Indian Journal of Biotechnology Vol 14, July 2015, pp 293-311

Heterologous expression of yeast and fungal phytases: Developments and future perspectives

Swati Joshi and T Satyanarayana*

Department of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi 110 021, India

Received 8 May 2014; revised 27 February 2015; accepted 10 March 2015

The phytases, which hydrolyse inositol hexaphosphate (IP6), are used as animal feed supplements for ameliorating phosphorus availability and assimilation, and mitigating anti-nutrient effects of phytates. Phytases have been reported from several yeasts and fungi. The phytase titres attainable from the wild yeast and fungal strains are very low and, therefore, attempts have been made to clone and overexpress them in heterologous hosts. The enzymes have also been improved by site directed mutagenesis and directed evolution to suit the applications. This review focuses on the progress made so far in cloning and expression of yeast and fungal phytases in heterologous hosts and the research needs.

Keywords: Acid stability, heterologous expression, inositol hexaphosphate, phytase, recombinant phytase, thermostability

Introduction

Phytate [inositol hexaphosphoric acid (IP6)] is an organic form of phosphorus found in plant products, such as, cereal grains, legumes, oilseeds and others, in which it represents more than two-thirds of the total Pi

1. Phytates are not digested by monogastric animals and human beings because of inadequate levels of phytase in their gastrointestinal tract and thus pass through faeces. Soil and aquatic microbes act on phytates and release inorganic phosphate that, in areas of intensive animal rearing, leads to eutrophication of aquatic bodies2,3. Due to the presence of six phosphate groups, phytic acid is a strong metal ion chelator (Fig. 1). Because of strong affinity to metal ions, phytates chelate various nutritionally important metal ions, such as zinc, iron, calcium, magnesium and others, and thus lower their bioavailability4,5 (Fig. 1). Phytate acts as an antinutrient that causes mineral deficiencies in populations where staple food is phytate-rich legumes and cereals6. In the past few decades, attempts have been made to mitigate the problems caused by phytates in foods and feeds by supplementing them with microbial phytases.

In order to fulfil their P requirement, animal feeds are supplemented with other forms of phosphorus like calcium salts of phosphate. Such supplementation

increases the overall feed cost and can culminate into a severe cause of water pollution. An effective solution to overcome poor phytate phosphorous utilization in monogastrics is supplementation of feeds and foods with exogenous phytase7.

The recent reviews on phytases have dealt with the production, characteristics and applications of native phytases8-12. This review focuses on the heterologous expression of fungal and yeast phytases and the characteristics of recombinant phytases, attempts made in ameliorating them by molecular approaches and their utility in dephytinization.

Phytate: A Major Form of Organic Phosphorous

Phosphorus is an essential nutrient for all living organisms and is the third most abundant mineral in

Fig. 1—Phytic acid (inositol hexaphosphate) complexed with metal ions, carbohydrate and protein.

—————— *Author for correspondence: Tel: +91-11-24112008; Fax: +91-11-24115270 tsnarayana@gmail.com

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the animal body. This versatile element is found in all cells and participates in every metabolic pathway. Phosphorus plays diverse functions in living systems being part of cell membrane structure in the form of phospholipids, in energy transfer as ATP, an important element in nucleic acids and as an important element of bone structure13. Life cannot be imagined without phosphorus14.

In soil, phosphorus exists both in inorganic and organic forms. Phosphorus is abundant in soils but it is not present in a form readily available for uptake by plants. According to Lopez-Bucio and coworkers15, 70% of the cultivable land in the world is either acid or alkaline, in which phosphorus tends to form compounds, which are not readily available to plants16. One of the most abundant forms of organic phosphorus in soil is inositol phosphates, which constitute up to 50% of the organic phosphorus17.

In mature seeds of the most traditional crops, about 75% of the total phosphorus is found in the form of phytic acid (phytate P)18, which is hexaphosphoric ester of the hexahydric cyclic alcohol meso-inositol. Phytic acid (IP6) is a major storage form of phosphorous in the plant tissues. Inositol pentaphosphate (IP5), tetra- (IP4) and triphosphate (IP3) are also called phytates. The molecular formula is C6H8O24P6 with molecular mass of 660.04 g mol-1. It represents a sum equivalent to about 50% of all phosphorus applied as fertilizer worldwide19, an amount that would satisfy a major fraction of animals’ dietary requirement for phosphorus.

The total phosphorus found as phytic acid phosphorus in grains fed to non-ruminants is simply wasted. During the last 15 years, phytases have attracted considerable attention from both scientists and entrepreneurs in the areas of nutrition, environmental protection and biotechnology. Undoubtedly the increasing public concern regarding the environmental impact of high phosphorus levels in animal excreta has driven the biotechnological development of phytase and its application in animal nutrition.

Effects of Phytic Acid The presence of phytates exerts both negative and

positive effects on nutritional and environmental health. Phytate-P is largely unavailable for assimilation by monogastrics due to lack of endogenous phytase, the enzyme that catalyzes the hydrolysis of phytate. Phytase secreted by microorganisms in the alimentary tract is effective in

hydrolysing phytate in ruminant animals20. Since microflora in monogastric animals and human beings is predominantly located in the large intestine, it can be understood that most of the vegetable phosphate liberated from phytate remains unabsorbed and excreted. Due to increase in the livestock population in various regions, manure applied to the soil is exceeding that leads to the accumulation of phosphate in soil21. Excess soil phosphorous could lead to eutrophication of waters bodies and leaching of phosphate into ground water could also occur22. Inositol hexaphosphate is reasonably reactive due to the presence of 12 dissociable protons with pKa values ranging from 1.5 to 1023.

