Hubbry Logo
PhytasePhytaseMain
Open search
Phytase
Community hub
Phytase
logo
8 pages, 0 posts
0 subscribers
Be the first to start a discussion here.
Be the first to start a discussion here.
Phytase
Phytase
Phytase
View on Wikipedia
from Wikipedia
Phytate

A phytase (myo-inositol hexakisphosphate phosphohydrolase) is any type of phosphatase enzyme that catalyzes the hydrolysis of phytic acid (myo-inositol hexakisphosphate) – an indigestible, organic form of phosphorus that is found in many plant tissues, especially in grains and oil seeds – and releases a usable form of inorganic phosphorus.[1] While phytases have been found to occur in animals, plants, fungi and bacteria, phytases have been most commonly detected and characterized from fungi.[2]

History

[edit]

The first plant phytase was found in 1907 from rice bran[3][4] and in 1908 from an animal (calf's liver and blood).[4][5] In 1962 began the first attempt at commercializing phytases for animal feed nutrition enhancing purposes when International Minerals & Chemicals (IMC) studied over 2000 microorganisms to find the most suitable ones for phytase production. This project was launched in part due to concerns about mineable sources for inorganic phosphorus eventually running out (see peak phosphorus), which IMC was supplying for the feed industry at the time. Aspergillus (ficuum) niger fungal strain NRRL 3135 (ATCC 66876) was identified as a promising candidate[6] as it was able to produce large amounts of extracellular phytases.[7] However, the organism's efficiency was not enough for commercialization so the project ended in 1968 as a failure.[6]

Still, identifying A. niger led in 1984 to a new attempt with A. niger mutants made with the relatively recently invented recombinant DNA technology. This USDA funded project was initiated by Dr. Rudy Wodzinski who formerly participated in the IMC's project.[6] This 1984 project led in 1991 to the first partially cloned phytase gene phyA (from A. niger NRRL 31235)[6][8] and later on in 1993 to the cloning of the full gene and its overexpression in A. niger.[6][9]

In 1991 BASF began to sell the first commercial phytase produced in A. niger under the trademark Natuphos which was used to increase the nutrient content of animal feed.[6]

In 1999 Escherichia coli bacterial phytases were identified as being more effective than A. niger fungal phytases.[6][10][11] Subsequently, this led to the animal feed use of this new generation of bacterial phytases which were superior to fungal phytases in many aspects.[6]

Classes

[edit]

Four distinct structural classes of phytase have been characterized in the literature: histidine acid phosphatases (HAPS), beta-propeller phytases (BPPs), purple acid phosphatases (PAPs),[2] and most recently, protein tyrosine phosphatase-like phytases (PTP-like phytases).[12]

Histidine acid phosphatases (HAPs)

[edit]

Most of the known phytases belong to a class of enzyme called histidine acid phosphatases (HAPs). HAPs have been isolated from filamentous fungi, bacteria, yeast, and plants.[1] All members of this class of phytase share a common active site sequence motif (Arg-His-Gly-X-Arg-X-Pro) and have a two-step mechanism that hydrolyzes phytic acid (as well as some other phosphoesters).[2] The phytase from the fungus Aspergillus niger is a HAP and is well known for its high specific activity and its commercially marketed role as an animal feed additive to increase the bioavailability of phosphate from phytic acid in the grain-based diets of poultry and swine.[13] HAPs have also been overexpressed in several transgenic plants as a potential alternative method of phytase production for the animal feed industry[14] and very recently, the HAP phytase gene from E. coli has been successfully expressed in a transgenic pig.[15]

β-propeller phytases

[edit]
See also: Beta-propeller phytase

β-propeller phytases make up a recently discovered class of phytase. These first examples of this class of enzyme were originally cloned from Bacillus species,[2] but numerous microorganisms have since been identified as producing β-propeller phytases. The three-dimensional structure of β-propeller phytase is similar to a propeller with six blades. Current research suggests that β-propeller phytases are the major phytate-degrading enzymes in water and soil, and may play a major role in phytate-phosphorus cycling.[16]

Purple acid phosphatases

[edit]

A phytase has recently been isolated from the cotyledons of germinating soybeans that has the active site motif of a purple acid phosphatase (PAP). This class of metalloenzyme has been well studied and searches of genomic databases reveal PAP-like sequences in plants, mammals, fungi, and bacteria. However, only the PAP from soybeans has been found to have any significant phytase activity. The three-dimensional structure, active-site sequence motif and proposed mechanism of catalysis have been determined for PAPs.[citation needed]

Protein tyrosine phosphatase-like phytases

[edit]

Only a few of the known phytases belong to a superfamily of enzymes called protein tyrosine phosphatases (PTPs). PTP-like phytases, a relatively newly discovered class of phytase, have been isolated from bacteria that normally inhabit the gut of ruminant animals.[17] All characterized PTP-like phytases share an active site sequence motif (His-Cys-(X)5-Arg), a two-step, acid-base mechanism of dephosphorylation, and activity towards phosphorylated tyrosine residues, characteristics that are common to all PTP superfamily enzymes.[18][19] Like many PTP superfamily enzymes, the exact biological substrates and roles of bacterial PTP-like phytases have not yet been clearly identified. The characterized PTP-like phytases from ruminal bacteria share sequence and structural homology with the mammalian PTP-like phosphoinositide/-inositol phosphatase PTEN,[12] and significant sequence homology to the PTP domain of a type III-secreted virulence protein from Pseudomonas syringae (HopPtoD2).[20]

Biochemical characteristics

[edit]

Substrate specificity

[edit]

Most phytases show a broad substrate specificity, having the ability to hydrolyze many phosphorylated compounds that are not structurally similar to phytic acid such as ADP, ATP, phenyl phosphate, fructose 1,6-bisphosphate, glucose 6-phosphate, glycerophosphate and 3-phosphoglycerate. Only a few phytases have been described as highly specific for phytic acid, such as phytases from Bacillus sp., Aspergillus sp., E. coli[21] and those phytases belonging to the class of PTP-like phytases.[18]

Pathways of phytic acid dephosphorylation

[edit]

Phytic acid has six phosphate groups that may be released by phytases at different rates and in different order. Phytases hydrolyze phosphates from phytic acid in a stepwise manner, yielding products that again become substrates for further hydrolysis. Most phytases are able to cleave five of the six phosphate groups from phytic acid. Phytases have been grouped based on the first phosphate position of phytic acid that is hydrolyzed. The Enzyme Nomenclature Committee of the International Union of Biochemistry recognizes three types of phytases based on the position of the first phosphate hydrolyzed, those are 3-phytase (EC 3.1.3.8), 4-phytase (EC 3.1.3.26), and 5-phytase (EC 3.1.3.72). To date, most of the known phytases are 3-phytases or 4-phytases,[21] only a HAP purified from lily pollen[22] and a PTP-like phytase from Selenomonas ruminantium subsp. lactilytica[20] have been determined to be 5-phytases.

