Current research and future perspectives of phytase bioprocessing

Abstract

Phosphorus is one of nature's paradoxes as it is life's bottleneck for subsistence on earth but at same time is detrimental in surplus quantities in an aquatic environment. Phosphorus cannot be manufactured, though fortunately it can be recovered and reused. The only way to avert a supply crisis is to implement the “3R's” of sustainability, “Reduce, Reuse and Recycle.” Phytase is likely to play a critical role in the dephosphorylation of antinutritional and indigestible phytate, a phosphorus locking molecule, to digestible phosphorus, calcium and other mineral nutrients in the coming years. Hence, efforts are required to produce cost effective phytase with fast upstream and economic downstream processing because the current available process is expensive and time consuming. This review summarizes the present state of methods studied for phytase bioprocessing. Production, extraction and purification incur a large cost in product development. In addition, the process has several limitations such as dilute concentration of enzyme, extensive downstream procedures and treatment of generated effluents. However, these methods are currently employed due to lack of alternative methods. Thus, there is a clear need for an efficient, scalable and economical process for phytase production and bioseparation, and improvements are especially required with regard to yield, purity, and energy consumption. Perspectives for an improved bioprocess development for phytase are discussed based on our own experience and recent studies. It is argued that the optimization of production techniques, strain improvement and liquid–liquid extraction deserves more attention in the future.

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1. Introduction

The biogeochemical cycling of nonrenewable and biocritical element phosphorus (P) is a very slow process in nature.¹ This vital mineral is important for bone and tissue growth, and is therefore the third most expensive component in poultry production subsequent to energy and protein. Despite its low abundance, the importance of P in biological systems is lucid. P reserves are present in few regions with others entirely dependent on import. India is the largest consumer of phosphate fertilizers and the demand continues to increase due to rising population and escalating demand for meat rich diets and bioenergy crops.²

Plants store P in the form of phytate (inositol 6-phosphate) carrying 6 phosphate groups. However, this bound P (60–70%) present in seed grain as phytate is unavailable to mono-gastric animals because they lack intrinsic phytase activity. Phytate being negatively charged chelates metal ions, reduces energy uptake and behaves as an antinutrient.³ Because P is an important macronutrient for growth, animal diets are customarily supplemented with surplus quantities of inorganic P supplements, which ultimately lead to nutrient enrichment in water bodies causing eutrophication. Thus, although P is a biocritical element, it is also a pollutant for living beings. The modern P cycle is atypical due to intertwined agricultural and human activities, which affect the ecosystem structure with impacts that are detrimental and hard to rescind.⁴ Only 10% of phosphorous is utilized in food production, while 90% is lost due to resource mismanagement. Measures for closing the loop in the broken P cycle involve strict legislation and norms for the discharging of P effluents, human interference and decomposition of underutilized phytate. However, at the current usage and extraction, a price hike in synthetic fertilizers is inevitable. These factors have currently led to the use of microbial phytase in animal feed.⁵

Use of phytase in animal feed will seize the anti-nutritional effects of phytate, decrease environmental pollution, increase availability of starch, protein, amino acids, calcium and P, and abolish the surplus addition of inorganic phosphate in animal feed. Phytases are also imminent candidates for the production of special isomers of different lower phosphate esters of myo-inositol, some of which are considered to be pharmacoactive and important intracellular secondary messengers.⁶

The FDA (The Food and Drug Administration) has approved a “generally recognized as safe (GRAS)” petition for the use of phytase in food, and it has been marketed as an animal feed enzyme in the US since 1996. All these factors have made P the third largest feed enzyme. Although a limited number of phytases have been reported and studied, our understanding of phytases is yet to provide a solution that meets the nutritional and environmental requirements of the real-world demands. The major hurdles hindering the exploitation of the repertoire of enzymatic processes are, in many cases, high production costs and low yields. Several reviews on phytase have focused on production, biochemical characteristics, biotechnological applications, crystal structure, directed evolution and protein engineering. This review describes the state of the art for upstream and downstream processing of phytase and its application. Upstream processing includes the type of fermentation, choice of strain, improvement of strain or process and bioreactor design followed by downstream processing, which involves separation, purification and formulation of the end product (Fig. 1).

Fig. 1 Phytase bioprocessing and applications.

1.1 Phosphorus paradox

Phosphorus (P), a nonmetal element from the nitrogen group (group 15) of the periodic table, is not found as a free element in nature due to its high reactivity. It is essential to all known life forms and is the second most abundant mineral present in the human body, surpassed only by calcium. P is life's bottleneck but ironically due to mismanagement and inadequate legislative norms it acts as pollutant, resulting in eutrophication leading to algal blooms (Fig. 2). Excess/less phosphate also leads to diarrhoea and calcification (hardening) of organs and soft tissue, hypophosphatemia, osteomalacia, anorexia and pica. Depleting P reserves (Peak P) by 2030 are suggested to occur due to the depletion of current high-grade reserves eventually increasing the cost of phosphate rock by 800% in 2008.⁷

Fig. 2 Phosphorous paradox.

P is a nonrenewable resource and cannot be produced, re-grown or regenerated, although fortunately, unlike oil, it can be recovered and reused over and over again. The global supply of phosphate rock is depleting at an alarming rate. This situation has many similarities with oil, yet unlike oil, there is no substitute for P in food production.⁸ Some developing countries, especially India which is the largest consumer, are entirely dependent on P import for food production (Fig. 3A). While all farmers need access to P, just 5 countries control around 90% of the world's remaining phosphate rock reserves; these countries include China, US and Morocco (which also controls the reserves of western Sahara) (Fig. 3B).⁹

Fig. 3 (A) World phosphate fertilizer consumption (% increase 2012). (B) World phosphate reserves.

Phosphate rock is one of the most highly traded commodities on the international market, and its crushed/processed fertilizer is generally used for food production. Phosphogypsum is a toxic, radioactive byproduct of P processing, which is a future threat to ground water contamination. Crushed/unprocessed P rock contains uranium and thorium, which contribute to soil radioactivity and is currently been done in European countries, India (largest P consumer) and Australia.¹⁰ There is a need of 3R's, i.e. Reduce, Reuse and Recycle, for maintaining the sustainability of P for future generations. The abovementioned reasons raise concern regarding the depleting phosphate reserves, thus current research should be directed toward reusing and recycling P. Phytase can provide an alternative option to reduce the use of phosphorous by hydrolyzing phytate, the P locking molecule.

