ca has been implicated in secondary synergistic effects. When two cations are present simultaneously they increase the proportion of the IP6metal complex that precipitates [19].
In addition to complexing with macro- and microminerals, intrinsic in seeds and those added to diets, IP6 may form complexes with proteins and starches. Proteins are able to bind directly with IP6 through electrostatic charges at low pH or through salt bridges formed at high pH, [20]. Starch binding occurs as a result of hydrogen bond formation [21]. The binding of IP6 with protein and starch may also reduce the availability of these nutrients from the diet. Furthermore, the complexing of IP6 with proteins or starches influences the degree of ionization of the complex and the ability of phytin to complex with minerals, potentially changing the efficacy of different phytases. From an applied poultry nutrition standpoint, understanding the interactions of IP6 with other nutrients in feeds and in the digesta is important because the physical state or solubility of IP6mineral complexes in the digesta dictates what the availability of PP will be. If the IP6-mineral complex is precipitated, then any phytases present in the small intestine, regardless of origin (i.e., added to the diet intrinsic in the GIT), will be unable to exert their hydrolytic effect resulting in a greatly reduced phytase efficacy. The increase in pH of the digesta as it moves distally along the GIT causes the PP molecule to be ionized and to more readily form complexes with divalent metal cations such as Zn, Ca, Mg, and Fe [12, 14]. The higher pH result in decreased solubility [16] of the complex and, therefore, decreased efficacy of phytase [14].
PHYTASES Phytases (myo-inositol hexaphosphate hydrolases) are phosphatase enzymes able to catalyze the hydrolysis of phosphate ester bonds. These enzymes are capable of hydrolyzing one or more phosphate groups from IP6 yielding Pi and a series of lower phosphoric esters [22, 23, 24]. Phytases are widely distributed in plants, animals, and microorganisms. The International Union of Biochemists [25] currently acknowledges two classes of phytase enzymes: a 3-phytase (EC 3.1.3.8) and a 6-phytase (EC 3.1.3.26), which initiate the dephosphorylation of IP6 at different positions on the inositol ring and produce different isomers of the lower inositol phosphates. The 3-phytases initiate the dephosphory-
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IP6 as a nutrient is primarily related to its P content [8]. In most cases of plant-based poultry diets, there are adequate levels of P to meet dietary needs, provided that the PP is released from the IP6 molecule as bioavailable Pi. In general, without some dietary modification, a large proportion of the PP is unavailable so that without added dietary Pi most poultry diets do not contain enough aP to meet bird requirements. Phytin is most commonly thought of as an antinutrient [9], because of its ability to complex or bind mineral cations in the seed or diet, rendering these bound cations as well as the PP partially or completely unavailable to the animal.
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pH optima of these enzymes as well as their thermostability [36] and resistance to inactivation by gastrointestinal proteolytic enzymes [32] make them more suitable as feed additives to increase PP availability than relying on plant phytases. Addition of microbial phytase to feed has led to increased PP utilization in turkeys [37, 38, 39] and chickens [38, 40, 41, 42, 43, 44, 45, 46]. The microflora in the GIT of animals are capable of hydrolyzing PP, but the impact of this hydrolysis has not been well documented and poultry research in this area, to the authors’ knowledge, has been minimal. Most of this research has been done in ruminants [8, 47, 48], rats [49], and swine [50]. Intestinal Phytase The first report of the presence of a phytatesplitting enzyme in the GIT was in rats in 1937 [51]. Since then numerous studies have demonstrated that extracts of the intestinal mucosa of ruminant and monogastric species are capable of hydrolyzing PP. Phytase activity has been detected in the small intestinal mucosa of chickens [18, 52, 53, 54, 55, 56, 57]. Even though phytase has been found in all segments of the small intestine, the highest levels have been found in the duodenum [53, 56]. There is controversy among scientists as to whether intestinal phytase activity is merely a manifestation of nonspecific acid or alkaline phosphatase activity, but this controversy is immaterial from a practical perspective. If small intestinal mucosa preparations are shown to be able to release Pi from IP6, it is irrelevant whether this is accomplished by a specific phytase or not. Published literature supports that an enzyme in the intestinal mucosa of poultry is capable of hydrolyzing PP [56, 58, 59]. This enzyme is affected by age of the animal, where higher levels have been found in mature versus young birds [18, 56], as well as genetics [18, 53] and nutrient levels in the diet. In addition to the effects of age and genetics, intestinal phytase activity is influenced by several nutritional factors. Citrate added to rat diets has been shown to decrease in vitro intestinal phytase activity [60, 61] but increase in vivo PP hydrolysis [56, 60]. This increase in PP hydrolysis in vivo may be due to the Ca-chelating ability
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lation of IP6 at the 3 position, yielding 1,2,4,5,6pentakisphosphate and Pi, whereas the 6-phytases initiate the dephosphorylation of IP6 at the 6 position, yielding 1,2,3,4,5-pentakisphosphate and Pi. The 3-phytases do not always completely dephosphorylate IP6, whereas the 6-phytases do [26]. It has been stated that microorganisms normally produce the 3-phytases and 6-phytases are normally found in plants [8, 22]. However, exceptions to this general rule have been reported. For example, an enzyme with 3-phytase activity has been reported in soybeans [27], and an enzyme with 6-phytase activity has been reported in Escherichia coli [28]. Microbial phytases tend to have pH optima in the range of 2 to 6, whereas plant phytases tend to have a pH optimum near 5 [26]. There are four possible sources of phytase that can be found in the digestive tracts of animals: 1) phytase present intrinsically in feed ingredients, 2) exogenous microbial phytases added to the diet, 3) phytases produced by microflora present in the GIT, and 4) intestinal mucosa membrane linked phytase. Phytase present in feed ingredients has been shown to improve PP utilization by monogastric animals [9]. High levels of phytase are found in wheat, rye, triticale, and barley [29, 30], yet there is a large variation of phytase activity within the seeds of the different varieties of these plants [31]. Phytase in feed ingredients may be inactivated by temperature during feed processing [26], by the low pH that occurs in the upper portion of the GIT [32], and by the action of pepsin on gastric secretions [32]. It has been shown that there is a decrease in phytase activity when pelleting temperatures exceed 75°C [33, 34], but this will be dependent on the form and type of the enzyme. The variability and instability of these natural phytases limit the use of plant ingredients as a reliable source of the enzyme in feeds. Many bacteria, yeasts, and fungi produce phytase [9]. Of all the microorganisms studied, phytase production is highest in the fungi Aspergilli with the highest levels of extracellular phytase produced by Aspergillus niger [35]. Aspergilli tend to produce two different phytases, one with pH optima of 5.5 and 2.5 (phyA) and one with a pH optimum of 2.0 (phyB) [26]. The
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PHYTIN PHOSPHORUS USE BY POULTRY Most evidence indicates that poultry are capable of utilizing at least a portion of dietary PP. However, there is wide variation in reported availability. Researchers working with poultry in the mid-1900s concluded that PP was virtually unavailable to chickens when supplied in its natural form or as the purified Ca or Na salt [69, 70, 71]. In later reports [71, 72], chickens were found to have a limited ability to utilize P from plant sources. Broiler chickens were found to inefficiently use PP, and the authors hypothesized that this was due to a minimal quantity of intestinal phytase [47], although intestinal phytase activity was not measured. As mentioned previously, there is evidence demonstrating that an enzyme in the intestinal mucosa can hydrolyze PP [14, 54]. The magnitude of the impact this endogenous phytase has on the hydrolysis of the PP molecule in vivo is still unknown. More recent studies with chickens indicate that PP utilization is variable and that dietary factors including the level of Ca, nPP, P, vitamin D3, and vitamin D3 metabolites, as well as feed processing and feed or ingredient particle size may influence PP hydrolysis in the GIT. Phytin P utilization by chickens is reported to be as low as 0 to 15% [43, 64, 73] and as high as 70 to 75% [41, 65, 74]. Hydrolysis of PP in 4-wkold female broilers ranged from 3 to 42% and depended primarily on dietary Ca level in the diet [63]. Three-week-old broilers were reported to utilize PP, and availability of P from plant feedstuffs varied between 16% in corn to 80% in lupin [75]. Availability of P from purified monosodium phytate was similar to that of feedgrade dicalcium phosphate or a reagent-grade monosodium phosphate, but P from Ca phytate was poorly available [76]. In general, the variation in the availability of PP observed in different research trials could be due to differences in experimental design, research methodologies, diet ingredient and nutrient composition, diet processing, analytical methods, age, species, and breed. Different ingredients have varying amounts of PP [77, 78] and different locations of PP within the plant [4]. Some feed ingredients such as wheat and rye have endogenous phytase [79], which might
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of citrate, making the Ca less available to form insoluble complexes with IP6. There is some indication that the level of IP6 may influence intestinal phytase activity; however, conflicting results have been reported. Neither purified sources of Ca phytate nor Na phytate added to the diet increased intestinal phytase activity in chicks according to one study [18], but another study [58] reported that increasing the level of IP6 in the diet of chicks decreased intestinal phytase activity. Dietary mineral levels influence intestinal phytase activity. High levels of Mg in the diet (0.4 and 0.6%) depress intestinal phytase activity of chicks [59]. Increasing the level of dietary Ca has also been shown to decrease PP hydrolysis in chickens [41, 42, 62, 63, 64, 65]. It has been suggested [12] that a diet with a molar ratio of Ca:IP6 greater than 6:1 leads to formation of insoluble Ca-IP6 complexes that are inaccessible to diet or intestinal phytases. High dietary Ca depresses intestinal phytase activity in chicks [58, 59]. Conversely, others have demonstrated that increased dietary Ca concentrations actually increase intestinal phytase activity in chickens [57]. Decreasing dietary Pi, from 0.48 to 0.16%, has been shown to double intestinal phytase activity in chicks [58, 59]. Conversely, others have reported no difference in the intestinal phytase activity of chickens or hens fed dietary Pi between 0.2 and 0.6% [18]. Altering the vitamin D3 level of the diet also influences the activity of intestinal phytase. An early study found that vitamin D3 added to a diet containing purified Ca phytate increased the utilization of PP by rats but did not increase the activity of intestinal phytase [66]. However, subsequent studies have demonstrated that phytase activity in chicken intestinal extracts is increased slightly as the vitamin D3 level of the diet is increased [58, 67]. The low dietary level of inorganic phosphate (0.25%) used in the early study [66] may have increased intestinal phytase activity so that the influence of vitamin D3 was masked. Recent studies have found mucosal phytase activity in chickens not to be influenced by dietary levels of 1α-hydroxycholecalciferol [68] or 25-hydroxycholecalciferol [57], although both compounds were shown to increase PP hydrolysis in vivo [57, 68].