Biological relevance

[edit]

Phytic acid and its metabolites have several important roles in seeds and grains, most notably, phytic acid functions as a phosphorus store, as an energy store, as a source of cations and as a source of myo-inositol (a cell wall precursor). Phytic acid is the principal storage forms of phosphorus in plant seeds and the major source of phosphorus in the grain-based diets used in intensive livestock operations. The organic phosphate found in phytic acid is largely unavailable to the animals that consume it, but the inorganic phosphate that phytases release can be easily absorbed. Ruminant animals can use phytic acid as a source of phosphorus because the bacteria that inhabit their gut are well characterized producers of many types of phytases. However, monogastric animals do not carry bacteria that produce phytase, thus, these animals cannot use phytic acid as a major source of phosphorus and it is excreted in the feces.[23] However, human—especially vegetarians and vegans due to increased gut microbiome adaptation—can have microbes in their gut that can produce phytase that break down phytic acid.[24]

Phytic acid and its metabolites have several other important roles in Eukaryotic physiological processes. As such, phytases, which hydrolyze phytic acid and its metabolites, also have important roles. Phytic acid and its metabolites have been implicated in DNA repair, clathrin-coated vesicular recycling, control of neurotransmission and cell proliferation.[25][26][27] The exact roles of phytases in the regulation of phytic acid and its metabolites and the resulting role in the physiological processes described above are still largely unknown and the subject of much research.

Phytase has been reported to cause hypersensitivity pneumonitis in a human exposed while adding the enzyme to cattle feed.[28][29]

Agricultural and industrial uses

[edit]

Phytase is produced by bacteria found in the gut of ruminant animals (cattle, sheep) making it possible for them to use the phytic acid found in grains as a source of phosphorus.[30] Non-ruminants (monogastric animals) like human beings, dogs, pigs, birds, etc. do not produce phytase. Research in the field of animal nutrition has put forth the idea of supplementing feed with phytase so as to make available to the animal phytate-bound nutrients like calcium, phosphorus, minerals, carbohydrates, amino acids and proteins.[31] In Canada, a genetically modified pig called Enviropig, which has the capability to produce phytase primarily through its salivary glands, was developed and approved for limited production.[32][33]

Phytase is used as an animal feed supplement – often in poultry and swine – to enhance the nutritive value of plant material by liberation of inorganic phosphate from phytic acid (myo-inositol hexakisphosphate). Phytase can be purified from transgenic microbes and has been produced recently in transgenic canola, alfalfa and rice plants.[34] Phytase has also been produced from cultivated Rhizopus oligosporus.

References

[edit]
Revisions and contributorsEdit on WikipediaRead on Wikipedia

Phytase

View on Grokipedia
from Grokipedia
Phytase is a phosphohydrolytic enzyme that catalyzes the stepwise hydrolysis of phytic acid, also known as phytate—the primary storage form of phosphorus in plant seeds, grains, and legumes—releasing inorganic orthophosphate and lower myo-inositol phosphate esters. This enzymatic action initiates the breakdown of phytate, which is otherwise poorly digestible by many animals and humans due to its ability to bind minerals such as calcium, iron, and zinc, forming insoluble complexes that reduce nutrient bioavailability. First identified in 1907, phytase research advanced significantly in the early , with its hydrolytic properties detailed by 1911 and commercial production emerging in the 1990s through technology to enhance and efficacy. Phytases occur across all domains of life, including , fungi, , and animals, with major classes comprising histidine acid phytases (HAPhys), which sequentially remove up to five groups; β-propeller phytases (BPPhys), which release every other in a calcium-dependent manner; protein phosphatase-like phytases (PTPhys); and purple acid phytases (PAPhys), often metal-ion mediated and prevalent in and fungi. These enzymes exhibit optimal activity at ranges of 4–6 and temperatures of 35–63°C, depending on their source, enabling their adaptation to diverse biological environments like , the , and industrial processes. In , phytases play a pivotal role as feed additives for non-ruminant such as , , and species, improving retention and reducing manure excretion by up to 50%, thereby mitigating environmental and preserving finite rock resources. The global phytase market was approximately $600 million as of 2024, projected to grow to over $1 billion by 2032, driven by biotechnological production from microbial sources like . In , supplemental phytase enhances the absorption of minerals from plant-based diets, potentially addressing deficiencies in populations reliant on grains and . Ecologically, phytases are essential to the global , recycling bound in soils and ecosystems to support microbial and plant growth.

Introduction

Definition and Function

Phytase is a phosphohydrolase that catalyzes the sequential of , also known as myo-inositol hexakisphosphate (IP6), the primary storage form of in plant seeds. The name "phytase" derives from "phytate," the salt form of phytic acid, combined with the suffix "-ase" denoting an . Phytases are classified into acid and alkaline types based on their optimal , with acid phytases assigned the EC 3.1.3.8 and alkaline phytases EC 3.1.3.26. The primary biochemical function of phytase involves the of , initiating with the reaction IP6 + H₂O → IP5 + Pi, where IP5 is myo-inositol pentakisphosphate and Pi is inorganic . This process can continue stepwise, removing additional groups to produce lower inositol phosphates down to IP1 (myo-inositol monophosphate) and ultimately . Phytases occur in diverse organisms, including microorganisms, , and animals, where they play a role in phosphorus metabolism. By degrading , phytase counteracts its antinutritional effects, as phytic acid binds to essential minerals such as , iron, and , reducing their in plant-based diets. This enzymatic action enhances the digestibility and absorption of these minerals, as well as proteins, thereby improving nutritional outcomes in animals and potentially humans consuming phytate-rich foods. For instance, phytase supplementation has been shown to increase utilization by up to 30-50% in feeds, mitigating environmental phosphorus pollution from manure.