1.2 Phytate

Phytate is the principal storage form of P, inositol, and a variety of minerals in plants, representing approximately 75–80% of the total P in plant seeds. Phytic acid bears six phosphate groups on one six-carbon molecule with a low molecular weight of 660 and molecular formula of C₆H₁₈O₂₄P₆. On the basis of Anderson's structure,¹¹ the systematic name for phytic acid is myo-inositol-1,2,3,4,5,6-hexakisphosphate. Phytate-P represents 50–82% of total P in cereals and oilseed meals.¹²

Phytate can exist in a metal-free form and in metal–phytate complex at acidic and neutral pH, in which the latter form binds with divalent metal cations, mostly Mg²⁺ and Ca²⁺.¹³ Table 1 presents an overview of the negative interactions of phytate with nutrients and the modes of action for the negative effects of phytate. The bioavailability of P and cations (Ca²⁺, Fe²⁺, Zn²⁺ and Mg²⁺⁾ is reduced due to phytate, which is a P locking molecule and a chelator. The after effects of unutilized phytate, i.e. eutrophication and algal blooms are more appalling.¹⁸

Table 1 Negative interaction of phytate and nutrients in food

Nutrients	Mode of action
Mineral ions (zinc, iron, calcium, magnesium, manganese and copper)	Formation of insoluble phytate–mineral complexes decreases mineral availability^14
Protein	Formation of nonspecific phytate–protein complex, not readily hydrolysed by proteolytic enzymes^15
Carbohydrate	Formation of phytate carbohydrate complexes making carbohydrate less degradable. Inhibition of amylase activity by complexing with Ca^2+ ion and decrease in carbohydrate degradation^16
Lipid	Formation of ‘lipophytin’ complexes may lead to metallic soaps in gut lumen, resulting in lower lipid availability^17

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Phytate hydrolysis is either enzymatic or non-enzymatic wherein the latter occurs under high temperature conditions. Phillippy et al.¹⁹ studied the hydrolysis of phytic acid and reported that at pH 1.0, 2.0, 4.0, 6.0, 8.0, and 10.8, the phytate decomposed up to 67.7, 76.8, 89.6, 81.9, 65.8, and 45.1%, respectively. Enzymatic phytate hydrolysis of phytase occurs by the sequential release of orthophosphate groups from the inositol ring of phytic acid to produce free inorganic P along with a series of intermediate myo-inositol phosphates (inositol pentaphosphate to inositol monophosphate). Phytase not only releases P from plant-based diets but also makes calcium, magnesium, protein and lipid available. Thus, by releasing bound P in the feed ingredients of vegetable origin, phytase makes more P available for bone growth and protects the environment against P pollution.²⁰

1.3 Phytase

In recent years, considerable efforts have been made to improve the nutritive value of animal feedstuff through supplementation with exogenous enzymes. The global market for feed enzymes is a promising segment in the enzyme industry. This market was estimated to be around $344 million in 2007, and is expected to reach $727 million in 2015. Currently used feed enzymes are divided into two main groups, the hemicellulases and phytases. Phytases (myo-inositol hexaphosphate phosphorhydrolase) hydrolyze phytic acid to myo-inositol and inorganic phosphates through a series of myo-inositol phosphate intermediates and eliminate its anti-nutritional characteristics.

Four sources, namely, plant phytase, microbial phytase (fungal and bacterial phytase), phytase generated by the small intestinal mucosa and gut-associated micro floral phytase are generally reported. However, the phytase activity of animals is negligible compared to their plant and microbial counterparts.²¹ Most of the scientific work has been conducted on microbial phytases, especially on those originating from filamentous fungi such as Aspergillus ficuum, Mucor piriformis and Cladosporium species. Although some plants, such as wheat and barley, are rich in intrinsic phytase because of a narrower pH spectrum of activity and low heat stability, their phytase activity is less effective than microbial phytases. In addition, the bio-efficacy of plant phytases is only 40% compared with that of microbial phytases. The International Union of Biochemists²² currently distinguishes between three classes of phytase enzymes depending on the position (3, 6 or 5) on the inositol ring where the dephosphorylation is initiated, as shown in Fig. 4.

Fig. 4 Classification of phytase.

However, there are some exceptions such as soybean phytase is a 3-phytase²³ and Escherichia coli phytase is a 6-phytase.²⁴ Histidine acid phosphatase (HAP) shows broad substrate specificity and hydrolyzes metal-free phytate at acidic pH, and produces myo-inositol monophosphate as the final product. Alkaline phytase exhibits strict substrate specificity for calcium–phytate complex and produces myo-inositol triphosphate as the final product. Alkaline phytases are not a subfamily of HAPs but are novel phytases, as can be seen in Table 2. Despite considerable differences between alkaline phytases and HAPs, only limited knowledge on the biochemical and catalytic properties of alkaline phytases is currently available. More focus has been given to acidic phytases because of their applicability in animal feed and broader substrate specificity than those of alkaline phytases. On the basis of their catalytic properties, phytases are classified as HAP, β propeller phytase (BPP), and purple acid phosphatases (PAP).²⁵ The finger print of phytases and relationship between motif and key active amino acids were investigated using MEME. It was found that plant phytases have a distinct mechanism for phytate utilization as compared to animals and microbes.²⁶

Table 2 Recombinant System for phytase

System	Advantages	Limitation
Plants	A. niger phyA gene successfully expressed in tobacco seeds or leaves and soybean cells	Thermostability is a major concern^49
Yeast	Conducted heterologous gene expression of bacterial and mold phytases in yeast expression systems	Few yeast phytases expressed^50
Bacteria	Inactive A. niger PhyA protein expressed intracellularly in E. coli and extracellularly in Streptomyces lividans	Glycosylation is a major concern with bacterial system to produce fungal phytase^51
Fungi	Phytase genes from A. niger, A. terreus, A. fumigatus, E. nidulans, and M. thermophila have all been expressed and secreted as active enzymes by A. niger	Fungal systems secrete active phytases but along with high level of undesired proteases. This requires further purification or inhibition of proteolysis that adds to the production cost

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1.4 Market trend and manufacture

Recent market trends have clearly shown that enzymes have emerged as big feed supplements. Feed enzymes (protease, xylanase, phytase, amylase, cellulase, lipase, β-glucanase) are the newest segment of the $5 billion animal nutrition market, which is growing fast. Currently, only about 6% of manufactured animal feed contains enzymes, against 80 ± 90% of vitamins, which is considered as the largest animal nutrition category. Gist Brocades introduced the first phytase product in the feed market in 1991, which is currently known as Natuphos and is available as a powder, granulate, or liquid.

