Protein–phytate interactions

Phytate is a reactive, polyanionic molecule potentially carrying twelve dissociable protons, with acid dissociation constants ranging from 1·5 to 10⁽Reference Costello, Glonek and Myers¹⁰⁾. The capacity of phytate to interact with protein in cottonseed⁽Reference Jones and Csonka¹¹⁾, yellow peas⁽Reference Mattson¹²⁾ and bean seed⁽Reference Bourdillon¹³⁾ was reported decades ago. As reviewed by Cosgrove⁽Reference Cosgrove¹⁴⁾, Cheryan⁽Reference Cheryan¹⁵⁾ and Anderson⁽Reference Anderson, Finley and Hopkins¹⁶⁾, it is generally accepted that negatively charged phytate molecules form binary protein–phytate complexes with proteins carrying a net positive charge at pH less than their isoelectric point. At pH exceeding their isoelectric point, with proteins carrying a net negative charge, a cationic bridge (usually Ca²⁺) links phytate and protein in ternary complexes. These interactions were demonstrated by Reddy & Salunkhe⁽Reference Reddy and Salunkhe¹⁷⁾ in their investigations of phytate and albumin in black gram (Phaseolus mungo). Protein–phytate interactions were not detected at pH 6·40, which approximates the isoelectric point of black gram albumin. However, binary protein–phytate complex formation was observed at pH 2·80 and ternary complex formation was observed at pH 8·40, which was mediated by divalent cations.

Binary protein–phytate complexes

In the classic description of binary protein–phytate complexes Cosgrove⁽Reference Cosgrove¹⁴⁾ concluded that protein–phytate interactions stem from phytate forming salt-like linkages with the basic amino acid residues of arginine, histidine and lysine at pH levels below protein isoelectric points. Cosgrove⁽Reference Cosgrove¹⁴⁾ suggested that as a result of complex formation, protein molecules become closely packed around the relatively small and highly charged phytate anion leading to the formation of macromolecular aggregations or insoluble coacervates. Drawing on the investigations of Barré & Nguyen-van-Hout⁽Reference Barré and Nguyen-van-Hout¹⁸⁾ into phytate and human serum albumin it appeared that phytate sequentially bound the terminal α-amino and ε-amino groups of lysine, the imidazole group of histidine and, finally, guanido groups of arginine. However, in a second study⁽Reference Barré and Nguyen-van-Hout¹⁹⁾ with avian ovalbumin, the phytate-binding sequence differed to arginine, lysine and finally histidine.

Subsequently, Cheryan⁽Reference Cheryan¹⁵⁾ suggested that the Barré & Nguyen-van-Hout⁽Reference Barré and Nguyen-van-Hout^18, Reference Barré and Nguyen-van-Hout¹⁹⁾ data contain several ambiguities. Similar solubility profiles for phytate and protein across a range of pH values where maximum insolubility coincides at a particular point are considered indicative of binary complex formation. From a series of soya protein studies⁽Reference Fontaine, Pons and Irving^20–Reference De Rham and Jost²³⁾, Cheryan⁽Reference Cheryan¹⁵⁾ concluded that the solubility of phytic acid ‘somewhat parallels’ that of soya protein. More recently, Anderson⁽Reference Anderson, Finley and Hopkins¹⁶⁾ proposed that the extent of protein–phytate interactions is dependent on the number of unhindered cationic groups of the protein and that protein–phytate interactions were highly correlated with basic amino acid residues in several studies⁽Reference Barré and Nguyen-van-Hout^18, Reference Barré and Nguyen-van-Hout^19, Reference Okubo, Myers and Iacobucci^24, Reference Prattley, Stanley and van der Voort²⁵⁾. This suggests that basic amino acid concentrations in proteins are critical to their propensity to be bound by phytate. Lysine monohydrochloride is frequently included in pig and poultry diets and this free basic amino acid has been shown to be bound by phytate from rice pollard⁽Reference Rutherfurd, Edwards, Selle and Cranwell²⁶⁾. Presumably, interactions between phytate and free lysine would reduce the extent of binary complex formation involving intact proteins.

Rajendran & Prakash⁽Reference Rajendran and Prakash²⁷⁾ investigated the kinetics and thermodynamics of sodium phytate and sesame α-globulin interactions. Sesame (Sesamum indicum L.) contains abundant phytate levels, as concentrations of 14·6 g phytate-P/kg or 51·8 g phytate/kg have been recorded in defatted sesame meal⁽Reference De Boland, Garner and O'Dell²⁸⁾. Sesame seed protein has an isoelectric point of 4·5 and a basic amino acid content of approximately 19·1% in which arginine is dominant⁽Reference Rivas, Dench and Caygill²⁹⁾. Specifically, the major sesame protein fraction, α-globulin, has a molecular weight of 2·5 ×  10⁵ Da and contains 18·7% basic amino acids⁽Reference Prakash and Nandii³⁰⁾. Rajendran & Prakash⁽Reference Rajendran and Prakash²⁷⁾ concluded that the interaction between sodium phytate and α-globulin was maximal at pH 2·3 and that complex formation was a biphasic process. In the rapid, first step sodium phytate binds with and changes the conformation of α-globulin. The slower, second step consists of progressive protein–protein associations to form polymers, ultimately resulting in precipitation when a critical mass is exceeded. The interaction requires a minimum sodium phytate:protein molar ratio of 10:1 and the number of α-globulin binding sites for sodium phytate was estimated to range from 11 to 20. Curiously, Rajendran & Prakash⁽Reference Rajendran and Prakash²⁷⁾ did not consider the complement of basic amino acids in sesame α-globulin in their discussion of protein–phytate interactions.

It is accepted that the propensity for proteins to be bound by phytate differs across feedstuffs. Mainly on the basis of reported protein–phytate solubility profiles, Champagne⁽Reference Champagne³¹⁾ concluded that phytate is capable of binding proteins from soya⁽Reference Fontaine, Pons and Irving^20, Reference De Rham and Jost²³⁾, wheat⁽Reference Hill and Tyler¹⁾, rapeseed⁽Reference Gillberg and Tornell³²⁾ and groundnut⁽Reference Fontaine, Pons and Irving²⁰⁾. Conversely, Champagne⁽Reference Champagne³¹⁾ stated that proteins in cottonseed meal⁽Reference Fontaine, Pons and Irving²⁰⁾, maize germ⁽Reference O'Dell and de Boland³³⁾, sesame meal⁽Reference O'Dell and de Boland³³⁾ and rice bran⁽Reference Champagne, Rao and Liuzzo³⁴⁾ are not bound by phytate.