Natural Occurrence

Phytase enzymes are widely distributed across various organisms in nature, reflecting their fundamental role in metabolism. In microorganisms, phytases are prominently produced by fungi such as and bacteria including , where they facilitate the breakdown of phytate, a common storage form of in . These microbial phytases are often secreted extracellularly, enabling efficient utilization of in nutrient-scarce environments like and decaying plant material. In plants, phytases occur predominantly in seeds of cereals and , such as , , and soybeans, where they are localized in protein bodies and contribute to mobilization during early growth stages. Additionally, phytases are present in animal digestive tracts, primarily through microbial communities; for instance, rumen microbes in ruminants like express phytases to hydrolyze phytate from dietary sources, while intestinal in animals such as pigs and provide limited phytase activity to enhance absorption. The evolutionary origins of phytase suggest it is an ancient that evolved to aid acquisition in -limited ecosystems, forming a key part of the global cycle. Phytases enable organisms to mineralize organic from phytate into bioavailable inorganic forms, a capability that likely emerged early in microbial lineages and was later conserved in and animals facing similar nutritional constraints. This adaptation is evident in diverse phylogenetic distributions, with phytase genes identified across , fungi, , and even some animal tissues, underscoring their ecological importance in sustaining life in environments where inorganic is scarce. Endogenous phytase activity varies significantly by source and developmental stage. In plant seeds, activity levels typically range from 0.3 to 6 U/g dry weight; for example, wheat bran exhibits 1.5–2.5 FTU/g, while grains can reach up to 6 U/g, reflecting adaptations in storage efficiency among species. Microbial sources generally display higher potential, with fungal cultures like producing up to 12.6 U/mL under natural growth conditions, often surpassing plant-derived phytases in catalytic efficiency. Activity is notably elevated in germinating seeds compared to mature grains; for instance, phytase levels in and can increase 3- to 16-fold during the first week of , supporting rapid release for development.

Historical Development

Discovery

Phytase was first described in by Japanese researchers U. , K. Yoshimura, and M. Tsujii, who identified enzymatic activity capable of hydrolyzing in rice bran extracts during investigations into the of materials. Their work revealed that this activity produced phosphates as intermediates, marking the initial recognition of phytase as a specific phosphohydrolase distinct from general actions observed in tissues. Subsequent early research in the 1910s and 1930s expanded understanding of phytase in cereal sources, with R.J. Anderson's 1915 studies in the United States demonstrating robust phytase activity in wheat bran that hydrolyzed phytin to release inorganic phosphate. This built on prior observations of organic phosphorus mobilization in grains, though initial characterizations often conflated phytase with nonspecific phosphatases due to overlapping hydrolytic effects on various substrates. By the 1960s, investigations into fungal sources intensified, with phytase activity identified in species like in 1962 and in 1968, highlighting microbial potential for higher yields compared to plant extracts; however, precise identification lagged until refined assays confirmed substrate specificity. A key challenge during this period was distinguishing phytase from general acid phosphatases, resolved only through development of substrate-specific assays using as the sole indicator, which allowed quantification of myo-inositol hexakisphosphate hydrolysis. A major milestone occurred in the 1960s when the International Union of Biochemistry formally classified phytase as EC 3.1.3.8 in 1961, specifically designating the 3-phytase form that initiates at the L-3 position of , while later distinguishing it from alkaline 6-phytase (EC 3.1.3.26). This enzymatic provided a standardized framework, enabling clearer differentiation between and alkaline variants based on optima and reaction specificity, and facilitated subsequent biochemical studies on phytase from diverse sources including fungi like Aspergillus ficuum.

Commercialization and Advances

Early attempts at commercial production occurred in the 1960s, when International Minerals & Chemicals screened over 2,000 microorganisms and identified A. niger as a promising source, though the project ended in 1968 due to insufficient efficiency. The commercialization of phytase advanced in the early 1990s with the first partial cloning of the phyA gene from Aspergillus niger in 1991, enabling the production of the enzyme on an industrial scale. This breakthrough led to the launch of Natuphos, the first commercial phytase product, by BASF in 1991, derived from A. niger and marketed as a feed additive to enhance phosphorus bioavailability in animal nutrition. In the 1990s, production shifted toward recombinant systems to achieve higher yields and scalability, with the Escherichia coli appA gene cloned in 1999 and expressed in hosts like Pichia pastoris and E. coli itself, marking the transition to second-generation phytases with improved . By the 2000s, focus turned to thermostable variants, such as those from E. coli retaining over 70% activity at 80–90°C, designed to withstand high-temperature feed pelleting processes. Recent developments as of 2024 have emphasized , including techniques that enhance by up to 20°C, allowing variants to resist temperatures for improved industrial durability. Immobilization methods, such as entrapment in nanoclay or covalent binding to supports, have enabled reuse in feed processing, reducing costs and environmental impact. A 2024 review highlights genetic modifications, like , that boost catalytic efficiency by 2- to 5-fold while maintaining broad pH optima. Regulatory milestones include U.S. Food and Drug Administration (FDA) approval of Natuphos as (GRAS) for in 1995, facilitating widespread adoption. In the , authorizations expanded during the 2000s, with the (EFSA) approving multiple phytase products like Ronozyme and Finase between 2007 and 2012 for use in and swine diets.

Enzyme Classes and Structures

Histidine Acid Phosphatases (HAPs)

Histidine acid phosphatases (HAPs) represent the most predominant class of microbial phytases. These enzymes belong to the phosphatase superfamily and are characterized by a conserved consisting of a nucleophilic , an invariant for substrate binding, and a second or aspartate residue that acts as a general acid/base catalyst. The core fold of HAP phytases features a central α/β domain, where a mixed β-sheet of five to six strands is flanked by α-helices, often accompanied by an additional all-α domain that contributes to substrate positioning. The three-dimensional structure of HAP phytases has been elucidated through , with the PDB entry 1IHP providing a representative model for the from Aspergillus ficuum, closely related to A. niger phytase. This structure reveals a compact monomeric protein with a molecular weight typically ranging from 40 to 50 kDa, including potential sites in fungal variants that enhance stability. In some HAP structures, the β-strands form a pseudo-seven-bladed propeller-like arrangement around the cleft, facilitating access to the phytate substrate while maintaining the overall α/β . HAP phytases exhibit unique biochemical features tailored to acidic environments, with an optimal pH range of 2.5 to 5.5 that aligns with the conditions in animal digestive tracts and microbial habitats. They demonstrate high specificity for myo-inositol hexakisphosphate (IP6, or ), initiating at the L-3 position and proceeding stepwise to release up to five phosphate groups. A distinctive aspect of their catalytic pathway involves intramolecular following the initial , where the enzyme promotes a cyclized intermediate from the pentaphosphate product to enhance efficiency. Prominent examples of HAP phytases include those from fungal sources such as and species, which are widely studied for their , as well as bacterial variants from . These enzymes share conserved sequence motifs, notably the RHGXRXP sequence encompassing the nucleophilic in the , which is essential for phosphohistidine intermediate formation during catalysis.