Only a few of the later products introduced from different companies are available as phytase preparations due to varied properties and efficacy (Fig. 5). The first phytases produced on a commercial scale were either derived from fungal strains mutated via standard procedures or by recombinant DNA technology.²⁷ However, effectiveness of these phytase supplements is low because of the lack of essential characteristic, and thus the quest for an ideal phytase continues. Phytase with desirable characteristics for application in the animal feed industry can be called an ‘ideal phytase’, which should be active in the stomach, stable during animal feed processing and storage, and easily processed by the feed manufacturer for its suitability as an animal feed additive. It should satisfy the following points:

Fig. 5 Development of phytase research.

1. Phytase should not be detected at the end of the small intestine. This is necessary because in this manner the phytase, which is produced by genetically modified organisms does not enter the environment.

2. It should be effective in releasing phytate-P in the digestive tract.

3. It should be stable to proteases (trypsin and pepsin).

4. It should be able to resist inactivation by heat during feed pelleting and storage.

5. Low cost of production.

Finally, phytase produced in high yield and purity by a relatively inexpensive system is attracting food industries worldwide. It is now realized that any single phytase may never be ‘ideal’ for all feeds and foods. For example, the stomach pH in finishing pigs is much more acidic than that of the weanling pigs.²⁸ Thus, phytase with optimum pH close to 3.0 will perform better in the former than in the latter. For poultry, an enzyme would be beneficial if it is active over a broad pH range, that is, acidic (stomach) to neutral (crop).²⁹ Phytases used for aquaculture applications require a lower temperature than those used for swine or poultry.³⁰ Therefore, the choice of an organism for phytase production and development is dependent upon the target application using directed evolution and protein engineering. All these features are not present within a single phytase, and therefore, based on the sequence of the available phytases, a consensus phytase could be designed.³¹ Genetic engineering techniques such as site directed mutagenesis could be employed for further ameliorating the properties. The strategies used for the designing and developing of an ideal phytase are as follows:

1. Immobilization of phytase for application in food, feed, pharmaceutical industry and biosensor.

2. Modification of active site for enhanced thermostability and efficient catalysis of phytase by incorporating vanadium in the active site for peroxidase activity.

3. Site directed mutagenesis for enhanced phytase thermo stability and protease resistance.

4. Transgenic expression in plants and animal for improving their nutrition and growth.

5. Protein engineering of phytase for enhanced thermostability and pH stability.

6. Scale up for economical and large scale phytase production.

7. Understanding the role of glycosylation in phytase stability.

2 Microbial production of phytase

2.1 Screening and assay

Several screening programmes have been carried out aiming at the isolation of the different groups of bacteria, yeast and fungi having extracellular phytase activity. Lissitskaya et al.³² screened microorganisms producing phytase using museum and soil samples, wherein it was found that moulds metabolized P more effectively than bacteria. Chen developed a bioassay method using the washed cells of Corynebacterium glutamicum as an indicator strain for the screening of extracellular phytase producing microorganisms.³³ Gargova et al. used a two-step procedure to screen 200 fungi producing phytase.³⁴ A simple and rapid method was described for determining microbial phytase by evaluating the inorganic orthophosphate released on the hydrolysis of sodium phytate at pH 5.5.³⁵ Bae et al. developed a method for detecting phytase activity on differential agar media, in which the disappearance of precipitated calcium or sodium phytate was an indication of enzyme activity.³⁶ However, this technique was unable to differentiate between phytase activity and acid production by ruminal bacteria.

The abovementioned assay is performed with phytate as a substrate and degradation of phytic acid to the amount of P released. However, the phytase screening media and assay has limitations. The traditional endpoint assay is time-consuming and is well known for its cumbersomeness in addition to its requirement of extra caution for the handling of toxic regents. However, this method does not give a very detailed picture of the actual mechanism of phytase action and other methods including chromatographic separation followed by quantification of the lower inositol phosphates are therefore sometimes employed making it time consuming.

Phytase kinetics is highly dependent on substrates and reaction conditions, making kinetic investigations of genuine substrates at physiologically relevant conditions an important concern. Thus, a simple, fast and nontoxic kinetic method was developed by Tran et al. for high throughput assaying of phytase, overcoming the limitations of traditional phytase assay methods. This assay is based on the principle that IP₆ forms stable turbid complexes with positively charged lysozyme in a wide pH range, and the hydrolysis of IP₆ in the complex is accompanied by a decrease in turbidity monitored at 600 nm.³⁷

2.2 Production technique

Phytase can be produced from a host of micro-organisms including bacteria, yeasts and fungi, and submerged (SmF) as well as solid state fermentation (SSF) have been employed for the production of phytases. SmF has largely been employed as a production technology for commercial phytases. However, in recent years solid state fermentation (SSF) has gained significant interest for the production of phytase. The type of strain, culture conditions, nature of the substrate and availability of the nutrients are critical factors affecting the yield and should be taken into consideration when selecting a particular production technique. For example, a filamentous fungus in SmF is exposed to hydrodynamic forces but in SSF the surface of the solid particles acts as the matrix for the culture.

Phytase production has been studied under SmF and SSF; and previous studies have reported that enzymatic production under SSF has several advantages in comparison to that of SmF. Varied substrates, such as wheat bran full-fat soybean flour, canola meal, cane molasses and oil cakes, are studied. Among them, the higher titers of enzyme production, extracellular nature of enzyme, and low protease production are observed.³⁸ SmF is the method of choice for phytase production due to ease of SmF operation, up scaling and less variability.³⁹

Several studies have compared phytase productivity values in different fermentation systems to explain how the fermentation system affects fungi physiology. In such comparisons, important aspects such as medium composition, A. niger morphology, and phytase production diffusion of nutrients, growth patterns, titers of enzymatic productivity culture conditions, type of strain, and nature of substrate have been included.⁴⁰ SmF and SSF processes have been compared for their suitability in Bacillus subtilis US417 phytase production.⁴¹

