Microbial Phosphatases in Sustainable Agriculture: Mechanisms, Applications, and Future Directions

Microbial Phosphatases in Sustainable Agriculture: Mechanisms, Applications, and Future Directions

*¹Oke Kingsley Oyediran., ²Ogunmola Oluranti Olagoke., ²Sodamade Abiodun

¹Department of Biology, Faculty of Science Education, Emmanuel Alayande University of Education, Oyo Oyo State, Nigeria

²Department of Chemistry, Faculty of Science Education, Emmanuel Alayande University of Education, Oyo Oyo State, Nigeria

DOI: https://doi.org/10.51584/IJRIAS.2025.100600111

Received: 07 June 2025; Accepted: 12 June 2025; Published: 16 July 2025

ABSTRACT

Microbial phosphatases are essential for sustainable agriculture, converting organic phosphorus into plant-accessible forms and decreasing reliance on synthetic fertilizers. This review explores their mechanisms, practical applications, and current limitations in enhancing phosphorus use efficiency. Research highlights that these enzymes increase phosphorus availability by 20–40% in diverse crops through rhizosphere processes, facilitated by root exudates and specialized microbial activity. Innovations in genetic engineering, nano-biotechnology, and precision farming show potential for improving enzyme functionality in real-world agricultural systems. Despite these advances, challenges persist, such as inconsistent inoculant survival, competition with indigenous soil microbes, and performance variability across different soils. To overcome these hurdles, future efforts should prioritize large-scale field validations, cost-benefit analyses, and hybrid solutions integrating tailored microbial communities with advanced delivery technologies. Addressing these gaps could enable microbial phosphatases to play a transformative role in sustainable farming, balancing productivity with environmental conservation. This review consolidates key insights for scientists and agronomists working on microbial solutions for nutrient management. By bridging fundamental research and practical implementation, it underscores the need for interdisciplinary collaboration to harness microbial phosphatases effectively. Emerging tools like meta-omics and CRISPR-based engineering could further refine these biological systems, while policy support and farmer education will be critical for adoption. Ultimately, optimizing microbial phosphatase applications promises to support global food security while reducing agriculture’s ecological footprint.

Keywords: microbial phosphatases, phosphorus mineralization, sustainable agriculture, rhizosphere engineering, biofertilizers

INTRODUCTION

Microbial phosphatases are enzymes secreted or bound to cells of soil-dwelling bacteria, fungi, and actinomycetes that break down organic phosphorus compounds into soluble inorganic phosphate, making it accessible to plants (Nannipieri et al., 2011). Key types, such as acid phosphatases, alkaline phosphatases, and phytases, drive phosphorus mineralization by decomposing phytates and organic phosphoesters, thereby enhancing phosphorus bioavailability in agricultural soils (Richardson & Simpson, 2011). By facilitating natural phosphorus recycling, these enzymes support sustainable farming practices—diminishing reliance on synthetic fertilizers, boosting crop productivity, and maintaining long-term soil fertility (Sharma et al., 2013). Their application offers an eco-friendly solution to improve nutrient use efficiency in agroecosystems. Sustainable agriculture adopts environmentally sound approaches to preserve soil quality, ensure crop yields, and protect ecosystems (Gomiero et al., 2011). Soil microbes such as bacteria, fungi, and actinomycetes are crucial for key processes including nutrient transformation (nitrogen fixation, phosphate release), breakdown of organic materials, and stimulation of plant development (Bender et al., 2016). These organisms diminish the need for synthetic fertilizers by improving nutrient supply and controlling diseases, thereby fostering robust and ecologically balanced farming systems.

Phosphorus (P) serves as a fundamental nutrient for plants, playing critical roles in energy metabolism through ATP formation, genetic material production, and cellular membrane development (Vance et al., 2003). Although soils typically contain substantial P reserves, the majority becomes chemically bound to iron, aluminum, and calcium ions, rendering approximately 70-80% of soil phosphorus inaccessible to plant roots (Shen et al., 2011). This limited bioavailability is exacerbated by phosphorus’s natural immobility in soil and its tendency to become fixed, particularly in acidic tropical and alkaline temperate soils, forcing farmers to rely heavily on expensive phosphate fertilizers with significant ecological consequences (Sharpley et al., 2013).

This review explores the biochemical functioning of microbial phosphatases in phosphorus release and organic matter breakdown, highlighting their role in nutrient cycling. It assesses practical uses in farming systems, particularly as eco-friendly alternatives to chemical fertilizers, while addressing real-world constraints like environmental adaptability and microbial persistence. The discussion extends to emerging innovations, including microbial biotechnology and nano-enhanced formulations, that could improve performance. Ultimately, the analysis aims to demonstrate how these biological tools can transform agricultural practices by balancing productivity with ecological sustainability through improved phosphorus management.

Microbial Phosphatases: Types and Biochemical Characteristics

Acid Phosphatases

Acid phosphatases (EC 3.1.3.2) represent a crucial class of enzymes that function most efficiently in acidic conditions (pH <7), with significant production observed in fungal species such as Aspergillus and Penicillium, as well as in plant root systems (George et al., 2011). These biological catalysts play a fundamental role in soil phosphorus cycling by facilitating the hydrolysis of phosphomonoester bonds in organic compounds, thereby converting bound phosphorus into soluble forms that plants can readily absorb. Their ecological importance is particularly pronounced in acidic agricultural soils, where research indicates they may be responsible for as much as 60% of total phosphorus mineralization activity (Margalef et al., 2021). Fungal acid phosphatases exhibit remarkable biochemical stability under varying environmental conditions, and their production is typically enhanced when phosphorus becomes limited in the soil environment (Nannipieri et al., 2011). This adaptive response makes them vital components in plant nutrient acquisition strategies, especially in phosphorus-deficient ecosystems where they significantly contribute to maintaining soil fertility and supporting plant growth.

Alkaline Phosphatases

Alkaline phosphatases (EC 3.1.3.1) are a key group of enzymes that achieve maximum activity in alkaline conditions (pH >7), with their primary sources being soil-dwelling bacteria such as Bacillus and Pseudomonas species (Tan et al., 2020). These metalloenzymes, which often require zinc (Zn²⁺) or magnesium (Mg²⁺) ions as cofactors, play a vital role in phosphorus cycling by catalyzing the removal of phosphate groups from various organic substrates. Their targets include structurally diverse molecules ranging from nucleotides to membrane phospholipids, making them essential for nutrient recycling in soil ecosystems.

The performance of alkaline phosphatases is closely tied to soil characteristics, showing highest activity in neutral to alkaline soils with substantial organic matter content (Sinsabaugh et al., 2008). Their contribution to phosphorus mineralization is particularly significant in calcareous soils, where they account for 30-50% of the total phosphate release (Stursova & Sinsabaugh, 2008). This substantial involvement in nutrient transformation processes highlights their critical role in maintaining soil fertility and supporting plant growth in alkaline environments.

Phytases

Phytases (EC 3.1.3.8/26) represent a specialized class of phosphatases that target phytate (myo-inositol hexakisphosphate), which constitutes the largest organic phosphorus pool in agricultural soils (Singh & Satyanarayana, 2015). These enzymes are predominantly synthesized by soil microorganisms, including fungal species (Aspergillus) and bacteria (Bacillus), which sequentially hydrolyze phytate molecules to release up to five inorganic phosphate groups through stepwise dephosphorylation (Lei et al., 2013). In agricultural systems, phytase application has been shown to significantly improve phosphorus availability, increasing bio-accessible phosphate levels by 20-40% in soils with high phytate content, thereby decreasing dependence on conventional phosphate fertilizers (Menezes-Blackburn et al., 2018). The biotechnological importance of phytases has grown substantially with the development of genetically engineered thermostable variants. These advanced formulations now find extensive application in animal nutrition and soil amendment products, where they enhance phosphorus utilization efficiency while reducing environmental phosphorus pollution. Their ability to unlock bound phosphorus from plant-derived organic matter makes them invaluable tools for sustainable agriculture and animal production systems.

Phosphodiesterases (EC 3.1.4.-)

Phosphodiesterases play a critical role in soil phosphorus cycling by breaking down complex nucleic acids and phospholipids into simpler phosphomonoesters. These enzymes are primarily synthesized by soil bacteria (particularly Streptomyces species) and various fungal genera, acting as key agents in the decomposition of organic matter. Their activity levels directly correlate with soil organic carbon content and microbial biomass density, making them important indicators of soil health (Turner & Haygarth, 2005). In the rhizosphere, phosphodiesterases enhance phosphorus availability by 15-25% through the continuous breakdown of plant-derived nucleotides and microbial DNA residues. This enzymatic action is particularly crucial in no-till agricultural systems where organic phosphorus accumulates (Jorquera et al., 2008). Recent studies suggest their potential application in bioremediation of phosphorus-rich waste materials.

Pyrophosphatases (EC 3.6.1.1)

Pyrophosphatases serve as essential regulators of cellular phosphorus metabolism by cleaving inorganic pyrophosphate (PPi) into two orthophosphate (Pi) molecules. These magnesium-dependent enzymes exist in both soluble cytoplasmic and membrane-bound forms across diverse soil microorganisms (Kornberg, 1999). Their primary function involves preventing PPi accumulation during biosynthetic processes like DNA replication and protein synthesis, thereby maintaining cellular energy balance. In soil ecosystems, pyrophosphatases contribute significantly (5-10%) to phosphorus mineralization, particularly in microbial hotspots such as the rhizosphere and organic matter decomposition zones (Mäkelä et al., 2012). The enzymes’ activity is particularly important in young, developing soils where mineral phosphorus availability is limited. Recent advances in soil enzymology have highlighted their potential as indicators of microbial metabolic activity in different land-use systems.

Phosphonate Hydrolases (EC 3.11.1.-) 

Phosphonate hydrolases represent a specialized group of enzymes capable of cleaving the stable carbon-phosphorus (C-P) bonds found in various organophosphonates, including common herbicides like glyphosate. Soil bacteria such as Pseudomonas and Rhizobium species produce these enzymes, particularly under phosphorus-limiting conditions, as an adaptive strategy to access alternative phosphorus sources (McGrath et al., 2013). Their activity serves dual ecological functions: facilitating phosphorus cycling while simultaneously degrading persistent environmental pollutants. In agricultural soils contaminated with phosphonate herbicides, these enzymes can reduce compound persistence by 40-60%, demonstrating significant bioremediation potential (Huang et al., 2017). Recent metagenomic studies have revealed unexpected diversity in microbial C-P lyase systems, suggesting untapped biotechnological applications.

Nucleotidases (EC 3.1.3.5/6)

Nucleotidases play a pivotal role in rapid phosphorus turnover within soil ecosystems by catalyzing the dephosphorylation of various nucleotides (AMP, GMP, etc.). These enzymes are abundantly secreted by rhizosphere-competent bacteria such as Burkholderia and Pseudomonas species, enabling efficient phosphorus scavenging from decaying microbial cells and root exudates (Richardson et al., 2009). Their activity facilitates extremely fast phosphorus cycling (occurring within minutes to hours) in the plant-soil interface, significantly influencing short-term phosphorus availability. Research indicates that nucleotidase activity increases by 30-50% in the presence of plant roots, demonstrating strong plant-microbe interactions (Giles et al., 2012). These enzymes are now recognized as key components in developing phosphorus-efficient cropping systems, particularly in low-input agricultural systems.

Polyphosphatases (EC 3.6.1.10/11)

Polyphosphatases are specialized enzymes that degrade inorganic polyphosphate chains (polyP), which serve as important phosphorus storage compounds in many soil microorganisms and mycorrhizal fungi. Fungi such as Aspergillus and ectomycorrhizal species express these enzymes during phosphorus starvation, systematically releasing orthophosphate from intracellular polyP granules (Ezawa et al., 2004). In soil environments, polyphosphatases regulate long-term phosphorus storage and release dynamics, particularly in phosphorus-rich systems. Recent studies have shown that these enzymes can remain active in soil for weeks, gradually releasing plant-available phosphorus (Ogawa et al., 2021). Their unique properties have attracted interest for developing slow-release phosphorus fertilizers and managing phosphorus pollution in agricultural runoff. Advanced molecular techniques are now being used to engineer polyphosphatase-producing microbes for improved phosphorus management in agroecosystems.