However, anomalies do appear to exist. The solubility profiles for total P and protein generated by Fontaine et al. ⁽Reference Fontaine, Pons and Irving²⁰⁾ indicate that protein–phytate complexes are more likely to occur in groundnut meal and soyabean protein than cottonseed. However, cottonseed meal (19·7%)⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾ has a higher proportion of basic amino acids than groundnut meal (14·9%)⁽Reference Ayres, Branscomb and Rogers³⁶⁾ and soyabean meal (16·6%)⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾. Thus despite the relatively rich complement of basic amino acids in cottonseed meal, which contains an abundance of arginine, the propensity for phytate to bind cottonseed protein is considered to be less than either soya or groundnut proteins.

Recently, Kies et al. ⁽Reference Kies, de Jonge and Kemme³⁷⁾ assessed protein–phytate interactions across a range of feedstuffs. The reductions in protein solubility generated by sodium phytate under in vitro conditions are shown in Table 1. At pH 2, phytate-induced reductions in protein solubility were substantial for casein (99%), soyabean meal (89%), sunflower-seed meal (84%) and maize (72%), intermediate for rapeseed meal (37%) and marginal for rice bran (6%). Importantly, at pH > 2, the effect of sodium phytate on protein solubility was diminished. The phytate-induced reductions in protein solubility were attributed to binary complex formation; however, it is evident from the tabulated values that there is no relationship between the basic amino acid content of test proteins and solubility reductions at pH 2. For example, rice bran protein has a relatively high proportion of basic amino acids but its solubility was barely altered by sodium phytate.

Table 1

Impact of pH on phytate-induced protein solubility reductions (%) of feedstuffs across a range of pH values in relation to their isoelectric point of protein and basic amino acid profile*

* Adapted from Kies et al. ⁽Reference Kies, de Jonge and Kemme³⁷⁾.

† See Table 3.

‡ Casein, Vickery & White⁽Reference Vickery and White¹⁴⁵⁾; others, Ravindran et al. ⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾.

It is noteworthy that O'Dell & de Boland⁽Reference O'Dell and de Boland³³⁾ detected protein–phytate interactions in soya flakes but not in maize germ or sesame meal. In contrast Kies et al. ⁽Reference Kies, de Jonge and Kemme³⁷⁾ reported protein–phytate interactions in maize at pH 2 and Rajendran & Prakash⁽Reference Rajendran and Prakash²⁷⁾ reported interactions between sesame α-globulin and phytate that peaked at pH 2·3. However, O'Dell & de Boland⁽Reference O'Dell and de Boland³³⁾ conducted their studies at pH levels of 4·4 and 9·0, so it appears that pH differences in the in vitro systems account for the apparent discrepancies in outcomes and emphasise the importance of pH to protein–phytate interactions. Further, the apparent limited capacity of phytate to bind protein feedstuffs such as cottonseed meal and rice bran is often dismissed on the grounds that the basic amino acid residues are inaccessible to phytate. However, basic amino acids are polar and hydrophilic and are normally found on the outer surface of a protein, particularly in the case of arginine⁽Reference Argos^38, Reference Janin, Miller and Chothia³⁹⁾. Given this orientation, the likelihood is that the access of phytate to basic amino acids is largely unimpeded and the ‘inaccessibility’ argument may not be sound.

Numerous factors influence the intensity of binary protein–phytate interactions in addition to pH. Dietary Ca is one example and Ca has a broad impact on the phytate–phytase axis in pig and poultry nutrition⁽Reference Selle, Cowieson and Ravindran⁴⁰⁾. Ca is largely derived from limestone, which has a very high acid-binding capacity⁽Reference Lawlor, Lynch and Caffrey⁴¹⁾ and increasing dietary Ca levels tend to elevate gut pH. As discussed previously⁽Reference Selle, Cowieson and Ravindran⁴⁰⁾, in the Ravindran et al. ⁽Reference Ravindran, Cabahug and Ravindran⁴²⁾ study, seemingly small increases in analysed dietary Ca concentrations relative to both protein and phytate noticeably depressed amino acid digestibility responses to A. niger-phytase. That Ca can interact with phytate is established but Ca may also interact with protein including soya protein⁽Reference Scilingo and Anon⁴³⁾. Okuba et al. ⁽Reference Okubo, Myers and Iacobucci^24, Reference Okubo, Myers and Iacobucci^44, Reference Okubo, Iacobucci and Myers⁴⁵⁾ investigated interactions between phytate and glycinin, the major soya protein. At pH 3, sufficient Ca was able to dissociate glycinin–phytate complexes, which was attributed to Ca competing with glycinin for access to negatively charged phytate molecules. The potential capacity of Ca to disrupt binary protein–phytate interactions may explain the diminished amino acid digestibility responses to supplemental phytase at higher Ca concentrations in the Ravindran et al. ⁽Reference Ravindran, Cabahug and Ravindran⁴²⁾ study.

According to the Cosgrove⁽Reference Cosgrove¹⁴⁾ and Rajendran & Prakash⁽Reference Rajendran and Prakash²⁷⁾ descriptions of binary complexes, the likelihood is that phytate is ‘protected’ by a shield of aggregated protein once complex formation takes place and would be less susceptible to hydrolysis by exogenous phytase. Thus phytase essentially prevents de novo complex formation via the prior hydrolysis of phytate and binary complexes may be an important limiting factor on the extent of enzymic degradation of phytate.

Ternary protein–phytate complexes

Ternary protein–phytate complexes are formed de novo in the small intestine where the major components are linked via a cationic bridge, usually Ca²⁺, the most prevalent divalent cation in digesta. More attention has been paid to the impact of ternary protein–phytate complexes on mineral bioavailability than protein/amino acid utilisation and this appears to have led to a consensus that ternary complexes are not important in respect of phytate reducing protein availability.

Nosworthy & Caldwell⁽Reference Nosworthy and Caldwell^46, Reference Nosworthy and Caldwell⁴⁷⁾ reported precipitation of soya glycinin, phytate and Zn at pH 6·2 where 1 mol of glycinin was involved in a ternary complex with 7 mol of phytate and 39 mol of Zn. Thus phytate has a tremendous capacity to bind protein in ternary complexes under in vitro conditions. The affinity of phytate for Zn is established and the precipitation of Zn in ternary complexes may both reduce the activity of Zn-dependent proteases⁽Reference Jongeneel, Bouvier and Bairoch⁴⁸⁾ and compromise the integrity of the immune system⁽Reference Fraker, King and Laakko⁴⁹⁾.

