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Phytase and Phytate: Engineering phytases for more efficient

breakdown of phytate

J. S. Sands P.W. Plumstead and A. J. Cowieson

Danisco Animal Nutrition, Marlborough, Wiltshire, SN8 1XN, UK

Introduction

Phosphorus (P) plays a vital role as metabolic intermediary and structural

component in the animals body. Thus it is essential to have an adequate supply

of dietary phosphorus to enable efficient growth and adequate bone development

of animals. In cereal-based commercial diets more than half of the ingested

phosphorus will be in the organic form referred to as phytin or phytate P. Due toinsufficient endogenous intestinal phytase activity, P in this form is largely

unavailable to monogastric species such as pigs and poultry. As a result,

depending on the feedstuffs used and the proportion of inorganic and organic

phosphorus available for digestion, only around 30- 50% of total dietary

phosphorus is retained by the pigs and poultry (Poulsen et al., 1999; Van Der Klis

and Versteegh). To account for this low availability of P, commercial livestock

feeds are supplemented with P from various sources including inorganic mineralphosphates, by-products from the meat processing industries and to fishmeal.

The continued application of manure and litter to soils has led to an increase in

soil test P concentrations in areas of intensive poultry and swine production

(Poulsen et al., 1999; Pautler and Sims, 2000). High P levels in soils have

substantial environmental implications as this was shown to increase P contained

in surface water runoff (Pautler and Sims, 2000; CAST, 2002). The environmental

implications of a high soil test P level have led to governmental regulations that limit

the amount of P from manure that can be applied to land (Environmental Protection

Agency, 2003). A consequence of the legislative environmental constraints that

have been imposed on pig and poultry production in certain countries is that the

cost of production has increased..

To improve the utilization of phytate P and reduce total P excreted in manure,

exogenous phytase enzymes of microbial origin have been developed(Angel et al.,

2002; Applegate et al., 2003). However, differences have subsequently been shown

To whom correspondence should be addressed

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to exist between commercial sources of phytase in their efficacy to hydrolyze phytate

in the digestive tract of the animal. These differences can impact the nutritional,

economic and environmental benefits of adding phytase to diets. Therefore, to be

able to obtain the most benefit from each class of phytase it is important that

nutritionists understand phytate and phytase chemistry and how this impacts P

utilization and was the objective of this paper.

Phytate an anti-nutrient

Phytate, also known as inositol phosphate-6 (IP6), or myo-inositol

1,2,3,4,5,6-hexakis dihydrogen phosphate, is ubiquitous in all plant-based feed

ingredients and serves as the main storage form of phosphorus in seeds.

Phytates, salts of phytic acid, constitute about 1 to 2% by weight of many cereals

and oilseeds, but can be as high as 3 to 6% (Cheryan, 1980). The concentration

of phytate varies considerably between feed ingredients, and also within feed

ingredients (Eeckhout and de Paepe 1991). For example, Cossa et al. (1997)

showed that the range of phytate phosphorus contained in 54 maize samples

varied almost two-fold from 1.92 mg/kg to 3.54 mg/kg, with an average of 2.66

mg/kg (SD of 0.34). The location of the phytate in grain also differs between

cereals. In maize, 90% of the phytate is concentrated in the germ portion of the

kernel, whereas in wheat and rice, most of the phytate is in the aleurone layers of

the kernel and outer bran (Angel et al., 2002). Therefore, not only can dietary

phytate vary widely within conventional maize-soybean-meal diets, but will also

vary with the inclusion of different ingredients.

The interaction between phytic acid, minerals and or proteins reduces the

bioavailability of minerals and to an extent, proteins. Key to the anti-nutritive

effect of phytate is its structure and chemistry. Under most conditions

encountered in feed phytic acid will be strongly negatively charged (Cheryan,1980). This characteristic indicates that phytic acid has very strong chelating

properties. Another property of phytate that is key to its anti-nutritive effects is its

solubility or lack of solubility. At around pH 5.5-6.0 (more or less depending on

the type and concentration of cations) the solubility of phytates rapidly declines.

This property of phytate has implications for its breakdown by phytase. The

stomach is the primary site of hydrolysis of phytate by phytase in the

gastrointestinal tract of pigs (Jonbloed et al., 1992, Kemme et al., 2006) will verylittle hydrolysis taking place in the intestine. The lack of hydrolytic activity in the

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lower gastrointestinal tract is likely due to the formation of insoluble, ternary

complexes with minerals and proteins (Cheryan, 1980).