The effect of light on fungal growth on solid media culture may also act as an index for mycelia fermentation. Understanding the effect of light on mycelia growth on plates may provide important information in working cultures, which are liquid cultures for the homogeneous growth of the fungus and the solid culture of photo fermentations. Examining the density and shapes of mycelia on plates would save time and reduce the costs of media selection, working culture and solid culture.⁴² Complexity in gene regulation helps an organism to grow in adverse conditions but at the same time this presents both problems and opportunities.⁴³ However, there is a complex relationship between the morphology of these microorganisms, transport phenomena, viscosity of the cultivation broth, and related productivity. The morphological characteristics vary between the freely dispersed mycelia and the distinct pellets of aggregated biomass with every growth form having a distinct influence on broth rheology. Hence, the advantages and disadvantages for mycelial or pellet cultivation need to be balanced out carefully. Because of the inadequate understanding of the morphogenesis of filamentous microorganisms, fungal morphology is often a bottleneck of productivity in industrial production.⁴⁴ There is abundant proof in the literature that the product spectrum from SSF is very different from that obtained in SmF. However, the mechanisms underlying these differences are not understood. Therefore, rational new designs of SSF processes to prepare new products and optimise the production of existing products is not possible.⁴⁵ Recently, significant advances were made in understanding the physical (process engineering) aspects of SSF but the information on physiology and molecular genetics is limited. To obtain an optimized production process, it is of significant importance to gain a better understanding of the molecular and cell biology of these microorganisms as well as the relevant approaches in biochemical engineering. Due to low productivities and lack of ideal characteristics, the quest for the discovery of new wild type phytases and improving the existing phytases continues.

2.3 Strategies employed for improved phytase production

The production levels of phytase in naturally occurring strains are very low to be economically viable. Improvement in phytase production is achieved mutually by developments in production technology and engineered phytases, as discussed below.

2.3.1 Classical mutagenesis. Strain improvement by mutagenesis and selection is a highly developed technique, and it plays a central role in the commercial development of microbial fermentation processes. Mutagenic procedures can be carried out in terms of type of mutagen and dose to obtain mutant types, which may be screened for improved phytase, as seen in A. niger, using physical and chemical mutagenesis.⁴⁶ Several bacterial strains (wild or genetically modified), such as Lactobacillus amylovorus, E. coli, B. subtilis, B. amyloliquefaciens, Klebsiella spp, etc., have been employed for phytase synthesis. In addition to good yield of phytase enzyme, A. niger CFR 335 produces large amounts of dark conidiospores that hamper the extraction of enzyme and cause health risks, such as allergic bronchopulmonary aspergillosis, if not handled properly. Therefore, a strain of A. niger CFR 335 with phytase overproduction and lower sporulation rate was developed through UV mutagenesis by Gunashree and Venkateshwaran.⁴⁷

Chelius and Wodzinski, during the strain improvement studies of A. niger NRRL 3135 by UV radiation, isolated a phytase catalytic mutant producing 3.3-fold higher phytase (phyA) than the wild type strain. The production of mutant phyA was highly repressed up to 60% by inorganic phosphate (0.006%, w/v); however, their approach was limited by the lack of specificity and sensitivity in discriminating between phytase and acid–phosphatase activity during primary screening process.⁴⁸

2.3.2 Genetic improvement via transgenic studies. Although phytases are widely distributed in nature, the production in wild-type organisms is far from being economically viable. Hence, cloning and expression of phytase genes in suitable host organisms is necessary to reach higher productivities. Because the cost effectiveness of phytase production is a major limiting factor for its application, different heterologous expression systems and hosts have been evaluated such as plants, bacteria, and fungi (including yeast). As expected, each system bears some unique advantages along with certain limitations, as seen in Table 2.

2.3.3 Protoplast fusion. Protoplast fusion has significant potential for strain improvement, and has been applied for various industrially important microorganisms. Protoplast fusion may be used to produce interspecific or even intergeneric hybrids, and is an important tool because it can overcome the limitations of conventional mating systems in gene manipulation.⁵² However, it is still an emerging area in phytase research with only few reports of interspecific protoplast fusion between two auxotrophic mutants, A. niger CFR 335 ala− and A. ficuum SGA 01 val−, isoleu. Hybrids have been obtained having high stability, delay in sporulation and enhanced phytase production.⁵³ Therefore, protoplast fusion has potential for strain improvement and for enhancing phytase production.

2.3.4 Response surface methodology. The conventional one variable at a time (OVAT) approach is time consuming and laborious because it involves the variation of a single variable while maintaining others at a constant level. The true optimum value is missed due to the lack of the interaction of components. An alternative to OVAT is response surface methodology (RSM) because it involves the systematic, efficient and simultaneous interaction of variables. Optimization is important for maximizing production and yield, and at the same time for minimizing the cost. Krishna and Nokes examined the effect of culture conditions, particularly inoculum age, media composition (wheat bran and full-fat soybean flour) and duration of SSF on phytase production by A. niger.⁵⁴ Bogar et al. reported phytase production by A. ficuum NRRL 3135, M. racemosus NRRL 1994 and R. oligosporous NRRL 5905 using various substrates such as canola meal, cracked corn, soybean meal, and wheat bran.⁵⁵ However, these reports are few because of the low productivities and difficulties associated with operating and up scaling of SSF conditions.⁵⁶ Sunitha et al. optimized the medium for recombinant phytase production by E. coli BL21 using response surface methodology. A 23 central composite experimental design was used to study the combined effects of the medium components, i.e. tryptone, yeast extract and NaCl. The optimized medium with glucose showed the highest phytase activity of 2250 U l⁻¹.⁵⁷ Phytase production using yeast cultures has generally been carried out in SmF systems. The strains used include Schwanniomyces castellii, Pichia, Arxula adeninivorans and Candida kruzei. Galactose and glucose were the preferred carbon sources. Phytase production from P. anomala has been extensively studied using response surface methodology.

The fermentation technique employed is SmF with glucose, and yeast extract is the main carbon and nitrogen source widely used. Sreemula et al. evaluated 19 strains of lactic acid-producing bacteria of the genera Lactobacillus and Streptococcus for the production of extra-cellular phytase. A number of them exhibited enzyme activity in the fermentation medium but Lactobacillus amylovorus B4552 produced the maximum amounts of phytase, ranging from 125 ± 146 U ml⁻¹ in SmF using glucose and inorganic phosphate.⁵⁸

2.3.5 Directed evolution. Engineering of enzymes using directed evolution is successful, especially in improving their thermostability and catalytic properties. This involves the construction of a mutant library through random mutagenesis or in vitro recombination techniques, followed by the selection of mutants with a desired characteristic using a high-throughput screening technique.⁵⁹ The desirable mutants are selected and identified by directional selection methods and excluding the mutants of non-interest.