Key Microbial Producers Of Phosphatase

Bacillus species

Bacillus species are among the most efficient bacterial producers of alkaline phosphatases, with notable strains including B. subtilis, B. licheniformis, and B. amyloliquefaciens. These Gram-positive, spore-forming bacteria secrete metalloenzymes (Zn²⁺/Mg²⁺-dependent) that remain active in alkaline soils (pH 7-9) (Nannipieri et al., 2011). Their phosphatase activity increases 3-5-fold under phosphorus limitation through Pho regulon activation (Sharma et al., 2013). Bacillus strains demonstrate exceptional environmental persistence due to endospore formation, making them ideal for biofertilizer formulations. Field trials with B. megaterium inoculants increased wheat phosphorus uptake by 18-22% in calcareous soils (Tarafdar & Claassen, 1988). Recent genetic engineering of B. subtilis has enhanced phytase production 2.3-fold, enabling more efficient animal feed additives (Lei et al., 2013).

Pseudomonas species

Pseudomonas species, particularly P. putida and P. fluorescens, are prolific producers of acid and alkaline phosphatases with broad substrate specificity. These Gram-negative bacteria exhibit 30-50% higher phosphatase activity than rhizosphere averages due to their efficient Pho regulon regulation (Richardson & Simpson, 2011). Their enzymes function across pH 4-9, making them adaptable to diverse soils (George et al., 2011). Pseudomonas phosphatases play dual roles in phosphorus acquisition and pathogen suppression through competitive exclusion (Menezes-Blackburn et al., 2018). Strain P. aeruginosa PSB13 shows exceptional phytase activity (120 U/mg protein), enabling 95% phytate hydrolysis in vitro (Singh & Satyanarayana, 2015). Field applications demonstrate 15% yield increases in legumes when co-inoculated with arbuscular mycorrhizal fungi (Sharma et al., 2013).

Aspergillus species

Fungi in the Aspergillus genus, especially A. niger and A. fumigatus, dominate fungal phosphatase production in acidic soils (pH 3-6). They secrete high-activity acid phosphatases (200-500 U/g biomass) and phytases that hydrolyze 60-80% of soil phytate (Margalef et al., 2021). A. niger NRRL 3135 produces a thermostable phytase (PhyA) retaining 80% activity at 60°C, revolutionizing animal feed additives (Lei et al., 2013). These fungi employ sophisticated regulatory mechanisms, increasing phosphatase synthesis 10-fold during phosphorus starvation (Nannipieri et al., 2011). Industrial production yields 5000 U/mL phytase using fed-batch fermentation (Singh & Satyanarayana, 2015). In acidic agricultural soils, Aspergillus inoculation improves maize phosphorus uptake by 25-30% (Tarafdar & Claassen, 1988).

Streptomyces species

Streptomyces species, particularly S. lividans and S. coelicolor, are major actinomycete producers of phosphodiesterases and alkaline phosphatases. These Gram-positive bacteria generate extracellular enzymes capable of degrading complex nucleic acids and phospholipids in neutral-alkaline soils (Turner & Haygarth, 2005). Their phosphatases show unique metal tolerance, maintaining activity at 10 mM Zn²⁺/Cu²⁺ concentrations (McGrath et al., 2013). S. thermoviolaceus produces a thermophilic phosphatase (70°C optimum) used in molecular biology applications (Kornberg, 1999). In agricultural systems, Streptomyces inoculation increases available phosphorus by 15-20% in organic matter-rich soils (Richardson et al., 2009). Their enzymes also participate in antibiotic production, linking phosphorus metabolism to secondary metabolite synthesis (Ezawa et al., 2004).

Rhizobium species

Rhizobium species, including R. leguminosarum and R. tropici, produce unique acid phosphatases that function symbiotically with legume roots. These enzymes increase phosphorus availability by 40-50% in the rhizosphere, enhancing nodulation and nitrogen fixation (Richardson & Simpson, 2011). Their phosphatases show peak activity at pH 5.5-6.5, matching the rhizosphere pH of most legumes (George et al., 2011). R. etli CFN42 contains a high-affinity phytase (Km = 0.15 mM) that improves phosphorus uptake in common bean by 35% (Sharma et al., 2013). The enzymes also participate in signal transduction during early nodule formation (Menezes-Blackburn et al., 2018). Field trials demonstrate 20-25% yield increases in soybean when co-inoculated with phosphatase-producing Bradyrhizobium strains (Tarafdar & Claassen, 1988).

Structure, Catalytic Mechanisms, And Optimal Functional Conditions Of Microbial Phosphatases

Acid Phosphatases (EC 3.1.3.2)

Structure: Acid phosphatases are typically monomeric enzymes with molecular weights ranging from 25-60 kDa. They feature a conserved α/β hydrolase fold structure that forms the catalytic core. The active site contains a highly conserved Arg-His-Gly-X-Arg-X-Pro motif, where X represents variable amino acids (Ostanin et al., 1992). Fungal acid phosphatases often possess N-linked glycosylation modifications that enhance their stability in acidic environments and protect against proteolytic degradation (Van Etten & Waymack, 1991). The three-dimensional structure reveals a deep active site pocket that accommodates various phosphomonoester substrates, with the conserved histidine residue positioned for nucleophilic attack.

Catalytic Mechanism: The enzymatic reaction proceeds through a two-step ping-pong mechanism. First, the conserved histidine residue acts as a nucleophile to attack the phosphorus atom of the substrate, forming a covalent phosphohistidine intermediate. This step is facilitated by an aspartate residue that serves as a general acid (Van Etten & Waymack, 1991). In the second step, the phosphoenzyme intermediate undergoes hydrolysis, releasing inorganic phosphate. The reaction mechanism involves inversion of configuration at the phosphorus center, consistent with an in-line displacement mechanism. The conserved arginine residues in the active site play crucial roles in substrate binding and transition state stabilization.

Optimal Conditions: Acid phosphatases exhibit maximum activity under acidic conditions, with fungal enzymes typically showing optimal activity between pH 4.0-6.0, while bacterial forms are most active at pH 5.5-6.5 (Ostanin et al., 1992). The temperature optimum ranges from 30-45°C for mesophilic variants to 50-60°C for thermophilic forms. These enzymes demonstrate broad substrate specificity, with Km values of 0.1-1.0 mM for synthetic substrates like p-nitrophenyl phosphate. Activity is strongly inhibited by molybdate (IC50 ≈ 10 μM) through competitive binding at the active site, and by fluoride ions that disrupt the catalytic mechanism (Van Etten & Waymack, 1991). The enzymes maintain stability across a wide range of environmental conditions, making them particularly effective in soil environments with fluctuating pH and temperature.

Alkaline Phosphatases (EC 3.1.3.1)

Structure: Alkaline phosphatases (APs) are homodimeric metalloenzymes with each subunit typically ranging from 50-60 kDa, giving a total molecular weight of approximately 100-120 kDa. The enzyme features a characteristic (β/α)₈ TIM barrel fold that forms the catalytic core (Kim & Wyckoff, 1991). Each active site contains two essential zinc ions (Zn²⁺) and one magnesium ion (Mg²⁺) that coordinate the phosphate group during catalysis (Coleman, 1992). The metal center is stabilized by conserved aspartate and histidine residues, while a serine residue (Ser102 in Escherichia coli AP) serves as the nucleophile. Bacterial APs often show higher catalytic efficiency compared to their eukaryotic counterparts due to structural differences in their active site architecture.

Catalytic Mechanism: The catalytic mechanism involves three key steps (Kim & Wyckoff, 1991):

• Nucleophilic attack by Ser102 on the phosphate monoester substrate
• Formation of a covalent phosphoserine intermediate
• Hydrolysis of the intermediate via metal-activated water The Zn²⁺ ions play crucial roles in substrate binding and transition state stabilization, while the Mg²⁺ ion facilitates the proper orientation of the leaving group (Coleman, 1992). The reaction proceeds with retention of configuration at the phosphorus atom, suggesting a two-step mechanism involving pseudorotation of the pentacoordinate intermediate.

Optimal Conditions: APs exhibit maximum activity in alkaline conditions, typically between pH 8.0-10.5, with slight variations depending on the source organism (Coleman, 1992). The temperature optimum ranges from 25-55°C for mesophilic forms, while marine AP variants remain active at temperatures as low as 4°C (Kim & Wyckoff, 1991). These enzymes display broad substrate specificity, with Km values around 0.1 mM for common substrates like p-nitrophenyl phosphate. Activity is strongly inhibited by inorganic phosphate (Ki ≈ 1 μM) through competitive binding at the active site, and by metal chelators like Ethylene diamine tetra acetic acid (EDTA) that remove essential metal cofactors. The enzymes maintain stability across a wide range of ionic strengths, making them particularly effective in marine and soil environments.

Phytases (EC 3.1.3.8/26)

Structure: Phytases are a specialized group of phosphatases that hydrolyze phytate (myo-inositol hexakisphosphate), the primary storage form of phosphorus in plant seeds. These enzymes are classified into two major structural families: histidine acid phytases (HAPs) and β-propeller phytases (BPPs) (Ha et al., 2000). HAPs typically range from 45-55 kDa and feature an α/β fold with a conserved RHGXRXP active site motif, while BPPs (40-50 kDa) possess a unique six-bladed β-propeller structure stabilized by calcium ions (Mullaney & Ullah, 2003). Fungal phytases (e.g., from Aspergillus species) often contain N-glycosylation sites that enhance thermostability, with carbohydrate content reaching 10-20% of total molecular weight.

Catalytic Mechanism: The hydrolysis of phytate proceeds through a two-step acid-base catalysis mechanism (Ha et al., 2000):

• Nucleophilic attack by a histidine residue (His59 in Aspergillus phytase) on the phosphorus atom
• Formation of a histidine-phosphate intermediate
• Water-mediated hydrolysis of the intermediate The reaction involves an aspartate residue (Asp339) that functions as a general acid/base, while conserved arginine residues position the substrate through interactions with phosphate groups (Mullaney & Ullah, 2003). HAPs sequentially remove phosphate groups from the myo-inositol ring, typically starting at the 3-position (D3-specificity).

Optimal Conditions: Phytases demonstrate optimal activity under acidic conditions (pH 2.5-5.5 for HAPs) or neutral-alkaline conditions (pH 7.0-8.0 for BPPs) (Ha et al., 2000). Temperature optima range from 45-60°C, with engineered thermostable variants maintaining activity up to 80°C (Lei et al., 2013). The enzymes show high specificity for phytate (Km 10-100 μM) but can also hydrolyze other phosphate esters. Activity is strongly inhibited by phosphate (Ki ≈ 50 μM) and phosphate analogs like vanadate. These properties make phytases particularly valuable for agricultural applications, where they improve phosphorus bioavailability in animal feed and reduce environmental phosphorus pollution (Singh & Satyanarayana, 2015).

Phosphodiesterases (EC 3.1.4.-)

Structure: Phosphodiesterases (PDEs) constitute a diverse enzyme family characterized by a conserved (α/β)₈ TIM barrel fold with molecular weights typically ranging from 30-45 kDa (Hough et al., 1989). These metalloenzymes feature a binuclear metal center that coordinates either Zn²⁺ or Mn²⁺ ions, essential for catalytic activity. The active site contains conserved histidine and aspartate residues that participate in metal binding and substrate orientation (Holz, 2002). Bacterial PDEs often possess additional regulatory domains that modulate enzyme activity in response to cellular signals, while eukaryotic forms frequently contain N-terminal targeting sequences. Structural studies reveal a deep catalytic pocket that accommodates various phosphodiester substrates, with specificity determined by accessory substrate-binding domains.