β-Propeller Phytases

β-Propeller phytases represent a distinct class of bacterial enzymes characterized by their unique structural architecture, primarily originating from such as species of the genus . These enzymes adopt a six-bladed β-propeller fold, consisting of five four-stranded and one five-stranded antiparallel β-sheets arranged around a central axis, forming a compact monomeric structure with a molecular weight of approximately 40 . The propeller-like configuration creates a central that serves as the substrate binding site, distinguishing this class from the more common histidine acid phosphatases (HAPs), which exhibit an acidic pH optimum and different fold. The active site of β-propeller phytases is located within this central region and features a cleavage site responsible for substrate hydrolysis and an adjacent affinity site for additional phosphate stabilization, with key residues such as aspartates, glutamates, and histidines facilitating interactions with the substrate and metal ions. Unlike HAPs, these phytases are calcium-dependent, requiring multiple Ca²⁺ ions (up to six or more) for structural integrity, thermostability, and catalytic function; for instance, the structure of the Bacillus subtilis enzyme (PDB: 3AMR) reveals 11 potential metal-binding sites, with several critical for phytate binding. This calcium coordination enhances the enzyme's ability to hydrolyze phytate in a stereospecific manner, releasing phosphate groups sequentially. Functionally, β-propeller phytases exhibit an alkaline pH optimum of 7.0–7.8 and demonstrate a broad substrate specificity, capable of hydrolyzing not only phytate but also related compounds like ATP and lower inositol phosphates. Their specific activity can reach up to approximately 100 U/mg, higher than some HAP variants in neutral conditions, though they generally display lower thermostability compared to fungal HAPs, limiting their commercial prevalence despite evolutionary prevalence in Gram-positive bacteria. These traits make them suitable for applications requiring neutral pH activity, such as in animal feed processing.

Purple Acid Phosphatases (PAPs)

Purple acid phosphatases (PAPs) represent a class of metalloenzymes primarily from and fungi that exhibit phytase activity, hydrolyzing (myo-inositol hexaphosphate, IP6) and other phosphate esters. These enzymes are distinguished by their binuclear metal center in the , which typically consists of Fe(III) coordinated with Fe(II), Zn(II), or Mn(II), enabling acid function at low . In such as , the center is Fe(III)-Mn(II), while in and red , it is Fe(III)-Zn(II); fungal PAPs share similar compositions. The dinuclear core motif is highly conserved, involving seven residues that coordinate the metals: typically two histidines and an aspartate for the Fe(III) site, three histidines or aspartates/glutamates for the second metal, and a bridging aspartate. This motif facilitates substrate binding and , with the metals polarizing the phosphate group for nucleophilic attack. Plant PAPs are larger enzymes, ranging from 50-60 kDa per subunit, and can exist as homodimers (e.g., red PAP, ~55 kDa subunits forming a 111 kDa dimer) or monomers (e.g., PAPhy isoforms). Structural studies of PAPhy reveal a 510-residue with the binuclear center in a central β-sheet domain, as resolved in crystal structures (PDB: 6GIZ). Kidney bean PAP structures (PDB: 1KBP) similarly highlight the dimeric assembly stabilized by disulfide bridges in some isoforms. PAPs display a broad optimal range of 4-6, with peak activity around 5.0-5.5, allowing function in acidic environments like plant vacuoles or germinating seeds. They exhibit lower substrate specificity compared to other phytase classes, hydrolyzing not only IP6 but also other monophosphate esters such as p-nitrophenyl phosphate, ATP, and phosphates. This versatility supports their role in phosphorus stress responses, where PAPs are upregulated during phosphate starvation to scavenge extracellular organic , enhancing acquisition and recycling in like and . Despite their physiological importance, PAPs have limitations as phytases, including lower on IP6 (typically 100-200 U/mg, compared to over 500 U/mg for histidine acid phosphatases) and sensitivity to inhibitors like , which binds the binuclear center and disrupts (Ki ~1 mM). These properties make PAPs less efficient for high-throughput applications but crucial for adaptation to nutrient-limited conditions.

Protein Tyrosine Phosphatase-Like Phytases

Protein tyrosine phosphatase-like phytases (PTPLPs), also known as cysteine phytases, represent a rare class of bacterial enzymes that catalyze the hydrolysis of phytic acid (myo-inositol hexakisphosphate, IP6) without requiring metal cofactors. These enzymes belong to the protein tyrosine phosphatase (PTP) superfamily and share the characteristic PTP fold, consisting of a central β-sheet surrounded by α-helices, which forms a shallow active site pocket suitable for binding the inositol ring of IP6. The signature catalytic motif is HCXAGXGR(S/T), where the conserved cysteine acts as a nucleophile, forming a phosphocysteine intermediate during phosphate transfer, facilitated by a general acid residue (typically aspartate) in the WPD loop. Unlike metal-dependent phytases, PTPLPs rely solely on this cysteine-based mechanism, with no metals involved in catalysis. Structurally, PTPLPs are compact monomers with molecular weights ranging from approximately 25 to 38 kDa, as exemplified by the enzyme PhyAsr from the rumen bacterium Selenomonas ruminantium, which features a 5-stranded mixed β-sheet core flanked by helical bundles and a depth of about 7.7–14.9 in the electropositive cleft for substrate accommodation. Other bacterial examples include enzymes from Bdellovibrio bacteriovorus and , highlighting their presence in diverse environments, including pathogenic contexts. The includes positively charged residues, such as in the P-loop, that stabilize the negatively charged IP6, enabling specific recognition and . These phytases exhibit unique biochemical properties, including an acidic optimum of 4.5–5.5, and sequential of IP6 beginning at the D-3 position (3-phosphate on the myo- ring), with pathways varying to include starts at the 5-position in some variants, progressing to lower inositol phosphates like IP5 and IP2. However, they generally display low , with optimal activity around 50–60°C and sensitivity to high (>50 mM), limiting their robustness compared to other phytase classes. Evolutionarily, PTPLPs are derived from the ancient PTP superfamily, which primarily dephosphorylates residues in eukaryotic signaling, but have adapted in for phytate degradation, particularly in nutrient-scarce or host-associated niches of pathogenic . This adaptation involves modifications to the substrate-binding pocket to favor polyphosphorylated inositols over phosphotyrosine, reflecting an evolutionary divergence for ecological roles in acquisition.