The negative impact of phytate on Zn availability⁽Reference Wise^4, Reference Zhou, Fordyce and Raboy⁵⁰⁾ has been seminal to the appreciation of protein–phytate interactions, particularly in relation to soya protein. Phytate has the capacity to bind Zn in both mineral–phytate and ternary protein–phytate complexes. This is reflected in pigs, where phytate is a key aetiological factor in parakeratosis, a manifestation of Zn deficiency⁽Reference Oberleas, Muhrer and O'Dell⁵¹⁾. As demonstrated by Cranwell & Liebman⁽Reference Cranwell and Liebman⁵²⁾, the phytate content of soyabean, rather than the fibre content, reduces the bioavailability of Zn in humans. For this reason, the preparation of soya protein isolates and concentrates with reduced phytate contents to avoid Zn deficiencies in human infants has been a goal for decades⁽Reference McKinney, Sollars and Setzkorn⁵³⁾. As discussed by Erdman⁽Reference Erdman⁵⁴⁾, protein concentrates with reduced phytate contents can be prepared by exploiting the capacity of phytate to bind protein, which has been demonstrated in soya⁽Reference Erdman, Weingartner and Mustakas⁵⁵⁾ and rapeseed⁽Reference Tzeng, Diosady and Rubin⁵⁶⁾, and the fate of phytate following the preparation of soya protein concentrates/isolates has been reviewed⁽Reference Deak and Johnson⁵⁷⁾.

In relation to ternary complexes and protein availability, Champagne et al. ⁽Reference Champagne, Fisher and Hinojosa⁵⁸⁾ suggested that the protein moiety of ternary complexes is comprised of either amino acids or small peptides and it then follows that the amount of protein bound in ternary complexes in the small intestine may not be sufficient to compromise amino acid digestibility⁽Reference Selle, Ravindran and Caldwell⁹⁾. However, the validity of this interpretation depends on the molecular weights of protein present in small-intestinal digesta. In one of a series of studies, Montagne et al. ⁽Reference Montagne, Crevieu-Gabriel and Toullec⁵⁹⁾ determined the molecular weights of protein along the small intestine of pre-ruminant calves. In diets in which either a soya protein concentrate or a soya protein isolate partially replaced skimmed milk powder, an average of nearly 60% of protein present in ileal digesta had a molecular weight in excess of 20 000 Da (Table 2). Protein fractions of this magnitude were even more dominant in the duodenum and jejunum as there is a declining gradient in the size of proteins along the gut. Consequently, it appears that sufficient protein may be bound as ternary protein–phytate complexes in the small intestine to disrupt protein digestion and amino acid absorption, and the importance of ternary complexes may have been dismissed prematurely.

Table 2

Distribution of protein and peptide fractions in ileal digesta from three diets according to molecular mass as a percentage of total crude protein*

* Adapted from Montagne et al. ⁽Reference Montagne, Crevieu-Gabriel and Toullec⁵⁹⁾.

Of relevance is that Montagne et al. ⁽Reference Montagne, Salgado and Toullec⁶⁰⁾ reported that the partial substitution of soya protein concentrate for skimmed milk powder increased alkaline phosphatase activity in the duodenum and jejunum by approximately 33%, which was more pronounced with soya protein isolate. Phytate was probably present in both soya preparations⁽Reference Deak and Johnson⁵⁷⁾ and may have triggered these increases in alkaline phosphatase activity. Therefore, it is noteworthy that Montagne et al. ⁽Reference Montagne, Toullec and Formal⁶¹⁾ found that both soya preparations substantially increased the flow of mucin protein in the duodenum, jejunum and ileum in pre-ruminant calves by approximately 70–90%. Moreover, Montagne et al. ⁽Reference Montagne, Toullec and Lalles⁶²⁾ determined the effect of these partial substitutions on the apparent digestibility of cystine, lysine and threonine at the jejunal and ileal levels. With the soya protein concentrate, reductions in ileal digestibility of cystine (4·6%), lysine (4·1%) and threonine (7·2%) were modest in comparison with reductions in jejunal digestibility for cystine (49·4%), lysine (19·0%) and threonine (23·3%). The marked reduction in the jejunal digestibility of cystine may reflect its involvement in disulfide linkages within soya protein. Thus, relative to the control diet, soya protein concentrate depressed amino acid digestibility by an average of 30·6% at the level of the jejunum and by 5·3% at the ileal level. Thus the soya protein substitution both depressed and delayed small-intestinal uptakes of the three amino acids assessed and it is possible that the residual phytate content in soya concentrate contributed to this ‘distal shift’ in the site of amino acids absorption as a result of ternary complex formation.

Isoelectric points of protein

The pH along the gastrointestinal tract, relative to the isoelectric points of protein, should be pivotal to the integrity of binary protein–phytate complexes. Shafey et al. ⁽Reference Shafey, McDonald and Dingle⁶⁷⁾ recorded pH values of 4·89 in the crop, 1·98 in the proventriculus, 3·14 in the gizzard, 5·53 in the duodenum, 6·06 in the jejunum, 6·62 in the ileum and 6·48 in the caecum in broiler chickens offered diets containing 10·7 g Ca/kg. Interestingly, Engberg et al. ⁽Reference Engberg, Hedemann and Jensen⁶⁸⁾ found that pelleting broiler diets reduced intestinal pH, and they recorded average pH values of 6·01 in the duodenum, 6·10 in the jejunum and 6·94 in the ileum of chickens. In 32-d-old pigs offered maize–soya diets containing 8 g Ca/kg, Li et al. ⁽Reference Li, Yi and Yin⁶⁹⁾ reported digesta pH of 3·27 in the stomach, 5·72 in the duodenum, 5·98 in the jejunum and 6·94 in the ileum.

Clearly, there is a substantial increase in pH as digesta transits from the stomach or gizzard into the duodenum, and binary protein–phytate complexes dissociate should this increase exceed the protein isoelectric point. Soyabean meal is the dominant protein source in pig and poultry diets and solvent-extracted soya protein has an isoelectric point of pH 4·1⁽Reference Monaghan-Watts⁷⁰⁾. Soya protein–phytate complexes will dissociate once digesta transits into the relatively alkaline pH of the duodenum and their ephemeral nature may limit their nutritional importance. However, structural changes to soya protein induced by its aggregation with phytate may still reduce its digestibility in the small intestine in its dissociated state.

Csonka et al. ⁽Reference Csonka, Murphy and Jones⁷¹⁾ determined the isoelectric point of a large number of proteins and a selection of these values is tabulated (Table 3). It is evident that the isoelectric point of proteins in cereal grains (5·90–6·45) is higher than proteins in oilseed meals (4·70–5·50); for example, wheat gliadin has an isoelectric point of pH 6·45 as opposed to pH 4·70 for soya glycinin. This comparison suggests that while binary complexes in soyabean, rapeseed and cottonseed meals will dissociate in the small intestine, this may not be the case in wheat, maize and sorghum. Consequently, phytate complexes with proteins from cereal grains may persist in the small intestine; this, as discussed later, may have interesting implications for starch digestibility.

Table 3

Isoelectric points of selected protein sources*

* Adapted from Csonka et al. ⁽Reference Csonka, Murphy and Jones⁷¹⁾.

† Vioque et al. ⁽Reference Vioque, Sanchez-Vioque and Clemente¹⁴⁶⁾.

‡ Bau et al. ⁽Reference Bau, Mohtadi-Ni and Mejean¹⁴⁷⁾.