In addition to reducing the availability of P, phytate has been shown to

reduce the ME and overall digestibility of dietary amino acids (Angel et al., 2002;

Ravindran et al., 2000). Selle et al. (2007) presents a comprehensive review of

phytase and AA digestibility in poultry, and offers three potential mechanisms by

which phytase can improve the AA value of a diet: firstly, the presence of

protein-phytate complexes in feedstuffs, reducing solubility and thereby

digestibility of dietary protein; secondly, the de novo formation of binary and

ternary complexes between protein, minerals and phytate in the gastro-intestinal

tract and finally the inhibition of proteolytic enzymes or their cofactors by phytate.

A final mechanism, reported by Cowieson and Ravindran (2007), is that phytase

elicits its positive effects on ME and AA digestibility, in part, via a reduction in

endogenous AA flow (Figure 1). The increased endogenous AA flow with the

ingestion of phytate and the reduced flow with phytase addition being credited to

changes in the secretion of mucin and endogenous enzymes, and also reflect

changes in the efficiency of re-absorption or both.

Phytase

Phytase (hexa-kisphosphate acid phosphorylase) catalyze the release of

orthophosphate from phytate and lower inositol phosphates. Phytase from plants

was described as early as 1907, whereas, the identification of fungal microbes as

sources of high phytase activity was reported in 1959 (Wodzinski and Ullah,

1996). According to the aforementioned review, the first commercial initiative to

develop phytase as a product was in 1962, but it wasnt until the year 1990 that

the first demonstration of the efficacy of a commercially produced microbial

phytase in poultry was reported (Simons et al., 1990). Until recently, limitations in

the thermal tolerance and costs of phytase restricted its use, except in regions

where its use as an option to abate environmental pollution was mandated by

legislation. Major shifts in the feed industry such as the ban of animal by-products

as feed ingredients and more recently, rapidly increasing cost of feed ingredients

and inorganic phosphate sources has also made phytase a more viable option to

feed producers.

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Even with the current levels of improvement in phytin P availability there is

room for substantial improvement in the effeciency of phytases. Recent advances

in industrial biotechnology, fermentation and feed application knowledge, has

enabled the development of new generation that are more efficient at hydrolyzing

phytate P and reducing P excretion. The differences in phytate hydrolysis

between classes of phytate are due to differences in the biophysical and

biochemical properties that determine their mode of action (Igbasan et al., 2000;

Wyss et al., 1999).

Biochemical properties determine the efficacy of phytases

There are three main characteristics that separate different classes of

phytases in their ability to hydrolyze phytate and reduce its anti nutritional effect

in the gastro-intestinal tract. These relate to differences in the protein structure of

the phytase enzyme that, in-turn, influence its enzyme kinetics i.e. the specificity

of the enzyme for phytate (IP6) versus lower order inositol phosphate esters, and

the rate at which the enzyme is able to cleave phosphate groups off the phytate

moiety. In addition, differences exist between phytases in the pH optimum at

which maximum phytase activity is reached. These differences in the pH optimum

of phytase determine the relative efficacy of phytases in different parts of the

gastrointestinal tract, and also determine the relative influence of other factors,

such as dietary calcium level, on the ability of different phytases to hydrolyze

phytate in the diet.

1. pH optimum.

The ability of phytase to hydrolyze phytate is dependent on the phytate being

soluble and not chelated to di- or tri-valent cations or dietary protein. Factors

affecting phytate solubility are complex and include interactions of pH, the molar

ratio of Ca:phytate and the presence of proteins and pepsin. In general, phytate

is soluble at a low pH. However, the pH optimum for phytate solubility is

decreased substantially by the addition of calcium to diets (Grynspan and

Cheryan, 1983). In practical maize-soybean meal diets that contain > 0.6%

calcium and ~0.26% phytate phosphorus, maximum solubility of phytate occurs at

or below a pH of 4.4 (Pontoppidan et al., 2007). The implication of this is that

hydrolysis of phytate by phytase must occur in the acid portion of the digestive

tract. Once feed enters the duodenum, and the pH of the digesta increases

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above pH 4.4, the phosphate groups on phytate become increasingly negatively

charged and attract more positively charged cations such as Ca2+, Zn2+, and

Fe2+. The formation of stable Ca-phytate or Zn-phytate complexes at the higher

pH of the proximal duodenum causes phytate to rapidly precipitate out of

solution, reducing the effectiveness of phytase enzymes to break down phytate.