The reactivity of inositol phosphate isomers depends on the conformation and configuration of the molecule and the pH of its environment23. The phosphorylated myoinositol (IP6) is a potent chelator of many mineral ions24,25. As a metal chelator, it forms insoluble salts at intestinal pH that results in reduction in the availability of minerals for absorption in the body fluids26-28. Furthermore, the availability of nutritionally important amino acids is also compromised in the presence of inositol hexaphosphate in the diet27,29. Some in vitro studies have shown that phytate-protein complexes are degraded very slowly by proteolytic enzymes30, and even some enzymes, such as, pepsin, amylopsin, and amylase, are inhibited by phytates. Phytates may also interfere with the digestibility of lipid and starch31.

The addition of exogenous phytase to the diets can improve the assimilation of dietary phosphate that leads to amelioration in performance parameters different from those associated with an increase in the phosphate utilization27,29. The interaction between protein and IP6 is dependent on pH. The binary complexes of protein-IP6 are formed at lower pH, while ternary complexes of protein-IP6- mineral form at pH near neutrality25,27,32. The complex formation of minerals due to chelating property of phytate can reduce their involvement as cofactors of enzymes, particularly under suboptimal dietary levels.

According to in vitro studies performed by Knuckles et al

33, the esters of inositol phosphate inhibit digestibility of casein by proteolytic enzyme pepsin and the inhibition is dependent on the degree of phosphorylation of the inositol ring. Positive effects of phytate includes protection from various types of cancers34, which is mediated through inhibition of the cellular signal transduction35,

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increase in activity of natural killer cells36, stimulation of cell differentiation genes37 and antioxidant properties38. Apart from this, phytate consumption also helps in the treatment of diabetes mellitus39,40, coronary heart disease41 and dental caries42. Phytates are known to reduce the chances of renal stone formation43, HIV-1 infection44 and heavy metal toxicity.

Methods of Reducing Phytic Acid Content in Foods and Feeds

Dephosphorylation of phytate improves nutritional quality of diets as the removal of anionic phosphate groups from the IP6 reduces mineral chelating strength of phytate that results in the improvement of assimilation of essential minerals45. Phytic acid content of food can be reduced by some physical (autoclaving, cooking & steeping), chemical (ion exchange & acid hydrolysis) and food processing (soaking, germination & fermentation) methods, but these methods decrease the nutritional value of foods. The reduction of phytic acid content in foods and feeds by enzymatic hydrolysis using phytase is desirable, since this does not affect their nutritional value and, thus, phytases find a very important application in nutrition. The dephytinizing capacities of phytases depend upon the origin due to differences in their inherent phytate-degrading activities46. The commercial phytase products have been mainly used as animal feed additives in diets, largely for swine and poultry, and to some extent for fish. In spite of its immense potential in processing and manufacturing of food for human consumption, there is no phytase product that has found its way to the market for human food application. Many researchers have reported a convincing improvement in food products by adding microbial phytase in food processing, for example, bread-making47,48, plant protein isolates49, corn wet milling50 and the fractionation of cereal bran51.

Phytase Global phosphate mineral deposits are non-

renewable. Phosphorous captured in the form of phytate is an optional phosphorous source, which can be efficiently converted to available phosphorous by phytases. Phytases (myo-inositolhexaphosphate phosphohydrolase) hydrolyze phosphomonoester bond of phytic acid and liberate myo-inositol and inorganic phosphates through various myo-inositol phosphate intermediates that mitigates its anti-nutritional properties. Phytases are synthesized in various microbes, plants and animals 11,52-54. Phytase is produced by a large number of microorganisms

including bacteria, filamentous fungi and yeasts. Various yeast species have been reported to produce phytase either intracellularly (Cryptococcus laurentii

strain AL255), cell bound (Candida krusei56

& Pichia

anomala57) or extracellularly (Arxula adeninivorans

58, Kluyveromyces lactis

59, P. anomala59, Rhodotorula

gracilis60, Saccharomyces cerevisiae

61, Schwanniomyces occidentalis

62 & Torulaspora

delbrueckii59). Phylogenetic relationship among

phytases of various yeast species is shown in Fig. 2. Fungi, such as Aspergillus niger

63,64, A. terreus64,

A. fumigatus65,66, A. oryzae

67, A. caespitosus68,

Emericella nidulans64, Myceliopthora thermophila

64 and Thermomyces lanuginosus

69, have been reported to produce phytases. The phylogenetic relationship among various fungal phytases is shown in Fig. 3.

Classification of Phytases

In consultation with the IUPAC-IUBMB Joint Commission on Biochemical Nomenclature (JCBN), Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB) has classified phytases into three categories. 3-Phytases (EC 3.1.3.8) constitute the first category, which first hydrolyse the ester bond at the 3 position of myo-inositol hexakisphosphate, and are mainly of microbial origin. Second category comprises 6-phytases (EC 3.1.3.26), which first hydrolyse the ester bond at the position 6 of

Fig. 2—Dendrogram showing phylogenetic relationship among yeast phytases. [Acc. no. of various phytase used for construction of the neighbour joining phylogenetic tree are: P. anomala

(GenBank ID: FN641803), P. pastoris (Swiss-Prot: P52291), Lodderomyces elognisporus (NCBI ref no.: XP001527604), C.

tropicalis (NCBI ref. no.: XP002546108), C. albicans (NCBI ref. no.: XP713452), Kodamaea ohmeri (GenBank ID: ABU53001), S. cerevisiae (GenBank ID: EDN64708), K. lactis (GenBank ID: CAA83964), Scheffersomyces stipitis CBS 6054 (NCBI ref. no.: XP001385108), Meyerozyma guilliermondii (GenBank ID: CAL69849.1), Blastobotrys adeninivorans (GenBank ID: CAJ77470.1), S. capriottii (GenBank ID: ABN04184.1)].