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size of the soluble IP6-mineral complex will impact the efficacy of the different phytases, and this impact will be specific to each type of phytase [81, 84]. Insoluble IP6-mineral complexes that form at the higher pH, where plant phytases are most active, may be resistant to hydrolysis by phytases of plant [84] and microbial origin [14]. When one looks at commercially available phytases, it is important to know what the pH optima is because that will dictate where the phytase will function along the GIT. For example, a commercially available microbial phytase from Aspergillus niger that has two pH optimas, one at 2.5 and the other at 5.5 [40], would be active in the gastric area where the pH is 2 [81]. At gastric pH, most of the IP6-mineral complex is in solution, making hydrolysis possible. Given the chemical properties of IP6 under GIT conditions it is not surprising that no appreciable PP hydrolysis was observed after the digesta exited the gizzard [84]. In a study [85] where a basal corn-soy diet with no added Ca, Pi, or microminerals and analyzed to contain 0.18% Ca, 0.40% P, and 0.29% PP was fed for 30 h to 20-d-old broiler chicks, PP disappearance (hydrolysis) in the GIT, up to the ileum, was 67.1%. When Ca carbonate was added to the basal diet to supply 0.5% Ca (analyzed total dietary Ca of 0.70%), PP hydrolysis was 18.9%. Thus, PP hydrolysis, from a cornsoy diet, in the GIT increased by a factor of 3.55 in the absence of added Ca. In this work, inclusion of an inorganic micromineral mix at average commercial levels had no effect on PP hydrolysis in the GIT, regardless of the presence or absence of Ca. Use of a commercially available micromineral-protein chelate had no effect on PP hydrolysis in the GIT versus when the inorganic micromineral source was used, and this may have been related to the ionization potential of the micro-mineral chelate in the GIT. The impact of Ca level in the diet on the efficacy of two different commercially available phytases was determined in a second study [86]. As in the first study, not adding Ca to a corn-soy diet resulted in a 2.5-fold increase in PP hydrolysis in the GIT vs. when 0.50% Ca was added. Two phytase sources were tested, a 3- and a 6-phytase, in which the researchers determined pH
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influence the extent of PP availability. The possible reason for the difference in PP availability observed when similar dietary ingredients were used could be related to experimental methodology differences or to the differences in PP level within the same ingredients depending on level of fertilization, growing conditions, and stage of maturity of the seed at harvest [80]. Researchers have demonstrated that as soil available P increases, total P and PP increase in the seed and that the increase in P is primarily due to an increase in PP [80]. The effect of high levels of diet Ca on GIT pH may be partly responsible for some of the deleterious effects that high dietary Ca has on PP hydrolysis [81, 82, 83]. Increasing dietary Ca from 1.07 to 2.53% resulted in higher pH in the crop (4.89 vs. 5.32, respectively) and ileum (6.62 vs. 7.39, respectively) [81]. IP6-mineral complexes are soluble at low pH (less than 3.5) with maximum insolubility occurring between pH 4 and 7 [81], and thus anything that alters intestinal pH will alter the efficacy of phytases and may affect different phytases in different ways. Size of the soluble IP6-mineral complexes also influences the availability of PP and IP6 complexed minerals. Smaller sizes of IP6-mineral complex have greater surface area available for the enzyme to attack, making the complex more easily hydrolyzed. The proportion of larger IP6-mineral complexes in the ileum was greater than in the duodenum, and as dietary Ca increased, the size of the IP6-mineral complexes increased, regardless of region of the intestine [81]. When considering this information, it is important to put it into the context of where in the GIT phytases may be active and where absorption of different nutrients occur. At the pH of the small intestine, not only are most of the IP6-mineral chelates precipitated, but when dietary Ca and P concentrations are increased, the proportion of IP6-mineral complexes precipitated also increases. The solubility of Ca was 16.9% when the Ca and aP were 1.53 and 0.43%, respectively, compared to a solubility of 8.3% when they were 2.26 and 0.83%, respectively [81]. Thus, increased dietary Ca and aP levels reduce the proportion of soluble minerals and thereby further decrease the availability of these minerals and of PP. The
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Potential for Future Applications As mentioned earlier dietary Ca has the highest practical implications because it is present in the highest concentration relative to other metal ions in the diet. Delivery of Ca to the birds in a manner that minimizes IP6-Ca complexing would result in improved phytase efficacy and increased PP availability. This is not a new concept [14, 63, 73], but until now there has been no economic incentive to develop a Ca chelate for animals. Developing a product that delivers Ca to the bird but does not complex with IP6 makes more economic sense given current and expected future environmental regulations. The inhibitory effect of Ca on PP hydrolysis can be prevented by a strong Ca chelator such as EDTA [14]. A concentration of EDTA greater than 5 mM is required to prevent the inhibitory effect of Ca on PP hydrolysis [14]. The mode of action of EDTA may be that it is competing for Ca ions. If a chelator of mineral ions is to improve PP hydrolysis in a diet, the affinity of the chelator would need to be sufficient to competitively bind to Ca, as PP is a stable and strong Ca chelator. The chelator to be developed must be either absorbed as a complex or be able to release Ca close to the absorptive surface in a way that minimizes potential IP6-Ca complexing. In addition, if the complex is absorbed intact, then the chelator must release Ca after the animal absorbs the complex. It is important to consider the potential other interactions of the chelator within the body. Based on the results of some of the work discussed in this review [85, 86], along with
what has been discussed previously in this paper about IP6 chemistry, a baseline of information is starting to emerge that provides some basis for understanding why different phytases act differently in the GIT and how changes in diet or intestinal pH can lead to large changes in enzyme efficacy. Recent work [85, 86] confirms and expands on previous research [14] and points to the possibility of developing new dietary feeding strategies or feed additives that will allow for the delivery of Ca into the bird in a manner that minimizes the formation of IP6-Ca complexes and allows the bird to use PP more efficiently through an increase in efficacy of diet and endogenous phytases.