A highly active and thermally improved bacterial Ymphytase has been obtained by directed evolution. Ymphytase represents an alternative to fungal phytases for monogastric feed products. A chemically more diverse SeSaM library yielded a thermally more resistant Ym phytase variant with five amino acid substitutions. Mutational analysis showed that the Ymphytase protein is highly robust towards mutations.⁶⁰

Similarly, the method of error-prone PCR was used to generate the mutant phytase with better catalytic efficiency than the original type by introducing several substitutions. The structural predictions indicated that the mutations generated by ep-PCR reorganized or remodeled the active site, which could lead to increased catalytic efficiency and higher specific activity (up to 61%).⁶¹

To explore the molecular determinants responsible for the thermostability of Bacillus phytases, structural analysis and site directed mutagenesis was employed,⁶² which will help rational protein engineering to develop effective phytases.

3 Downstream processing of phytase

Downstream processing, involving recovery and formulation, incurs 70% of the overall production cost of enzyme due to the complexity of the system and the need to maintain biological activity. Phytase technology for separation and purification, employing a chromatographic process, has evolved slowly as compared to production. Most of these approaches were employed for analytical purposes especially for biochemical, molecular and structural characterization. Phytase is susceptible towards inactivation, thus for enhanced stability, phytase enzymes are often formulated as solid-state proteins produced by spray drying, lyophilization or granulation. A dry formulation greatly reduces the likelihood of chemically and biologically mediated inactivation. Thus, there is a growing interest for fast and economic processes, which will stimulate research to unlock new insights in phytase down streaming technology. Conventional methods for phytase separation and purification involve pretreatment and chromatographic methods.

3.1 Pretreatment and concentration

Several different concentration and purification steps are required to reach the final end step quality product. Certain pretreatments are required because phytases may be intracellular and extracellular. Depending on the location of the cell bound enzyme, various permeabilization treatments including organic solvents, enzymes, detergents and physical methods are used.⁶³

Solid liquid separation techniques, such as centrifugation and decant, are usually used for extracellular phytase separation. The culture filtrate is concentrated by salt precipitation, acetone precipitation and ultrafiltration for various phytases from plants, bacteria and fungi.

3.2 Chromatography process

Further purification of phytases includes gel filtration, ion-exchange chromatography, affinity chromatography and hydrophobic interaction. One major problem in the purification of phytate-degrading enzymes, especially from plants, is the separation of phytate-degrading enzymes from contaminating nonspecific acid phosphatases.⁶⁴

The recovery and purification of phytase has been achieved through several steps using different techniques. Boyce and Walsh purified phytase from Mucor hiemalis, utilizing five steps (ultrafiltration, diafiltration, ion exchange, gel filtration and hydrophobic interaction) and achieved 51% recovery with a purification factor of 14.1.⁶⁵ Azek et al. obtained two phytases from Rhizopus oligosporus in five steps (Acetone Fractionation, Mono-S HR 5/50 Cationic-Exchange Chromatography, 16/60 Sephacryl S-200 HR chromatography, Mono-S HR 5/50 Cationic-Exchange Chromatography, Mono-Q HR 5/5 Anionic-Exchange Chromatography) with recovery: phytase 1 (1.3%) and phytase 2 (1.6%) and purification factor (75, 46).⁶⁶ Debaryomyces castellii phytase was purified to homogeneity in a single step by hydrophobic interaction chromatography. Its molecular mass is 74 kDa with 28.8% glycosylation. Its activity was optimal at 60 °C at pH 4.0. The Km value for sodium phytate was 0.532 mM.⁶⁷

Phytase generated on citric pulp fermentation by A. niger FS3 was purified by cationic-exchange, anionic exchange chromatography and chromatofocusing steps with 6.35% yield.⁶⁸ Previous work from Caseys' lab indicated that extracellular phytase from A. niger ATCC 9142 was purified with a purification factor of 24.89-fold and a 26% yield.⁶⁹ A phytase from Bacillus was purified 124-fold from the culture broth with 15.4% yield, and exhibited an activity of 36.0 U mg⁻¹.⁷⁰ Li et al. reported an extracellular phytase from a marine yeast with a purification factor of 7.2-fold and a 10.4% yield.⁷¹

Three phytases were purified about 14 200-fold (LP11), 16 000-fold (LP12), and 13 100-fold (LP2) from germinated 4 day-old lupine seedlings to apparent homogeneity with recoveries of 13% (LP11), 8% (LP12), and 9% (LP2), referring to the phytase activity in the crude extract. They behave as monomeric proteins of molecular mass of about 57 kDa (LP11 and LP12) and 64 kDa (LP2). The purified proteins are acid phytases and exhibit a single pH optimum at 5.0. Optimal temperature for the degradation of sodium phytate is 50 °C.⁷²

An extracellular phytase from A. niger 11T53A9 was purified about 51-fold to apparent homogeneity with a recovery of 20.3% referred to the phytase activity in the crude extract. Purification was achieved by ammonium sulphate precipitation, ion chromatography and gel filtration. The purified enzyme behaved as a monomeric protein with a molecular mass of about 85 kDa and exhibited maximal phytate-degrading activity at pH 5.0. The optimum temperature for the degradation of phytate was 55 °C.⁷³

3.3 Liquid–liquid extraction

The application of single step aqueous two-phase extraction (ATPE) for the downstream processing of phytase from A. niger NCIM 563, produced under SSF, has been studied and compared with the traditional multi-step procedure, involving salt precipitation and column chromatography. High phytase recovery (98.5%) within a short time (3 h) and improved thermostability was attained by ATPE in comparison to 20% recovery in 96 h by the chromatography process. The ATPE system consisting of the combination of polyethylene glycol (PEG) 6000 and 8000 (10.5%) and sodium citrate (20.5%) resulted in a one-sided partitioning of phytase in the bottom phase with a purification factor of 2.5.⁷⁴

The partition and recovery behavior of phytase, produced by solid-state cultivation utilizing citrus pulp as substrate, was determined in an ATPE composed of PEG–citrate. The highest partition coefficient (14.42) was observed within a 26% (w/w) PEG 400 (g mol⁻¹) and a 20% (w/w) sodium citrate at pH 6.0. The independent variables, which greatly influenced the partition coefficient and recovery were citrate concentration and PEG mass molar.⁷⁵ These results suggest that PEG–citrate ATPE is an interesting and efficient alternative to traditional chromatographic methods.