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myo-inositol hexakisphosphate. These are typical for plants, although this has recently been reported in some Basidiomycete fungi70. Third category contains 5-phytases, which start phytate hydrolysis at the D-5 position and, hence, are classified as 5-phytases. So far, only one enzyme has been reported from this class, which is an alkaline phytate-degrading enzyme from Lily pollen71.

Based on the pH for enzymatic activity, phytases have been broadly classified into two major classes: acid phytases and alkaline phytases. Acidic phytases have been explored more compared to alkaline phytases because of their applicability in foods and animal feeds, their broader substrate specificity and acid stability. Besides two systems of classification, depending on their structure and catalytic mechanism, phytases are also classified as HAP (histidine acid phosphatase), BPP (β-propeller phytase), CP (cysteine phosphatase) and PAP (purple acid phosphatase)72.

HAPs (EC 3.1.3.8) are the most commonly studied phytases, which share a common active site motif (RHGXRXP), a catalytically active dipeptide (HD)

and employ a two-step mechanism consisting of a nucleophilic attack on the phosphorous atom by the histidine of the active site motif, followed by hydrolysis of the resulting phospho-histidine intermediate73. The positive charge of the guanido group of the arginine residue in the conserved tripeptide RHG interacts directly with the phosphate group in the substrate, thus making it more susceptible to nucleophilic attack; while the histidine residue serves as a nucleophile in the formation of covalent phosphohistidine intermediate. The aspartic acid residue (from the C-terminal HD-motif) protonates the group, leaving the substrate. The substrate specificity of all known HAPs for phytic acid varies from one to another64.

BPPs (EC 3.1.3.8), first identified in Bacillus species, have six-bladed β-propeller structures. BPPs show a novel mechanism for hydrolyzing its substrate. Crucial amino acid moieties, which are involved in the conformation of two phosphate binding sites and six calcium binding sites, are conserved in all BPPs. This class of enzymes is mainly useful in aquaculture due to activity at neutral pH74,53. Most freshwater monogastric and agastric fishes have a digestive tract with a neutral environment; this includes cyprinid fishes, a group of commercially important freshwater species. Thus, a phytase useful in aquaculture should have high activity at neutral gut pH and at the lower body temperature of fish (20-28°C)75.

A novel class of phytases, cysteine phosphatases, is predominantly produced by anaerobic bacteria of the rumen76-78. In contrast to monogastrics, ruminants can utilize phytate from a plant-based diet more efficiently because of the presence of the bacteria in their rumen, which can hydrolyse phytate very efficiently79,80. Yanke et al

80 have identified an anaerobic rumen bacterium Selenomonas

ruminantium capable of producing phytase. The structure and proposed catalytic mechanism of this phytase suggested it to be a member of the cysteine phosphatase (CP) superfamily76. Its deduced amino acid sequence contains the active site motif HCXXGXXR (T/S) and other substantial similarities with protein tyrosine phosphatase (PTP), a member of the CP group. Cysteine phytases are active at acidic pH, mostly at pH 4.576,81. These are, therefore, classified as acid phytases (pH optima of 4.0-5.0)81.

Purple acid phosphatases (PAP) are binuclear metallohydrolases that are found in plants, animals

Fig. 3—Dendrogram showing phylogenetic relationship among fungal phytases. [Acc. no. of various phytase used for construction of the neighbour joining phylogenetic tree are: A. terreus (GenBank ID: AAB52507), A. nidulans (GenBandk ID: AAB96871), A. ficuum (GenBank ID: AAG40885), A. niger

(GenBank ID: BAA74433), A. usamii (GenBank ID: ABA42097.1), A. oryzae (GenBank ID: AAT12504), A. japonicus

(GenBank ID: ACE79228), A. flavus (NCBI ref. no.: XP002376973), Penicillium sp. Q7 (GenBank ID: ABM92788), P. oxalicum (GenBank ID: AAL55406), P. chrysogenum (NCBI ref. no.: XP002561094), Myceliophthora thermophila (GenBank ID: AAB52508), Neurospora crassa (GenBank ID: AAS94253), Trichoderma reesei (GenBank ID: EGR47873), Trametes

pubescens (GenBank ID: CAC48234), Agrocybe pediades

(GenBank ID: CAC48160), Peniophora lycii (GenBank ID: CAC48195).

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and fungi82,83. PAPs harbour a binuclear metal centre of Fe(III)–M(II), where M can be Fe, Zn or Mn. A tyrosine to Fe(III) charge transfer transition imparts PAPs their characteristic purple colour84. The type of bivalent metal ion is dependent on the type species and is Fe2+ in animals and Zn2+ or Mn2+ in plants (a bacterial PAP has not yet been characterized)85,86. Hegeman and Grabau87 isolated PAPs from the cotyledons of germinating soybeans (GmPhy). GmPhy has the active site motif of a PAP. The mol wt of GmPhy (70-72 kDa) is in the range of mol wt of other plant PAPs87. The GmPhy is the only known PAP reported to have significant phytase activity. A. niger NRRL 3135 PAP has been reported to display only a minimum ability to utilize phytate as a substrate88.

Heterologous Expression of Microbial Phytases Heterologous expression systems offer several

advantages in protein expression for industrial applications. The choice of a heterologous host depends on the objective of the study. Among various systems, bacteria (Escherichia coli, Streptomyces sp. & Lactobacillus lactis), yeast (S. cerevisiae & P. pastoris), fungi (Aspergillus spp.), animal cells [insect cell lines, e.g., baculovirus in Autographa

californica (moth); mammalian cell lines, e.g., Chinese hamster ovary (CHO) and baby hamster kidney (BHK) cell line] and plant systems can be used for heterologous protein expression. Microbes are known to produce phytases with broad substrate specificity, but most of them exhibit low specific activities64. In the enzyme industry, low yield and high cost of enzyme production have been constraints in the direct use of theses enzymes in animal diets. With the advancements in heterologous microbial expression systems, large-scale phytase production is now possible at an affordable cost.