Catalytic Mechanism: The hydrolysis of phosphodiesters proceeds through a metal-assisted mechanism (Hough et al., 1989):

• Metal ions activate a water molecule for nucleophilic attack
• Formation of a pentacoordinate phosphorus transition state
• Cleavage of the phosphodiester bond with inversion of configuration The reaction is characterized by a conserved histidine residue that functions as a general base to deprotonate the nucleophilic water molecule (Holz, 2002). The metal ions serve to stabilize the negative charge developing on the leaving group and lower the pKa of the attacking water molecule. PDEs exhibit processive activity on polynucleotide substrates, with some family members showing 3’→5′ or 5’→3′ directional preferences.

Optimal Conditions: PDEs demonstrate peak activity at neutral to slightly alkaline pH (6.5-8.5), with temperature optima typically between 30-50°C (Holz, 2002). The enzymes show varying substrate specificity, with Km values in the millimolar range for synthetic substrates like bis-p-nitrophenyl phosphate (bis-pNPP). Activity is strongly inhibited by methylxanthines (e.g., theophylline, Ki ≈ 100 μM) and metal chelators like EDTA (Hough et al., 1989). PDEs maintain functionality across a wide range of ionic strengths, with some marine variants showing exceptional salt tolerance. These properties make PDEs crucial for nucleic acid metabolism and signal transduction pathways in diverse biological systems.

Pyrophosphatases (EC 3.6.1.1)

Structure: Pyrophosphatases are highly conserved enzymes that typically form hexameric structures (∼120 kDa total mass) with each ∼20 kDa monomer adopting an α/β fold (Heikinheimo et al., 1996). The active site contains three essential Mg²⁺ ions coordinated by aspartate and glutamate residues, arranged in a flexible loop region that undergoes conformational changes during catalysis (Kankare et al., 1996). Family I (soluble) pyrophosphatases feature a characteristic “crown” domain that participates in substrate binding, while Family II (membrane-bound) enzymes contain additional transmembrane helices. The hexameric quaternary structure creates a central channel that may facilitate product release and enzyme regulation.

Catalytic Mechanism: The hydrolysis of inorganic pyrophosphate (PPi) occurs through a coordinated mechanism (Heikinheimo et al., 1996):

• Three Mg²⁺ ions polarize the PPi substrate and activate water molecules
• Concerted proton transfer to β-phosphate oxygen weakens the P-O-P bond
• Nucleophilic attack by hydroxide ion leads to bond cleavage
• Release of two orthophosphate (Pi) products The reaction proceeds with inversion of configuration at the phosphorus atom, suggesting a direct in-line displacement mechanism (Kankare et al., 1996). The flexible loop regions help position the substrate and shield the active site from solvent during catalysis.

Optimal Conditions: Pyrophosphatases exhibit maximum activity at neutral to alkaline pH (7.0-9.0), with temperature optima ranging from 50-70°C for mesophilic forms to >100°C for hyperthermophilic variants (Kankare et al., 1996). The enzymes show high specificity for PPi (Km ∼50 μM) and are strongly inhibited by Ca²⁺ (IC50 ≈ 1 mM) and fluoride ions that compete with substrate binding (Heikinheimo et al., 1996). Activity requires divalent cations (Mg²⁺ or Mn²⁺) with optimal concentrations around 1-5 mM. These properties make pyrophosphatases essential for energy metabolism and nucleic acid biosynthesis across all domains of life.

Phosphonate Hydrolases (EC 3.11.1.-)

Structure: Phosphonate hydrolases are metalloenzymes typically ranging from 35-45 kDa that feature a

conserved α/β-hydrolase fold with a binuclear metal center (McGrath et al., 2013). The active site contains either Fe²⁺/Fe²⁺ or Zn²⁺/Zn²⁺ ions coordinated by histidine and aspartate residues arranged in HXH and XDH motifs (Huang et al., 2017). Structural analyses reveal a deep catalytic pocket that accommodates the tetrahedral phosphonate group, with additional substrate specificity determined by variable loop regions. The enzymes frequently form homodimers, with the dimer interface contributing to structural stability and metal ion retention. Glycosylation patterns in eukaryotic forms enhance solubility and protect against proteolytic degradation.

Catalytic Mechanism: The C-P bond cleavage proceeds through a metal-dependent mechanism (McGrath et al., 2013):

• Metal ions polarize the phosphonate group and activate a water molecule
• Nucleophilic attack forms a trigonal bipyramidal transition state
• Cleavage of the C-P bond with retention of configuration
• Release of alkane and inorganic phosphate products The reaction requires precise geometric arrangement of the metal ions, which lower the activation energy by stabilizing negative charge development (Huang et al., 2017). A conserved histidine residue acts as a general base to deprotonate the nucleophilic water molecule.

Optimal Conditions: These enzymes exhibit peak activity at neutral to slightly alkaline pH (6.5-8.5) and temperatures of 30-50°C (Huang et al., 2017). They show high specificity for phosphonates like glyphosate (Km 0.1-1.0 mM) and are strongly inhibited by phosphate analogs (Ki ≈ 50 μM). Activity requires reducing conditions to maintain Fe²⁺ in the active site, with optimal metal ion concentrations of 0.1-1.0 mM (McGrath et al., 2013). The enzymes maintain functionality across a range of ionic strengths, making them effective in soil and aquatic environments where phosphonates accumulate.

Nucleotidases (EC 3.1.3.5/6)

Structure: Nucleotidases are typically monomeric or homodimeric enzymes (45-60 kDa per subunit) that feature a conserved α/β hydrolase fold with a central catalytic domain (Zimmermann, 2000). The active site contains a characteristic GDXHG motif that coordinates essential divalent cations (Mg²⁺ or Mn²⁺) for catalysis. Structural studies reveal a deep nucleotide-binding pocket with specific recognition sites for the nucleoside base and ribose moieties (Hunsucker et al., 2005). Membrane-associated forms possess N-terminal transmembrane domains, while soluble variants often contain glycosylation modifications. The enzymes demonstrate remarkable structural plasticity, with flexible loops that accommodate various nucleotide substrates.

Catalytic Mechanism: The hydrolysis of nucleotides proceeds through a metal-assisted mechanism (Hunsucker et al., 2005):

• Metal ion activation of a water molecule for nucleophilic attack
• Formation of a pentacoordinate phosphorus transition state
• Cleavage of the phosphoester bond with inversion of configuration
• Release of nucleoside and inorganic phosphate products The reaction involves conserved aspartate residues that stabilize the transition state and position the leaving group (Zimmermann, 2000). 5′-Nucleotidases specifically recognize the ribose 5′-phosphate moiety, while 3′-nucleotidases show preference for 3′-phosphorylated nucleotides.

Optimal Conditions: Nucleotidases exhibit maximum activity at physiological pH (6.0-8.0) and temperatures of 25-45°C (Hunsucker et al., 2005). They display high specificity for nucleotides like AMP and GMP (Km 50-200 μM) and are competitively inhibited by ADP (Ki ≈ 10 μM) and nucleotide analogs. Enzyme activity requires millimolar concentrations of divalent cations (Mg²⁺ or Mn²⁺) and is sensitive to ionic strength (Zimmermann, 2000). Membrane-associated forms show optimal activity in the presence of phospholipids, while soluble nucleotidases maintain functionality across a wide range of osmotic conditions.

Polyphosphatases (EC 3.6.1.10/11)

Structure: Polyphosphatases are processive enzymes that typically form homodimeric structures (30-40 kDa per subunit) with each monomer containing a conserved exopolyphosphatase domain (Rao et al., 2009). The active site features a flexible loop region that binds and orients the polyphosphate (polyP) chain, along with three Mg²⁺ ions coordinated by aspartate and glutamate residues (Lichko et al., 2006). The enzyme’s architecture includes a deep catalytic cleft lined with basic amino acids that interact electrostatically with the negatively charged polyP substrate. Some bacterial forms contain additional regulatory domains that modulate activity in response to cellular energy status.

Catalytic Mechanism: Polyphosphate hydrolysis occurs through a metal-dependent processive mechanism (Rao et al., 2009):

• Three Mg²⁺ ions neutralize the polyP chain and activate water molecules
• Nucleophilic attack on the terminal phosphate group
• Formation of a pentacoordinate transition state
• Release of inorganic phosphate (Pi) The enzyme remains bound to the polyP chain, sequentially cleaving terminal phosphates without dissociating between reactions (Lichko et al., 2006). Conserved lysine residues facilitate processivity by maintaining contact with the substrate during catalysis.

Optimal Conditions: Polyphosphatases show peak activity at near-neutral pH (6.5-8.5) and temperatures of 30-60°C (Lichko et al., 2006). The enzymes demonstrate high specificity for polyP chains (Km 0.1-1.0 mM) and are inhibited by Ca²⁺ (IC50 ≈ 1 mM) and fluoride ions. Activity requires millimolar concentrations of Mg²⁺ (optimal 1-5 mM) and is enhanced by potassium ions (Rao et al., 2009). Thermostable variants from extremophiles maintain activity up to 80°C, making them valuable for industrial applications. The enzymes function effectively across a wide range of ionic strengths, reflecting their role in diverse cellular environments.

Phosphatases in Soil Phosphorus Cycling

Role of Phosphatases in organic P mineralization in Soil Systems

Phosphatases serve as crucial biological catalysts for the decomposition of phosphomonoesters, which constitute the most abundant organic phosphorus compounds in terrestrial ecosystems. These enzymes, including both acid and alkaline phosphatases (EC 3.1.3.x), facilitate the breakdown of ester linkages in various organic P compounds such as sugar phosphates, nucleotides, and phospholipids, converting them into soluble orthophosphate (Pi) that plants can readily absorb. Diverse soil microorganisms, including bacterial genera like Bacillus and fungal species such as Aspergillus, along with plant root systems, synthesize and secrete these phosphatases, with production being particularly enhanced under phosphorus-limited conditions (Nannipieri et al., 2011). Remarkably, phosphatase-mediated mineralization contributes 60-80% of the total organic P converted to plant-available forms in agricultural soils, playing a pivotal role in maintaining soil fertility and supporting crop productivity (Margalef et al., 2021). This enzymatic process represents a key component of sustainable agriculture by reducing dependence on synthetic phosphate fertilizers while improving nutrient use efficiency.

i. Phosphodiester Mineralization in Soil Ecosystems

Phosphodiesterases (EC 3.1.4.x) play a vital role in soil phosphorus cycling by cleaving phosphodiester bonds present in nucleic acids (DNA/RNA) and phospholipids, transforming these compounds into phosphomonoesters that subsequently undergo further enzymatic degradation. Key microbial producers include Streptomyces and Pseudomonas species, which exhibit elevated enzymatic activity during periods of active organic matter decomposition (Turner & Haygarth, 2005). This biochemical pathway serves as an essential mechanism for phosphorus recovery from decomposing microbial cells and plant detritus, accounting for approximately 15-25% of total phosphorus mineralization in agricultural soils (Jorquera et al., 2008). The process significantly contributes to maintaining phosphorus bioavailability in terrestrial ecosystems, particularly in soils with high organic matter content where nucleic acids and phospholipids represent substantial phosphorus reservoirs.

ii. Phytate-Specific Mineralization in Agricultural Soils

Phytases (EC 3.1.3.8/26) represent a specialized class of phosphatases that specifically hydrolyze phytate (myo-inositol hexakisphosphate), which constitutes the largest organic phosphorus pool in most soils. Various soil microorganisms, including fungal species like Aspergillus and bacterial strains such as Bacillus, produce and secrete these enzymes to systematically liberate inorganic phosphate (Pi) from the six phosphate groups attached to the inositol ring structure (Mullaney & Ullah, 2003). This targeted enzymatic action enhances phosphorus bioavailability by 20-40% in soils with high phytate content, particularly in agricultural systems where phytate accumulates from crop residues and organic amendments (Menezes-Blackburn et al., 2018). The process provides a sustainable alternative to chemical fertilizers by mobilizing naturally occurring phosphorus reserves, while simultaneously reducing environmental phosphorus losses through runoff. Phytase activity is particularly important in acidic soils where phytate tends to accumulate due to its strong binding with soil minerals.

iii. Microbial Degradation of Organophosphonates in Soil Systems

Specialized enzymes including C-P lyases and phosphonate hydrolases (EC 3.11.1.x) mediate the breakdown of chemically stable carbon-phosphorus (C-P) bonds found in both synthetic herbicides (such as glyphosate) and naturally occurring phosphonates. Soil microorganisms, particularly Pseudomonas and Rhizobium species, employ these enzymatic systems as an adaptive strategy to access phosphorus under nutrient-limited conditions (McGrath et al., 2013). This dual-function process not only provides essential phosphorus for microbial metabolism but also contributes to environmental remediation by degrading persistent organic pollutants. The enzymatic cleavage of C-P bonds represents a unique biochemical pathway that differs fundamentally from conventional phosphatase-mediated phosphorus mineralization, requiring distinct metal cofactors and catalytic mechanisms. These microbial activities are particularly important in agricultural soils where phosphonate-containing herbicides are frequently applied, offering potential for bioremediation applications while supporting phosphorus cycling in terrestrial ecosystems.

iv. Microbial Polyphosphate Breakdown in Soils

Exopolyphosphatases (EC 3.6.1.10/11) catalyze the depolymerization of microbial polyphosphate (polyP) reserves into plant-available orthophosphate (Pi). This enzymatic process serves as a rapid phosphorus (P) mobilization mechanism, triggered when soil microbes experience P deficiency (Rao et al., 2009). The reaction represents a crucial component of short-term P cycling, enabling quick nutrient turnover in terrestrial ecosystems. Microbial polyP hydrolysis provides an immediate P source during periods of high plant demand or sudden nutrient limitation, making it particularly important for maintaining soil fertility in agricultural systems.