Research by Wyss et al. (1999) has shown that the relative activity of phytases

over the pH range of 2.2 to 4.5 was 50% higher for E. coli vs. A. niger phytases

(Figure 2). The implication of these differences in the enzyme pH optima is that

the relative activity of E. coli phytases in the acidic portion of the stomach

(proventriculus/ gizzard in poultry) will be greater than that of fungal phytases. In

this pH range (2.2-4.5) the formation of stable Ca-phytate complexes does not

occur, meaning the phytate stays in solution and can be hydrolyzed by phytase.

Therefore, the lower pH optimum of E. coli phytases versus fungal phytases

implies that their ability to hydrolyze dietary phytate may be less affected by the

high dietary Ca2+ levels of >0.6% that occur in commercial pig and poultry diets

(Plumstead, 2007).

2. Phytase enzyme specificity.

In contrast to Ca-phytate complexes that form at the higher pH of the proximal

duodenum, the formation of insoluble protein-phytate complexes occurs at a low

pH (2.0-4.0) in the gizzard/ proventriculus. The formation of insoluble protein-

phytate complexes in this acidic part of the digestive tract is increased by the

presence of phytate, but is decreased by secretion of pepsin (Pontoppidan et al.,

2007). Therefore, to improve the solubility of protein in the presence of phytate,

pepsin secretion in the proventriculus increases in response to increased

concentrations of insoluble protein-phytate complexes. The increased secretion

of pepsin and other digestive secretions such as pancreatic enzymes, bile, and

mucin in response to phytate was thought to be one of the mechanisms whereby

phytate increased endogenous protein and energy losses at the terminal ileum

(Cowieson and Ravindran, 2007). As the formation of insoluble protein-phytate

complexes is proportional to the amount of phytate in the in the acid part of the

digestive tract, the speed at which phytases remove phytate from the digesta

may be influential in preventing the formation of protein-phytate complexes and

their negative effect on increased pepsin secretion and endogenous nutrient

losses.

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Substantial differences exist between E. coliand fungal phytases in their affinity

to bind to phytate (IP6) vs. binding lower order IP esters. A greater affinity for

phytate, in combination with a low optimum pH will increase the rate of phytate

hydrolysis in the acidic portion of the digestive tract (pH

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vitrosimulation; 2) in vivodigestbility assays and 3) growth performance assays.

In vitro orthophosphate release assays are indirect and relatively cheap, but only

provide an estimate of the amount of P release under practical feeding

conditions. In vitro assays that attempt to simulate the gastrointestinal conditions

will provide more relevant results, but still provides only an approximation. If

comparing phytases in vitro, the substrate (phytate or naturally occurring phytin),

pH and time are but three of several factors that should be controlled. The

difficulty lies in the inability to adequately simulate the dynamics of

gastrointestinal conditions. If a phytase from of a given source releases

orthophosphate at a higher rate than the other, but the incubation time is

extended, then both phytase products will appear equally efficient at releasing P

from phytate.

Growth assays are also an indirect method of assessing the efficiency of

phytases. If body weight gain is the only criteria measured, maximum growth

may be achieved, but this may not be the optimal biological dose that meets

requirement for adequate bone strength. Using bone strength or bone ash as an

index to estimate P requirements and differences in phytase efficacy is preferred,

and has been shown to be a more sensitive criterion. Bone ash is commonly

assessed as an index of bone strength, because it is a well established

procedure and equipment required is less expensive. Onyango et al., (2003)

demonstrated that bone ash and bone strength are highly correlated therefore the

former can be used as an index of the latter. Digestibility assays are the most

direct and effective way to evaluate the efficiency of a phytase releasing P and

other nutrients that are bound to phytate. Measuring the ileal digestibility of P

provides a good estimate of the amount of P released by phytase. In contrast,

total tract digestibility assays present an estimate of the P retained by the animal

and are useful when comparing phytases in terms of their ability to reduce P

excretion in manure.

However, the results of biological efficacy assays must be interpreted with

caution. It is essential that when evaluating the response of an animal to phytase,

that P is the limiting nutrient in the negative control diet. Studies that compare the

efficacy of two phytases under conditions in which P has not been sufficiently

reduced may produce results that suggest both enzymes to be equally

efficacious, when in fact they are not. Choosing the right test model is also critical

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for evaluating efficacy. Younger animals have a higher proportionate requirement

than older animals, thus provide a more sensitive test model. The length of the

test period is also an influential factor in bioefficacy assays. The test period

should be appropriate to the dietary nutrient levels, the animals adaptive

mechanisms and appropriate welfare standards. As bioefficacy assays are used

to generate P replacement values, it is crucial the tests are conducted in such a

way that the results are a close estimate of the actual amount of P release by

phytase.