Presently in India, dicalcium phosphate (DCP) is used in animal feeds to fulfil phosphorous requirement. The substitution of DCP with phytase could replace 50-60% DCP. It has been estimated that 1 kg DCP can be replaced with 25 g of phytase and, thus, the demand for phytase in animal feed industry could be around 4000 tonnes per year89. In order to reduce the phytase production cost and substituting phytase produced by microbial fermentation with that of plant-produced phytase, a number of attempts have been made to express bioactive phytases in plants90-93. Phytases from different sources have been expressed in various transgenic plants, such as tobacco leaf90, tobacco seed94, alfalfa leaf95, Arabidopsis roots96,97,

rice98,99 and wheat grains100, and many others (Table 1). Fungal and yeast phytases with the desirable properties have also been cloned and expressed in various microbial expression systems (Table 2).

Conserved Amino Acid Sequences in Phytases Primer design is greatly assisted if conserved

sequences and sequences exhibiting similarity to that of known phytase can be identified. Histidine acid phytases have conserved heptapeptide sequence (RHGXRXP) and a dipeptide (HD) in the primary structure of phytase protein. Figs 4 and 5 show the positions of these conserved heptapeptide and dipeptide sequences in fungal and yeast phytases. Acid phosphatases from both prokaryotes and eukaryotes have been shown to share two regions of primary sequence similarity, each centred around a conserved histidine residue110. According to SWISS-PROT protein domain data base, there are two regions: [LIVM]-X(2)-[LIVMA]-X(2)-[LIVM]-X-R-H- [GN]-X-R-X-[PAS] and [LIVMF]-X-[LIVMFFAG]-X(2)-[STAGI]-H-D-[STANQ]-X- [LIVM]-X(2)-[LIVMFY]-X(2)-[STA]. Disulphide bridges increase both stability and heat tolerance in proteins. Mullaney and Ullah126 have reported that fungal histidine acid phosphatases exhibiting phytase activity contained a conserved eight cysteine motif, which appears to participate in disulphide bond formation. Fig. 6 compares the position of eight cysteine residues in A. niger HAP with HAP of other yeast and fungal phytases.

Approaches Used For Amplification of Phytase

Genes The cloning of eukaryotic genes is difficult as

compared to that of prokaryotic genes as most of the former are interrupted by introns, which are removed during RNA processing128,129. The cloning of a eukaryotic gene with an intron will alter the reading frame of the gene and will result in the expression of a non-functional protein. To get rid of introns, the researchers have used different strategies for cloning and expression of eukaryotic genes. Lassen and coworkers70 constructed and screened cDNA libraries of four basidiomycete fungi (Agrocybe pediades, Ceriporia sp., Peniophora lycii & Trametes

pubescens) for the selection of phytase encoding cDNA. The libraries were constructed in E. coli DH10B and then S. cerevisiae was transformed by the recombinant plasmids, and the clones were subsequently screened for phytase activity. Finally all

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Table 1—Yeast and fungal phytases expressed in plants

Source organism Host plant Tissue Vector used Remarks References

Phytases from yeast

Schwanniomyces

occidentalis Rice Leaf pOriF, pModF, pOriT &

pModT Chlorophyll a/b binding protein

(cab) promoter 92

Phytases from fungi

A. niger Tobacco Leaf pTZ117 Expression under control of constitutive CaMV 35S promoter

90

A. niger Tobacco Seed - - 94 A. niger Tobaco Root pPLEX502 CaMV35S promoter and octopine

synthase (OCS) terminator was used

101

A. niger Alfalfa Leaf pTZ117 CaMV35S promoter 95 A. niger Arabidopsis Root (secreted) pART7, pPLEX502 CaMV 35S promoter 96, 97 A. niger Trifolium

subterraneum Root (secreted) - 77-fold increase in exuded phytase

activity in 5 recombinant than wild-type controls

91

A. niger Sesame Root (secreted) pMOG413 CaMV 35S promoter 102 A. niger Soybean Callus

(secreted) pHYG2 Dual-enhanced CaMV 35S promoter

and the seed-specific soybean P-conglycinin a'-subunit

promoter were used

103

A. niger Canola Seed pMOG425 Seed-specific cruciferin (CruA) promoter

104

A. fumigatus Rice Grains pKS1, pCAMBIA 1390 CaMV35S promoter and the CaMV polyadenylation sequence

98, 105

A. niger Wheat Grains pUBARN-1dIII-HI α-Amylase signal peptide sequence was used for secretion of recombinant

protein

100

A. niger Maize Seed pLPL-phyA Murine immunoglobulin leader peptide sequence, rice glutelin-1 seed-specific

promoter

93

A. niger Glycin max Cell-suspension culture

pRTL2 Dual-enhanced CaMV35S promoter and two under the control of a

seed-specific soybean P-conglycinin a'-subunit promoter

103

A. niger Oryza sativa L. variety

Guanglinxiangjin (GLXJ)

Seeds and the leaf

pYF7035 - 106

A. ficuum (niger) potato Leaves - - 107 A. niger, P. lycii Nicotiana tabacum

(var. W38) Roots PLEX502 Expression was under control of

CaMV35S promoter but all the constructs were modified for

extracellular secretion using an extracellular targeting sequence from

the carrot extension (ex) gene

108

phytases were expressed in A. oryzae. Berka et al69

constructed genomic DNA library using bacteriophage cloning vector λZipLox and E. coli Y1090ZL cells. The screening of the library of recombinant plaques was done by plaque hybridization method using the radio-labelled phytase probe. The identified phytase gene was cloned in pDM181 expression vector for phytase expression in