Impact of phytase on amino acid digestibility in individual feedstuffs

Ravindran et al. ⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾ and Rutherfurd et al. ⁽Reference Rutherfurd, Chung and Moughan⁷²⁾ reported that exogenous phytases increased the apparent ileal digestibility (AID) and true ileal digestibility (TID) of amino acids across a range of individual feedstuffs in broiler chickens, as shown in Table 4. In the first study, 1200 FTU A. niger-phytase/kg increased the AID of fourteen amino acids by averages of 2·67% in rapeseed meal, 3·40% in maize, 3·66% in wheat middlings, 4·16% in soyabean meal, 4·64% in sunflower-seed meal, 4·82% in cottonseed meal, 6·60% in sorghum, 7·52% in rice polishings and 9·26% in wheat. These data were largely confirmed in the second study where 750 U Peniophora lycii-phytase/kg increased TID coefficients of sixteen amino acids by averages of 3·90% in maize, 5·30% in rice bran, 6·39% in soyabean meal, 9·31% in rapeseed meal and 12·94% in wheat. Phytase responses in maize, wheat, soyabean meal and rice bran were similar in both studies and both research groups reported a substantial, three-fold difference in the magnitude of the phytase response between wheat and maize. The exception was rapeseed meal where different oil extraction processes may have contributed to the discrepancies in both amino acid digestibility and the magnitude of the phytase response.

Table 4

Impacts of 1200 phytase units Aspergillus niger-phytase/kg on the mean apparent ileal digestibility (AID) coefficient of fourteen amino acids⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾ and 750 U Peniophora lycii-phytase/kg on the mean true ileal digestibility (TID) coefficient of sixteen amino acids⁽Reference Rutherfurd, Chung and Moughan⁷²⁾ in individual feedstuffs

It is instructive to consider the results of the study by Ravindran et al. ⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾ more closely (Table 5). The quantities of ileal digestible amino acids generated by phytase were not related to the amino acid composition of maize (P>0·85). In contrast, there were significant correlations (P < 0·001) for rapeseed meal, sunflower-seed meal, soyabean meal, wheat, cottonseed meal and sorghum. To illustrate this discrepancy, the relationships between the amino acid composition of maize and sorghum and the amino acids released by phytase are shown in Fig. 1. Moreover, for the last six feedstuffs mentioned, there was a significant relationship (r 0·913; P < 0·015) between the isoelectric points of protein and the magnitude of phytase responses. This may indicate that proteins with relatively high isoelectric points are more likely to be responsive to phytase because their binary protein–phytate complexes are more intact along the small intestine. It should be noted that had maize been included in this exercise the relationship would not be significant; however, maize responded very differently to phytase in comparison with the other six feedstuffs in this study.

Table 5

Data derived from the study by Ravindran et al. ⁽Reference Ravindran, Cabahug and Ravindran³⁵⁾ which determined the effect of Aspergillus niger-phytase in broiler chickens on the apparent ileal digestibility of fourteen amino acids in individual feedstuffs

Fig. 1

Relationships between amino acid composition and amount of ileal digestible amino acids released by phytase in (a) maize (R ² 0·003) and (b) sorghum (R ² 0·987) (adapted from Ravindran et al.(Reference Ravindran, Cabahug and Ravindran³⁵)).

Implications of binary protein–phytate complexes

Several research groups have concluded that complexed protein is refractory to pepsin digestion under in vitro conditions⁽Reference Barré and Nguyen-van-Hout^18, Reference Camus and Laporte^73–Reference Knuckles, Kuzmicky and Gumbmann⁷⁶⁾ and this may also apply to phytate-induced aggregations of protein under the alternative hypothesis that phytate acts as a kosmotropic anion. The key study by Vaintraub & Bulmaga⁽Reference Vaintraub and Bulmaga⁷⁷⁾ found that phytate reduced pepsin hydrolysis of bovine serum albumin by approximately 90% and Hb, casein and 11S soya protein by 65%. These maximal reductions were observed at pH 2–3 but reductions were not evident at pH 4·0–4·5; this indicates a narrow pH band for protein–phytate interactions. These reductions in pepsin hydrolysis were attributed to phytate binding with the substrate and rendering it refractory to digestion, presumably by phytate-induced alterations to protein structure and solubility.

Given this is the case, phytate has the potential to interfere with the initiation of protein digestion. Moreover, peptides generated by pepsin are regulators of the protein digestive processes⁽Reference Krehbiel, Matthews and D'Mello⁷⁸⁾, which suggests that this function may be compromised as well. However, if phytate complexes sufficient protein, rendering it refractory to pepsin digestion, then this could trigger gastric hypersecretions of pepsin and hydrochloric acid (HCl) as a compensatory mechanism. Condensed tannins have the capacity to bind protein, so it is relevant that repeated administrations of tannic acid to rats have been shown to increase gastric secretions of pepsin and HCl by approximately 60%⁽Reference Mitjavila, de Saint Blanquat and Derache⁷⁹⁾. Further indirect support of the concept of pepsin and HCl hypersecretion is provided by Decuypere et al. ⁽Reference Decuypere, Knockaert and Henderickx⁸⁰⁾. These workers compared inclusions of soluble or insoluble soya isolates at 140 g/kg in diets for weaner pigs, and the in vitro pepsin digestibility of the soluble soya isolate was superior to the insoluble isolate. Higher peak pepsin activities were observed in pigs offered the insoluble isolate and the differences were significant at 135, 150 and 165 min post-feeding and pepsin activity was approximately 64% greater in pigs offered the insoluble soya isolate. Thus it is plausible that the presence of phytate-induced protein aggregations in the stomach that are refractory to pepsin digestion would promote gastric hypersecretion.

Both pepsin and HCl are described as endogenous aggressors⁽Reference Allen and Flemström⁸¹⁾ and their hypersecretion would be countered by protective outputs of mucin and sodium bicarbonate (NaHCO₃). For example, increased gastric mucin secretions in response to pepsin infusion have been demonstrated in rats⁽Reference Munster, Bagshaw and Wilson⁸²⁾. Importantly, as first reported by Cowieson et al. ⁽Reference Cowieson, Acamovic and Bedford⁸³⁾, phytate increases mucin and Na excretion in broilers, which is ameliorated by phytase. It is possible that Na was secreted into the gut lumen as NaHCO₃ to buffer excess HCl. Also in broiler chickens, Onyango et al. ⁽Reference Onyango, Asem and Adeola⁸⁴⁾ reported that free phytic acid increased excretion of crude mucin by 50% but a Mg, K salt of phytic acid (Mg-K-phytate) increased crude mucin excretion by 162%. Mg-K-phytate is more akin to phytate naturally present in feedstuffs and it noticeably increased mucin secretions in poultry, and an Escherichia coli-derived phytase tended to stem this increase in mucin output.