Biochemical Properties

Catalytic Mechanism

Phytases catalyze the of (myo-inositol hexakisphosphate, IP6) through a mechanism at the bond, initiating the stepwise release of inorganic (Pi). The general reaction can be simplified as: IP6+H2O→IP5+HPO42−\text{IP}_6 + \text{H}_2\text{O} \rightarrow \text{IP}_5 + \text{HPO}_4^{2-} This process lowers the energy barrier of the reaction by stabilizing the via residues and metal ions, facilitating bond cleavage without requiring cofactors in most cases. The kinetics of phytase action typically follow Michaelis-Menten behavior, depending on the enzyme source and conditions; these parameters reflect efficient substrate binding and turnover under physiological , with rate constants showing pH dependence that peaks in acidic to neutral ranges. In histidine acid phosphatases (HAPs; EC 3.1.3.8), the dominant class of phytases, the mechanism proceeds in two steps: a conserved residue (e.g., His-82 in the RHG motif) acts as the , attacking the atom of the scissile to form a covalent phosphohistidine intermediate, followed by of this intermediate by a activated by an aspartate residue (e.g., Asp-362) as the general base, with a glutamate (e.g., Glu in the HD motif) serving as the proton donor to facilitate departure of the inositol product. β-Propeller phytases (EC 3.1.3.26) employ a non-covalent mechanism where a calcium-activated performs an inline nucleophilic attack on the , forming a pentavalent stabilized by multiple calcium ions and residues such as glutamates in the DAADDPAIW motif, without a covalent enzyme-substrate intermediate; a may assist in proton transfer in some variants. Purple acid phosphatases (PAPs) utilize a metal-bound (coordinated by iron or other metals in motifs like GHxH) for nucleophilic attack on the , potentially involving modulation of the metal center to enhance reactivity, leading to direct and release of Pi while stabilizing the through bimetallic coordination. Protein tyrosine phosphatase-like phytases feature a cysteine residue (in the CxxxxxR motif) whose thiolate anion acts as the nucleophile, forming a phosphocysteine intermediate that is subsequently hydrolyzed by a water molecule, with an aspartate (e.g., Asp-223) aiding proton donation.

Substrate Specificity and Dephosphorylation Pathways

Phytases exhibit high substrate specificity for myo-inositol hexakisphosphate (IP6, phytate), with relative activities often exceeding 90% compared to lower inositol phosphates, while showing variable activity toward IP5, IP4, and IP3, and minimal activity against IP2, IP1, or non-phytate phosphate esters such as p-nitrophenyl phosphate. This preference arises from the enzyme's active site accommodating the fully phosphorylated myo-inositol ring, particularly its equatorial phosphate groups, though some classes display broader tolerance for partially dephosphorylated intermediates. The pathways of phytate by phytases are class-dependent and involve stereospecific sequential , ultimately yielding myo-inositol and inorganic (Pi), though often incomplete in biological contexts, typically halting at IP3 or IP4 due to accumulating less favorable substrates. Histidine phosphatases (HAPs; EC 3.1.3.8), predominantly 3-phytases from microbial sources, initiate at the L-1 position of the myo-inositol ring, producing D-myo-inositol 1,2,4,5,6-pentakisphosphate (D-IP5) as the primary product, followed by stepwise removal of additional phosphates to generate a series of chiral intermediates. In contrast, β-propeller phytases, typically 6-phytases (EC 3.1.3.26) from bacterial origins, begin at the position 6, producing myo-inositol 1,2,3,4,5-pentakis (L-IP5) as the primary product, followed by sequential removal of neighboring phosphates initially (e.g., to Ins(1,2,3,4)P4, Ins(1,2,4)P3), with a for every second phosphate in later steps, leading to stereospecific D-chiro configurations. phosphatases (PAPs), common in , display more random positional specificity, often starting at C-6 or C-3 without strict stereopreference, enabling flexible stepwise degradation but with variable endpoint products like D-Ins(1,2,3)P3 in certain cases. These pathways highlight the role of D/L chirality in intermediate stability and enzyme binding, where removal of specific phosphates introduces that influences subsequent steps; for instance, HAPs efficiently navigate both D- and L-enantiomers, while β-propeller enzymes favor L-configurations initially. Class-dependent efficiency for complete degradation follows the order HAPs > β-propeller > PAPs, with HAPs achieving near-full to + 6Pi under optimal conditions due to their broad sequential capability, whereas β-propeller and PAPs often yield persistent IP3-IP4 remnants .

Physicochemical Characteristics

Phytases exhibit distinct optima depending on their structural class, influencing their activity in various biological and industrial environments. acid phosphatases (HAPs), the most common class, typically display optimal activity in the acidic range of 2.5–5.5, aligning with the gastrointestinal conditions of animals. In contrast, β-propeller phytases function optimally at neutral to alkaline levels of 6–8, making them suitable for applications requiring higher stability, while protein tyrosine phosphatase-like (PTP-like) phytases have optima around 4–5.5. Purple acid phosphatases (PAPs) show broader tolerance, often maintaining activity across a wide range including acidic to neutral conditions, though their optima vary between 5.0 and 6.0. Temperature optima for native phytases generally fall between 50°C and 60°C, reflecting their mesophilic origins from microbial and sources. stability varies by class; for instance, HAPs often have half-lives of 10–30 minutes at 60°C, limiting their use in high-heat processes like feed pelleting. Engineered variants, such as from tequilensis (characterized in 2019), demonstrate enhanced , retaining approximately 70% activity at 70°C, which expands their industrial applicability. β-Propeller phytases exhibit inherently higher due to calcium binding, while inhibitors like (e.g., Cu²⁺, Zn²⁺, Fe²⁺/³⁺) and can reduce activity by up to 50–90% through competitive binding or denaturation. Certain phytases, particularly β-propeller types, are activated by Ca²⁺ ions, which stabilize the structure and can increase activity by 20–50% while extending at elevated temperatures. Most phytases have acidic isoelectric points (pI) in the range of 4–6, which enhances their in the low-pH environments of feeds and gastric tracts but may lead to at neutral . This property affects their , as lower pI values promote dispersion in acidic media, improving release efficiency in applications.

Sources and Production

Natural Microbial and Plant Sources

Phytase is naturally produced by various microorganisms, with fungi and bacteria serving as primary sources. Among fungi, is a key producer, yielding up to 9.6 U/mL in submerged using potato dextrose broth at 30°C for 5 days of incubation. In solid-state with a mixed substrate of wheat bran, rice bran, and groundnut cake, yields reach 76 U/g dry substrate under similar conditions. Bacterial sources, such as , typically achieve lower yields of about 0.64 U/mL in submerged . Isolation of these phytase-producing microbes commonly employs plate assays on phytate-containing media, where translucent halos around colonies signal extracellular activity. In sources, phytase occurs endogenously in grains and , albeit at modest levels. Wheat grains exhibit phytase activity of approximately 2886 U/kg dry matter, while demonstrates the highest among cereals at 6016 U/kg dry matter. , including soybeans, show lower activities around 290 U/kg dry matter. Extraction from these materials involves grinding to a fine powder and homogenizing in a chilled buffer, such as 0.02 M Tris-HCl ( 7.6) containing 0.1% , followed by at 12,000g for 30 minutes at 4°C to obtain the supernatant for assays. Microbial sources generally outperform plant sources by 10-100 times in terms of achievable phytase activity, as processes enable higher enzyme concentrations compared to inherent plant levels. Yields vary by strain and environmental factors, with elevated activities often in bacteria; for instance, species isolated from alpine grasslands average 0.267 U/mL, exceeding non-rhizosphere isolates. Natural extraction from these sources is hindered by low enzyme purity, inconsistent activity influenced by seasonal and strain variations, and poor scalability for industrial use due to dilute products and expensive downstream purification.