Fusarium venenatum under F. oxysporum trypsin gene promoter. Phytase genes of E. nidulans and Talaromyces thermophilus were also identified by genomic DNA library construction and southern hybridization122. Genomic DNA library was constructed using pBluescriptII KS (-) and E. coli

TG-1 cells and confirmed the presence of an intron in phytase genes by sequence analysis. Xiong et al

125

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Table 2—Heterologous expression of yeast and fungal phytases in microbial hosts

Source of the gene Gene Host used Vector used Temp.Opt. (°C)

pHOpt Specific activity ( U mg-1)

References

Phytases from yeasts

Debaryomyces castellii

CBS 2923

PhytDc P. pastoris X33 pPICZα B & pGAPZα B

60 4–4.5 182 109

Kodamaea ohmeri BG3 Phy1 E. coli BL21 (DE3)

pET24a (+) 65 5 16.5 110

P. anomala Pphy S. cerevisiae,

A. adeninivorans

&

Hansenula

polymorpha,

pYES2.1-V5-His-TOPO, Xplor1-URA3-SwARS-FMD, pBS-TEF-

PHO5

60 4 1.56 111

P. anomala PPHY P. pastoris pPICZαA 60 4 - 112

Phytases from fungi

A. ficuum - E. coli pIAβ 8 & pGAPZαA

50 2.5 - 113

A. ficuum (BCRC 32870) phyA P. pastoris X33 pPICZα A - - - 114 A. fumigatus WY-2 - P. pastoris

GS115

pPIC9K 55 5.5 51 115

A. niger PhyA S. cerevisiae pYEGPD - - - 116 A. niger - S. cerevisiae pYES2 - - - 117 A. niger 113 phyI1 P. pastoris pPIC9 60 2.0, 5.0 9.5 118 A. niger 963 phyA2 P. pastoris pPIC9 - - - 119 A. niger XP - P. pastoris X33 pPICZαA - - - 120 A. niger sp. phyA P. pastoris pPICZα A 55 5.5 503 121 E. nidulans, Talaromyces

thermophilus

- - pBluescript II KS (-)

- - - 122

Neosartorya spinosa

BCC 41923

- P. pastoris

KM71

pPIC9K 50 5.5 38.62 123

Obesumbacterium

proteus

phyA E. coli. pET22b (+) 40–45 4.9 310 124

P. lycii 6-phytase P. pastoris pPIC9K 50 4.5 - 125 P. lycii, Agrocybe

pediades, Ceriporia sp. & Trametes pubescens

- A. oryzae pHD414 - - - 70

synthesized a gene sequence based on nucleotide sequence of the native phytase of P. lycii; 20 oligonucleotide sequences were used to synthesize complete gene in a single PCR reaction. This synthetic gene was subsequently amplified by using end primers and used for expression in P. pastoris. In an attempt to express PhyA gene of A. fumigatus and A. awamori in A. awamori, Martin et al

130

employed pGEXA expression vector. The genomic DNA was used for the amplification of PhyA genes from both the fungi. The mRNA was captured using a poly (dT) column and prepared cDNA by RT-PCR for amplifying the phytase gene of A. niger 113. While Li et al

110 used Rapid Amplification of C-DNA Ends- (RACE-) PCR for amplifying the full-length

phytase gene from Kodamaea ohmeri BG3. For amplification of P. anomala phytase gene (PPHY), specific primers (which were designed based on consensus sequence of phytases) were used to amplify partial gene sequence, which was followed by restriction ligation of genomic DNA to use reverse primers for full length gene amplification111. Heterologous Expression of Fungal and Yeast Phytases

The addition of microbial phytases to food and feed is a usual practice to mitigate environmental and nutritional problems caused by phytates. This practice requires production of high titres of phytase by the microbial sources. Heterologous expression offers solution to this problem by generating recombinant

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Fig. 4—Multiple sequence alignment (MSA) of amino acid sequences of fungal phytases [phytase of Thielavia heterothallica (ID PHYA_THIHE, AC O00107), T. thermophilus (IDvO00096_TALTH, AC O00096), A. niger (ID PHYA_ASPNG, AC P34752), A. fumigatus (ID PHYA_ASPFU, AC O00092)] have been retrieved from Uniprot (Universal Protein Resource database, website: http://www.uniprot.org). The conserved heptapeptide and dipeptide are shown in black boxes.

microbes capable of producing very high enzyme titres as compared to their wild type counterparts. Watanabe et al

131 reported that a recombinant Pichia strain could produce 23 g L-1 of recombinant phytase. The yield of A. fumigatus phytase expressed in Pichia was 729 mg L-1 of purified protein, exhibiting specific activity of 43 U mg-1 132. P. pastoris offers several advantages over other microbial expression hosts, such as, genomic integration of the transgenes, availability of expression vectors where cheap methanol can be used both as inducer and carbon source, no requirement of antibiotic once the stable integrant has been constructed and very high culture densities can be achieved for high protein

expression133. This budding yeast offers utilization of various alternative promoters, such as, pAOX1, pFLD1, pEPEX8, pYPT1, pDHAS, pICL1, pTEF and pGAP, which are regulated by the carbon sources incorporated in the production medium134-136 and a few, such as, pFLD1, are regulated by nitrogen source137.