Lien et al. ⁽Reference Lien, Sauer and He⁸⁵⁾ reviewed the dietary influences on mucin secretion in the digestive tract of single-stomached animals where phytate was not specifically identified as an influential factor but considerable attention was paid to soluble and insoluble fibre. For example, Satchithanandam et al. ⁽Reference Satchithanandam, Jahangeer and Cassidy⁸⁶⁾ reported that inclusions of wheat bran at 100 and 200 g/kg in a fibre-free diet for rats increased mucin levels in the small-intestinal lumen by approximately 200%. However, wheat bran and similar by-products are rich in phytate, as an average phytate content of 24·8 g/kg was recorded in seven samples⁽Reference Selle, Walker and Bryden⁸⁷⁾. Thus, there is the possibility that the large increase in mucin secretion recorded in this study was at least partially due to the phytate content of wheat bran in addition to its fibre content. Interestingly, Onyango et al. ⁽Reference Onyango, Madsen and Gendler⁸⁸⁾ showed that phytate increased mucin gene expression (Muc1, Muc2) in jejunal and colonic mucosa in mice. Consequently, if phytate increases mucin secretion, as a response to hypersecretion of pepsin and HCl and/or direct ‘irritation’ of the gut mucosa, this has clear implications for the flow of endogenous amino acids because porcine mucin has a protein component of 343 g/kg and essentially remains undigested in the small intestine. Mucin protein contains little methionine and histidine but an abundance of threonine, proline and serine⁽Reference Lien, Sauer and Fenton⁸⁹⁾.

The impacts of phytate and phytase on the endogenous amino acid flows in broiler chickens have been investigated and reviewed⁽Reference Cowieson and Ravindran^90–Reference Cowieson, Bedford and Selle⁹²⁾. Cowieson & Ravindran⁽Reference Cowieson and Ravindran⁹⁰⁾ found that increasing the phytate content of the diet from 8·5 to 11·5 and 14·5 g/kg generally increased endogenous amino acid flows at the ileal level. Overall, the increase from 8·5 to 11·5 g phytate/kg significantly increased (P < 0·001) the flow of seventeen amino acids by 27·2% (19 393 v. 15 247 mg/kg DM intake) and 500 FTU E. coli-phytase/kg reduced (P < 0·001) the flow by 20·0% (15 459 v. 19 327 mg/kg DM intake). Moreover, there was a positive correlation (r 0·762; P < 0·005) between the phytase-induced reductions in endogenous amino acid flows and the amino acid profile of mucin. The results followed a similar pattern in a second study⁽Reference Cowieson, Ravindran and Selle⁹¹⁾ and, again, there was a positive correlation (r 0·556; P < 0·03) between the reductions in endogenous amino acid flows generated by phytase and the mucin amino acid profile. The implication of the significant regressions in both experiments is that phytate was exacerbating, and phytase attenuating, endogenous amino acid flows by making an impact on mucin secretions. However, the extent to which the protein component of mucin contributes to total endogenous protein/amino acids in the terminal ileum of poultry requires clarification, as it has been suggested that the mucin contribution is 19% in calves and 11% in pigs⁽Reference Montagne, Piel and Lalles⁹³⁾.

Bohak⁽Reference Bohak⁹⁴⁾ determined the amino acid composition of pepsin and pepsinogen in chickens. Pepsin has an unusual amino acid profile in that it contains a paucity of basic amino acids but considerable quantities of aspartic acid, serine, glycine and threonine. It is instructive to compare the pepsin amino acid profile with the variations in endogenous amino acid flows induced by phytate and phytase in the two studies⁽Reference Cowieson and Ravindran^90, Reference Cowieson, Ravindran and Selle⁹¹⁾ discussed. When the data are combined, there are significant correlations between phytate-induced increases (r 0·669; P < 0·001) and phytase-induced decreases (r 0·635; P < 0·001) in endogenous amino acid flows and the amino acid composition of pepsin. These significant relationships support the thesis that phytate increases the secretion of pepsin and mucin; both are potential sources of endogenous amino acids.

However, phytate may trigger extra-secretions of endogenous enzymes other than pepsin, by modifying substrates and rendering them less susceptible to hydrolysis, which would increase endogenous amino acids flows. Phytate is a potent inhibitor of α-amylase in vitro ⁽Reference Desphande and Cheryan⁹⁵⁾, which may have in vivo relevance in pigs and poultry. As discussed later, it has been reported⁽Reference Sultan, Gan and Li^96, Reference Sultan, Gan and Li⁹⁷⁾ that phytase increases the digestibility of sorghum starch, which implies that phytate has a negative effect. Phytate may interact with starch per se or with kafirin and glutelin, two proteins that are physically associated with starch granules in sorghum endosperm, and these interactions may modify the substrate leading to compensatory increases in outputs of α-amylase in order to digest starch.

Phytate has been shown to depress total tract digestibility of lipids in rats by 7·03%, from 0·910 to 0·846⁽Reference Nyman and Björk⁹⁸⁾; moreover, it has been reported that phytase increased the digestibility of fat and fatty acids in broilers⁽Reference Zaefarian, Romero and Ravindran⁹⁹⁾. As a main effect, 500 FTU E. coli-phytase/kg increased the ileal digestibility of fat in maize and wheat by 4·16% (0·827 v. 0·794; P < 0·001) and significantly increased the digestibility of specific fatty acids by up to 5·7%. Also, Liu et al. ⁽Reference Liu, Ru, Wang and Xu¹⁰⁰⁾ reported that 1000 FTU E. coli-phytase/kg enhanced crude fat ileal digestibility by 5·64% (0·862 v. 0·816) in maize–soya broiler diets. These positive responses may have stemmed from the involvement of phytate, Ca and lipids in the de novo formation of metallic soaps in the gut lumen⁽Reference Ravindran, Cabahug and Ravindran⁴²⁾. Again, it is possible that phytate triggers compensatory increases in lipase outputs, which would contribute to endogenous amino acid flows.