INFORMATION NEEDED FOR INTERPRETATION OF PHYTASE EFFICACY RESEARCH To minimize the limitations we have as scientists in interpreting published research, key information should be included. Knowledge of the chemical characteristics of a key constituent of seed-based ingredients such as phytin and its interactions with other nutrients in the digest as it moves along the GIT makes this even more important. One of the limitations in interpreting published research is that numerous papers lack the information necessary for comparison with other work as well as for interpretation of the information published. Most work published on phytase use in animal diets does not include information on actual PP content of the diet and relies on book values of PP in the ingredients used. Also, in numerous papers, the amount of phytase added to diets is based on addition of a quantity of commercial product where the addition to the diet is based on the label claim of that product. Few papers have actually analyzed the commercial product to be used. Analytical procedures are available and accurate for analysis of phytase premixes. Few report analyzed levels of the enzyme in the final diets, which is not surprising since the analytical procedure for diet phytase is currently not very accurate. Other key information that often is missing from papers is the final weight, breed, strain, or sex of the animals used in the study and analyzed values for key nutrients being studied, especially those that have an important effect on the nutrient being studied.
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optimas of 3 and 5 for the 3-phytase and between 4 and 4.5 for the 6-phytase. When the 6-phytase was tested, PP hydrolysis in the GIT of broiler chicks was 0.093 and 0.055% per unit of analyzed dietary phytase when the chicks were fed diets containing 0 and 0.50% added Ca from Ca carbonate, respectively, for 30 h. When 3phytase was tested, PP hydrolysis in the GIT was higher per unit of analyzed phytase regardless of Ca level in the diet (0.137 and 0.110% in diets with 0 and 0.50% added Ca, respectively). Phytase level in the diet was formulated based on analyzed premix levels to be 500 U/kg. These diets contained no added Pi but had average commercial levels of inorganic microminerals.
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the response to the additive is beyond the response curve range. Appropriate selection of response criteria is also important. For example, when the treatment being studied has an effect on gain, use of percentage bone ash as the response criteria will lead to predictions that are not correct. How published data is interpreted is ultimately in the hands of the person wishing to use the information. Thus, readers must use their own judgment and should be cognizant of experimental design and the pitfalls inherent in the work. For example when comparing efficacy between studies it is important to note what the source of P used to determine the response curve is. The reason this information is so important is that the availability of the P source will impact measured efficacy and different P sources have different availabilities.
CONCLUSIONS AND APPLICATIONS 1. The strong chelating properties of phytic acid (IP6) present in plant-based diets result in — formation of IP6-mineral complexes that are soluble at proventriculus and gizzard pH but insoluble at the pH of the small intestine. — a large reduction in the efficacy of phytase when the IP6-mineral complexes are present, regardless of source of phytase. — varying sizes of soluble IP6-mineral complexes, with larger complexes reducing phytase efficacy. 2. Calcium, rather than other mineral ions in the diet, is the metal ion most involved in the IP6mineral complexes that reduce PP availability. 3. It is possible to develop feed management tools that maximize innate abilities of birds to use PP and thus alter the need for supplemental Pi and phytase.
REFERENCES AND NOTES 1. Lott, J. N. A. 1984. Accumulation of seed reserves of phosphorus and other minerals. Pages 139–166 in Seed Physiology. Vol. 1. D. R. Murray, ed. Academic Press, Sydney, Australia.
7. Cosgrove, D. J., and G. C. J. Irving. 1980. In Inositol Phosphates: Their chemistry, Biochemistry and Physiology. Elsevier/ North Holland, Inc., New York.
2. National Research Council. 1994. Nutrient Requirements of Domestic Animals. Nutrient Requirements of Poultry. 9th rev. ed. Natl. Acad. Sci., Washington, DC.
8. Reddy, N. R., S. K. Sathe, and D. K. Saunkhe. 1982. Phytates in legumes and cereals. Adv. Food Res. 28:1–92.
3. National Research Council. 1984. Nutrient Requirements of Domestic Animals. Nutrient Requirements of Poultry. 8th rev. ed. Natl. Acad. Sci., Washington, DC.
9. Pallauf, J., and G. Rimbach. 1997. Effect of supplemental phytase on mineral and trace element bioavailability and heavy metal accumulation in pigs with different type diets. In Phytase in Animal Nutrition and Waste Management. M. B. Coehlbo and E. T. Kornegay, ed. BASF Publication DC9601. BASF Corporation, Mount Olive, NJ.
4. Ravindran, V., W. L. Bryden, and E. T. Kornegay. 1995. Phytin: Occurrence, bioavailability and implications in poultry nutrition. Poult. Avian Biol. Rev. 6:125–143. 5. O’Dell, B. L., and A. R. De Boland. 1976. Complexation of phytin with proteins and cations in corn germ and oilseed meals. J. Agric. Food Chem. 24:804–808.
10. Costello, A. J. R., T. Glonek, and T. C. Myers. 1976. 31Pnuclear magnetic resonance-pH titrations of myo-inositol hexaphosphate. Carbohydr. Res. 46:159–165.
6. Cheryan, M. 1980. Phytic acid interactions in food systems. CRC Crit. Rev. Food Sci. Nutr. 13:297–335.
11. Erdman, J. W., Jr. 1979. Oilseed phytates: Nutritional implications. J. Am. Oil Chem. Soc. 56:736–741.
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In the case of phytase studies, the substrate (PP) that the enzyme is working on should be analyzed, as well as P and Ca. Vitamin D formulated values as well as source should also be provided. Precise ingredient description and inclusion levels should be listed. In efficacy studies researchers will sometime use response curves derived based on treatment means rather than based on pen (replicate) measured levels to calculate efficacy. This overestimates the accuracy of the calculated efficacy value and results in the use of an equation that is not correct. Response curves should be determined based on treatment replicate mean information. Also of concern is the prediction of efficacy based on response curves that have poor correlation coefficients. It is also incorrect to assume that the relationships found in the response curve extend beyond the response curve levels studied when
ANGEL ET AL: INFORMAL NUTRITION SYMPOSIUM 12. Wise, A., 1983. Dietary factors determining the biological activities of phytase. Nutr. Abstr. Rev. 53:791–806. 13. Persson, H., M. Turk, M. Nyman and A. S. Sandberg. 1998. Binding of Cu2+, Zn2+, and Cd2+ to inositol Tri-, tetra-, penta-, and hexaphosphate. J. Agric. Food Chem. 46:3194–3200. 14. Maenz, D. D., C. M. Engle-Schaan, R. W. Newkirk, and H. L. Classen. 1999. The effect of minerals and mineral chelators on the formation of phytase-resistant and phytase susceptible forms of phytic acid in solution and in a slurry of canola meal. Anim. Feed Sci. Technol. 81:177–192. 15. Wise, A. 1995. Phytate and zinc bioavailability. Int. J. Food Sci. Nutr. 46:53–63.
17. Vohra, P., A. Gray, and F. H. Kratzer. 1965. Phytic acid metal complexes. Proc. Soc. Exp. Biol. Med. 120:447. 18. Maddaiah, V. T., A. A. Kurnick, and B. L. Reid. 1964. Phytic acid studies. Proc. Soc. Exp. Biol. Med. 115:391. 19. Simpson, C. J., and A. Wise. 1990. Binding of zinc and calcium to inositol phosphates (phytate) in vitro. Br. J. Nutr. 64:225–232. 20. Graf, E. 1986. Chemistry and applications of phytic acid: an overview. Pages 173–194 in Phytic Acid: Chemistry and Applications. E. Graf, ed. Pilatus Press, Minneapolis, MN. 21. Thompson, L. U. 1986. Phytic acid: a factor influencing starch digestibility and blood glucose response. Pages 173–194 in Phytic Acid: Chemistry and Applications. E. Graf, ed. Pilatus Press, Minneapolis, MN. 22. Nayini, N. R., and P. Markakis. 1986. Phytases. Pages 101– 118 in Phytic Acid: Chemistry and Applications. E. Graf, ed. Pilatus Press, Minneapolis, MN. 23. Lasztity, R., and L. Lasztity. 1990. Phytic acid in cereal technology. Pages 309–371 in Advances in Cereal Science and Technology. Vol. 10. Y. Pomeranz, ed. Amer. Assoc. of Cereal Chem., Inc., St. Paul, MN.