3.4 Immobilization

Immobilization of phytase on natural supports, such as allophone, was studied using E. coli and A. niger phytase. The residual activity of immobilized phytase on allophanic and montmorillonite nanoclay supports was higher under acidic conditions and led to a higher thermal stability and resistance to proteolysis.⁷⁶

The production of myo-inositol phosphate isomers is a budding area but is hampered by lack of stability under processing conditions and difficulties in recovery from reaction mixtures. However, this has been overcome by the immobilization of phytases onto Fe₃O₄-magnetic nanoparticles with high operational stability.⁷⁷

The major constraint in the application of phytase in animal feed is its reduced thermostability at the pelleting process. Pelleting stability is improved by protected formulation and thermostability coatings to some extent. Protein or enzyme stabilizers include the use of non reducing sugars, organic and inorganic salts and polyols. Granulation involves the use of water soluble polymers, fat coating, organic salts and stabilizers for the encapsulation of the biologically active part to prevent inactivation at high temperature. However, the inactivation of phytase at high temperature still requires further investigation.

4 Biotechnological applications of phytase

Since the first commercial phytase product Natuphos® launch in 1991, the market volume has reached ca. 150 million Euros, and is likely expand with new applications. The main application is still using feed supplements to improve P bioavailability in plant feed-stuffs via the enzyme-mediated hydrolysis of phytate. Most importantly, the improved utilization of the phosphate deposits in the feed results in a substantial reduction in the phosphate content in animal manure, and hence decreases phosphate load on the environment in areas of intensive animal agriculture. High dietary P bioavailability reduces the need for supplemental inorganic P such as mono- and dicalcium-phosphate (MCP, DCP).

Because of the strong economic growth in China and India along with the price hike of oil, the supply and cost of MCP and DCP has become a practical concern. Furthermore, inorganic phosphate is a non-renewable resource, and it has been estimated that the easily-accessible phosphate on earth will be depleted in the next 50 years. Thus, phytase is an effective tool for natural resource management of P on a global scale.

The ban of dietary supplementation of meat and bone meal, as a cheap source of feed P, in Europe to prevent possible cross-species transfer of diseases such as BSE has led to a profound change in the feed P management. This has given phytase a new socio-economic impact as a cost effective alternative to ensure that animals obtain adequate available P from plant-based diets. As the major storage form of P in seeds, plant phytate was produced in 2000 at a global yield of >51 million metric tons. This amount accounts for approximately 65% of the elemental P sold worldwide as fertilizers.⁷⁸ Apparently, phytase can turn plant phytate into a very valuable P resource by improving its bioavailability for animal nutrition. Denmark and the Netherlands have imposed regulations to promote the use of microbial phytases.

Organic P (Po) hydrolysis by microbial phytases has been extensively considered in diverse biotechnological applications, including environmental protection and agricultural, animal and human nutrition.⁷⁹ Because of the potential value of phytases in improving the efficiency of P use, biotechnology has led the rapid development of the field to its current state. With the development of heterologous gene expression, large amounts of enzymes can be produced at relatively low cost. The importance of phytases as potential biotechnological tools has been recognized in various fields (Table 3). However, only a limited number of phytases have been reported and studied, and our knowledge of the mechanisms and factors regulating phytase activity is limited. Further research for developing new technologies and identifying the most efficient phytases must continue, and should be directed towards application oriented research.

Table 3 Potential applications of phytases

Application	Role and effect	Properties	Challenges
Feed industry	Increased P utilization, metal bioavailability, decreased P conc. in excrement, substitutes expensive di-calcium phosphate	Resistance to low pH, active in the stomach, stable during animal feed processing and storage, low cost of production and easily processed by the feed manufacturer	Lack of desirable properties, high cost of production
Food industry	Increased P utilization, metal bioavailability, technical improvement of food processing	—	It will be a challenge to minimize the negative effect of phytate on iron and zinc nutrition without losing its potential health benefits
Myoinositol phosphate	Myoinositol phosphate intermediates used as enzyme stabilizers, enzyme inhibitors, Potential drugs, chiral building blocks	—	Further intensive investigations, using diverse phytases, need to be undertaken for designing and producing pharmacologically important lower myo-inositol phosphates
Aquaculture	Substitute for expensive protein source such as menhaden fish meal and maintains the acceptable levels of P in water	Phytase active at low temperature and broad pH optima is required	Effects of phytase supplementation or various physiological and endocrine parameters like secretion of other enzymes, bile salts on the on the immune response, hormone levels including growth hormone, thyroid hormone, insulin etc needs to be studied
Soil amendment	Plant growth stimulation by mobilization of soil phytate into inorganic P	Phytase with broad pH optima and catalytic activity	Needs more research on phytase supplementation for boosting the productivity in agriculture and horticulture
Hydroxyapatite formation	Simple biocatalytic process	Cost effective as compared to commercial process	Needs more efforts for product development
Nanoparticles	Hollow metal nanocapsules formed using phytase andionic liquid	Implications in biocatalyst and drug delivery	Needs more research for exploring its possibility in biomedical application

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4.1 Phytases in animal nutrition

Monogastric animals, such as swine, fish, and poultry, show negligible or no phytase activity in the digestive tract. Consequently, phytates can not be metabolized by animals, thus creating a need to enhance phosphate and mineral bioavailability via phytase supplementation of animal feed. Of late, phytases are also viewed as environmental friendly products, which can reduce the level of phosphate pollution in intensive livestock management by avoiding the addition of exogenous phosphate.⁸⁰ Undigested phytate of monogastric manure is washed off the farmland, which imperils adjacent waterways by eutrophication.⁸¹ The effect of feeding phytase to animals on pollution has been quantitatively determined. If phytase were used in the feed of all the monogastric animals reared in the U.S., it would release phosphorus amounting to 168 million U.S. dollars and would preclude 8.23 × 104 tonnes of phosphate from entering the environment per annum. The use of phytase as a feed additive has been approved in 22 countries by the FDA.⁸²

During the past two decades, there has been significant increase in the use of phytases as feed additive in pig, poultry, and fish diets. In numerous studies, the efficacy of microbial phytases to release phytate-bound P has been demonstrated in various animals. Phytases were also found to enhance the utilization of different minerals. Phytases from different sources have been evaluated individually and in combination for their efficacy as feed additives in poultry.^83–85 Use of both bacterial and fungal phytases together as feed additive would be another promising alternative in improving the phosphorus utilization and alleviation of mineral deficiency due to their synergistic activities throughout the gastrointestinal tract of the animals. The use of phytase as a feed enzyme sets certain demands on the properties of the enzyme. In particular, the enzyme should withstand high temperatures. This is because poultry and pig feed is commonly pelleted, which ensures that the animals have a balanced diet and facilitates the preservation of enzyme-containing product in the feed industry. During the pelleting process the temperatures may temporarily reach 90 °C. The first commercial phytase product, which became commercially available 10 years ago, offered animal nutritionists the tool to drastically reduce the phosphorus excretion of monogastric animals by replacing inorganic phosphates with microbial phytase. Depending on diet, species, and the level of phytase supplementation, P excretion can be reduced between 25% and 50%.⁸⁶