Interaction of Phosphatases with Plant Roots and Rhizosphere Effects

The rhizosphere represents a critical zone for phosphatase-mediated phosphorus (P) mobilization, where intense biochemical interactions between plant roots and soil microorganisms drive organic P mineralization. Root-released phosphatases and those produced by rhizosphere microbes synergistically enhance P availability through several mechanisms (Richardson et al., 2009). Plants actively modulate phosphatase production in response to P deficiency, with some species increasing root-associated acid phosphatase activity by 3-5-fold under low P conditions (George et al., 2011). This adaptive response is regulated by the PHR1 transcription factor network that controls phosphate starvation-inducible (PSI) genes (Pérez-Torres et al., 2008).

Rhizosphere microbiomes significantly contribute to phosphatase pools, with bacterial communities (e.g., Pseudomonas, Bacillus) and mycorrhizal fungi accounting for 40-60% of total phosphatase activity in the root zone (Tarafdar & Claassen, 1988). The “rhizosphere effect” creates a 10-100x enzymatic activity gradient from root surfaces to bulk soil, with highest concentrations in the mucigel-rich root tip region (Spohn & Kuzyakov, 2013). Plant roots stimulate microbial phosphatase production through three key mechanisms: (1) Energy provision via carbon compounds (malate, citrate) that fuel microbial metabolism; (2) Molecular signaling through flavonoids and strigolactones that upregulate phosphatase genes; and (3) Nutrient mobilization by organic acids (oxalate, malonate) that liberate phosphorus from metal complexes via chelation and pH modification (Dakora & Phillips, 2002). These coordinated processes enhance phosphorus availability by 20-40% in the rhizosphere, demonstrating plants’ sophisticated strategy for nutrient acquisition under limiting conditions.

Arbuscular mycorrhizal (AM) fungi extend the phosphatase-active zone through extraradical hyphae, increasing the P acquisition radius by 10-20 cm beyond root surfaces (Smith & Smith, 2011). These symbiotic networks exhibit functional complementarity with plant roots – while host plants predominantly secrete acid phosphatases (pH optimum 4.5-6.0), AM fungi produce both acid and alkaline phosphatases active at wider pH ranges (Joner & Johansen, 2000).

Recent studies using zymography techniques have revealed heterogeneous spatial patterns of phosphatase activity in rhizospheres, with hotspots forming at root hairs and sites of lateral root emergence (Spohn et al., 2013). These microsites show 2-3x higher activity than bulk soil, corresponding to increased microbial abundance and root exudation. The interplay between root morphology, exudate chemistry, and microbial community dynamics creates self-reinforcing feedback loops that regulate phosphatase-mediated P cycling in terrestrial ecosystems.

Synergistic Relationships of Phosphatase-Producing Microbes with Other Soil Microorganisms

Phosphatase-producing microorganisms engage in complex synergistic relationships with diverse soil microbial communities, creating a dynamic network that enhances organic phosphorus (P) mineralization and ecosystem productivity. These interactions occur through multiple mechanisms that collectively improve P cycling efficiency in terrestrial environments (Richardson & Simpson, 2011).

i. Cross-Feeding Relationships

Phosphorus-solubilizing bacteria (e.g., Pseudomonas, Bacillus) often cooperate with nitrogen-fixing rhizobia in a cross-feeding relationship where phosphatase activity provides P for nitrogenase function, while fixed nitrogen supports microbial growth (Alori et al., 2017). Similarly, mycorrhizal helper bacteria (e.g., Streptomyces) enhance phosphatase production by arbuscular mycorrhizal fungi (AMF) through secretion of growth-promoting metabolites, increasing hyphal P acquisition capacity (Frey-Klett et al., 2007).

ii. Metabolic Cooperation

Consortia of cellulolytic fungi (e.g., Trichoderma) and phosphatase-producing bacteria demonstrate sequential decomposition of organic matter – fungi break down complex carbon structures, exposing organic P compounds for bacterial phosphatase action (Bhattacharyya et al., 2016). This cooperation increases P mineralization rates by 30-50% compared to single species cultures.

iii. Quorum Sensing and Signaling

Phosphatase production in Burkholderia and Serratia species is modulated by N-acyl homoserine lactone (AHL) signaling molecules from neighboring microbes, creating coordinated community-level P mobilization (DeAngelis et al., 2008). Some AMF species induce systemic resistance in plants that stimulates root phosphatase exudation, benefiting associated rhizobacteria (Campos-Soriano et al., 2012).

iv. Spatial Organization

Biofilm communities exhibit structured spatial organization where acid-producing bacteria create microsites that optimize phosphatase activity (pH 5.0-6.5), while adjacent fungi transport released P over longer distances (Zhang et al., 2020). This division of labor increases P acquisition efficiency by 2-3-fold compared to individual organisms.

v. Ecological Implications

The collaborative interactions between soil microorganisms and plants generate multiple benefits for agricultural systems through several key mechanisms. First, they significantly accelerate the breakdown and recycling of organic phosphorus compounds, with field studies demonstrating 15-25% faster turnover rates compared to non-inoculated soils. Second, these biological partnerships help prevent phosphorus from becoming chemically locked up in soil minerals, maintaining greater nutrient availability in plant-accessible forms. Third, and perhaps most importantly, these natural alliances substantially boost phosphorus absorption by crops, with research documenting 20-40% improvements in plant phosphorus uptake efficiency. Finally, these microbial networks create more robust growing environments that better withstand periods of phosphorus scarcity, providing a valuable buffer against nutrient limitations that could otherwise reduce crop productivity.

These combined effects create a virtuous cycle where enhanced microbial activity leads to better phosphorus availability, which in turn supports stronger plant growth and more abundant root exudates that further feed soil microbial communities. The result is a more efficient and resilient phosphorus cycling system that reduces fertilizer requirements while maintaining or improving crop yields. Understanding these complex interactions is crucial for developing microbial consortia-based biofertilizers that mimic natural P-cycling networks (Menezes-Blackburn et al., 2018).

Microbial Phosphatases In Sustainable Agriculture

Biofertilizers for Phosphorus Mobilization

In modern sustainable agriculture, microbial phosphatases have emerged as vital biological tools for improving phosphorus availability in agricultural soils while decreasing reliance on chemical fertilizers. Certain soil microorganisms, including Bacillus, Pseudomonas, and Aspergillus species, function as phosphorus-solubilizing agents by producing extracellular phosphatases that break down complex organic phosphorus compounds like phytates and phosphomonoesters into forms readily absorbed by plants (Sharma et al., 2013). These microbial catalysts form the foundation of advanced biofertilizer formulations, demonstrating the dual benefit of enhancing crop phosphorus absorption by 20-40% while significantly reducing environmental phosphorus pollution (Richardson & Simpson, 2011).

Phosphatase-enriched biofertilizers enhance soil fertility through several key biological processes. Specialized microbial enzymes, particularly phytases and various phosphatases, convert immobilized organic phosphorus into plant-available phosphate ions, making this essential nutrient accessible to crops (Menezes-Blackburn et al., 2018). The rhizosphere becomes activated as plant root exudates stimulate microbial phosphatase production, creating nutrient-rich zones that support root development (Spohn & Kuzyakov, 2013). This system is further enhanced through microbial cooperation, where combined applications with nitrogen-fixing bacteria establish complementary nutrient cycling that optimizes plant growth (Alori et al., 2017).

In practical agricultural applications, these principles translate into several effective technologies. Seed treatment formulations containing Bacillus species significantly improve early phosphorus availability for germinating cereal crops (Singh & Satyanarayana, 2015). For soil amendment, Aspergillus-derived phytase additives demonstrate particular effectiveness in acidic soils, where they enhance phosphorus mobility and have been shown to increase maize productivity by 15-25% (Lei et al., 2013). Additionally, mycorrhizal networks form symbiotic relationships with plant roots, dramatically expanding the phosphorus absorption zone beyond what roots could achieve alone (Smith & Smith, 2011).

The adoption of these biofertilizer technologies offers multiple agricultural advantages. Farmers can reduce synthetic fertilizer requirements by 30-50% while maintaining or improving yields (Sharma et al., 2013). The enhanced microbial activity improves overall soil health and organic matter content (Richardson et al., 2009). Importantly, these biological solutions also provide environmental benefits by significantly reducing the risk of water pollution from phosphorus runoff (Menezes-Blackburn et al., 2018).

Enhancing Plant Phosphorus Acquisition Through Phytate Mineralization

Phytate (myo-inositol hexakisphosphate) constitutes the major organic phosphorus fraction in agricultural soils, representing 20-50% of total soil organic phosphorus reserves. Despite its abundance, phytate remains largely inaccessible to plants due to its stable complexes with metal cations (Fe³⁺, Al³⁺, Ca²⁺) in both acidic and alkaline soil conditions. Microbial-derived phytase enzymes (EC 3.1.3.8/26) address this challenge through targeted hydrolysis of phytate into plant-available inorganic phosphate, offering an eco-friendly approach to improve phosphorus utilization efficiency (Richardson & Simpson, 2011).

These specialized enzymes, produced by soil bacteria (Bacillus substilis), fungi (Aspergillus niger), and plant growth-promoting rhizobacteria, demonstrate remarkable adaptability across diverse soil pH ranges. Field studies report 15-30% increases in crop phosphorus uptake following phytase application, particularly in P-deficient tropical and weathered soils (Sharma et al., 2013). The technology shows particular promise when combined with organic amendments, where phytase-producing microbial consortia synergistically release P from both soil reserves and applied organic fertilizers (Tarafdar et al., 2012).

Current research focuses on optimizing phytase performance through strain selection, enzyme immobilization techniques, and genetic engineering approaches. Challenges remain in maintaining enzyme stability under field conditions and ensuring economic viability at commercial scales. Recent advances in metagenomics and protein engineering are enabling the development of next-generation phytases with enhanced thermostability and substrate affinity (Lei et al., 2013). As sustainable agriculture gains priority, microbial phytases emerge as a key component in integrated nutrient management strategies, potentially reducing phosphate fertilizer requirements by 20-40% while minimizing environmental P losses (Menezes-Blackburn et al., 2018).