33. Pointillart, A., 1993. Importance of phytates and cereal phytases in the feeding of pigs. Pages 192–198 in Enzymes in Animal Nutrition. C Wenk, ed. ETH-Zurich, Zurich. 34. Hughes, K. P., and J. H. Soares Jr. 1998. Efficacy of phytase on phosphorus utilization in practical diets fed to striped bass, Morone saxatilis. Aquacult. Nutr. 4:133–140. 35. Shieh, T. R., and J. H. Ware. 1968. Survey of microorganisms for the production of extracellular phytase. Appl. Microbiol. 16:1348–1351. 36. Wyss, M., L. Pasamontes, R. Remy, J. Kohler, E. Kusznir, M. Gadient, F. Muller, and A. P. G. M. Van Loon. 1998. Comparison of the thermostability properties of three acid phosphatases from molds: Aspergillus fumigatus phytase, A. niger phytase, and A. niger pH 2.5 acid phosphatase. Appl. Environ. Microbiol. 64:4446–4451. 37. Qian, H., E. T. Kornegay, and D. M. Denbow. 1996. Phosphorus equivalence of microbial phytase in turkey diets as influenced by calcium to phosphorus ratios and phosphorus levels. Poult. Sci. 75:69–81. 38. Yi, Z., E. T. Kornegay, V. Ravindran, and D. M. Denbow. 1996. Improving phytate phosphorus availability in corn and soybean meal for broilers using microbial phytase and calculation of phosphorus equivalency values for phytase. Poult. Sci. 75:240–249. 39. Zyla, K., D. R. Ledoux, M. Kujawski, and T. L. Veum. 1996. The efficacy of an enzymatic cocktail and a fungal mycelium in dephosphorylating corn-soybean meal based feeds fed to growing turkeys. Poult. Sci. 75:381–387. 40. Simons, P. C. M., H. A. J. Versteegh, A. W. Jonbloed, P. A. Kemme, P. Slump, K. D. Bos, M. G. E. Wolters, R. F. Beudeker, and G. J. Verschoor. 1990. Improvement of phosphorus availability by microbial phytase in broilers and pigs. Br. J. Nutr. 4:525–540. 41. Mitchell, R. D., and H. M. Edwards, Jr. 1996. Effects of phytase and 1,25dihydroxycholecalciferol on phytate utilization and the quantitative requirement for calcium and phosphorus in young broiler chickens. Poult. Sci. 75:95–110. 42. Qian, H., E. T. Kornegay, and D. M. Denbow. 1997. Utilization of phytate phosphorus and calcium as influenced by microbial phytase, cholecalciferol, and the calcium:total phosphorus ratio in broiler diets. Poult. Sci. 76:37–46.
24. Harland, F. B., and E. R. Morris, 1995. Phytin: A good or a bad food component. Nutr. Res. 15:733–754.
43. Carlos, A. B., and H. M. Edwards, Jr. 1998. The effects of 1,25-Dihydroxycholecalciferol and phytase on the natural phytate phosphorus utilization by laying hens. Poult. Sci. 77:850–858.
25. International Union of Biochemists (IUB). 1979. Enzyme nomenclature. Page 247 in Recommendations of the Nomenclature Committee of the International Union of Biochemistry. Academic Press, New York.
44. Rao, S. V. R., V. R. Reddy, and V. R. Reddy. 1999. Nonphytin requirements of commercial broilers and White Leghorn layers. Anim. Feed Sci. Technol. 80:1–10.
26. Wodzinski, R. J., and A. H. J. Ullah. 1996. Phytase. Pages 263–302 in Advances in Applied Microbiology. Vol. 42. Academic Press, Inc., New York.
45. Waldroup, P. W., J. H. Kersey, E. A. Saleh, C. A. Fritts, F. Yan, H. L. Stilborn, R. C. Crum, Jr., and V. Raboy. 2000. Nonphytate phosphorus requirement and phosphorus excretion of broilers fed diets composed of normal or high available phosphate corn with and without microbial phytase. Poult. Sci. 79:1451–1459.
27. Phillippy, B. Q., M. R. Johnston, S. H. Tao, and M. R. S. Fox. 1988. Inositol phosphates in processed foods. J. Food Sci. 53:496–499. 28. Greiner, R., U. Konietzny, and K. D. Jany 1993. Purification and characterization of two phytases from Escherichia coli. Arch. Biochem. Biophys. 303:107–113. 29. Eeckhout, W., and M. De Paepe. 1994. Total phosphorus, phytase phosphorus and phytase activity in plant feedstuffs. Anim. Feed Sci. Technol. 47:19–29. 30. Viveros, A., C. Centeno, A. Brenes, R. Canales, and A. Lozano. 2000. Phytase and acid phosphatase activities in plant feedstuffs. J. Agric. Food Chem. 48:4009–4013. 31. Nys, Y., D. Frapin and A. Pointillart. 1996. Occurrence of phytase in plants, animals microorganisms. Pages 213–226 in Phytase in Animal Nutrition and Waste Management, M. B. Coelho and E. T. Kornegay, ed. BASF Corporation, Mount Olive, NJ. 32. Phillippy, B. Q. 1999. Susceptibility of wheat and Aspergillus niger phytases to inactivation by gastrointestinal enzymes. J. Agric. Food Chem. 47:1385–1388.
46. Yan, F., J. H. Kersey, and P. W. Waldroup. 2001. Phosphorus requirements of broiler chicks three to six weeks of age as influenced by phytase supplementation. Poult. Sci. 80:455–459. 47. Nelson, T. S., L. B. Daniels, J. R. Hall, and L. G. Shields. 1976. Hydrolysis of natural phytate phosphorus in the digestive tract of calves. J. Anim. Sci. 42:1509–1512. 48. Morse D., H. H. Head, and C. J. Wilcox. 1992. Disappearance of phosphorus in phytate from concentrates in vitro and from rations fed to lactating dairy cows. J. Dairy Sci. 75:1979–1986. 49. Lopez, H. W., F. Vallery, M. A. Levrat-Verny, C. Coudray, C. Demigne, and C. Remesy. 2000. Dietary phytic acid and wheat bran enhance mucosal phytase activity in rat small intestine. J. Nutr. 130:2020–2025. 50. Sandberg, A. S., T. Larsen, and B. Sandstrom. 1993. High dietary calcium level decreases colonic phytate degradation in pigs fed a rapeseed diet. J. Nutr. 123:559–566. 51. Patwardhan, V. N. 1937. The occurrence of a phytin-splitting enzyme in the intestine of albino rats. Biochem. J. 31:560–564.
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16. Kaufman, H. W., and I. Kleinberg. 1971. Effect of pH on calcium binding by phytic acid and its inositol phosphoric acid derivatives and on the solubility of their calcium salts. Arch. Oral Biol. 16:445–460.
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52. Spitzer, R. R., and P. H. Phillips 1945. Enzymatic relationships in the utilization of soybean oil meal phosphorus by the rat. J. Nutr. 30:183–192.
69. Lowe, J. T., H. Steenbock, and C. H. Kreiger. 1939. Cereals and rickets. XII. The availability of phytin-P to the chicks. Poult. Sci. 18:40–41.
53. Bitar, K., and J. G. Reinhold. 1972. Phytase and alkaline phosphatase activities in intestinal mucosa of rat, chicken, calf, and man. Biochim. Biophys. Acta 268:442–452.
70. Krieger, C. H., R. Bunkfeldt, and H. Steenbock. 1940. Cereals and rickets. X. The availability of phytic acid phosphorus. J. Nutr. 20:7–14.
54. Davies, M. I., and I. Motzok, 1972. Properties of chick intestinal phytase. Poult. Sci. 51:494–501.
71. Gillis, M. B., L. C. Norris, and G. F. Heuser. 1953. Phosphorus metabolism and requirements of hens. Poult. Sci. 32:977–984.
55. Biehl, R. R., D. H. Baker, and H. F. De Luca 1995. 1 alphahydroxylated cholecalciferol compounds act additively with microbial phytase to improve phosphorus, zinc, and manganese utilization in chicks fed soy-based diets. J. Nutr. 125:2407–2416.
72. McGinnis, J., L. C. Norris, and G. F. Heuser. 1944. Poor utilization of phosphorus in cereals and legumes by chicks for bone development. Poult. Sci. 23:157–159.