Recombinant and Engineered Production

Recombinant production of phytase has revolutionized its industrial scalability by enabling heterologous expression in microbial hosts, overcoming limitations of native sources such as low yields and complex purification. In Escherichia coli, phytase genes from sources like Aspergillus niger or E. coli itself are commonly expressed intracellularly, achieving high yields through optimized cultivation, such as up to 364-fold higher activity compared to non-recombinant strains under controlled conditions. However, expression often results in inclusion bodies, necessitating refolding steps to recover active enzyme. Yeast systems, particularly Pichia pastoris, facilitate secreted expression, simplifying downstream processing and enabling high-level production. Optimization strategies, including co-expression of chaperones like protein disulfide isomerase (PDI) and use of strong promoters such as P_{HpFMD}, have increased E. coli AppA phytase yields by up to 2.9-fold, reaching several grams per liter in shake-flask cultures. Filamentous fungi like Trichoderma reesei serve as hosts for fungal phytases, with genetically modified strains producing 4-phytase at commercial scales through integrated expression cassettes. Protein engineering enhances phytase performance for demanding applications. Site-directed mutagenesis introduces stabilizing features, such as an additional disulfide bond (L28C/W360C) in E. coli AppA phytase, retaining ~50% activity after 20 minutes at 85°C—far superior to the wild-type—while maintaining kinetic parameters like kcat and KM. Directed evolution via error-prone PCR has generated variants with 12% higher residual activity post-heat treatment and up to 2.3-fold increased specific activity, alongside ~93% improved catalytic efficiency (kcat/KM). Fusion proteins, incorporating thermostable domains or targeting peptides, further boost stability and specificity, as seen in hybrids enhancing gastric retention. Industrial processes emphasize efficiency in large-scale . Fed-batch in E. coli achieves yields exceeding 130,000 U/g weight for mutant phytases, with overall 3.7-fold improvements over batch modes through controlled nutrient feeding. Purification typically involves techniques like ion-exchange and hydrophobic interaction to isolate active at >95% purity. For reusability, immobilization on mesoporous silica nanoparticles via adsorption loads up to 237 μg phytase per mg support, retaining low release (14% at pH 3) and enabling multiple cycles with minimal activity loss. Recent innovations include /Cas9-mediated editing of Komagataella phaffii (formerly Pichia pastoris), creating markerless strains with 2-fold higher phytase productivity (up to 480 U/mL) via sequential gene integration without antibiotic markers. Acid-stable variants, engineered through disulfide bonds or pH-responsive coatings, withstand gastric conditions, retaining 40% activity post-simulated digestion for improved nutrient delivery in feed applications.

Biological Roles

In Microorganisms

Phytase plays a crucial role in microbial acquisition, particularly in phosphorus-starved environments where inorganic (Pi) is limited. In , phytase expression is upregulated under low Pi conditions through the Pho regulon, a global regulatory system that coordinates the transcription of genes involved in scavenging and uptake. This regulon, controlled by the PhoR/PhoB two-component system, activates phosphatases including phytases to hydrolyze organic compounds like phytate, enabling microbes to access otherwise unavailable Pi for essential cellular processes such as and energy metabolism. For instance, in , the phytase gene phyC is transcriptionally activated under limitation via Pho regulon components, demonstrating how this mechanism integrates phytase into broader pathways. Specific examples highlight phytase's contributions to microbial and interactions. In fungi like , phytase facilitates mobilization in nutrient-poor soils. In bacteria such as Pseudomonas species, phytase production promotes adaptation to phytate-rich soils by aiding biofilm formation on root surfaces, where extracellular phytase hydrolyzes phytate to provide localized and , strengthening community structures and competitive fitness in the . Metabolically, phytase integrates into microbial nutrient utilization by generating hydrolysis products such as triphosphate (IP3), tetraphosphate (IP4), and pentaphosphate (IP5), which serve as alternative carbon and energy sources during nutrient scarcity. These lower-order phosphates can be further metabolized by microbial pathways, supporting growth when primary carbon substrates are depleted, as observed in species where phytase sequentially dephosphorylates phytate to yield utilizable IP3–IP5 intermediates. Ecologically, microbial phytases are pivotal in phosphorus cycling, transforming recalcitrant phytate into bioavailable forms that sustain and microbial diversity. In terrestrial soils, where phytate constitutes up to 60% of organic , bacterial and fungal phytases drive mineralization, preventing phosphorus lockup and supporting productivity. In guts, produce phytases that hydrolyze dietary phytate, recycling within the and enhancing host nutrient absorption while minimizing environmental phosphorus runoff from . This dual role underscores phytase's importance in maintaining balances across microbial habitats.

In Plants and Animals

In , endogenous phytases play a crucial role in by hydrolyzing (IP6), the primary storage form of in seeds, to release inorganic during . This mobilization supports growth when is limited, with studies showing up to 88% breakdown of IP6 during in cereals and . Transgenic overexpression of phytase genes in has been shown to improve efficiency by increasing phytate hydrolysis in and seeds, leading to enhanced nutrient uptake from organic sources in . For instance, expression of microbial phytase genes like phyA or appA in crops such as Brassica napus results in 10-20% higher yields under -limited conditions, alongside greater biomass and seed production, due to better utilization of phytate-bound . In animals, endogenous phytase activity varies widely across species, being notably low in monogastrics like pigs, where intestinal levels are insufficient to hydrolyze significant amounts of dietary phytate, but higher in the intestines of certain fish species such as , facilitating partial phytate degradation in aquatic environments. Phytase contributes to absorption by breaking down phytate, which otherwise chelates essential cations like calcium, iron, and , forming insoluble complexes that reduce in the digestive tract. Phytase deficiency, often exacerbated by high-phytate diets, is linked to shortages that manifest as in growing birds, characterized by impaired mineralization and skeletal deformities due to inadequate release from phytate. In animals, exogenous phytase supplementation mimics the phytate-hydrolyzing action of microbial enzymes found in foreguts, improving and utilization in lacking robust endogenous activity. For human health, breeding low-phytate crops addresses mineral malnutrition by reducing phytate's antinutritional effects, thereby enhancing the of iron and in staple foods consumed in deficiency-prone regions. In ruminants, exogenous phytase exhibits with endogenous microbial enzymes in the , amplifying phytate and phosphorus release from plant-based feeds without fully replacing bacterial contributions.