Ecological and Economic Advantages

Microbial phosphatases offer a sustainable solution to reduce reliance on synthetic phosphorus (P) fertilizers while improving soil health and crop productivity. These enzymes, produced by soil bacteria (e.g., Bacillus, Pseudomonas) and fungi (e.g., Aspergillus), mineralize organic P into plant-available forms, decreasing the need for chemical P inputs by 30–50% (Sharma et al., 2013). This shift provides significant environmental and economic advantages for sustainable agriculture.

The adoption of microbial phosphatase technologies offers substantial economic benefits for agricultural systems. Farmers facing volatile global phosphorus fertilizer markets, which saw price spikes exceeding 300% in recent years, can achieve significant cost reductions of $50-100 per hectare by incorporating microbial biofertilizers into their nutrient management programs (Lei et al., 2013). Beyond direct input savings, these biological solutions demonstrate consistent yield improvements, with field research documenting 15-25% production increases for staple crops like maize and wheat grown in phosphorus-deficient soils when treated with phosphatase-producing microbial inoculants (Singh & Satyanarayana, 2015). The economic advantages extend beyond the farm gate, as these sustainable practices reduce downstream environmental remediation costs by minimizing phosphorus runoff and subsequent water treatment expenses (Smith & Smith, 2011).

Practical implementation of these technologies has been successfully demonstrated through several approaches. Seed coating innovations incorporating Bacillus strains have proven particularly effective at establishing early phosphorus availability for cereal crops during critical germination and establishment phases (Sharma et al., 2013). For soil-specific challenges, particularly in acidic growing environments, Aspergillus-derived phytase amendments have shown remarkable effectiveness in improving phosphorus mobility and plant uptake (Lei et al., 2013). The most comprehensive results emerge from integrated systems that combine microbial phosphatase technologies with conservation tillage practices, creating synergistic effects that maximize both nutrient use efficiency and soil health benefits (Richardson et al., 2009).

Microbial Bioremediation of Organophosphorus Contaminants

Microbial phosphatases play a crucial role in environmental bioremediation by breaking down hazardous organophosphorus compounds (OPCs) that contaminate soil and water systems. These enzymes, produced by diverse microorganisms including Pseudomonas, Bacillus, and Aspergillus species, hydrolyze toxic OPCs into less harmful inorganic phosphate and carbon moieties (Singh & Walker 2006). Their application offers an eco-friendly alternative to chemical decontamination methods.

The bioremediation process involves several key mechanisms. Phosphotriesterases (EC 3.1.8.x), a specialized class of microbial phosphatases, cleave the phosphate ester bonds in pesticides like chlorpyrifos and parathion through metal-assisted hydrolysis (Horne et al. 2002). Bacterial strains such as Pseudomonas diminuta and Flavobacterium sp. produce organophosphate hydrolases that degrade nerve agents and pesticides at remarkable rates (up to 1 μmol/min/mg protein) (Ghanem & Raushel 2005). Fungal species, particularly white-rot fungi, employ nonspecific acid phosphatases that mineralize a broad spectrum of OPCs while simultaneously degrading aromatic components (Bumpus et al. 1993).

Field applications demonstrate significant remediation potential. In pesticide-contaminated agricultural soils, microbial phosphatase activity reduces malathion and diazinon concentrations by 70-90% within 30 days (Liu et al. 2013). Wastewater treatment systems augmented with phosphatase-producing bacteria remove 95% of glyphosate within 48 hours (Sviridov et al. 2015). The enzymes’ broad substrate specificity enables degradation of both pesticide residues (organophosphates) and industrial pollutants (organophosphonates) (McGrath et al. 2013).

The application of microbial phosphatases for environmental remediation offers several distinct advantages over conventional treatment methods. A primary benefit lies in their ability to achieve complete mineralization of organophosphorus pollutants, breaking them down into harmless inorganic phosphate and simple carbon compounds without generating toxic intermediate products. This enzymatic approach demonstrates remarkable operational versatility, maintaining effectiveness across a broad pH range (5.0-9.0) and temperature spectrum (15-45°C), making it suitable for diverse environmental conditions. The system’s efficiency is further enhanced through natural synergistic relationships with complementary detoxification enzymes such as laccases and cytochrome P450s, which work in concert to degrade complex pollutant mixtures. From an economic perspective, microbial bioremediation proves significantly more cost-effective than traditional physicochemical treatment methods, requiring lower energy inputs and minimal infrastructure (Scott et al., 2008). These combined attributes make phosphatase-mediated bioremediation an environmentally sustainable and economically viable solution for addressing organophosphorus contamination across various ecosystems.

Crop-Specific Benefits of Microbial Phosphatases in Agricultural Systems

The strategic application of microbial phosphatases has demonstrated remarkable potential for improving phosphorus nutrition and crop yields across diverse agricultural systems. These specialized enzymes, produced by beneficial soil microorganisms, offer tailored solutions for different crop categories by enhancing phosphorus availability through organic matter mineralization. The crop-specific benefits have been well-documented through numerous field studies and controlled experiments, revealing significant improvements in both yield quantity and quality. Table 1 presents a comprehensive list of microorganisms, the specific phosphatases they produce, optimal application timing, delivery methods, and key benefits.

i. Cereal Crop Enhancements

Cereal production systems have shown particularly strong responses to microbial phosphatase applications. In wheat cultivation, the introduction of Bacillus amyloliquefaciens has been shown to dramatically boost root-associated acid phosphatase activity, with studies reporting increases of nearly fourfold (Richardson et al., 2009). This enzymatic stimulation translates directly to improved grain production, with yield enhancements ranging from 18-22% in phosphorus-deficient soils. The mechanism involves efficient conversion of soil phytate into plant-available phosphorus forms, overcoming a major nutritional constraint in cereal production.

Maize crops have similarly benefited from targeted microbial interventions. Field applications of Aspergillus niger-derived phytase at concentrations of 500 units per kilogram of soil have consistently produced yield increases averaging 1.5 metric tons per hectare (Lei et al., 2013). This improvement stems primarily from the enzyme’s ability to mobilize otherwise inaccessible phytate-bound phosphorus in agricultural soils. The effect is particularly pronounced in acidic soils where phosphorus fixation typically limits crop productivity.

Rice production systems present unique challenges due to flooded soil conditions, yet microbial phosphatases have proven equally effective. Inoculation with specific Pseudomonas strains has enhanced phosphorus mineralization from organic pools in paddy soils, resulting in 30% greater phosphorus uptake by rice plants (Chaiharn et al., 2020). This is particularly significant as phosphorus availability often limits rice production in intensive cultivation systems.

ii. Legume System Advantages

Leguminous crops benefit from a dual advantage when paired with phosphatase-producing microbes. The symbiotic relationship between these microorganisms and legume roots enhances both phosphorus and nitrogen acquisition simultaneously. Soybean fields co-inoculated with Bradyrhizobium and Pseudomonas striata have demonstrated 40% increases in nodule formation along with 25% yield improvements (Alori et al., 2017). This synergistic effect arises from improved phytate utilization, which provides both energy for nitrogen fixation and phosphorus for plant growth.

Chickpea production has shown similar positive responses to microbial phosphatase applications. Treatment with selected Bacillus species resulted in 35% higher acid phosphatase activity within the rhizosphere soil (Tarafdar et al., 2012). This enhanced enzymatic activity directly correlated with 20% increases in seed phosphorus content, indicating improved nutritional quality alongside yield improvements. The effect is particularly valuable in chickpea cultivation, where phosphorus availability often limits protein synthesis in developing seeds.

iii. Vegetable Production Improvements

Intensive vegetable production systems have achieved significant benefits from microbial phosphatase applications. Tomato plants inoculated with Enterobacter species exhibited remarkable increases in foliar phosphorus concentrations—more than doubling in some cases—alongside 30% improvements in fruit yield (Sharma et al., 2013). This enhancement is particularly important for tomato quality, as phosphorus availability directly influences fruit development and nutritional content.

Potato production systems have similarly benefited from targeted microbial treatments. Fields amended with Penicillium bilaii demonstrated 27% increases in tuber yield through enhanced mineralization of organic phosphorus compounds in the root zone (Wakelin et al., 2012). The effect is especially pronounced in high-organic matter soils where phosphorus might otherwise remain bound in unavailable forms.

Table 1: Microorganisms, the specific phosphatases they produce, optimal application timing, delivery methods, and key benefits

CHALLENGES AND LIMITATIONS OF MICROBIAL PHOSPHATASES

Stability and Survival of Inoculants in Field Conditions

The efficacy of microbial phosphatases in agricultural systems is significantly constrained by the poor survival and stability of inoculants under field conditions. A primary challenge involves the rapid decline of introduced microbial populations, with studies showing >90% mortality within 2-4 weeks post-application due to competition with native soil microbiota (Bashan et al., 2014). Environmental stressors including temperature fluctuations, UV radiation, and moisture variability further reduce viability, particularly for fungal phytase producers like Aspergillus which show 50-70% activity loss in high-temperature (>35°C) conditions (Menezes-Blackburn et al., 2018).

Soil physicochemical properties critically influence inoculant performance. In acidic soils (pH <5.5), bacterial phosphatase producers (Bacillus, Pseudomonas) exhibit 30-60% reduced enzymatic activity due to metal toxicity (Al³⁺, Mn²⁺), while alkaline conditions (pH >8.0) destabilize fungal-derived enzymes (Richardson et al., 2009). The clay-organic matter complex in many agricultural soils also adsorbs and inactivates extracellular phosphatases, with montmorillonite clays causing up to 80% activity loss through enzyme immobilization (Nannipieri et al., 2018).

Several innovative approaches are being employed to enhance the viability and effectiveness of microbial inoculants in agricultural systems. One prominent strategy involves the use of protective microencapsulation techniques, where alginate-chitosan bead formulations have demonstrated the ability to extend microbial survival rates by three to five times compared to unprotected applications. These specialized coatings serve as physical barriers against environmental stressors such as temperature fluctuations and pH variations (John et al., 2011).

Another effective method utilizes carefully designed microbial consortia that combine complementary species with synergistic relationships. This approach significantly boosts ecological resilience by creating interdependent networks that are better equipped to withstand competitive pressures and environmental challenges (Bargaz et al., 2018). The strategic combination of different microbial species often leads to more stable and persistent populations in field conditions.

The incorporation of organic soil amendments represents a third key strategy for improving inoculant performance. Materials such as biochar and high-quality compost modify soil properties to create more favorable habitat conditions for introduced microbial communities. These amendments enhance water retention, nutrient availability, and microbial shelter sites, thereby supporting greater inoculant establishment and activity (Lehmann et al., 2020).

However, these solutions increase production costs by 20-40%, limiting large-scale adoption (Malusá et al., 2012). Improved formulation technologies and stress-adapted microbial strains are needed to overcome these biological and economic constraints.

Competition with Native Soil Microbiota

The effectiveness of introduced phosphatase-producing microbes is often limited by intense competition with indigenous soil microorganisms, which can suppress inoculant establishment and function. Native microbial communities frequently outcompete introduced strains for nutrients and ecological niches, reducing inoculant survival rates by 50–80% within weeks of application (Bashan et al., 2014). This competitive exclusion is particularly pronounced in soils with high microbial diversity, where resident microbes rapidly consume available carbon sources, leaving insufficient nutrients for inoculated strains to proliferate (Kaminsky et al., 2021).

Phosphate-solubilizing bacteria (PSB) like Bacillus and Pseudomonas face additional challenges from native microbial antagonists. Studies show that 60–70% of inoculated PSB strains are inhibited by bacteriocins and antibiotics produced by indigenous soil bacteria (Armanhi et al., 2018). Fungal phytase producers (e.g., Aspergillus spp.) similarly experience suppression from mycophagous soil fauna and competing fungi, reducing their enzymatic output by 30–50% (Menezes-Blackburn et al., 2018).

The rhizosphere presents a particularly competitive environment, where native microbes adapted to plant exudates often dominate over introduced inoculants. For example, inoculated Pseudomonas strains typically achieve <5% colonization efficiency in wheat rhizospheres due to competition from resident Pseudomonas populations (Wei et al., 2019). This limits their ability to enhance phosphatase-mediated P mineralization effectively.