57. Applegate, T. J., C. R. Angel, H. L. Classen, R. W. Newkirk, and D. D. Maenz. 2000. Effect of dietary calcium concentration and 25-hydroxycholecalceferol on phytate hydrolysis and intestinal phytase activity in broilers. Poult. Sci. 79(Suppl. 1):21. (Abstr.) 58. Davies, M. I., G. M. Ritcey and I. Motzok. 1970. Intestinal and alkaline phosphatase of chicks: Influence of dietary calcium, inorganic and phytate phosphorus and vitamin D3. Poult. Sci. 49:1280–1286. 59. McGuaig, L. W., M. I. Davies, and I. Motzok. 1972. Intestinal alkaline phosphatase and phytase of chicks: Effect of dietary magnesium, calcium, phosphorus and thyroactive casein. Poult. Sci. 51:526–530. 60. Pileggi, V. J., H. F. De Luca, J. W. Cramer, and H. Steenbock. 1956. Citrate in the prevention of rickets in rats. Arch. Biochem. Biophys. 60:52–57. 61. Roberts, A. H., and J. Yudkin. 1961. Effect of phytate and other dietary factors on intestinal phytase and bone calcification in the rat. Br. J. Nutr. 15:457–471. 62. Edwards, H. M., Jr. 1982. Phosphorus. 1. Effect of breed and strain on utilization of suboptimal levels of phosphorus in the ration. Poult. Sci. 62:77–84. 63. Ballam, G. C., T. S. Nelson, and L. K. Kirby. 1984. Effect of fiber and phytate source and of calcium and phosphorus level on phytate hydrolysis in the chick. Poult. Sci. 63:333–338. 64. Scheideler, S. E., and J. L. Sell. 1987. Utilization of phytate phosphorus in laying hens as influenced by dietary phosphorus and calcium. Nutr. Rep. Int. 35:1073–1081. 65. Mohammed, A., M. J. Gibney, and T. G. Taylor. 1991. The effects of dietary levels of inorganic phosphorus, calcium and cholecalciferol on the digestibility of phytate-P by the chick. Br. J. Nutr. 66:251–259. 66. Spitzer, R. R., G. Maruyama, L. Michaud, and P. H. Phillips. 1948. The role of vitamin D in the utilization of phytin phosphorus. J. Nutr. 35:185–193. 67. Steenbock, H., C. H. Krieger, W. G. Wiest, and V. J. Pileggi, 1953. Vitamin D and intestinal phytase. J. Biol. Chem. 205:993–999. 68. Biehl, R. R., and D. H. Baker. 1997. 1-alpha-hydroxycholecalciferol does not increase the specific activity of intestinal phytase but does improve phosphorus utilization in both cecectomized and sham-operated chicks fed cholecalciferol-adequate diets. J. Nutr. 127:2054–2059.
73. Nelson, T. S. 1976. The hydrolysis of phytin phosphorus by chicks and laying hens. Poult. Sci. 55:2262–2264. 74. Edwards, H. M., Jr. 1993. Dietary 1,25-hydroxycholecalciferol supplementation increases natural phytate phosphorus utilization in chickens. J. Nutr. 123:567–577. 75. Van der Klis, J. D., and H. A. J. Versteegh. 1996. Phosphorus nutrition of poultry. Pages 71–83 in Proc. Nottingham Feed Mfgr. Conf, Nottingham, UK. 76. Waldroup, P. W., C. B. Ammerman, and R. H. Harms. 1964. The availability of phytic acid phosphorus for chicks. 2. Comparison of phytin phosphorus sources. Poult. Sci. 43:426–431. 77. Nelson, T. S., L. W. Ferrara, and N. L. Storer. 1968. Phytin phosphorus content of feed ingredients derived from plants. Poult. Sci. 47:1372–1374. 78. Nelson, T. S. 1967. The utilization of phytin phosphorus by poultry—A review. Poult. Sci. 46:862–869. 79. Barrier-Guillot, B., P. Casado, P. Maupetit, C. Jondreville, and F. Gatel. 1996. Wheat phosphorus availability: 2—In vivo study in broilers and pigs; relationship with endogenous phytase activity and phytic phosphorus content in wheat. J. Sci. Food Agric. 70:69–74. 80. Barrier-Guillot, B., P. Casado, P. Maupetit, C. Jondreville, and F. Gatel. 1996. Wheat phosphorus availability: 1—In vitro study; factors affecting endogenous phytasic activity and phytic phosphorus content. J. Sci. Food Agric. 70:62–68. 81. Shafey, T. M., M. W. McDonald, and J. G. Dingle. 1991. Effects of dietary Ca and available phosphorus on digesta pH and on the availability of Ca, iron, magnesium and zinc from the intestinal contents of meat chicken. Br. Poult. Sci. 32:185–194. 82. Selle, P. H., V. Ravindran, R. A. Caldwell, and W. L. Bryden. 2000. Phytate and phytase: Consequences for protein utilization. Nutr. Res. Rev. 13:255–278. 83. Champagne, E. T. 1988. Effects of pH on mineral-phytate, protein-mineral-phytate, and mineral-fiber interactions. Possible consequences of atrophic gastritis on mineral bioavailability from highfiber foods. J. Am. Coll. Nutr. 7:499–508. 84. Sooncharernying, S., and H. M. Edwards, Jr. 1993. Phytate content of excreta and phytate retention in the gastrointestinal tract of young chickens. Poult. Sci. 72:1906–1916. 85. Tamim, N. M., and R. Angel. 2002. Effect of dietary calcium level and micro mineral source on phytin phosphorus hydrolysis in broilers. Poult. Sci. 81(Suppl. 1):75. (Abstr.) 86. Tamim, N. M., and R. Angel. 2002. Effect of calcium level and phytase on phytin phosphorus hydrolysis in broilers. Poult. Sci. 81(Suppl. 1):75. (Abstr.)
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56. Maenz, D. D., and H. L. Classen. 1998. Phytase activity in the small intestinal brush border membrane of the chicken. Poult. Sci. 77:557–563.
Phytic Acid Chemistry: Influence on Phytin-Phosphorus Availability and Phytase Efficacy1 R. Angel,*,2 N. M. Tamim,* T. J. Applegate,† A. S. Dhandu,* and L. E. Ellestad*
Primary Audience: Feed Formulators, Researchers SUMMARY Poultry diets are primarily composed of seed-based ingredients and contain a high proportion of their P in the phytic acid molecule, making this P poorly available. Phytic acid (IP6) is a highly reactive acidic compound that readily binds mineral cations, and in this complexed form is called phytin. The chemical characteristics of IP6 influence exogenous (diet) and intestinal phytase efficacy and the availability of both phytin-P (PP) and any minerals bound to IP6. Research has shown that minimizing IP6-mineral complexes in the digesta of poultry can lead to increased PP availability in the absence of dietary phytase, as well as increased efficacy of dietary phytases. Understanding these binding interactions and how they influence the efficacy of different phytases can provide a valuable tool in choosing when to use phytase and what phytase to use under different situations. This understanding can provide the foundation for developing new methods, as well as minimizing the cost and maximizing efficacy of current methods, to reduce the amount of P excreted by poultry. Key words: phytin-phosphorus, phytase, availability, phosphorus excretion 2002 J. Appl. Poult. Res. 11:471–480
DESCRIPTION OF PROBLEM The accumulation of P in soils and the threat to surface water quality that may result from P losses to waterways, due to runoff or leaching, are major challenges facing the poultry industry today. Worldwide concerns regarding P content in poultry litter have risen to the forefront among legislators, the public, poultry growers, and integrators. In specific regions, recent legislation has been implemented that limits the use of litter 1
application to soil (based partly on soil and litter P content). Legislation in the state of Maryland requires that all poultry feed contain phytase or other feed additives that decrease excreted P. Given limited land for litter application in areas where the greatest concentration of poultry production exist, the poultry industry must find strategies that reduce P in litter with minimal impact on the economics of producing sellable product. Why is P high in relation to other nutrients in poultry excreta? The primary constituents of
Presented as part of the Informal Nutrition Symposium “Making Sense of Scientific Research and Applying It Properly” at the 91st Annual Meeting of the Poultry Science Association, Newark, Delaware, August 11–14, 2002. 2 To whom correspondence should be addressed: [email protected].