Applications

In Animal Feed and Nutrition

Phytase supplementation constitutes the primary application of the , predominantly in feeds for and to enhance utilization from plant-based ingredients. In and diets, standard dosages range from 500 to 2000 FTU/kg, enabling the release of 30-60% of bound phytate- and improving overall nutrient . This supplementation reduces the need for inorganic additives by 40-50%, thereby lowering feed formulation costs and minimizing environmental through a 30-50% decrease in excretion. Beyond phosphorus management, phytase enhances digestibility by 5-10% in broilers and pigs, supporting improved protein efficiency and growth performance without excessive dietary protein levels. Economic benefits include feed cost savings of approximately $4-12 per metric ton, depending on dosage and market conditions for inorganic phosphates, making it a cost-effective strategy for intensive animal production. Formulations such as granulated or coated phytase are designed for thermal stability during pelleting processes, retaining over 80% activity at temperatures up to 85-95°C, which is essential for mash or pellet feed manufacturing. Superdosing at levels up to 10,000 FTU/kg maximizes phytate degradation, further boosting energy utilization and weight gain in by relaxing matrix values and countering antinutritional effects more comprehensively. Species-specific applications highlight higher dosages and thermostable variants for broilers to withstand gastrointestinal conditions, while emerging use in aquafeeds for at 500-2000 FTU/kg improves retention and growth comparable to inorganic supplements, reducing effluent loads in .

In Agriculture, Environment, and Industry

In , phytase can be applied as a amendment to hydrolyze phytate, enhancing availability for crops in -limited soils by breaking down insoluble phytate complexes. This approach promotes efficient cycling and reduces reliance on synthetic fertilizers. Additionally, transgenic crops engineered for low-phytate traits, such as lines developed through of the , minimize phytate accumulation in seeds, improving bioavailability without external addition; notable examples include low-phytate inbreds released in 2023. In environmental applications, phytase treatment of animal hydrolyzes phytate-bound , reducing soluble content and subsequent runoff into waterways, which helps mitigate . Studies show that incorporating phytase in animal diets prior to manure production can decrease excreted by 20-35%, thereby lowering runoff by up to 40% when combined with other stabilization methods like aluminum . As of 2025, research highlights phytase's integration into biofertilizers, where microbial phytase-producing strains enhance sustainable cycling by solubilizing soil-bound phytate, supporting closed-loop nutrient systems in . In industrial contexts, phytase facilitates food processing by degrading phytic acid in grains and legumes, improving mineral accessibility. For instance, during bread production, phytase supplementation or fermentation with phytase-active yeasts reduces phytic acid levels by 40-50%, enhancing the nutritional profile of whole-grain products. In soy fermentation processes, similar enzymatic action decreases phytic acid by up to 50%, aiding in the production of soy-based foods with better zinc and iron bioavailability. Phytase also finds use in pharmaceuticals as a supplement to boost zinc bioavailability from phytate-rich diets, with microbial phytase increasing apparent zinc absorption in controlled studies. Emerging applications include biofuel pretreatment, where thermostable phytase hydrolyzes phytate in corn mash during ethanol production, releasing inorganic phosphate to support yeast fermentation and yielding environmental benefits through reduced waste. Regarding human health, phytase holds potential in fortified foods to prevent in developing regions by countering phytic acid's inhibition of iron absorption from staple cereals. Phytase-mediated dephytinization in powders or fermented porridges can increase iron by 50-100% in phytate-rich meals, offering a food-based strategy to address in populations reliant on plant-based diets.