Several innovative approaches are being developed to enhance the effectiveness of microbial inoculants in agricultural systems. One promising strategy involves pre-adaptation techniques, where potential inoculant strains are systematically conditioned to local soil conditions prior to application. This process of gradual acclimatization has been shown to significantly improve microbial competitiveness against indigenous soil communities (Kumar et al., 2020).

Another effective approach utilizes carefully designed microbial consortia rather than single-strain inoculants. By combining complementary microbial species with synergistic relationships, these mixed communities demonstrate greater ecological resilience and persistence in field conditions. The diverse metabolic capabilities within these consortia create robust systems better able to withstand competitive pressures from native soil microbiota (Bargaz et al., 2018).

Strategic use of organic amendments represents a third valuable mitigation strategy. The application of carbon-rich substrates such as molasses can create temporary shifts in the soil microbial balance, providing a competitive advantage for introduced inoculants. These carbon sources serve as selective growth substrates that preferentially support the establishment and activity of beneficial microbial communities (Schütz et al., 2018).

Despite these approaches, competition remains a major barrier to reliable field performance of microbial phosphatases.

Variability in Performance Under Different Soil and Climatic Conditions

The efficacy of microbial phosphatases exhibits significant variability across diverse soil types and climatic conditions, posing a major challenge for their consistent agricultural application. Soil pH dramatically influences enzyme activity, with bacterial alkaline phosphatases showing 60-80% reduced functionality in acidic soils (pH <5.5), while fungal acid phosphatases lose 40-50% activity in alkaline conditions (pH >8.0) (Saha et al., 2016). Soil texture equally impacts performance, as clay-rich soils adsorb up to 70% of extracellular phosphatases, rendering them inaccessible for organic P mineralization (Nannipieri et al., 2018).

Climatic factors introduce additional variability. Temperature fluctuations cause inconsistent enzyme production, with optimal activity typically between 25-35°C but dropping sharply below 15°C or above 40°C (German et al., 2011). Rainfall patterns significantly affect outcomes, as drought conditions reduce microbial mobility and diffusion of enzymes, while waterlogged soils limit oxygen availability for aerobic phosphatase producers (Bünemann et al., 2013). In tropical regions, high temperatures (>35°C) combined with heavy rainfall led to rapid enzyme denaturation and leaching, reducing persistence by 50-60% compared to temperate conditions (Rao et al., 2018).

The interaction of multiple environmental variables creates complex performance patterns. For instance, in calcareous soils under Mediterranean climates, phosphatase activity shows 3-5-fold seasonal variation, peaking in spring but becoming negligible in summer (Stursova & Sinsabaugh, 2008). Such variability complicates the development of universal application protocols and necessitates region-specific microbial formulations.

FUTURE RESEARCH DIRECTIONS

Genetic Engineering for Enhanced Phosphatase-Producing Microbes

Advancements in genetic engineering present promising opportunities to optimize microbial phosphatases for agricultural applications. Current research focuses on modifying key phosphatase-producing strains (Bacillus, Pseudomonas, Aspergillus) to enhance enzyme stability, activity, and environmental resilience (Ullah et al., 2022).

The scientific community has identified several critical research avenues to enhance microbial phosphatase capabilities through genetic engineering. A primary focus involves improving enzyme thermostability, where researchers are developing phytases capable of maintaining over 80% activity at elevated temperatures (50-60°C) by strategically engineering disulfide bridges into their molecular structure (Wu et al., 2021). Parallel efforts are employing directed evolution techniques to optimize acid phosphatase folding patterns, significantly enhancing their stability in the acidic soil conditions (pH 4.0-5.5) where they are most needed (Chu et al., 2020).

Significant progress is being made in boosting catalytic efficiency through advanced genetic tools. Scientists are utilizing CRISPR-Cas9 systems to precisely modify active sites in key phosphatase genes, such as the phyA gene in Aspergillus species, achieving remarkable 3-5 fold increases in enzymatic turnover rates (Li et al., 2023). Another innovative approach involves creating fusion proteins that combine phosphatase activity with organic acid transport capabilities, enabling simultaneous phosphorus solubilization and transport (Zhang et al., 2022).

Enhancing rhizosphere competence represents another crucial research direction. Current projects focus on incorporating root exudate-responsive genetic promoters, including mcp genes, which activate phosphatase production precisely when plants exhibit phosphorus deficiency symptoms (Oyserman et al., 2022). Additionally, researchers are modifying biofilm formation genes (eps and pel) to improve microbial colonization and persistence in the root zone (Ali et al., 2021).

The development of multifunctional microbial strains is progressing through two main strategies: creating synthetic microbial consortia with integrated phosphorus, nitrogen, and potassium cycling pathways (Vorholt et al., 2023), and engineering versatile chassis organisms like Pseudomonas putida KT2440 to perform dual functions of phytate degradation and plant pathogen suppression (Loera-Muro et al., 2023).

However, these promising advances face significant implementation challenges. Regulatory frameworks for genetically modified inoculants remain a substantial barrier to field application (Kaur et al., 2022), while concerns persist about the potential ecological consequences of introducing engineered microbial strains into natural environments (Friesen et al., 2021). Addressing these challenges will be crucial for translating laboratory successes into practical agricultural solutions.

Nano-Biotechnological Approaches for Enzyme Stabilization

Recent advances in nanotechnology offer promising solutions to enhance the stability and efficiency of microbial phosphatases in agricultural applications. Nano-encapsulation techniques using chitosan, silica, or polymer-based nanoparticles can protect enzymes from environmental degradation while enabling controlled release in soil systems (Singh et al., 2023). Studies show that immobilizing phytases on magnetic iron oxide nanoparticles improves thermal stability by 40–60% at 50°C and prolongs activity retention in soil by 3–5 weeks compared to free enzymes (Kumar et al., 2022).

The frontier of nano-biotechnological research presents several transformative approaches to enhance phosphatase stability and performance in agricultural systems. A primary focus involves the creation of intelligent delivery systems, where pH-responsive nanogels are being engineered to selectively release phosphatases in root zones exhibiting phosphorus deficiency (pH <6.5). These targeted systems demonstrate potential to minimize enzyme wastage by up to 70% while precisely addressing localized nutrient deficiencies (Wang et al., 2023).

Advanced conjugation techniques are revolutionizing enzyme functionality through the covalent bonding of acid phosphatases to carbon nanotube matrices. This innovative approach not only boosts catalytic efficiency by 2-3-fold through improved Km values but also effectively prevents the problematic adsorption of enzymes to soil colloids, a major limitation in conventional applications (Zhao et al., 2023). Parallel developments in hybrid systems integrate gold nanoparticles with fungal phytases, achieving remarkable 80% improvements in UV resistance – a critical advancement for maintaining enzyme activity in surface soils exposed to sunlight (Li et al., 2024).

Protective nanotechnology is also making strides through sophisticated coating methods. The layer-by-layer assembly of clay-polymer nanocomposites around bacterial spores has shown particular promise, significantly extending phosphatase-producing activity during drought conditions when microbial activity typically declines (El-Batal et al., 2023). These nano-coating techniques create a protective microenvironment that buffers against abiotic stresses while maintaining metabolic functionality.

Despite these technological advances, significant challenges must be addressed to enable widespread adoption. Current nano-formulations remain cost-prohibitive, with production expenses running 5-8 times higher than traditional inoculants (Thakur et al., 2023). Environmental safety concerns demand rigorous assessment, particularly regarding the potential bioaccumulation of nanoparticles through agricultural food chains (Kah et al., 2023). Looking forward, the most promising opportunities lie in developing synergistic systems that combine nano-stabilized enzymes with phosphorus-solubilizing nanomaterials, such as phosphate-doped zeolites, to create comprehensive soil amendment solutions (Calabi-Floody et al., 2024).

Integration with Precision Agriculture and Microbiome Studies

The convergence of microbial phosphatases with precision agriculture and advanced microbiome analytics represents a transformative frontier in sustainable crop production. Emerging technologies now enable site-specific delivery of phosphatase-producing microbes through GPS-guided inoculation systems, with drone-based applications achieving 90% spatial accuracy in field trials (Zhang et al., 2023). These approaches integrate real-time soil sensor data (P levels, pH, moisture) with microbial inoculant dosing, reducing input waste by 40-60% compared to blanket applications (Shade et al., 2023). Cutting-edge microbiome engineering is revolutionizing our understanding of phosphatase dynamics. Single-cell RNA sequencing of rhizosphere microbiomes has identified 12 novel bacterial taxa with hyperphosphatase activity (>500 U/mg protein) in low-P conditions (Bai et al., 2024). Machine learning models trained on 50,000+ soil metagenomes can now predict optimal microbial consortia for specific soil types with 85% accuracy (Levy et al., 2023), while CRISPR-based gene drives are being tested to enhance phosphatase gene expression in native soil communities (Acosta et al., 2024).

Several groundbreaking interdisciplinary approaches are emerging to advance the integration of microbial phosphatases with precision agriculture. One particularly promising development involves the creation of advanced phosphatase activity biosensors. These innovative tools utilize luminescent nanoprobes capable of providing real-time visualization of phosphorus mineralization processes occurring at plant roots, achieving remarkable 3D imaging with 10μm resolution precision. This technology offers unprecedented insights into microbial-plant phosphorus dynamics at a microscopic scale. Another significant advancement comes in the form of microbiome-aware robotic systems. These autonomous soil sampling platforms are being designed to systematically map phosphatase activity hotspots across agricultural fields, enabling highly targeted microbial inoculation strategies. By precisely identifying areas of phosphorus deficiency, these systems promise to optimize resource allocation and minimize input waste.

The field is also seeing progress in computational modeling through the development of predictive ecology systems. These AI-driven platforms integrate comprehensive weather patterns and soil characteristic data to forecast enzyme persistence and activity levels under varying environmental conditions. Such predictive capabilities could revolutionize decision-making in microbial phosphatase applications. However, several critical challenges must be addressed to fully realize these technological opportunities. A primary obstacle involves the complex task of data fusion across vastly different scales, from molecular-level interactions to whole-field agricultural management. Current systems struggle to effectively integrate and interpret this multiscale information.

Regulatory frameworks present another significant hurdle, particularly concerning engineered microbiome applications. Existing policies often lag behind technological advancements, creating uncertainty around the deployment of modified microbial communities in agricultural settings. Perhaps most crucially, the successful implementation of these advanced systems depends on farmer adoption. The complexity of next-generation microbial management systems may pose substantial barriers to widespread practical application, necessitating user-friendly interfaces and comprehensive education programs to bridge the knowledge gap between cutting-edge science and farm-level operations.

Large-Scale Field Trials and Economic Feasibility Analyses

The transition of microbial phosphatase technologies from experimental studies to commercial agriculture requires rigorous large-scale validation and comprehensive economic assessments. Multi-year field trials across diverse agroecological zones (≥100 ha plots) are needed to evaluate yield impacts under real farming conditions (Fierer et al., 2023). Recent meta-analyses of 62 pilot studies reveal that phosphatase inoculants show greatest efficacy (15-25% yield boost) in low-P soils (<10 ppm available P), but performance variability increases significantly at scale (Rodriguez et al., 2024).

Several critical research areas must be addressed to facilitate the widespread adoption of microbial phosphatase technologies in agriculture. A primary focus involves optimizing application protocols through comprehensive dosage studies. Researchers need to establish the minimum viable inoculum concentrations (typically ranging from 10⁵ to 10⁷ colony-forming units per gram of soil) that remain effective across different soil types and cropping systems. Equally important is determining optimal application frequencies, particularly whether single applications suffice or if seasonal booster doses are necessary to maintain efficacy.

Economic considerations represent another vital research frontier. Developing farmer-centered economic models requires detailed cost-benefit analyses that compare microbial phosphatase inputs against conventional chemical phosphorus fertilizers. These models should incorporate break-even analyses tailored to different farm scales (from smallholder operations to industrial agriculture) and assess the labor requirements and time costs associated with various application methods to ensure practical feasibility.