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*4131 Animal and Avian Sciences, Department of Animal and Avian Sciences, University of Maryland, College Park, Maryland 20742; and †1151 Lilly Hall, Room 2-114, Department of Animal Science, Purdue University, West Lafayette, Indiana 47907
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DEFINITION OF TERMS Before delving into the chemistry of IP6, it is important to define terms. Myo-inositol 1,2,3,4,5,6-hexakis dihydrogen phosphate (IP6), an organic phosphate, is a phosphorylated cyclic sugar alcohol. In plants, IP6 exists in its anionic form, phytate. In mature seeds, IP6 is found as a complex salt of Ca, Mg, and K, and in some cases it is bound to proteins and starches. This complexed or chelated molecule of IP6 is known as phytin [1]. Confusion exists related to the terms used for the different forms of P. Total P is generally referred to as P and encompasses any and all forms of P in the diet. Available P (aP) refers to the P that is absorbed from the diet by the animal (i.e., feed P minus P in the distal ileum). Retained P refers to the P that stays in the body (i.e., feed P minus excreta P). As mentioned earlier, most of the P in plant ingredients of seed origin is present in the phytin molecule and is referred to as PP. Any P that is
not bound to the phytin molecule is referred to as nonphytin P (nPP). This nPP can be chemically determined by subtracting analyzed PP from analyzed P. A key difference between aP and nPP is that the term aP includes all absorbed forms of P and will include inorganic P (Pi) and organic P (including PP), whereas nPP excludes any PP available to the animal. The 1994 NRC [2] nPP recommendations for broilers are based mostly on research done using book values of aP for different feed ingredients. NRC states that the nPP requirement for chicks from hatch to 3 wk of age remain unchanged from those published in the previous NRC [3], even though the previous (1984) recommendations were given as aP. Thus, it appears that the most recent NRC [2] used aP and nPP interchangeably, yet aP and nPP are not equivalent as per previous definitions.
PHYTIN Phytin is found primarily in seeds [4], with its location within the seed differing among plants. Ninety percent of the phytin in corn is found in the germ portion of the kernel, whereas in wheat and rice most of the phytin is in the aleurone layers of the kernel and the outer bran [5]. In most oilseeds and grain legumes, phytin is associated with protein and concentrated within subcellular inclusions called globoids that are distributed throughout the kernel; however, in soybean seeds, there appears to be no specific location for phytin [4]. Location of phytin within the seed and its chemical associations with other nutrients influence its availability. Phytin constitutes between 1 and 3%, by weight, of many of the cereals and oilseeds used in animal feeds [6]. In seeds, the role of phytin is as follows: 1) a P reserve, 2) an energy store, 3) a competitor for adenosine triphosphate during the rapid biosynthesis of phytin near seed maturity when seed metabolism is inhibited and dormancy is induced, 4) an immobilizer of divalent cations needed for the control of cellular processes and that are released during germination upon the action of intrinsic plant phytases, and 5) a regulator of readily available seed Pi level [7]. The IP6 molecule contains 28.2% P and is the main source of P in seed-based poultry diets. Phytin accounts for approximately 50 to 80% of the P in seed feedstuffs [4]. The importance of
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poultry diets are seed-based ingredients. Most of the P stored in seeds is present as phytin-P (PP) a form poorly available to poultry. The availability of PP varies within and among ingredients. Therefore, P from inorganic sources, such as deflourinated or dicalcium phosphate, is added to poultry diets to meet requirements and results in diets with a much higher total P than that needed by the bird. This excess P, mainly from poorly available PP, is excreted. To solve the challenge of reducing P in poultry litter in the most economical way, it is important to understand the role the P form most prevalent in poultry diet, PP, has in determining phytase efficacy, PP availability to the bird and the amount of P in excreta. Understanding the interactions that occur in the gastrointestinal tract (GIT) between phytic acid (IP6) with other seed, diet, and digesta nutrients and how these interactions affect efficacy of the different phytases can provide a valuable tool in the applied decision-making process of which dietary phytase to use, when to use it, and how much to use. This understanding may explain why an enzyme capable of hydrolyzing P from phytin, present in the intestinal mucosa of poultry appears to have very limited efficacy and how this efficacy could be improved.
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Chemistry of Phytic Acid Phytic acid has 12 replaceable protons or reactive sites. Six are strongly acidic, with pK of 1.5 to 2.0; two are weakly acidic with a pK of approximately 6.0, and four are very weakly acidic with pK of 9.0 to 11.0 [10, 11]. This means that at pH normally encountered in feeds and in the digestive tract, phytin will carry a strong negative charge and is capable of binding di- and trivalent cations such as Zn2+, Cu2+, Ni2+, Co2+, Mn2+, Fe2+, and Ca2+ in very stable complexes [12, 13, 14, 15, 16, 17, 18], thus reducing the availability of these complexed minerals as well as that of the PP to the animal [9]. The order of potency of selected minerals to inhibit PP hydrolysis by phytase at pH 7 was Zn2+ >> Fe2+ > Mn2+ > Fe3+ > Ca2+ > Mg2+ [14]. The ability of the different metal ions to inhibit PP hydrolysis was related to the stability of the complex, the pH of the solution, and the phytate to mineral molar ratio. Despite the fact that the IP6-Ca complex has a lower stability than other IP6-metal ion complexes, it may be dietary Ca that plays the most critical role in minimizing the effectiveness of both exogenous and endogenous phytases. The level of Ca added to poultry diets is eightto 40-fold higher than that of Zn, and the impact of the much higher concentrations of Ca may be more powerful than its lower affinity due to simple mass action. The impact of Ca may be exacerbated by the presence of other cations, as Ca has been implicated in secondary synergistic effects. When two cations are present simultaneously they increase the proportion of the IP6metal complex that precipitates [19].
In addition to complexing with macro- and microminerals, intrinsic in seeds and those added to diets, IP6 may form complexes with proteins and starches. Proteins are able to bind directly with IP6 through electrostatic charges at low pH or through salt bridges formed at high pH, [20]. Starch binding occurs as a result of hydrogen bond formation [21]. The binding of IP6 with protein and starch may also reduce the availability of these nutrients from the diet. Furthermore, the complexing of IP6 with proteins or starches influences the degree of ionization of the complex and the ability of phytin to complex with minerals, potentially changing the efficacy of different phytases. From an applied poultry nutrition standpoint, understanding the interactions of IP6 with other nutrients in feeds and in the digesta is important because the physical state or solubility of IP6mineral complexes in the digesta dictates what the availability of PP will be. If the IP6-mineral complex is precipitated, then any phytases present in the small intestine, regardless of origin (i.e., added to the diet intrinsic in the GIT), will be unable to exert their hydrolytic effect resulting in a greatly reduced phytase efficacy. The increase in pH of the digesta as it moves distally along the GIT causes the PP molecule to be ionized and to more readily form complexes with divalent metal cations such as Zn, Ca, Mg, and Fe [12, 14]. The higher pH result in decreased solubility [16] of the complex and, therefore, decreased efficacy of phytase [14].
PHYTASES Phytases (myo-inositol hexaphosphate hydrolases) are phosphatase enzymes able to catalyze the hydrolysis of phosphate ester bonds. These enzymes are capable of hydrolyzing one or more phosphate groups from IP6 yielding Pi and a series of lower phosphoric esters [22, 23, 24]. Phytases are widely distributed in plants, animals, and microorganisms. The International Union of Biochemists [25] currently acknowledges two classes of phytase enzymes: a 3-phytase (EC 3.1.3.8) and a 6-phytase (EC 3.1.3.26), which initiate the dephosphorylation of IP6 at different positions on the inositol ring and produce different isomers of the lower inositol phosphates. The 3-phytases initiate the dephosphory-
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IP6 as a nutrient is primarily related to its P content [8]. In most cases of plant-based poultry diets, there are adequate levels of P to meet dietary needs, provided that the PP is released from the IP6 molecule as bioavailable Pi. In general, without some dietary modification, a large proportion of the PP is unavailable so that without added dietary Pi most poultry diets do not contain enough aP to meet bird requirements. Phytin is most commonly thought of as an antinutrient [9], because of its ability to complex or bind mineral cations in the seed or diet, rendering these bound cations as well as the PP partially or completely unavailable to the animal.