Apart from microbial production of phytases, the recombinant plants harbouring phytase gene offer an added advantage as these genetically modified plants can be used as feed material without any need to add phytase from external source. Maize seeds are a major ingredient of commercial feed used in pork and poultry138. Recombinant maize has been produced by

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Fig. 5—Multiple sequence alignment of amino acid sequences of different yeast phytases [phytases of S. cerevisiae FostersO (GenBank Acc. no.: EGA63258.1), S. capriottii (GenBank Acc. no.: ABN04184.1), D. castellii (PDB: 2GFI_A), Cyberlindnera fabianii (GenBank: BAH58739.1), Wickerhamomyces anomalus (GenBank: FN641803.1) Yarrowia lipolytica CLIB122 (GenBank: CR382129.1)]. Conserved heptapeptide and dipeptide sequences are shown in black boxes.

a group of Chinese researchers that produces phytase 2200 U kg-1 of maize seeds138. This titre was 50-fold higher than the native maize plant and was stable across at least four generations138. Similarly the construction of a recombinant microalga Chlamydomonas reinhardtii has

been reported producing phytase of E. coli (AppaA)

in its chloroplasts139. Microalgal cell hydrolysate can directly be used as feed additive. It has been observed that it contains 10 U g-1 dw. Upon feeding recombinant microalga based diets to young broiler chicks, the excretion of faecal phytate was reduced with increase in inorganic phosphate139.

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Biochemical and Molecular Attributes of

Recombinant Phytases

pH and Temperature Optima The phytase of A. niger 113 expressed in

P. pastoris exhibited two pH optima, at pH 2.0 and pH 5.0118. Similarly, in another study, PhyA of A. niger had two optimal peaks at pH 2.5 and 5.5, and it was optimally active at 60°C140. However, the optimum pH and temperature for A. fumigatus phytase were 5.5 and 60°C, respectively132. Further, the phytase of Debaryomyces castellii CBS 2923 expressed in P. pastoris exhibited pH optima of 4.0, similar to the native phytase141. This phytase was optimally active at 60°C. The recombinant phytase (PHY1) of K. ohmeri BG3 was optimally active at pH 5.0 and 65°C110; while the recombinant phytase of P. anomala was optimally active at pH 4.0 and 60°C like that of the native phytase111. On the other hand, the phytase coexpressed with a xylanase in P. pastoris was optimally active at 55°C and pH 5.0142.

Kinetic Characteristics of Recombinant Phytases

The recombinant phytase of P. anomala expressed in Hansenula polymorpha has Km and kcat of 0.25 mM (phytate) and 18133 s-1, respectively111. While the phytase of A. fumigatus expressed in P. pastoris displayed Km of 1.38 mM for pNPP and 2.84 mM for sodium phytate132. Similarly high values of turnover rate (kcat) and substrate specificity constant (kcat/Km) for pNPP have been reported for A. fumigatus

recombinant enzyme. Recently Joshi and Satyanarayana112 reported heterologous expression of P. anomala phytase in P. pastoris. The recombinant phytase exhibited high substrate specificity for calcium phytate as evident by its low Km (0.2 mM) with Vmax of 78 n moles mg-1 s-1. Molecular Mass and Glycosylation of Recombinant Phytases

Molecular mass of the recombinant A. niger 113 phytase was 66-80 kDa121, which was larger than the actual molecular mass of A. niger 113 phytase. The molecular mass of the recombinant enzyme was more compared to the native due to glycosylation141. Ragon et al

140 expressed D. castelli CBS 2923 phytase in P. pastoris and reported difference in glycosylated and deglycosylated recombinant phytases. Phytase phyA2 expressed in P. pastoris had a molecular mass of 85 kDa and on deglycosylation, the molecular mass changed to 64 kDa119. Similarly, Xiong et al

125 reported that molecular mass of the recombinant phytase of P. lycii ranged between ~70 kDa and ~110 kDa, and upon deglycosylation, its molecular mass on SDS-PAGE was 60 kDa. Deglycosylation reduced molecular mass of phyA of A. niger from 95 to 55 kDa141. Li et al

110 found that phytase gene ORF of K. ohmeri BG3 consisted of 1389 bp. The calculated molecular mass of this phytase was 51.9 kDa, which had a putative signal peptide of 15 amino acid residues. When Li et al

110 expressed this phytase in E. coli, the molecular mass of the recombinant

Fig. 6—Conserved cysteine moieties in A. niger NRRL 3135 phyA [AN] and non-filamentous ascomycetes HAPs (A) and basidiomycetes HAPs (B). NCBI locus number of the sequence and microbe are: (A). [CA] EAK94299, C. albicans; [DO] CAB70441, D. occidentalis; [Pho1] NP596847, Schizosaccharomyes pombe pho1; [Pho4] NP595181 S. pombe pho4; [Pho3] CAA85045, S.

cerevisiae pho3. (B). [AP] CAC48160 Agrocybe pediades; [CER] CAC48164 Ceriporia sp. CBS 100231; [PL] CAC48195, P. lycii; and [TA] CAC48234, Trametes pubescens. Position of the1st aa of each sequence is denoted in parentheses. The * denotes the total number of cysteines in the sequence. AA indicates the number of amino acids in each sequence.

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phytase was 51 kDa that differed from that of the native phytase. The molecular mass of native K.

ohmeri BG3 phytase was 92.9 kDa, which can be attributed to many N-glycosylation sites present in the primary sequence of the protein as well as other processes during secretion. Monomeric recombinant phytase from P. anomala exhibited molecular mass of ~ 70 kDa, while deglycosylated form exhibited a molecular mass of ~ 53 kDa. Thus, the excess molecular mass is due to N-linked glycans112.

Thermostability

A. niger 113 phytase expressed in P. pastoris retained 70% residual activity after 5 min heating at 90°C, while the recombinant enzyme exhibited 75.2, 74.1 and 68.8% residual activities after 15 min exposure to 80, 85 and 90°C, respectively121. Synthetic 6-phytase gene of P. lycii, which was heterologously expressed in P. pastoris, retained only 25% of its initial activity after 10 min exposure to 80°C125. Han et al

141 reported that the deglygosylated PHY1 of K. ohmeri BG3 retained 93% residual activity after the treatment at 60°C for 1 h, and it lost total activity upon treatment at 80°C for 1 h110. A phytase (PhyA-2A) coexpressed with xylanase showed optimum activity at 55°C. PhyA-2A retained 70% residual activity after exposure to 90°C for 5 min142. Thermostability of recombinant phytase of A. fumigatus was dependent on pH and buffer used as it exhibited higher thermostability at 65 and 90°C, when acetate buffer was used as compared to that in citrate buffer132.