4.2 Phytases in human nutrition

Mineral deficiency of diets, caused by radical changes in food habits, is a major concern in developing countries. The processing and manufacturing of human food is also a possible application for phytases. Up to now, no phytase product for a relevant food application is available in the market. Research in this field focuses on better mineral absorption or technical improvement of food processing. Phytate present in cereal-based and legume-based complementary foods has been found to inhibit mineral absorption.⁸⁷ The small intestine of humans has limited ability to digest undegraded phytates, resulting in adverse nutritional consequences with respect to metabolic cation imbalances. Phytic acid (PA)—containing 12 dissociable protons with pK_a values ranging from ∼1.5 to 10—is a highly reactive and potent chelator of several mineral ions such as Ca²⁺, Mg²⁺, Zn²⁺, and Fe²⁺. Phytic acid forms insoluble salts, at normal acidity (pH 3.0–6.8), in the human digestive tract, thereby reducing the bioavailability of these critical mineral nutrients for absorption.⁸⁸ Mucosal phytase and alkaline phosphatases, even if present in the human small intestine, do not play a significant role in the phytate digestion, while dietary phytase serves as an important factor in phytate hydrolysis.⁸⁹ Haros et al. investigated the possible use of phytase in bread making. Different amounts of fungal phytase were added in whole wheat breads and it was shown that phytase is an excellent bread-making improver. The main achievement of this activity was the shortened fermentation period without affecting the pH of the bread dough. An increase in bread volume and an improvement in crumb texture were also observed.⁹⁰

Application of immobilized E. coli phytase and fusion protein in the dephytinization of soy milk led to a 10% increase in the release of inorganic phosphate at 50 °C relative to free fusion protein.⁹¹ The lowest phytic acid concentration and highest zinc bioavailability index were achieved when S. cerevisiae, L. plantarum, and Leu. mesenteroides were used at 30.0% dough replacement with sourdough. In this study, effects of 8 different sourdough starters prepared with Saccharomyces cerevisiae, Lactobacillus plantarum, L. acidophilus, and Leuconostoc mesenteroides were investigated on the phytic acid level and the mole ratio of phytic acid to zinc in a traditional Iranian bread (sangak).⁹²

Vitamin C, selenium, zinc and iron are deficient in the diet of lactating women in rural central Mexico, albeit moderate pulque drinking appears to ameliorate iron and zinc deficiencies by the presence of phytase from live bacteria in the latter.⁹³

4.3 Phytases in aquaculture

A major concern in aquaculture is the utilization of dietary phosphates, which critically affects fish growth as well as the aquatic environment. An efficient utilization of feed leading to optimum fish growth serves as a benchmark of successful aquaculture worldwide. Studies using phytase as a feed additive in aquaculture have established that phytase supplementation could enhance the bioavailability of P, nitrogen, and other minerals, thereby decreasing phosphorus-load in aquatic environment.^94,95

The enzyme from phytase producing intestinal bacteria of Atlantic cod can stimulate intracellular head kidney leukocyte activities but not the production of extracellular substances, which are involved in antibacterial response. These have implications on the potential use of bacterial phytase as feed supplement to boost cellular immune response of the fish and could be employed as a health management strategy in culture systems.⁹⁶ Thus, these may have significant impact on the development of feed supplements and health management in aquaculture systems.

4.4 Role of phytases in soil amendment

Phosphorus is an essential plant nutrient that limits agricultural production on a global scale. Approximately 30–80% of the total P in soils is bound in organic form.⁹⁷ Phytate constitutes ∼50% of the total organic P pool in the soil and is poorly utilized by plants. Extracellular phytase activities have been reported under phosphate stress conditions in diverse plant species, namely, tobacco,⁹⁸ barley,⁹⁹ tomato, alfalfa,¹⁰⁰ etc. The ability of plants to use phosphorus from low phosphate or phytate containing media and/or from soil is improved when soil/media are inoculated with microorganisms, possessing the ability to exude phytase or when a purified phytase is added.

Root physiological adaptations (i.e. rhizosphere carboxylate content and P-uptake rate) are more important than morphological adaptations (i.e. root length and diameter) to enhance the uptake of P and cations.¹⁰¹

4.5 Phytase in plant growth promotion

Novel Enterobacter cancerogenus MSA2 is a plant growth promoting gamma-proteobacterium that was isolated from the rhizosphere of Jatropha curcas, which is a potentially important biofuel feed stock plant. MSA2 is the first plant growth-promoting bacterium, which produces ACC deaminase enzyme and promotes plant growth with Jatropha curcas.¹⁰² The effect of fungal phytase on plant growth at pot and tray level, compared with commercial fertilizers pertaining to chemical and physiological parameter and as soil amendment, was studied. Phytase was efficient in reducing the phytic acid content of soil by about 30% while simultaneously increasing the phytate phosphate availability by 1.18-fold.¹⁰³

4.6 Budding applications

Lower phosphoric esters of myo-inositol (mono, bis, tris, and tetrakisphosphates) play a crucial role in transmembrane signaling processes and in calcium mobilization from intracellular store in animal as well as in plant tissues.¹⁰⁴ Research interest in this field prompted the need for various inositol phosphate preparations; however, chemical synthesis is difficult. In contrast, enzymatic synthesis has the advantage of high stereospecifity and mild reaction conditions. The use of phytase has been shown to be very effective in producing different inositol phosphate species.

Different isomers of myo-inositol phosphates have shown pharmacological effects in the prevention of diabetic complications, anti-inflammatory effects,¹⁰⁵ and antiangiogenic and antitumor effects.¹⁰⁶ Myo-inositol phosphates are also known to ameliorate heart disease conditions by controlling hypercholesterolemia and atherosclerosis,¹⁰⁷ and also prevent renal stone formation.¹⁰⁸

A single step rapid biocatalytic process of hydroxyapatite and myoinositol intermediates synthesis has several advantages such as stereo specificity, mild reaction conditions and cost effectiveness as compared to chemical process.¹⁰⁹

Self-assembly of phytase molecules in ionic liquid leading to the formation of enzyme capsules is being studied. These capsules act as soft functional templates for the in situ reduction and decoration of metal salts.¹¹⁰

5 Future perspectives and new insights

There is a large gap between the metabolic and bioprocessing level of microbes especially in the case of fungi. There are several reports on phytase production and purification in different fermentation systems, which affect microbial physiology and productivity. This includes various aspects such as media composition, morphology, fermentation system, type of strain and substrate used, as shown in Table 4. However, there is no information regarding structural differences among phytase produced under both systems. Complex microbes, especially fungi, exploit a wide range of environmental conditions but morphology under varied fermentation system is often a bottleneck in productivity of industrially important desired products. There is abundant proof in literature that the product spectrum from SSF is very different from that obtained in submerged fermentation (SmF). However, the mechanisms underlying these differences are not understood.