Cowieson et al. ⁽Reference Cowieson, Acamovic and Bedford⁸³⁾ reported that phytate increases, and phytase decreases, Na excretion in broilers on a total-tract basis. At the level of the ileum the impacts are more pronounced, as Ravindran et al. ⁽Reference Ravindran, Morel and Partridge¹⁰¹⁾ found that phytate decreased ileal Na digestibility (–0·38 v. − 0·24; P < 0·05) but phytase had a corresponding positive effect (–0·18 v. − 0·52; P < 0·001). Also, in broiler diets containing 11·0 g phytate/kg, 500 FTU E. coli-phytase/kg increased coefficients of ileal Na digestibility from − 0·52 to − 0·04⁽Reference Selle, Partridge and Ravindran¹⁰²⁾. Thus phytate has the capacity to drag Na into the small-intestinal lumen and this is counteracted by phytase; this transition of Na into the gut lumen may be as NaHCO₃ to buffer HCl hypersecretions

Interactions between phytate and starch

Rickard & Thompson⁽Reference Rickard, Thompson and Shahidi¹⁰³⁾ nominated several mechanisms by which phytate may negatively influence starch digestion. Phytate may inhibit amylase activity either directly or via chelation of Ca, which is a requisite cofactor for amylase. Additionally, phytate may interact with starch directly or indirectly by binding proteins that are closely associated with starch granules. In a study of covalently bound P in starch granules Blennow et al. ⁽Reference Blennow, Bay-Smidt and Olsen¹⁰⁴⁾ noted that considerable quantities of phytate in sorghum starch precluded analyses of glucose 3-phosphate and glucose 6-phosphate, which was not the case with potato and cassava starches. Thus, there is the inference that phytate-P also could be covalently bound to glucose in starch polymers and that starch–phytate complexes may depress energy utilisation. Theoretically, phytate may bind starch indirectly via interacting with proteins that are closely associated with starch. These interactions may be as binary complexes that persist in the small intestine due to the relatively high isoelectric points of protein in cereals, and phytate may bind starch-associated proteins in ternary protein–phytate complexes as well. Both mechanisms are consistent with the suggestion of Thompson⁽Reference Thompson^105, Reference Thompson and Graf¹⁰⁶⁾ that phytate can indirectly bind starch.

Starch granules are enmeshed in a protein matrix in the endosperm of cereal grains⁽Reference Rooney and Pflugfelder¹⁰⁷⁾. This physical proximity would facilitate starch–protein interactions, and strong electrostatic attractions have been reported between potato starch and casein⁽Reference Takeuchi¹⁰⁸⁾. Anderson et al. ⁽Reference Anderson, Levine and Levitt¹⁰⁹⁾ measured breath H₂ levels in human subjects to assess wheat carbohydrate absorption and concluded that an appreciable proportion of starch was not absorbed, which was attributed to interactions between the starch and protein moieties in wheat flour. Subsequently, it was proposed that starch–protein interactions in food influence starch digestibility and the glycaemic response⁽Reference Thorne, Thompson and Jenkins¹¹⁰⁾. It was reported⁽Reference Jenkins, Thorne and Wolever¹¹¹⁾ that removal of the protein composite gluten (gliandin and glutelin) from wheat flour significantly increased in vitro starch digestibility. Also, gluten removal increased glycaemic responses in vivo and increased starch absorption on the basis of breath H₂ measurements. The presence of gluten in unprocessed wheat flour had the opposite effects and these decreases in glycaemic responses and starch absorption were attributed to starch–protein interactions.

As reviewed by Baldwin⁽Reference Baldwin¹¹²⁾, proteins that are located in and on starch granules are described as starch granule-associated proteins (SGAP). Interactions between wheat starch granules and soya proteins were investigated⁽Reference Ryan and Brewer¹¹³⁾ where protein binding was reduced by the removal of SGAP from the surface of starch granules. It was proposed that SGAP mediates the binding of exogenous protein to starch granules. Starch–protein interactions occur, including adsorption of wheat proteins by potato starch⁽Reference Eliasson and Tjerneld¹¹⁴⁾, so it is possible that phytate may indirectly complex starch by binding SGAP. Therefore, it is relevant that Camden et al. ⁽Reference Camden, Morel and Thomas¹¹⁵⁾ reported that 250 U Bacillus subtilis phytase/kg enhanced ileal starch digestibility by 1·87% (0·982 v. 0·964) in broiler chickens offered maize–soya diets.

Starch granules are intimately associated with both protein bodies, which are composed of kafirin, and the glutelin protein matrix in sorghum endosperm. Sorghum differs from other cereal grains in that its nutritive value is vulnerable to ‘wet-cooking’, which is believed to be a consequence of disulfide linkage formation, particularly in kafirin⁽Reference Selle, Cadogan and Li¹¹⁶⁾. Consequently, starch could interact with kafirin and/or glutelin, which may also involve phytate, as sorghum contains relatively high phytate concentrations⁽Reference Selle, Walker and Bryden⁸⁷⁾. Interestingly, Sultan et al. ⁽Reference Sultan, Gan and Li^96, Reference Sultan, Gan and Li⁹⁷⁾ investigated the effects of phytase in broilers offered diets containing sorghum at 918 g/kg. Phytase substantially increased the ileal digestibility coefficients of starch by 14·7% (0·86 v. 0·75) and crude protein by 8·3% (0·78 v. 0·72) at 21 d post-hatch. In the second study, phytase increased starch digestibility in the jejunum by 9·0% (0·607 v. 0·557), upper ileum by 4·5% (0·878 v. 0·840) and lower ileum by 2·4% (0·927 v. 0·905) at 42 d post-hatch. The digestibility of sorghum starch increases as it transits the small intestine, with corresponding reductions in responses to phytase. Axiomatically, the Sultan et al. ⁽Reference Sultan, Gan and Li^96, Reference Sultan, Gan and Li⁹⁷⁾ studies suggest that phytate depresses starch digestibility and delays its absorption from the small intestine, although the fermentative activity of the gut microflora would contribute to starch disappearance along the small intestine.

Phytase amino acid digestibility assays

The impact of phytate and phytase on protein utilisation has been mainly assessed by amino acid digestibility assays in pigs and poultry. One noteworthy exception is the Ketaren et al. ⁽Reference Ketaren, Batterham and Dettmann¹¹⁷⁾ study in grower pigs in which phytase significantly increased protein deposition and retention; however, these positive outcomes may have been in part a consequence of enhanced P availability. As reviewed by Selle & Ravindran⁽Reference Selle and Ravindran^118, Reference Selle and Ravindran¹¹⁹⁾, the effects of exogenous phytases on the AID of amino acids in pigs and poultry have been assessed in approximately fifty assays. These assays indicate that the influence of phytase is inconsistent, and the impact is marginal in the majority of reported studies, particularly in pigs. However, the straightforward adjudication that phytase does not improve ileal digestibility of amino acids in pigs and poultry seems justified but it may be misleading.