Resistance to Proteolytic Action

In in vivo conditions, the actual functional site of phytase is the stomach143,144, where pH instantly increases to 5.5 just after food intake and, thereafter, it decreases to pH 2.0. Pepsinogen is secreted from stomach cell lining, which activates pepsin by the release of HCl in the stomach. An ideal phytase used as a food and feed additive needs to possess a strong resistance to proteolysis by digestive tract proteases. Additionally, it must retain high enzymatic activity at the body temperature and low pH145. Phytases from fungal and bacterial origin exhibit different levels of resistance to trypsin and pepsin146,147, and the former appears to have a higher resistance to proteolysis than the latter148. Site-directed mutagenesis has been used to block protease-sensitive regions of phytases, which are normally found in exposed loops at the molecular surface64. In vitro studies indicated phytases from A. niger (r-PhyA) and E. coli (r-AppA) to exhibit

differential resistance to trypsin and pepsin149. In the presence of pepsin or pancreatin, A. niger phytase was more stable than the wheat phytase150. The recombinant phytase from A. fumigatus was found to be resistant to proteolysis132. When treated with gastric enzymes, trypsin and pepsin at pH 2.5 and 37°C and at different ratios of digestive enzymes and phytase (0.001 to 0.1), a partial loss of phytase activity was observed 132. On treatment with higher ratios of proteolytic enzymes, the phytase activity was lost.

Crystal Structure of Phytases

Crystal structures of phytases from different sources have been solved. The crystal structure of a protein can reveal unexplored truths about functioning of the proteins and give answers for their special biophysical characters. The phytases from A. niger and A. fumigatus have 66% sequence identity, but they differ in their thermostability151; this was attributed to the removal of repulsive ion pair interaction due to substitutions with polar residues along with the formation of hydrogen bonds. Apart from hydrogen bonding, a helical capping at C-terminal also contributed to the stability of A. fumigatus phytase151. Crystal structure of A. fumigates phytase was solved at 1.5 Å resolution. Electron density experiment revealed the presence of six predicted N-glycosylation sites in the phytase. Xiang et al

151 observed a partially phosphorylated His 59 residue in this phytase that indicated two step catalytic mechanism.

Based on the crystallographic studies, thermostable and thermolabile enzymes can be modified in vitro by rational design. Phytases from different microbial sources have been crystallized. As far as fungal and yeast phytases are concerned, only one yeast phytase and two fungal phytases have been crystallized (http://www.rcsb.org/pdb); the yeast phytase comes from D. castellii CBS 2932152. Crystal structure of the phytase from D. castellii (PhytDc) was solved at 2.3 Å resolution that belongs to space group P6522. The three dimensional structural analysis revealed that this is a dimer containing five NAG molecules along with 628 water molecules. D. castellii CBS 2923 phytase (PhytDc) is a glycoprotein that exhibits thermostability up to 65.85°C, which cleaves six IP6-bound phosphate groups140. The PhytDc belongs to histidine acid phosphatases since it contains the characteristic sequence motif RHGXRXP. The structure of PhytDc can be divided into 2 parts: a small α-domain and a large α/β-domain with a

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6-stranded β-sheets140. The primary amino acid sequence of PhytDc exhibits 36% identity with acid phosphatase of A. niger (PAAn), 25 % with the phytase A of A. niger (PhytAn) and 23% with phytase of A. fumigatus phytase A (PhytAf)140. The N-terminal regions of PhytDC and PAAn are extended (Fig. 7A), which are involved in dimer and tetramer formation. A consensus phytase designed based on 13 fungal phytases have also been crystallized153 (Fig. 7B). This consensus phytase exhibits normal catalytic activity but unexpectedly shows 15-22°C increase in unfolding temperature; its structure was resolved at 2.9 Å. The substrate specificity of this novel consensus phytase is similar to that of A. fumigatus and E. nidulans

153. Lehman and coworkers153 observed stabilization of a surface loop in the consensus phytase, which is one of the highly variable phytases but exhibits consensus sequence as T249, S250, D251, A 252, T253. Liu et al

154 studied the crystal structure of A. fumigates phytase both at pH 5.5 and 7.5. The A. fumigatus phytase structure was resolved at higher than 1.7 Å. The presence of a larger catalytic pocket explains the broader substrate specificity of A. fumigatus phytase than to that of E. coli. Liu and coworkers observed three water molecules at the substrate binding site, which were considered to be responsible for pH dependent activity of A. fumigatus phytase (Fig. 7C).