For example, the efficicacy of an E. coli-derived phytase, expressed in

Schizosaccharomyces pombewas compared to that of a fungal-derived phytase

expressed in Aspergillus niger in a trial with 21-day-old turkey poults. Nicholas

Large White female turkey poults were fed a maize-based diet reduced in P and

Ca levels compared to requirements. The feed was supplemented with phytases

at 0, 250, 500 and 1000 FTU/kg feed. Performance and bone mineralization were

measured at 21 days of age and percent retention of P and Ca calculated

between 15 and 19 days of age. Phytase addition to the P and Ca deficient diet

led to a significant growth improvement and a bone mineralization (expressed as

percent toe and tibia ash) improvement due to an increase in P and Ca retention.

Response curves were plotted using an exponential decay curve fit. A difference

in efficacy between E. coli and A. niger phytases was observed in this trial,

especially with lower doses (250 and 500 FTU/kg feed) where dietary P level is

below requirement of the birds. Biochemical characteristics of E. coli-derived

phytase confers a wider pH activity profile and greater resistant to breakdown by

gastrointestinal proteases. These two properties are essential for high activity in

the upper portion of the gastrointestinal tract where phytates are soluble and

therefore accessible to phytase. These results clearly demonstrate that phytases

from different sources possess different structural and biochemical properties that

determine their bioefficacy.

Phytase and nutrient digestibility

Recent work by Cowieson and Ravindran (2007) showed that a reduction

in apparent nutrient digestibility due to the presence of phytate was, in part,

mediated by an increase in the loss of endogenous amino acids and energy from

the terminal ileum. Importantly, the negative effect of phytate on energy and

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amino acid utilization was proportional to the concentration of phytate in the feed

and was significantly reduced by the addition of an E. coli phytase to the diet

(Figure 4a&b). Other research has also demonstrated that the improvement in

dietary energy and amino acid utilization from phytase increased with increased

phytate phosphorus concentrations from 0.23 to 0.33% in the feed (Shan &

Davis, 1994; Ravindran et al., 2006), and was also dependent on the age of the

broiler (Sands et al., 2004).

In dietary ingredients phytate exists as a salt of K and Mg and is relatively

unreactive (Lott et al., 2000). However, when feed is exposed to the low pH

conditions in the proximal gut, phytate becomes soluble as H+ ions replace K and

Mg (Cosgrove, 1966). Though protonated, phytate still carries a net negative

charge and can react electrostatically with basic amino acid residues in dietary

protein. The extent of this reaction depends on the concentration and solubility of

phytate, ambient pH, the isoeletric point of the protein, and also its tertiary and

quaternary structure (i.e. the degree of steric hindrance between reactive amino

acids and phytate). These phytate-protein complexes are variably refractory to

solubilisation with HCl and digestion by pepsin (Rajendran & Prakash, 1993),

leading to an increase in the secretion of these by the animal. On gastric

emptying, the distal gut is faced with a luminal challenge to maintain favourable

conditions for optimal functioning of the pancreatic and brush border enzymes,

and for a satisfactory ion balance for nutrient transport. Hyper-secretion of mucin

and sodium bicarbonate, corresponding to variation in proteolytic and H+

antagonism follows, which increases the presence of endogenous amino acids

and sodium in the lumen and presumably alters maintenance requirements.

Further, the loss of endogenous protein from the ileum has a direct effect on the

digestible energy value of the diet, depending on the amino acid composition of

the protein leaving the terminal ileum. This direct energetic cost has been

estimated to be as much as 0.1MJ/kg DM intake for every 1g/kg dietary phytate-P

(Figure 1; Cowieson & Ravindran, 2007b), without including effects on net energy

associated with protein synthesis and turnover. Thus, the ingestion of dietary

phytate influences endogenous loss indirectly via a reduction in the solubility of

dietary protein with a subsequent cascade altering intestinal dynamics via

secretive and absorptive mechanisms.

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A comprehension of mode of action for the extra-phosphoric effects of

phytase is not only scientifically enlightening but also assists in explaining why

published phytase matrix values for amino acids and energy vary so significantly.