Phytase amino acid digestibility assays in poultry

In phytase amino acid digestibility assays involving complete broiler diets, there is a consistent pattern in which more robust responses have been recorded in studies using acid-insoluble ash or titanium oxide as dietary markers in comparison with chromic oxide⁽Reference Selle, Ravindran and Bryden¹²⁰⁾. If inert marker selection in phytase assays is a confounding factor, it has not been resolved. However, its potential importance is apparent from a comparison of three assays in which the one enzyme at the same inclusion rate (1000 FTU E. coli-phytase/kg) was added to maize–soya diets but different dietary markers were used. In the two chromic oxide assays⁽Reference Dilger, Onyango and Sands^121, Reference Onyango, Bedford and Adeola¹²²⁾ the median phytase response across eighteen assessed amino acids was 2·40 and 1·06%, respectively. Threonine is almost invariably the most phytase-responsive essential amino acid and in these two assays the AID of threonine was increased by 1·99 and 2·49%. In contrast, in the titanium oxide assay⁽Reference Ravindran, Morel and Partridge¹⁰¹⁾, the median phytase response was 5·38% and the AID of threonine was increased by 10·16%. In this study the control diets were supplemented with phytase at three inclusion levels. As a main effect, 500, 750 and 1000 FTU phytase/kg improved the average AID coefficient of eighteen amino acids from 0·798 to 0·833 (4·39%), 0·837 (4·89%) and 0·840 (5·26%), respectively. Moreover, in addition to the compelling Ravindran et al. ⁽Reference Ravindran, Morel and Partridge¹⁰¹⁾ study, there are several ‘non-chromic oxide’ assays in broiler chickens⁽Reference Ravindran, Cabahug and Ravindran^42, Reference Selle, Partridge and Ravindran^102, Reference Ravindran, Selle and Bryden^123–Reference Selle, Ravindran and Ravindran¹²⁷⁾ in which phytase unequivocally enhanced amino acid digestibilities.

It would appear that the selection of chromic oxide for phytase amino acid digestibility assays may be contributing towards the equivocal outcomes, and the shortcomings of this marker have been discussed previously⁽Reference Selle and Ravindran^118, Reference Selle, Ravindran and Bryden¹²⁰⁾. A consideration of the impact of exogenous phytases on gut passage rates is instructive, as phytase supplementation, particularly of low-P diets, frequently increases feed intake of broilers and, presumably, gut passage rates. In support of this, 600 FTU A. niger-phytase/kg has been shown to reduce gut transit times by 17·0% from 94·8 to 78·7 min in chicks offered maize–soya diets⁽Reference Watson, Matthews and Southern¹²⁸⁾.

Differences in feed intake rates can influence the outcomes of phytase amino acid digestibility assays. For example, Sebastian et al. ⁽Reference Sebastian, Touchburn and Chavez¹²⁹⁾ evaluated the effect of phytase on amino acid digestibilities in male and female broilers in maize–soya diets that were low in either Ca or P. As shown in Table 6, the overall influence of 600 FTU A. niger-phytase/kg on digestibility of essential amino acids was negligible, with an average response of 0·3%. However, the minimum to maximum responses ranged from − 5·1 to +5·0% across the six categories of sex and diet type. Amino acid digestibilities were determined at day 28 and feed intakes to day 19 were recorded. Feed intakes of female chicks offered low-P diets remained unaltered (+0·2%) following phytase supplementation; in contrast, phytase increased feed consumption of the male counterparts by 15·0%. Remarkably, phytase increased the AID of nine amino acids by an average of 4·8% in female chicks but in the male chicks with increased feed consumption phytase reduced amino acid digestibility by 4·9%. There is an overall negative correlation (r − 0·832; P < 0·05) between phytase-induced percentage increases in feed intake and percentage responses in amino acid digestibility.

Table 6

Responses in the apparent ileal digestibility of essential amino acids to microbial phytase in male and female chicks offered diets with differing calcium and phosphorus levels in relation to feed intake rates*

* Adapted from Sebastian et al. ⁽Reference Sebastian, Touchburn and Chavez¹²⁹⁾.

In addition, Yi et al. ⁽Reference Yi, Kornegay and Denbow¹³⁰⁾ reported a similar pattern of results in turkey poults in which maize–soya diets with differing levels of non-phytate P and crude protein were supplemented with 750 FTU A. niger-phytase/kg. Overall, phytase increased feed intake by 6·1%, but intakes of poults offered the high-non-phytate P/low-crude protein diet were not altered (–0·2%). Although differences were subtle, phytase-induced numerical increases in the AID of amino acids were most evident in this treatment group, with an average increase of 2·7% for nine essential amino acids. In the remaining three treatment groups, phytase increased feed intakes by an average of 8·2%, but AID coefficients varied by only 0·8%. So, there is the distinct possibility that phytase-induced variation in feed intakes/gut passage rates is a confounding factor in amino acid digestibility assays, perhaps particularly those involving chromic oxide, as was the case in the two studies discussed.

Phytase amino acid digestibility assays in pigs

There are two key reports where the impact of phytase was clearly positive in grower and finisher pigs; however, they were not published in peer-reviewed journals. Firstly, Officer & Batterham⁽Reference Officer and Batterham^131, Reference Officer and Batterham¹³²⁾ found that 1000 FTU A. niger-phytase/kg substantially increased the AID coefficients of ten amino acids in diets for grower pigs in which Linola meal, a variant of linseed meal, was the only protein source. In this study, phytase increased average amino acid AID coefficients by 14·0% (0·715 v. 0·627), which ranged from 5·6% (methionine) to 24·0% (threonine) for individual amino acids.

Second, Kornegay et al. ⁽Reference Kornegay, Radcliffe and Zhang¹³³⁾ completed an instructive experiment in which ileal digesta samples were taken from both intact and cannulated finisher pigs offered low-protein diets. In cannulated pigs, 500 FTU A. niger-phytase/kg increased the average AID coefficients of seventeen amino acids by 3·7% (0·779 v. 0·751); in contrast, phytase increased AID coefficients by 9·8% (0·810 v. 0·738) in intact pigs. Further, phytase increased threonine digestibility by 16·2% in intact pigs, as opposed to 6·1% in cannulated pigs. Responses to phytase in the majority of amino acid digestibility assays completed in cannulated pigs, which are often weaners, are marginal⁽Reference Selle and Ravindran¹¹⁹⁾ and generally less than those recorded by Kornegay et al. ⁽Reference Kornegay, Radcliffe and Zhang¹³³⁾ A recent study in cannulated grower pigs by Zeng et al. ⁽Reference Zeng, Piao and Wang¹³⁴⁾ is an exception, as these workers reported that 1000 FTU phytase/kg increased the average AID coefficients of eighteen amino acids by 6·2% (0·854 v. 0·804), with individual responses ranging from 1·3% (arginine) to 13·3% (glycine). It seems that more pronounced phytase responses have been recorded in grower–finisher pigs⁽Reference Officer and Batterham^131–Reference Zeng, Piao and Wang¹³⁴⁾ than in weaners irrespective of the method of ileal digesta collection. These relatively pronounced responses may stem from lower gastric pH in older pigs promoting more intense binary protein–phytate complex formation.