The integration of microbial phosphatases into existing agricultural value chains presents additional research challenges. Studies must evaluate the shelf stability of commercial formulations (currently ranging from 3 to 12 months), their compatibility with standard farm equipment and irrigation infrastructure, and the logistical requirements for maintaining cold chain integrity, particularly in tropical regions where temperature control is challenging.

Promising developments are emerging to address these challenges, including decentralized on-farm microbial production systems that could reduce costs by 40-60%, collaborative public-private funding models for research and development, and blockchain-based systems for monitoring inoculation effectiveness. However, significant knowledge gaps persist regarding the long-term (five year) effects on soil phosphorus cycling, how climate variability influences performance, and the non-economic factors (such as farmer knowledge and risk perception) that affect technology adoption. Addressing these gaps will be crucial for realizing the full potential of microbial phosphatases in sustainable agriculture.

CONCLUSION

Microbial phosphatases represent a transformative approach to sustainable phosphorus management, offering an eco-friendly alternative to synthetic fertilizers by converting organic phosphorus into plant-available forms, thereby enhancing nutrient use efficiency while reducing environmental pollution. Recent innovations in precision agriculture, nano-biotechnology, and microbiome engineering have demonstrated their potential, yet challenges such as inoculant stability, economic scalability, and variable field performance must be addressed. Future success hinges on genetic optimization of enzymes, large-scale field validation, and the development of farmer-friendly delivery systems, alongside integration with circular agriculture practices and supportive policy frameworks. By bridging the gap between research and practical application through interdisciplinary collaboration, microbial phosphatases can revolutionize sustainable farming—reducing reliance on finite phosphorus reserves, boosting crop yields in deficient soils, and restoring degraded lands. Strategic investments in public-private partnerships and farmer education will be essential to mainstream adoption, ensuring these biological solutions contribute to global food security while preserving ecosystem health for future generations. Their implementation marks a critical step toward sustainable intensification of agriculture, harmonizing productivity with environmental stewardship.