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pH optima of these enzymes as well as their thermostability [36] and resistance to inactivation by gastrointestinal proteolytic enzymes [32] make them more suitable as feed additives to increase PP availability than relying on plant phytases. Addition of microbial phytase to feed has led to increased PP utilization in turkeys [37, 38, 39] and chickens [38, 40, 41, 42, 43, 44, 45, 46]. The microflora in the GIT of animals are capable of hydrolyzing PP, but the impact of this hydrolysis has not been well documented and poultry research in this area, to the authors’ knowledge, has been minimal. Most of this research has been done in ruminants [8, 47, 48], rats [49], and swine [50]. Intestinal Phytase The first report of the presence of a phytatesplitting enzyme in the GIT was in rats in 1937 [51]. Since then numerous studies have demonstrated that extracts of the intestinal mucosa of ruminant and monogastric species are capable of hydrolyzing PP. Phytase activity has been detected in the small intestinal mucosa of chickens [18, 52, 53, 54, 55, 56, 57]. Even though phytase has been found in all segments of the small intestine, the highest levels have been found in the duodenum [53, 56]. There is controversy among scientists as to whether intestinal phytase activity is merely a manifestation of nonspecific acid or alkaline phosphatase activity, but this controversy is immaterial from a practical perspective. If small intestinal mucosa preparations are shown to be able to release Pi from IP6, it is irrelevant whether this is accomplished by a specific phytase or not. Published literature supports that an enzyme in the intestinal mucosa of poultry is capable of hydrolyzing PP [56, 58, 59]. This enzyme is affected by age of the animal, where higher levels have been found in mature versus young birds [18, 56], as well as genetics [18, 53] and nutrient levels in the diet. In addition to the effects of age and genetics, intestinal phytase activity is influenced by several nutritional factors. Citrate added to rat diets has been shown to decrease in vitro intestinal phytase activity [60, 61] but increase in vivo PP hydrolysis [56, 60]. This increase in PP hydrolysis in vivo may be due to the Ca-chelating ability
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lation of IP6 at the 3 position, yielding 1,2,4,5,6pentakisphosphate and Pi, whereas the 6-phytases initiate the dephosphorylation of IP6 at the 6 position, yielding 1,2,3,4,5-pentakisphosphate and Pi. The 3-phytases do not always completely dephosphorylate IP6, whereas the 6-phytases do [26]. It has been stated that microorganisms normally produce the 3-phytases and 6-phytases are normally found in plants [8, 22]. However, exceptions to this general rule have been reported. For example, an enzyme with 3-phytase activity has been reported in soybeans [27], and an enzyme with 6-phytase activity has been reported in Escherichia coli [28]. Microbial phytases tend to have pH optima in the range of 2 to 6, whereas plant phytases tend to have a pH optimum near 5 [26]. There are four possible sources of phytase that can be found in the digestive tracts of animals: 1) phytase present intrinsically in feed ingredients, 2) exogenous microbial phytases added to the diet, 3) phytases produced by microflora present in the GIT, and 4) intestinal mucosa membrane linked phytase. Phytase present in feed ingredients has been shown to improve PP utilization by monogastric animals [9]. High levels of phytase are found in wheat, rye, triticale, and barley [29, 30], yet there is a large variation of phytase activity within the seeds of the different varieties of these plants [31]. Phytase in feed ingredients may be inactivated by temperature during feed processing [26], by the low pH that occurs in the upper portion of the GIT [32], and by the action of pepsin on gastric secretions [32]. It has been shown that there is a decrease in phytase activity when pelleting temperatures exceed 75°C [33, 34], but this will be dependent on the form and type of the enzyme. The variability and instability of these natural phytases limit the use of plant ingredients as a reliable source of the enzyme in feeds. Many bacteria, yeasts, and fungi produce phytase [9]. Of all the microorganisms studied, phytase production is highest in the fungi Aspergilli with the highest levels of extracellular phytase produced by Aspergillus niger [35]. Aspergilli tend to produce two different phytases, one with pH optima of 5.5 and 2.5 (phyA) and one with a pH optimum of 2.0 (phyB) [26]. The
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PHYTIN PHOSPHORUS USE BY POULTRY Most evidence indicates that poultry are capable of utilizing at least a portion of dietary PP. However, there is wide variation in reported availability. Researchers working with poultry in the mid-1900s concluded that PP was virtually unavailable to chickens when supplied in its natural form or as the purified Ca or Na salt [69, 70, 71]. In later reports [71, 72], chickens were found to have a limited ability to utilize P from plant sources. Broiler chickens were found to inefficiently use PP, and the authors hypothesized that this was due to a minimal quantity of intestinal phytase [47], although intestinal phytase activity was not measured. As mentioned previously, there is evidence demonstrating that an enzyme in the intestinal mucosa can hydrolyze PP [14, 54]. The magnitude of the impact this endogenous phytase has on the hydrolysis of the PP molecule in vivo is still unknown. More recent studies with chickens indicate that PP utilization is variable and that dietary factors including the level of Ca, nPP, P, vitamin D3, and vitamin D3 metabolites, as well as feed processing and feed or ingredient particle size may influence PP hydrolysis in the GIT. Phytin P utilization by chickens is reported to be as low as 0 to 15% [43, 64, 73] and as high as 70 to 75% [41, 65, 74]. Hydrolysis of PP in 4-wkold female broilers ranged from 3 to 42% and depended primarily on dietary Ca level in the diet [63]. Three-week-old broilers were reported to utilize PP, and availability of P from plant feedstuffs varied between 16% in corn to 80% in lupin [75]. Availability of P from purified monosodium phytate was similar to that of feedgrade dicalcium phosphate or a reagent-grade monosodium phosphate, but P from Ca phytate was poorly available [76]. In general, the variation in the availability of PP observed in different research trials could be due to differences in experimental design, research methodologies, diet ingredient and nutrient composition, diet processing, analytical methods, age, species, and breed. Different ingredients have varying amounts of PP [77, 78] and different locations of PP within the plant [4]. Some feed ingredients such as wheat and rye have endogenous phytase [79], which might
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of citrate, making the Ca less available to form insoluble complexes with IP6. There is some indication that the level of IP6 may influence intestinal phytase activity; however, conflicting results have been reported. Neither purified sources of Ca phytate nor Na phytate added to the diet increased intestinal phytase activity in chicks according to one study [18], but another study [58] reported that increasing the level of IP6 in the diet of chicks decreased intestinal phytase activity. Dietary mineral levels influence intestinal phytase activity. High levels of Mg in the diet (0.4 and 0.6%) depress intestinal phytase activity of chicks [59]. Increasing the level of dietary Ca has also been shown to decrease PP hydrolysis in chickens [41, 42, 62, 63, 64, 65]. It has been suggested [12] that a diet with a molar ratio of Ca:IP6 greater than 6:1 leads to formation of insoluble Ca-IP6 complexes that are inaccessible to diet or intestinal phytases. High dietary Ca depresses intestinal phytase activity in chicks [58, 59]. Conversely, others have demonstrated that increased dietary Ca concentrations actually increase intestinal phytase activity in chickens [57]. Decreasing dietary Pi, from 0.48 to 0.16%, has been shown to double intestinal phytase activity in chicks [58, 59]. Conversely, others have reported no difference in the intestinal phytase activity of chickens or hens fed dietary Pi between 0.2 and 0.6% [18]. Altering the vitamin D3 level of the diet also influences the activity of intestinal phytase. An early study found that vitamin D3 added to a diet containing purified Ca phytate increased the utilization of PP by rats but did not increase the activity of intestinal phytase [66]. However, subsequent studies have demonstrated that phytase activity in chicken intestinal extracts is increased slightly as the vitamin D3 level of the diet is increased [58, 67]. The low dietary level of inorganic phosphate (0.25%) used in the early study [66] may have increased intestinal phytase activity so that the influence of vitamin D3 was masked. Recent studies have found mucosal phytase activity in chickens not to be influenced by dietary levels of 1α-hydroxycholecalciferol [68] or 25-hydroxycholecalciferol [57], although both compounds were shown to increase PP hydrolysis in vivo [57, 68].