For example, the amino acid composition of mucin and pepsin are highly

correlated with the phytase-induced improvements in ileal amino acid digestibility

(Cowieson & Ravindran, 2007a). This is instructive as it explains why the

improvement in threonine, serine and cysteine digestibility with phytase is

typically superior to that of methionine (endogenous proteins being essentially

devoid of methionine). It can be concluded that exogenous microbial phytase

can beneficially influence amino acid and energy digestibility and that a

significant proportion of these effects may be explained by a reduction in the

antinutritive effects of phytate. A greater understanding of the physiological

consequences of changes in endogenous amino acid and mineral secretion and

resorption is justified, as maintenance requirements, immune function and

microbial-host dynamics may be expected to be involved.

The importance of these reports is two-fold. In the first instance, the anti-

nutritional effects of phytate extend beyond a simple reduction in phosphorus and

calcium availability and include negative effects of phytate on endogenous

nutrient losses, which reduce the overall efficiency of energy and amino acid

utilization. Secondly, these extra-phosphoric effects of phytate are dependant on

the concentration of phytate in the feed and the age of the broiler. The implication

of these findings is that the nutrient contribution from phytase is by no means a

fixed entity, but will be dependent on the initial phytate concentration in the diet,

the amount of phytase added to the feed and, most importantly, the rate and

extent of phytate hydrolysis from phytase in the early part of the gastro-intestinal

tract. This latter factor is likely the most influential in describing the large

differences in phytase efficacy that exist between traditional fungal sources of

phytase and the new generation of E. coli phytases.

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Table 1. Resistant an E. coli*phytase and two fungal phytases (A. niger and P.lycii)phytases to breakdown by pancreatic proteases.

Remaining phytase activity (%) after treatment with pepsin,trypsin or chymotrypsin in vitro

E. coliphytase - XP Fungal phytase - N Fungal phytase - R

Pepsin 76.7a 31.4b 5.42c

Trypsin 23.0a 0.45b 1.25b

Chymotrypsin 65.8a 2.95b 5.77b

abcP

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0

2 0

4 0

6 0

8 0

1 0 0

1 2 0

0 2.4 3.2 4.0

Dietary Phytate P (g/kg)

No Phytase

+ E.coliphytase1

GEfromEndogenousProtein

(kcal/kgDM

intake)

0

20

40

60

80

100

2 3 4 5 6 7 8

RelativeActivity(%)

pH

A.nigerPhytase

E.coliPhytase

Figure 2. The relative activity of E. coliphytases over pH range of 2.0 to 4.5 in theproximal digestive tract is 50% higher than A. nigerphytases (Wyss et al., 1999)

Figure 1. Effect of phytate and an E. coliphytase (500 FTU/kg feedfrom Phyzyme XP, Danisco Animal Nutrition) on the gross energylost from endogenous protein exiting the terminal ileum of broilers(Cowieson & Ravindran, 2007)

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Figure 3 a,b. The specific activity of E. coliphytases for phytate (IP6) is 8 x greaterthan that of A. nigerphytase (3a) and results in a 3-fold decrease in the time required

to completely hydrolyze phytate to lower inositol phosphate esters (3b) (Wyss et al.,1999)

0

2 0 0

4 0 0

6 0 0

8 0 0

1 0 0 0

SpecificActivityforIP6

(U/mg)

A.niger E.coli

(a)

0

20

40

60

80

100

0 5 10 15 20

Rema

iningPhytate(%)

Time (min)

A.nigerPhytase

E.coliPhytase

(b)

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17

R2 = 0.998

R2 = 0.999

0 20 0 40 0 60 0 80 0 1000

Phytase FTU/kg

30

35

40

45

50

Tibia Ash % E coli

A. niger

Figure 4. Effect of E. coli () and A niger () phytases on percent tibia ash of

turkey poults fed low-phosphorus and calcium maize-soy-based diets from day 1-

21 of age.

E. coli y = 31.6 + 11.5*(1-exp(-0.008*x))A. niger y= 31.6 + 11.4*(1-exp(-0.003 *x))

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18

8.37.0 7.3

21.7

6.14.5 4.3

9.89.1

10.5

18.5

7.0

9.2

4.6 4.8 4.8

2.6

7.0

3.0

4.7

1.6 2.2 1.7

3.7

2.0 2.11.3

3.42.4

1.5 1.8 1.72.3

2.0

0

1000

2000

3000

4000

5000

Endogenousflow

(mgAA/kgDM

intake)

Basal IP6

Asp Thr Ser Glu Pro Gly Ala Val Ile Leu Tyr Phe His Lys Arg Cys Met

Basal + 4g/kg PP

Total AA flow 87.6% N flow 78% Ser 152%; Thr 135% Glu 49%; Ala 29%

P