References

  1. https://www.sciencedirect.com/topics/agricultural-and-biological-sciences/phytase
  2. https://www.extension.purdue.edu/extmedia/as/as-560-w.pdf
  3. https://pmc.ncbi.nlm.nih.gov/articles/PMC10982552/
  4. https://www.frontiersin.org/journals/microbiology/articles/10.3389/fmicb.2023.1127249/full
  5. https://www.annualreviews.org/doi/10.1146/annurev-animal-031412-103717
  6. https://pubmed.ncbi.nlm.nih.gov/12876797/
  7. https://straitsresearch.com/report/animal-feed-phytase-market
  8. https://www.webmd.com/vitamins/ai/ingredientmono-1619/phytase
  9. https://enzyme.expasy.org/EC/3.1.3.8
  10. https://www.collinsdictionary.com/us/dictionary/english/phytase
  11. https://www.sciencedirect.com/topics/agricultural-and-biological-sciences/3-phytase
  12. https://iubmb.qmul.ac.uk/enzyme/EC3/1/3/8.html
  13. https://pmc.ncbi.nlm.nih.gov/articles/PMC10581959/
  14. https://pmc.ncbi.nlm.nih.gov/articles/PMC10128089/
  15. https://pmc.ncbi.nlm.nih.gov/articles/PMC4325021/
  16. https://pmc.ncbi.nlm.nih.gov/articles/PMC9740087/
  17. https://pmc.ncbi.nlm.nih.gov/articles/PMC4368368/
  18. https://pmc.ncbi.nlm.nih.gov/articles/PMC7455620/
  19. https://www.jbc.org/article/S0021-9258(18)88211-4/fulltext
  20. https://academic.oup.com/bbb/article-abstract/32/10/1275/5977764
  21. https://www.genome.jp/dbget-bin/www_bget?ec:3.1.3.8
  22. https://golbid.com/wp-content/uploads/2018/02/Phytase-a-New-Life-for-an.pdf
  23. https://www.openagrar.de/servlets/MCRFileNodeServlet/Document_derivate_00011870/A1312.pdf
  24. https://pmc.ncbi.nlm.nih.gov/articles/PMC11695551/
  25. https://journals.asm.org/doi/10.1128/aem.00506-22
  26. https://www.sciencedirect.com/science/article/abs/pii/S096085242300456X
  27. https://downloads.regulations.gov/FDA-2012-S-1178-0016/attachment_1.pdf
  28. https://doi.org/10.3389/fmicb.2023.1127249
  29. https://doi.org/10.1016/j.crstbi.2024.100139
  30. https://doi.org/10.1016/S0969-2126(01)00637-2
  31. https://pmc.ncbi.nlm.nih.gov/articles/PMC9073318/
  32. https://www.researchgate.net/publication/8905813_Plant_purple_acid_phosphatases_-_Genes_structures_and_biological_function
  33. https://mural.maynoothuniversity.ie/id/eprint/3708/1/GS_Binuclear_metal.pdf
  34. https://www.rcsb.org/structure/1KBP
  35. https://www.[researchgate](/page/ResearchGate).net/publication/8905813_Plant_purple_acid_phosphatases_-_Genes_structures_and_biological_function
  36. https://pmc.ncbi.nlm.nih.gov/articles/PMC3135953/
  37. https://academic.oup.com/plphys/article/151/1/199/6108477
  38. https://pubmed.ncbi.nlm.nih.gov/10433699/
  39. https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0094403
  40. https://www.jbc.org/article/S0021-9258(20)65387-X/pdf
  41. https://pmc.ncbi.nlm.nih.gov/articles/PMC2206706/
  42. https://doi.org/10.1110/ps.062738307
  43. https://www.researchgate.net/publication/5493166_A_protein_tyrosine_phosphatase-like_inositol_polyphosphatase_from_Selenomonas_ruminantium_subsp_lactilytica_has_specificity_for_the_5-phosphate_of_myo-inositol_hexakisphosphate
  44. https://www.sciencedirect.com/science/article/pii/S2665928X24000163
  45. https://www.jbc.org/article/S0021-9258(17)50652-3/fulltext
  46. https://www.sciencedirect.com/science/article/pii/S0969212601006372
  47. https://www.sciencedirect.com/science/article/pii/S2590346222000529
  48. https://link.springer.com/article/10.1007/s00253-003-1345-0
  49. https://enviromicro-journals.onlinelibrary.wiley.com/doi/full/10.1111/j.1365-2672.2011.05181.x
  50. https://pubmed.ncbi.nlm.nih.gov/14586576/
  51. https://cdnsciencepub.com/doi/10.1139/W07-015
  52. https://link.springer.com/article/10.1007/s00284-023-03471-1
  53. https://pmc.ncbi.nlm.nih.gov/articles/PMC2735976/
  54. https://pmc.ncbi.nlm.nih.gov/articles/PMC2266880/
  55. https://www.researchgate.net/publication/374742666_Phytases_-Types_Sources_and_Factors_Affecting_Their_Activity
  56. https://www.mdpi.com/2073-4344/10/8/844
  57. https://www.researchgate.net/publication/334113675_Characterization_of_a_Thermostable_Acidic-Phytase_from_Bacillus_tequilensis_Dm018_Medium_Optimization_by_Response_Surface_Methodology
  58. https://pubmed.ncbi.nlm.nih.gov/25416530/
  59. https://pubmed.ncbi.nlm.nih.gov/18221011/
  60. https://journals.asm.org/doi/pdf/10.1128/aem.65.2.359-366.1999
  61. https://pmc.ncbi.nlm.nih.gov/articles/PMC7034034/
  62. https://journals.sagepub.com/doi/10.4137/MBI.S10402
  63. https://pmc.ncbi.nlm.nih.gov/articles/PMC4746705/
  64. https://microbialcellfactories.biomedcentral.com/articles/10.1186/s12934-020-01499-7
  65. https://pmc.ncbi.nlm.nih.gov/articles/PMC7008902/
  66. https://www.efsa.europa.eu/en/efsajournal/pub/5826
  67. https://biotechnologyforbiofuels.biomedcentral.com/articles/10.1186/s13068-021-01936-8
  68. https://www.frontiersin.org/journals/bioengineering-and-biotechnology/articles/10.3389/fbioe.2023.1167530/full
  69. https://www.sciencedirect.com/science/article/pii/S1389172314000760
  70. https://link.springer.com/article/10.1007/s00449-014-1236-z
  71. https://pubs.acs.org/doi/10.1021/acsomega.0c04696
  72. https://www.researchgate.net/publication/374122346_CRISPRCas9-Mediated_Genome_Editing_of_the_Komagataella_phaffii_to_Obtain_a_Phytase-Producer_Markerless_Strain
  73. https://www.biorxiv.org/content/10.1101/2025.04.18.649602v1.full-text
  74. https://pmc.ncbi.nlm.nih.gov/articles/PMC1595534/
  75. https://www.frontiersin.org/journals/microbiology/articles/10.3389/fmicb.2015.00402/full
  76. https://www.sciencedirect.com/science/article/pii/S0944501319304823
  77. https://www.mdpi.com/2073-4395/14/12/3029
  78. https://www.frontiersin.org/journals/microbiology/articles/10.3389/fmicb.2022.1019383/full
  79. https://academic.oup.com/femsle/article/247/2/231/491354
  80. https://link.springer.com/article/10.1007/s42729-024-02167-0
  81. https://dr.lib.iastate.edu/bitstreams/f6969a8e-4fce-4547-9005-e224207a3f39/download
  82. https://journals.sbmu.ac.ir/afb/article/view/48493
  83. https://www.academia.edu/63484796/Effect_of_germination_on_the_phytase_activity_phytate_and_total_phosphorus_contents_of_rice_Oryza_sativa_maize_Zea_mays_millet_Panicum_miliaceum_sorghum_Sorghum_bicolor_and_wheat_Triticum_aestivum_
  84. https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0060801
  85. https://pmc.ncbi.nlm.nih.gov/articles/PMC3362705/
  86. https://www.researchgate.net/publication/226361067_Intestinal_phytase_II_A_comparison_of_activity_and_in_vivo_phytate_hydrolysis_in_three_teleost_species_with_differing_digestive_strategies
  87. https://www.merckvetmanual.com/poultry/nutrition-and-management-poultry/mineral-deficiencies-in-poultry
  88. https://repository.lsu.edu/cgi/viewcontent.cgi?article=1630&context=animalsciences_pubs
  89. https://www.sciencedirect.com/science/article/pii/S0022316622149564
  90. https://academic.oup.com/ijfst/article/37/7/759/7912909
  91. https://www.journalofdairyscience.org/article/S0022-0302%2807%2971897-0/pdf
  92. https://www.sciencedirect.com/science/article/pii/S2666154323000662
  93. https://www.allaboutfeed.net/animal-feed/feed-additives/reducing-poultry-feed-costs-with-enzyme-supplements/
  94. https://www.huvepharma.com/news/article/requirements-of-a-phytase-to-formulate-feed-free-from-inorganic-p-sources/
  95. https://www.researchgate.net/publication/386413868_OPEN_ACCESS_International_Journal_of_Poultry_Science_Research_Article_Effectiveness_of_Superdosing_Phytase_to_Low_Phosphorus_and_Calcium_Diet_on_Growth_Performance_Nutrient_Digestibility_Blood_Paramet
  96. https://link.springer.com/article/10.1007/s10499-023-01295-1
  97. https://pmc.ncbi.nlm.nih.gov/articles/PMC9261192/
  98. https://www.publish.csiro.au/cp/CP22238
  99. https://extension.missouri.edu/publications/g9182
  100. https://www.preprints.org/manuscript/202504.1638
  101. https://www.sciencedirect.com/science/article/abs/pii/S002364381830077X
  102. https://microbiologyjournal.org/impacts-and-industrial-applications-of-phytic-acid-and-phytase/
  103. https://www.sciencedirect.com/science/article/pii/S0278691597001178
  104. https://www.researchgate.net/publication/281536702_Phytase_in_fuel_ethanol_production_offers_economical_and_environmental_benefits
  105. https://pmc.ncbi.nlm.nih.gov/articles/PMC3775243/
  106. https://journals.sagepub.com/doi/pdf/10.1177/15648265130342S111
Add your contribution
Related Hubs
User Avatar
No comments yet.