Table 4 Summary of various fermentation systems used for the production and down streaming of phytase

Type Microbial strain	Substrate	pH	Temp (°C)	Activity	Mol wt (kDa)	Purification	Recovery (%)	Reference
Achromobacter sp PB-01	WB RSM			7.05 IU ml^−1				111
Apergillus oryzae SBS 50	RSM					Protease resistant phytase		112
Apergillus niger NCIM 563	Semisynthetic	2.5	55	40 IU ml^−1	264	HIC, GF	30.24	130
A. niger NCIM 563	Semisynthetic	5	55	10 IU ml^−1	66	HIC, GF	26.55	130
A. niger NCIM 563	RB RSM	2.5		268 IU ml^−1				46
Bacillus nealsonii ZJ0702		7.5	55		43		5.7	133
B. subtilis MJA	Sodium phytate	5	37		36			113
K. pneumoniae 9-3B	MM9	4	50		45	Salt ppt, CC	15.8	114
Lactobacillus plantarum	Synthetic medium	3.4	120	984.5 U ml^−1	46			115
Nocardia sp MB 36	Starch, beef extract			0.4 IU ml^−1				116
	RSM
SMF
Paecilomyces variotii	Orange pomace			350 IU ml^−1		Tannase and phytase for detoxifocation of castor beans		117
Pichia anomala	Cane molasses RSM			1780 U g^−1		Permeabilization		118
P. pastoris recomb	MSGW RSM			441 U ml^−1				119
Rhizopus oligosporous (DSMZ 1964)	Rice flour	3,4.5	60		45	Salt ppt, CC	1.3	66
	Rice flour	3,5	55		45	Salt ppt, CC	1.6	66
Saccharomyces cerevisiae	Bactopeptone, starch			165 U ml^−1	80	Phytase and amylase		120
Shigella sp CD2	Sodium phytate	5.5	60		43	Salt ppt, IEC	41	121
Sporotrichum thermophile	Starch	5	60					131
Streptomyces spp	Glycerol	5	55			Fust repot		132
Coniochaeta spp						Fust repot		122
Recombinant A. niger phytase in E coli		6.5	50		92	Chaperonin co-expression		129
Bacillus subtilis US 417	WB			112 U g^−1				41
Absidia blakesleeana URM5604	Citrus pulp					LLE	115	128
A. ficuum	Lentils RSM			32 U g^−1				123
A. ficuumNTG-23	Waste vinegar	1.3	67		65.5	IEC, GF	23.8	124
A. niger11T53A9	WB	5	55		85	Salt ppt, GF	20.3	73
A. nigerFS3	Citric by products	5	60		108	CC	10	68
A. nigerNCIM 563	WB RSM	5.6	60	154 U g^−1	85	CC and LLE	CC(20) LLE (98.5)	38 and 75
A. nigerNCIM 563	WB RSM	2.5		250 IU g^−1	120	CC		110
SSF
A. oryzae	Soy meal 2 temp design			58.7 U g^−1				125
B. subtilis US 417	WB	7.5	55	85 U g^−1	41	Heat treatment, salt ppt, FPLC		41
R. oryzae	Linseed cake RSM	5	45	149 U g^−1	36	Salt, IEC	26	126
Schizophyllum commune	WB RSM	5	50	113.7 U g^−1		LLE	367 (Partial)	127
S. thermophile	Sesame oil cake	5	60	282 IU g^−1				131

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There is only report on the structural differences among phytase produced under SSF and SmF by A. niger, and this study provides the basis for the explanation of stability and catalytic differences observed for these three phytase. In fact, only two reports on the comparative production of phytase by these two fermentation processes are available for fungal and bacterial (Table 4).

More powerful and automated image analysis techniques will aid in morphology engineering and will provide new insights to the existing “black box” of SSf/SmF biotechnology for phytase production. Strategies such as microparticle addition and osmolality variation will aid in the targeted engineering of fungal morphology.

Along with microbial production, downstream processing is an essential aspect of phytase bioprocessing. Rapid and economic methods such as liquid–liquid extraction are imminent promising alternatives, as can be seen from Table 4. More efforts are required for the development of efficient, scalable and economical process for phytase bioseparation to overcome the techno-economic limitations of conventional downstream processes.

The core aim of a viable process is to retain the activity during storage and use. Limitations related to phytase formulation and stabilization is a major bottleneck in its industrial application. Thus, techniques such as immobilization and application targeted research will help in solving the problem to some extent.

A focused platform for microbial production, downstream processing and application oriented research will help in developing a integrated technological solution to phytase production. This will present new insights in the biological and engineering facets of phytase producing microbes and reveal a new era in phytase biotechnology.

6 Conclusion

P is an indispensable resource, which has been mismanaged to the point that we are jeopardizing our long-term food and water security. As the need to conserve global phosphate reserves increases, the role of phytase will broaden. Phytases are now being recognized for their beneficial environmental role in reducing the P levels in manure and minimizing the need to supplement P in diets. The conventional methods for phytase production and purification are economically unviable due to various shortcomings. Hence, there is a need for additional and improved strategies that will help in developing a robust system for the phytase production. Further application oriented efforts are required to design versatile “second-generation” phytases with wider applicability. Modification and upgradation of enzymatic properties can be achieved by the adoption of genetic and protein engineering methods. The combination of fungal and bacterial phytases as feed additives might improve the bioavailability of P and minerals due to their synergistic activity in animal digestive system. Further insights in the development of application oriented phytases will open a new era in its bioprocessing and widen the horizons of its applicability and efficiency. New market segments such as aquaculture and agriculture will provide new opportunities for phytase.

Acknowledgements

The author, Ms Kavita Bhavsar thanks the Council of Scientific and Industrial Research, Government of India for financial assistance. The authors also gratefully acknowledge support and facilities provided by the Center of Excellence in Scientific Computing, National Chemical Laboratory, India.

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