Oakley et al155 have crystallized both native and

partially deglycosylated A. niger phytase (PhyA) in complex with potent inhibitor myo-inositol-1,2,3,4,5,6-hexakis sulfate. The structural analysis

revealed that PhyA prefers 3′-phosphate group of phytic acid. The structure of A. niger phytase consisted of an α/β-domain and α-domain and one extension at N-terminal. Out of the eight expected glycosylation sites, only four sites (N82, N184, N316 & N353) were observed with N-acetylglucosamine residues. Other than minor shifts in the orientation of carbohydrate bearing asparagine residues, no major structural differences were observed in fully and partially glycosylated PhyA155. DALI search resulted in identification of structural homologues of PhyA, which included PhyA of A. fumigatus, PhytDc of D. castellii CBS 2923, a phosphatase from A. niger, acid phosphatises of human and rat acid glucose-1-phosphatase (Agp) 3-phytase (AppA) of E. coli. This information provides insight into important regions of catalysis and substrate binding155. Molecular Approaches for Ameliorating Properties of

Phytases

Although various phytases have been reported from different microbial sources, none of them possess all the properties of an ideal phytase. An ideal phytase suitable as food and feed additive should be thermostable to withstand the high temperature during feed pelletting process, and should be adequately acid-stable and protease resistant to withstand high acidic pH of the stomach and proteases, respectively. Biotechnologists have tried to tailor phytases for the acquisition of important properties. Kim et al

156 introduced mutations in the phytase of A. niger and could identify one mutein, which exhibited shift in the

Fig.7(A-C)—3-D structures of yeast [A, adopted from ref. 152] and fungal phytases [B & C, adopted from ref. 153 and 154]. [A] Cartoon structure of D. castelli phytase (blue colour) overlapped with A. niger (grey colour) phytase sequence where their N-termini are coloured pink and cyan, respectively. [B] Consensus phytase sequence. [C] Reaction product phosphate groups attached to A. fumigatus active site.

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pH optima towards acidic range. Similarly mutagenesis was used to increase the thermostability of phytase of A. niger by Lehman and colleagues153. They increased thermostability by 7°C along with other favourable changes. A few mutant phytases have made their way to market. An E. coli phytase Phy9X, a product of genetic engineering, has exhibited improved properties and made its way to the market. This enzyme is being sold under the brand name of QuantumTM 605157. In one of the mutagenesis studies of phytases, Mullaney et al

158 performed mutation in A. niger phytase for alteration of disulphide bridges. Five disulphide bridges (DB) present in the phytase were subjected to mutation forming cysteine residues, and heterologous expression of the mutated gene was carried out in P. pastoris. The effect of alteration of every DB was studied in terms of Topt, pHopt and substrate specificity (Km). Alteration in DBs completely abolished the activity, while altering disulphide bond 1, 3/4 and 5 lowered the Topt to 53, 37 and 42°C, respectively. The pH profile of the mutated phytases was changed in such a way that a peak at 2.5 and 5 was abolished on removing DB at 1, 3 and 5. The Km of mutant phytases did not change significantly, but the turnover of the muteins was significantly lowered.

Utility of Recombinant Phytases in Dephytinization

Last few decades have witnessed increased utilization of microbial phytases in the field of animal nutrition, where plant based diets are used. Phytases have also found application in various other fields, such as, aquaculture159, agriculture160 and environmental protection145. The recombinant phytases have also made their entry for these applications. As mentioned earlier, Quantum TM 605 is one of the recombinant phytases available in the market153. Several other unaltered microbial phytases are also sold in the market under different brand names, such as, Finase (source: A. awamori; production strain: Trichoderma reesei), SP (source and production strain: A. oryzae), Allzyme phytase (source and production strain: A. niger), Natuphos (source and production strain: A. niger), AMAFERM (source and production strain: A. oryzae), Bio-feed Phytase (source: P. lycii; production strain: A. oryzae), Phyzyme (source and production strain: A. oryzae), Avizyme (source: A. awamori; production strain: T. reesei), ROVABIO (source: Penicillium

simplicissimum; production strain: P. funiculosum), Roxazyme (source and production strain:

A. oryzae)161-163. Phytic acid complexes with carbohydrates164, proteins165 and other dietary elements166-168 alter their properties. Joshi and Satyanarayana112 have demonstrated dephytinization of phytic acid complexed with soy proteins by recombinant phytase of P. anomala (rPPHY). Two of the soy protein components, which are known to be highly allergenic in nature, exhibit differences in biophysical properties when they are dephytinized. Utilizing this observation, they have successfully fractionated β-conglycinin from glycinin. By using rPPHY in whole wheat bread preparation, reduction in phytic acid content up to 75% has recently been reported169. A recombinant β-propeller phytase from B. licheniformis exhibited digestion efficiency for commercial feed and soybean meal170. Sanz-Penella et al

171 have shown the utilization of microbial phytase in dephytinizing infant cereals for the first time. This observation suggests the potential use of phytases in human nutrition too.

Future Perspectives

The worldwide interest on phytases has led to the development of recombinant microbial phytase producers and plant bioreactors. Being a low value and high volume product, low cost of phytase production is required. Many of the native phytases lack either one or the other requisite property. The in vitro manipulations in microbial phytases and recombinant DNA technology have led to the generation of phytases with improved properties. A phytase with all the desirable characteristics, such as, thermostability, acid stability, protease insensitivity, adequate activity at animal body temperature and high specific activity in a single phytase is still a challenge. Phytases have been explored for their utility as a food and feed additive, but other applications, such as in soil amendment, environmental protection and enzymatic synthesis of lower inositol phosphate isomers are yet to be adequately explored.

Conclusion

The reduction of phytates in foods and feeds is of a major concern for monogastrics like humans, pigs, poultry birds and fishes because they mainly depend on the plant based diets and do not produce adequate levels of phytate-degrading enzymes in their alimentary tracts. As phytates act as anti-nutrients, the reduction in phytates is an essential step in the processing foods and feeds. Although a number of yeast and fungal phytases have been cloned and

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expressed in different heterologous systems, the requirements for an ideal phytase have not yet been fulfilled. A few attempts have also been made to evolve phytases with the desirable characteristics by site-directed mutagenesis or directed evolution. Further developments in microbial phytases are needed in order to evolve an ideal phytase.

Acknowledgements We wish to thank the University Grants

Commission (UGC) and Department of Biotechnology, Government of India, New Delhi for the financial assistance and to Council of Scientific and Industrial Research (CSIR), New Delhi for awarding a fellowship to SJ, while writing this review.

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