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size of the soluble IP6-mineral complex will impact the efficacy of the different phytases, and this impact will be specific to each type of phytase [81, 84]. Insoluble IP6-mineral complexes that form at the higher pH, where plant phytases are most active, may be resistant to hydrolysis by phytases of plant [84] and microbial origin [14]. When one looks at commercially available phytases, it is important to know what the pH optima is because that will dictate where the phytase will function along the GIT. For example, a commercially available microbial phytase from Aspergillus niger that has two pH optimas, one at 2.5 and the other at 5.5 [40], would be active in the gastric area where the pH is 2 [81]. At gastric pH, most of the IP6-mineral complex is in solution, making hydrolysis possible. Given the chemical properties of IP6 under GIT conditions it is not surprising that no appreciable PP hydrolysis was observed after the digesta exited the gizzard [84]. In a study [85] where a basal corn-soy diet with no added Ca, Pi, or microminerals and analyzed to contain 0.18% Ca, 0.40% P, and 0.29% PP was fed for 30 h to 20-d-old broiler chicks, PP disappearance (hydrolysis) in the GIT, up to the ileum, was 67.1%. When Ca carbonate was added to the basal diet to supply 0.5% Ca (analyzed total dietary Ca of 0.70%), PP hydrolysis was 18.9%. Thus, PP hydrolysis, from a cornsoy diet, in the GIT increased by a factor of 3.55 in the absence of added Ca. In this work, inclusion of an inorganic micromineral mix at average commercial levels had no effect on PP hydrolysis in the GIT, regardless of the presence or absence of Ca. Use of a commercially available micromineral-protein chelate had no effect on PP hydrolysis in the GIT versus when the inorganic micromineral source was used, and this may have been related to the ionization potential of the micro-mineral chelate in the GIT. The impact of Ca level in the diet on the efficacy of two different commercially available phytases was determined in a second study [86]. As in the first study, not adding Ca to a corn-soy diet resulted in a 2.5-fold increase in PP hydrolysis in the GIT vs. when 0.50% Ca was added. Two phytase sources were tested, a 3- and a 6-phytase, in which the researchers determined pH
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influence the extent of PP availability. The possible reason for the difference in PP availability observed when similar dietary ingredients were used could be related to experimental methodology differences or to the differences in PP level within the same ingredients depending on level of fertilization, growing conditions, and stage of maturity of the seed at harvest [80]. Researchers have demonstrated that as soil available P increases, total P and PP increase in the seed and that the increase in P is primarily due to an increase in PP [80]. The effect of high levels of diet Ca on GIT pH may be partly responsible for some of the deleterious effects that high dietary Ca has on PP hydrolysis [81, 82, 83]. Increasing dietary Ca from 1.07 to 2.53% resulted in higher pH in the crop (4.89 vs. 5.32, respectively) and ileum (6.62 vs. 7.39, respectively) [81]. IP6-mineral complexes are soluble at low pH (less than 3.5) with maximum insolubility occurring between pH 4 and 7 [81], and thus anything that alters intestinal pH will alter the efficacy of phytases and may affect different phytases in different ways. Size of the soluble IP6-mineral complexes also influences the availability of PP and IP6 complexed minerals. Smaller sizes of IP6-mineral complex have greater surface area available for the enzyme to attack, making the complex more easily hydrolyzed. The proportion of larger IP6-mineral complexes in the ileum was greater than in the duodenum, and as dietary Ca increased, the size of the IP6-mineral complexes increased, regardless of region of the intestine [81]. When considering this information, it is important to put it into the context of where in the GIT phytases may be active and where absorption of different nutrients occur. At the pH of the small intestine, not only are most of the IP6-mineral chelates precipitated, but when dietary Ca and P concentrations are increased, the proportion of IP6-mineral complexes precipitated also increases. The solubility of Ca was 16.9% when the Ca and aP were 1.53 and 0.43%, respectively, compared to a solubility of 8.3% when they were 2.26 and 0.83%, respectively [81]. Thus, increased dietary Ca and aP levels reduce the proportion of soluble minerals and thereby further decrease the availability of these minerals and of PP. The
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Potential for Future Applications As mentioned earlier dietary Ca has the highest practical implications because it is present in the highest concentration relative to other metal ions in the diet. Delivery of Ca to the birds in a manner that minimizes IP6-Ca complexing would result in improved phytase efficacy and increased PP availability. This is not a new concept [14, 63, 73], but until now there has been no economic incentive to develop a Ca chelate for animals. Developing a product that delivers Ca to the bird but does not complex with IP6 makes more economic sense given current and expected future environmental regulations. The inhibitory effect of Ca on PP hydrolysis can be prevented by a strong Ca chelator such as EDTA [14]. A concentration of EDTA greater than 5 mM is required to prevent the inhibitory effect of Ca on PP hydrolysis [14]. The mode of action of EDTA may be that it is competing for Ca ions. If a chelator of mineral ions is to improve PP hydrolysis in a diet, the affinity of the chelator would need to be sufficient to competitively bind to Ca, as PP is a stable and strong Ca chelator. The chelator to be developed must be either absorbed as a complex or be able to release Ca close to the absorptive surface in a way that minimizes potential IP6-Ca complexing. In addition, if the complex is absorbed intact, then the chelator must release Ca after the animal absorbs the complex. It is important to consider the potential other interactions of the chelator within the body. Based on the results of some of the work discussed in this review [85, 86], along with
what has been discussed previously in this paper about IP6 chemistry, a baseline of information is starting to emerge that provides some basis for understanding why different phytases act differently in the GIT and how changes in diet or intestinal pH can lead to large changes in enzyme efficacy. Recent work [85, 86] confirms and expands on previous research [14] and points to the possibility of developing new dietary feeding strategies or feed additives that will allow for the delivery of Ca into the bird in a manner that minimizes the formation of IP6-Ca complexes and allows the bird to use PP more efficiently through an increase in efficacy of diet and endogenous phytases.
INFORMATION NEEDED FOR INTERPRETATION OF PHYTASE EFFICACY RESEARCH To minimize the limitations we have as scientists in interpreting published research, key information should be included. Knowledge of the chemical characteristics of a key constituent of seed-based ingredients such as phytin and its interactions with other nutrients in the digest as it moves along the GIT makes this even more important. One of the limitations in interpreting published research is that numerous papers lack the information necessary for comparison with other work as well as for interpretation of the information published. Most work published on phytase use in animal diets does not include information on actual PP content of the diet and relies on book values of PP in the ingredients used. Also, in numerous papers, the amount of phytase added to diets is based on addition of a quantity of commercial product where the addition to the diet is based on the label claim of that product. Few papers have actually analyzed the commercial product to be used. Analytical procedures are available and accurate for analysis of phytase premixes. Few report analyzed levels of the enzyme in the final diets, which is not surprising since the analytical procedure for diet phytase is currently not very accurate. Other key information that often is missing from papers is the final weight, breed, strain, or sex of the animals used in the study and analyzed values for key nutrients being studied, especially those that have an important effect on the nutrient being studied.
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optimas of 3 and 5 for the 3-phytase and between 4 and 4.5 for the 6-phytase. When the 6-phytase was tested, PP hydrolysis in the GIT of broiler chicks was 0.093 and 0.055% per unit of analyzed dietary phytase when the chicks were fed diets containing 0 and 0.50% added Ca from Ca carbonate, respectively, for 30 h. When 3phytase was tested, PP hydrolysis in the GIT was higher per unit of analyzed phytase regardless of Ca level in the diet (0.137 and 0.110% in diets with 0 and 0.50% added Ca, respectively). Phytase level in the diet was formulated based on analyzed premix levels to be 500 U/kg. These diets contained no added Pi but had average commercial levels of inorganic microminerals.
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the response to the additive is beyond the response curve range. Appropriate selection of response criteria is also important. For example, when the treatment being studied has an effect on gain, use of percentage bone ash as the response criteria will lead to predictions that are not correct. How published data is interpreted is ultimately in the hands of the person wishing to use the information. Thus, readers must use their own judgment and should be cognizant of experimental design and the pitfalls inherent in the work. For example when comparing efficacy between studies it is important to note what the source of P used to determine the response curve is. The reason this information is so important is that the availability of the P source will impact measured efficacy and different P sources have different availabilities.
CONCLUSIONS AND APPLICATIONS 1. The strong chelating properties of phytic acid (IP6) present in plant-based diets result in — formation of IP6-mineral complexes that are soluble at proventriculus and gizzard pH but insoluble at the pH of the small intestine. — a large reduction in the efficacy of phytase when the IP6-mineral complexes are present, regardless of source of phytase. — varying sizes of soluble IP6-mineral complexes, with larger complexes reducing phytase efficacy. 2. Calcium, rather than other mineral ions in the diet, is the metal ion most involved in the IP6mineral complexes that reduce PP availability. 3. It is possible to develop feed management tools that maximize innate abilities of birds to use PP and thus alter the need for supplemental Pi and phytase.
REFERENCES AND NOTES 1. Lott, J. N. A. 1984. Accumulation of seed reserves of phosphorus and other minerals. Pages 139–166 in Seed Physiology. Vol. 1. D. R. Murray, ed. Academic Press, Sydney, Australia.
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In the case of phytase studies, the substrate (PP) that the enzyme is working on should be analyzed, as well as P and Ca. Vitamin D formulated values as well as source should also be provided. Precise ingredient description and inclusion levels should be listed. In efficacy studies researchers will sometime use response curves derived based on treatment means rather than based on pen (replicate) measured levels to calculate efficacy. This overestimates the accuracy of the calculated efficacy value and results in the use of an equation that is not correct. Response curves should be determined based on treatment replicate mean information. Also of concern is the prediction of efficacy based on response curves that have poor correlation coefficients. It is also incorrect to assume that the relationships found in the response curve extend beyond the response curve levels studied when
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24. Harland, F. B., and E. R. Morris, 1995. Phytin: A good or a bad food component. Nutr. Res. 15:733–754.
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