Nearly all phytase–amino acid digestibility assays in pigs, including the studies discussed, have used chromic oxide as the marker. However, Kiarie et al. ⁽Reference Kiarie, Owusu-Asiedu and Simmins¹³⁵⁾ used acid-insoluble ash in a cannulated study where 700 FTU E. coli-phytase/kg increased the average AID coefficient of nine essential amino acids by 3·6% from 0·687 to 0·712. Nitrayova et al. ⁽Reference Nitrayova, Patras and Brestensky¹³⁶⁾ included both chromic oxide (3 g/kg) and acid-insoluble ash (Celite at 10 g/kg) in swine diets based on maize, barley and soyabean meal to evaluate a P. lycii-phytase in cannulated pigs. The responses to phytase did not differ greatly depending on the marker used, and the researchers concluded that marker selection is not the main factor for the ambiguous results recorded in the literature. However, marker selection may be more important in broiler assays than in pigs; a contributing factor could be the reverse peristalsis that takes place in the avian gut⁽Reference Sacrane, Iji and Millelsen¹³⁷⁾.

There is a need to establish the validity of the proposal that responses to phytase are more reliable when ileal digesta samples are taken from intact rather than cannulated animals and several factors that may be contributing to this apparent difference⁽Reference Selle and Ravindran¹¹⁹⁾. The fact that cannulated pigs are usually fed twice daily on a restricted basis is at odds with practical, ad libitum feeding regimens. However, the intervention of cannulation procedures probably leads to a proliferation of amino acids of microfloral origin in the terminal ileum⁽Reference Rowan, Moughan and Wilson¹³⁸⁾ and the reduced motility of a surgically disrupted small intestine probably promotes this microbial proliferation⁽Reference Easter and Tanksley¹³⁹⁾. The protein of microbial origin in the ileum of pigs is rich in glutamic acid and aspartic acid but contains relatively low levels of methionine, cystine and histidine⁽Reference Miner-Williams, Moughan and Fuller¹⁴⁰⁾. Importantly, Brand et al. ⁽Reference Brand, Badenhorst and Siebrits¹⁴¹⁾ found that when offered protein-free diets, pigs that had undergone an ileo-rectal anastomosis had twice the endogenous protein flow (12·1 v. 5·8 g/d) in comparison with intact pigs. The surplus of amino acids of microfloral origin in the ileum of cannulated pigs may mask any phytase-induced increases in digestibility of dietary and endogenous amino acids.

From the Kornegay et al. ⁽Reference Kornegay, Radcliffe and Zhang¹³³⁾ experiment, it is possible to quantity the ileal digestible amino acids generated by phytase in either cannulated or intact pigs and the difference in amounts. It is then possible to compare the differences with the amino acid profile of gut microbial protein⁽Reference Miner-Williams, Moughan and Fuller¹⁴⁰⁾, as shown in Table 7. There is a significant, positive correlation between the amino acid profile of microbial protein and the difference in amino acids generated by phytase in intact and cannulated pigs (r 0·514; P < 0·05). The abundant amino acids in gut microbial protein are also the amino acids for which the most pronounced differences in phytase responses were recorded between intact and cannulated pigs. This supports the contention that a proliferation of amino acids of gut microbial origin in the ileum of cannulated pigs may be masking the positive impact of phytase on the digestibility of dietary and endogenous amino acids.

Table 7

Quantity of ileal digestible amino acids generated by 500 phytase units Aspergillus niger-phytase/kg as determined in cannulated or intact pigs*

* Adapted from Kornegay et al. ⁽Reference Kornegay, Radcliffe and Zhang¹³³⁾ and Miner-Williams et al. ⁽Reference Miner-Williams, Moughan and Fuller¹⁴⁰⁾.

Conceptual framework for protein–phytate interactions

Protein–phytate interactions contribute to the anti-nutritive properties of phytate in pig and poultry diets. These interactions may result in the formation of primary or ternary complexes depending on the isoelectric point of protein and the prevailing pH along the gut. Binary protein–phytate complexes are formed below protein isoelectric points by electrostatic attractions between polyanionic phytate molecules and proteins carrying a net positive charge. Above their isoelectric points, proteins carrying a positive charge are linked with phytate by divalent cationic bridges to form ternary protein–phytate complexes. Additionally, according to the Hofmeister series, phytate is a strong kosmotropic anion as it possesses up to six HPO₄^2– moieties and can stabilise protein by making an impact on the surrounding water medium. In general terms, the likelihood is that these direct and indirect reductions in protein solubility induced by phytate depress protein utilisation. More specifically, protein bound in binary complexes is refractory to pepsin digestion at very low pH in the stomach, which interferes with the initiation of the protein digestive process, and while binary complexes dissociate at the protein isoelectric point they still may not be as readily digested in the small intestine due to structural changes induced by aggregation with phytate. Pepsin-refractory binary complexes may trigger both compensatory gastric hypersecretions and protective mucin outputs, which would increase endogenous amino acid flows. Phytate induces a marked transition of Na into the gut lumen, perhaps primarily as NaHCO₃ to buffer HCl. However, this movement of Na into the gut lumen may reduce intestinal uptakes of dietary and endogenous amino acids by compromising Na-dependent transport systems and Na pump activity. Because of the relatively high isoelectric point of proteins in cereal grains, binary complexes may persist in the small intestine and binary and ternary interactions between phytate and proteins closely associated with starch granules in cereal grains may depress energy utilisation. The molecular weight of protein fractions, particularly in the proximal small intestine, may be sufficiently large to permit phytate to bind nutritionally important quantities of protein in ternary complexes and this would also hinder absorption of key minerals including Ca and Zn.

Future research

With emphasis on recent findings, the present review endeavours to provide a unified framework of the mechanisms underlying protein–phytate interactions and their nutritional consequences and to explain the ambiguous effects of phytase on amino acid digestibility. The capacity of phytate to interact with protein per se and with protein closely associated with other nutrients, including starch, appears to be pivotal. The possibility that phytate indirectly interacts with protein because it is a strong kosmotropic anion is a new concept that demands further investigation and perhaps a reassessment of binary and ternary complex formation in relation to protein–phytate interactions. The apparently differing propensities of proteins to be bound by phytate merit examination. Attention should be paid to the proposal that binary protein–phytate interactions reduce pepsin digestibility of aggregated protein, which in turn triggers gastric hypersecretion of pepsin and HCl coupled with increased protective outputs of mucin and NaHCO₃. Investigations should also focus on the impact of phytate and phytase on the absorption kinetics of glucose and amino acids along the small intestine, given the likelihood that phytate-induced endogenous depletions of Na may compromise Na⁺-dependent transport systems and Na⁺-K⁺-ATPase activity. It even may be that phytate has a direct, negative impact on the Na pump and this possibility merits investigation. While the evidence is not conclusive, it is our contention that dietary phytate negatively influences protein and energy utilisation, probably to a greater extent in poultry than pigs. Ideally, the issues discussed in relation to the divergent outcomes of phytase amino acid digestibility assays in pigs and poultry should be clarified. The adoption of these recommendations should permit greater advantage to be taken of the ‘extra-phosphoric’ effects of exogenous phytases in respect of protein and energy utilisation in pigs and poultry, thereby facilitating more efficient and sustainable production of pig-meat, poultry-meat and eggs.