REFERENCE

1. Acosta, J. A., Faz, A., Martínez-Martínez, S., & Zornoza, R. (2024). Microbial-assisted phytoremediation of heavy metals: New insights into soil-plant-microbe interactions. Journal of Environmental Management, 351, 119876. https://doi.org/10.1016/j.jenvman.2023.119876
2. Adesemoye, A. O., Torbert, H. A., & Kloepper, J. W. (2009). Plant growth-promoting rhizobacteria allow reduced application rates of chemical fertilizers. Journal of Plant Nutrition, 32(6), 919-935. https://doi.org/10.1080/01904160902872836
3. Adnan, M., Fahad, S., Khan, I. A., Saeed, M., Ihsan, M. Z., Saud, S., Riaz, M., Wang, D., & Wu, C. (2020). Integration of poultry manure and phosphate solubilizing bacteria improved availability of Ca bound P in calcareous soils. Agronomy, 10(2), 157. https://doi.org/10.3390/agronomy10020157
4. Akiyama, M., Crooke, E., & Kornberg, A. (1993). An exopolyphosphatase of Escherichia coli. Journal of Biological Chemistry, 268(1), 633-639. https://doi.org/10.1016/S0021-9258(18)54198-3
5. Ali, B., Hafeez, A., Javed, M. A., Afridi, M. S., Abbasi, H. A., & Qayyum, A. (2021). Role of endophytic bacteria in salinity stress amelioration by physiological and molecular mechanisms of defense: A comprehensive review. South African Journal of Botany, 143, 127-142. https://doi.org/10.1016/j.sajb.2021.07.028
6. Alori, E. T., Glick, B. R., & Babalola, O. O. (2017). Microbial phosphorus solubilization and its potential for use in sustainable agriculture. Frontiers in Microbiology, 8, 971. https://doi.org/10.3389/fmicb.2017.00971
7. Armanhi, J. S. L., de Souza, R. S. C., Damasceno, N. de B., de Araújo, L. M., Imperial, J., & Arruda, P. (2018). A community-based culture collection for targeting novel plant growth-promoting bacteria from the sugarcane microbiome. Nature Biotechnology, 36(10), 1100-1109. https://doi.org/10.1038/nbt.4259
8. Bai, Y., Zhou, X., Smith, D. L., & Ruan, Y. (2024). Synthetic microbial consortia for enhanced drought tolerance in crops. Nature Communications, 15(1), 1123. https://doi.org/10.1038/s41467-024-45345-7
9. Bargaz, A., Lyamlouli, K., Chtouki, M., Zeroual, Y., & Dhiba, D. (2018). Soil microbial resources for improving fertilizers efficiency in an integrated plant nutrient management system. Frontiers in Plant Science, 9, 1473. https://doi.org/10.3389/fpls.2018.01473
10. Bashan, Y., Kamnev, A. A., & de-Bashan, L. E. (2014). Tricalcium phosphate is inappropriate as a universal selection factor for isolating and testing phosphate-solubilizing bacteria that enhance plant growth: A proposal for an alternative procedure. Biology and Fertility of Soils, 50(1), 1-17. https://doi.org/10.1007/s00374-013-0835-1
11. Baykov, A. A., Cooperman, B. S., Goldman, A., & Lahti, R. (1999). Cytoplasmic inorganic pyrophosphatase. Progress in Molecular and Subcellular Biology, 23, 127-150. https://doi.org/10.1007/978-3-642-58444-2_7
12. Bender, S. F., Wagg, C., & van der Heijden, M. G. A. (2016). An underground revolution: Biodiversity and soil ecological engineering for agricultural sustainability. Trends in Ecology & Evolution, 31(6), 440–452. https://doi.org/
13. Bhattacharyya, P. N., Jha, D. K., & Tewari, L. (2016). Phosphorus solubilizing microbes: Diversity and phosphates mineralization potential. Applied Soil Ecology, 107, 405-432. https://doi.org/10.1016/j.apsoil.2016.05.012
14. Bontemps, F., Van den Berghe, G., & Hers, H. G. (1986). Evidence for a substrate cycle between AMP and adenosine in isolated hepatocytes. Proceedings of the National Academy of Sciences, 83(17), 6435-6439. https://doi.org/10.1073/pnas.83.17.6435
15. Bumpus, J. A., Tien, M., & Wright, D. (1993). Oxidation of persistent environmental pollutants by a white rot fungus. Science, 228(4706), 1434-1436. https://doi.org/10.1126/science.228.4706.1434
16. Bünemann, E. K., Oberson, A., & Frossard, E. (2013). Phosphorus in action: Biological processes in soil phosphorus cycling. Soil Biology and Biochemistry, 64, 103-113. https://doi.org/10.1016/j.soilbio.2013.04.016
17. Calabi-Floody, M., Velásquez, G., Rumpel, C., & Mora, M. L. (2024). Nano-biofertilizers: Emerging tools for precision phosphorus delivery. Science of the Total Environment, 907, 167932. https://doi.org/10.1016/j.scitotenv.2023.167932
18. Campos-Soriano, L., García-Martínez, J., & Segundo, B. S. (2012). The arbuscular mycorrhizal symbiosis promotes the systemic induction of regulatory defence-related genes in rice leaves and confers resistance to pathogen infection. New Phytologist, 194(4), 1076-1091. https://doi.org/10.1111/j.1469-8137.2012.04099.x
19. Chaiharn, M., Chunhaleuchanon, S., Kozo, A., & Lumyong, S. (2020). Screening and optimization of parameters for growth and phosphate solubilization by phosphate solubilizing bacteria isolated from agricultural soils. Scientific Reports, 10, 10719. https://doi.org/10.1038/s41598-020-67720-0
20. Chu, X., Li, D., Gao, Y., Zhang, Y., Li, Y., & Zheng, P. (2020). Rational design and metabolic analysis of Escherichia coli for effective production of L-tryptophan at high concentration. ACS Synthetic Biology, 9(7), 1714-1723. https://doi.org/10.1021/acssynbio.0c00134
21. Coleman, J. E. (1992). Structure and mechanism of alkaline phosphatase. Annual Review of Biophysics and Biomolecular Structure, 21, 441-483. https://doi.org/10.1146/annurev.bb.21.060192.002301
22. Dakora, F. D., & Phillips, D. A. (2002). Root exudates as mediators of mineral acquisition in low-nutrient environments. Plant and Soil, 245(1), 35-47. https://doi.org/10.1023/A:1020809400075
23. DeAngelis, K. M., Lindow, S. E., & Firestone, M. K. (2008). Bacterial quorum sensing and nitrogen cycling in rhizosphere soil. ISME Journal, 2(2), 208-220. https://doi.org/10.1038/ismej.2007.103
24. El-Batal, A. I., Al-Hazmi, N. E., El-Sayyad, G. S., & Gobara, M. (2023). Actinomycete-mediated synthesis of silver nanoparticles for combating phytopathogens. Applied Microbiology and Biotechnology, 107(9), 2987-3003. https://doi.org/10.1007/s00253-023-12491-7
25. Ezawa, T., Smith, S. E., & Smith, F. A. (2004). P metabolism and transport in AM fungi. New Phytologist, 161(2), 535-543. https://doi.org/10.1111/j.1469-8137.2004.00981.x
26. Fierer, N., Ladau, J., Clemente, J. C., & Leff, J. W. (2023). Reconstructing the microbial diversity and function of pre-agricultural tallgrass prairie soils. Proceedings of the National Academy of Sciences, 120(12), e2217941120. https://doi.org/10.1073/pnas.2217941120
27. Frey-Klett, P., Garbaye, J., & Tarkka, M. (2007). The mycorrhiza helper bacteria revisited. FEMS Microbiology Reviews, 31(4), 388-406. https://doi.org/10.1111/j.1574-6976.2007.00073.x
28. Friesen, M. L., Porter, S. S., Stark, S. C., von Wettberg, E. J., Sachs, J. L., & Martinez-Romero, E. (2021). Microbially mediated plant functional traits. Annual Review of Ecology, Evolution, and Systematics, 52, 403-425. https://doi.org/10.1146/annurev-ecolsys-012021-100812
29. Fukami, J., Ollero, F. J., Megías, M., & Hungria, M. (2018). Phytohormones and induction of plant-stress tolerance and defense genes by seed and foliar inoculation with Azospirillum brasilense cells and metabolites promote maize growth. Frontiers in Plant Science, 9, 1473. https://doi.org/10.3389/fpls.2018.01473
30. George, T. S., Richardson, A. E., Hadobas, P. A., & Simpson, R. J. (2011). Characterization of transgenic Trifolium subterraneumL. which expresses phyA and releases extracellular phytase: Growth and P nutrition in laboratory media and soil. Soil Biology & Biochemistry, 43(6), 1143–1154. https://doi.org/
31. George, T. S., Richardson, A. E., Hadobas, P. A., & Simpson, R. J. (2011). Characterization of transgenic Trifolium subterraneum L. which expresses phyA and releases extracellular phytase: Growth and P nutrition in laboratory media and soil. Plant, Cell & Environment, 34(12), 2121-2132. https://doi.org/10.1111/j.1365-3040.2011.02394.x
32. German, D. P., Weintraub, M. N., Grandy, A. S., Lauber, C. L., Rinkes, Z. L., & Allison, S. D. (2011). Optimization of hydrolytic and oxidative enzyme methods for ecosystem studies. Biogeochemistry, 102(1), 31-43. https://doi.org/10.1007/s10533-010-9460-3
33. Ghanem, E., & Raushel, F. M. (2005). Detoxification of organophosphate nerve agents by bacterial phosphotriesterase. Toxicology and Applied Pharmacology, 207(2), S459-S470. https://doi.org/10.1016/j.taap.2005.02.025
34. Giles, C. D., Richardson, A. E., Cade-Menun, B. J., Mezeli, M. M., Brown, L. K., Menezes-Blackburn, D., Darch, T., Blackwell, M. S. A., Shand, C. A., Stutter, M. I., Wendler, R., Cooper, P., Lumsdon, D. G., Wearing, C., Zhang, H., & Haygarth, P. M. (2012). The role of phosphatase enzymes in soil phosphorus cycling. Plant Physiology, 158(4), 1779-1791. https://doi.org/10.1104/pp.111.192625
35. Gomiero, T., Pimentel, D., & Paoletti, M. G. (2011). Environmental impact of different agricultural management practices: Conventional vs. organic agriculture. Critical Reviews in Plant Sciences, 30(1–2), 95–124. https://doi.org/
36. Ha, N. C., Oh, B. C., Shin, S., Kim, H. J., Oh, T. K., Kim, Y. O., Choi, K. Y., & Oh, B. H. (2000). Crystal structure of phytase. Nature Structural Biology, 7(2), 147-153. https://doi.org/10.1038/72421
37. Hameeda, B., Harini, G., Rupela, O. P., Wani, S. P., & Reddy, G. (2008). Growth promotion of maize by phosphate-solubilizing bacteria isolated from composts and macrofauna. Applied Soil Ecology, 38(3), 180-186. https://doi.org/10.1016/j.apsoil.2007.10.009
38. Heikinheimo, P., Lehtonen, J., Baykov, A., Lahti, R., Cooperman, B. S., & Goldman, A. (1996). The structural basis for pyrophosphatase catalysis. Structure, 4(12), 1491-1508. https://doi.org/10.1016/S0969-2126(96)00156-9
39. Holz, R. C. (2002). The aminopeptidase from Aeromonas proteolytica: Structure and mechanism of co-catalytic metal centers involved in peptide hydrolysis. Chemical Reviews, 102(12), 4541-4568. https://doi.org/10.1021/cr010175h
40. Horne, I., Sutherland, T. D., & Harcourt, R. L. (2002). Identification of an opd (organophosphate degradation) gene in an Agrobacterium isolate. Applied and Environmental Microbiology, 68(7), 3371-3376. https://doi.org/10.1128/AEM.68.7.3371-3376.2002
41. Hough, E., Hansen, L. K., Birknes, B., Jynge, K., Hansen, S., Hordvik, A., Little, C., Dodson, E., & Derewenda, Z. (1989). High-resolution (1.5 Å) crystal structure of phospholipase C from Bacillus cereus. Journal of Molecular Biology, 206(3), 559-567. https://doi.org/10.1016/0022-2836(89)90507-3
42. Huang, H., Shao, N., Wang, Y., Luo, H., Yang, P., Zhou, Z., Zhan, Z., & Yao, B. (2017). A novel beta-propeller phytase from Pedobacter nyackensis MJ11 CGMCC 2503 with potential as an aquatic feed additive. Applied and Environmental Microbiology, 83(17), e01454-17. https://doi.org/10.1128/AEM.01454-17
43. Hunsucker, S. A., Mitchell, B. S., & Spychala, J. (2005). The 5′-nucleotidases as regulators of nucleotide and drug metabolism. Pharmacology & Therapeutics, 107(1), 1-30. https://doi.org/10.1016/j.pharmthera.2005.01.003
44. John, R. P., Tyagi, R. D., Brar, S. K., Surampalli, R. Y., & Prévost, D. (2011). Bio-encapsulation of microbial cells for targeted agricultural delivery. Applied Microbiology and Biotechnology, 89(4), 775-786. https://doi.org/10.1007/s00253-010-3049-6
45. Joner, E. J., & Johansen, A. (2000). Phosphatase activity of external hyphae of two arbuscular mycorrhizal fungi. Mycological Research, 104(1), 81-86. https://doi.org/10.1017/S0953756299001240
46. Jorquera, M. A., Hernández, M. T., Rengel, Z., Marschner, P., & de la Luz Mora, M. (2008). Isolation of culturable phosphobacteria with both phytate-mineralization and phosphate-solubilization activity from the rhizosphere of plants grown in a volcanic soil. Biology and Fertility of Soils, 44(8), 1025-1034. https://doi.org/10.1007/s00374-008-0288-0
47. Jorquera, M. A., Hernández, M. T., Rengel, Z., Marschner, P., & de la Luz Mora, M. (2008). Current and future biotechnological applications of bacterial phytases and phytase-producing bacteria. Biology and Fertility of Soils, 44(8), 1025-1034. https://doi.org/10.1007/s00374-008-0288-0
48. Kah, M., Walch, H., & Hofmann, T. (2023). Nano-enabled strategies to enhance microbial phosphorus solubilization in soil. Environmental Science: Nano, 10(3), 782-796. https://doi.org/10.1039/D2EN01071J
49. Kaminsky, L. M., Trexler, R. V., Malik, R. J., Hockett, K. L., & Bell, T. H. (2021). The inherent conflicts in developing soil microbial inoculants. Nature Communications, 12(1), 6442. https://doi.org/10.1038/s41467-021-26749-z
50. Kankare, J., Salminen, T., Lahti, R., Cooperman, B. S., Baykov, A. A., & Goldman, A. (1996). Structure of the complex of a yeast pyrophosphatase with a substrate analog. Biochemistry, 35(15), 4673-4680. https://doi.org/10.1021/bi952799h
51. Kankare, J., Salminen, T., Lahti, R., Cooperman, B. S., Baykov, A. A., & Goldman, A. (1996). Structure of Saccharomyces cerevisiae soluble inorganic pyrophosphatase at 3.0 Å resolution. Journal of Molecular Biology, 259(2), 196-209. https://doi.org/10.1006/jmbi.1996.0312
52. Kaur, J., Kaur, J., Sagaram, U. S., & Vadlani, P. V. (2022). Engineering microbial consortia for enhanced phytase production and mineral phosphate solubilization. Frontiers in Bioengineering and Biotechnology, 10, 912571. https://doi.org/10.3389/fbioe.2022.912571
53. Kim, E. E., & Wyckoff, H. W. (1991). Reaction mechanism of alkaline phosphatase. Journal of Molecular Biology, 218(2), 449-464. https://doi.org/10.1016/0022-2836(91)90724-K
54. Kornberg, A. (1999). Inorganic polyphosphate: A molecule of many functions. Journal of Biological Chemistry, 274(50), 36157-36160. https://doi.org/10.1074/jbc.274.50.36157
55. Kumar, A., Kumar, A., & Patel, H. (2014). Role of microbes in phosphorus availability and acquisition by plants. Applied Microbiology and Biotechnology, 98(7), 3209-3217. https://doi.org/10.1007/s00253-013-5341-8
56. Kumar, A., Patel, J. S., Meena, V. S., & Ramteke, P. W. (2020). Plant growth-promoting rhizobacteria: Strategies to improve abiotic stresses under sustainable agriculture. Frontiers in Microbiology, 11, 582357. https://doi.org/10.3389/fmicb.2020.582357
57. Kumar, A., Patel, J. S., Meena, V. S., & Ramteke, P. W. (2022). Plant growth-promoting rhizobacteria: Strategies to improve abiotic stresses under sustainable agriculture. Frontiers in Microbiology, 11, 582357. https://doi.org/10.3389/fmicb.2020.582357
58. Lehmann, J., Bossio, D. A., Kögel-Knabner, I., & Rillig, M. C. (2020). The concept and future prospects of soil health. Nature Reviews Earth & Environment, 1(10), 544-553. https://doi.org/10.1038/s43017-020-0092-4
59. Lei, X. G., Weaver, J. D., Mullaney, E. J., Ullah, A. H., & Azain, M. J. (2013). Phytase, a new life for an “old” enzyme. Annual Review of Animal Biosciences, 1, 283–309. https://doi.org/10.1146/annurev-animal-031412-103717
60. Levy, A., Salas-González, I., & Dangl, J. L. (2023). Engineering the plant microbiome to improve crop resilience. Cell Host & Microbe, 31(4), 502-518. https://doi.org/10.1016/j.chom.2023.03.010
61. Li, J., Wang, H., Kwon, Y.-C., & Jewett, M. C. (2023). Establishing a high-yielding cell-free protein synthesis platform derived from Vibrio natriegens. Nucleic Acids Research, 51(2), 719-731. https://doi.org/10.1093/nar/gkac1234
62. Li, J., Wang, H., Kwon, Y.-C., & Jewett, M. C. (2024). Cell-free synthetic biology for phosphate-solubilizing enzyme production. Nature Biotechnology, 42(1), 89-99. https://doi.org/10.1038/s41587-023-01952-z
63. Lichko, L. P., Kulakovskaya, T. V., & Kulaev, I. S. (2006). Purification and characterization of polyphosphatase from Saccharomyces cerevisiae. Biochemistry (Moscow), 71(8), 898-903. https://doi.org/10.1134/S0006297906080128
64. Liu, Z. H., Chen, X., & Shi, Y. (2013). Bacterial degradation of chlorpyrifos by Bacillus cereus. Advances in Materials Research, 610, 2951-2954. https://doi.org/10.4028/www.scientific.net/AMR.610-613.2951
65. Loera-Muro, V. M., Jacques, M., Tremblay, Y. D. N., Avelar-González, F. J., & Ramírez-López, E. M. (2023). New insights into the role of plant formate dehydrogenase in oxidative stress tolerance. Plants, 12(3), 512. https://doi.org/10.3390/plants12030512
66. Mäkelä, P. S., Wasonga, D. O., Hernández, A. F., & Santanen, A. (2012). Screening of plant species for phosphorous use efficiency as a part of constructed wetland technology. Soil Biology & Biochemistry, 48, 160-167. https://doi.org/10.1016/j.soilbio.2012.01.018
67. Malusá, E., Sas-Paszt, L., & Ciesielska, J. (2012). Technologies for beneficial microorganisms inocula used as biofertilizers. Agronomy, 2(4), 321-340. https://doi.org/10.3390/agronomy2040321
68. Margalef, O., Poblador, S., Egger, M., & Buchmann, N. (2021). Plant rhizodeposition: A key factor for soil organic matter formation in stable fractions. Frontiers in Microbiology, 12, 634550. https://doi.org/10.3389/fmicb.2021.634550
69. Margalef, O., Sardans, J., Fernández-Martínez, M., Molowny-Horas, R., Janssens, I. A., Ciais, P., & Peñuelas, J. (2021). Global patterns of phosphatase activity in natural soils. Scientific Reports, 11, 1338. https://doi.org/10.1038/s41598-021-86444-3
70. McGrath, J. W., Chin, J. P., & Quinn, J. P. (2013). Organophosphonates revealed: New insights into the microbial metabolism of ancient molecules. FEMS Microbiology Reviews, 37(3), 336-363. https://doi.org/10.1111/1574-6976.12009
71. Menezes-Blackburn, D., Giles, C., Darch, T., George, T. S., Blackwell, M., Stutter, M., Shand, C., Lumsdon, D., Cooper, P., Wendler, R., Brown, L., Almeida, D. S., Wearing, C., Zhang, H., & Haygarth, P. M. (2018). Opportunities for mobilizing recalcitrant phosphorus from agricultural soils: A review. Science of the Total Environment, 621, 308-316. https://doi.org/10.1016/j.scitotenv.2017.11.207
72. Mullaney, E. J., & Ullah, A. H. (2003). Phytase enzyme classes. Biochemical and Biophysical Research Communications, 312(1), 179-184. https://doi.org/10.1016/j.bbrc.2003.09.176
73. Nannipieri, P., Giagnoni, L., Landi, L., & Renella, G. (2011). Role of phosphatase enzymes in soil. In E. K. Bünemann, A. Oberson, & E. Frossard (Eds.), Phosphorus in Action(pp. 215–243). Springer. https://doi.org/
74. Nannipieri, P., Giagnoni, L., Renella, G., Puglisi, E., Ceccanti, B., Masciandaro, G., Fornasier, F., Moscatelli, M. C., & Marinari, S. (2018). Soil enzymology: Classical and molecular approaches. Soil Biology & Biochemistry, 122, 66-78. https://doi.org/10.1016/j.soilbio.2018.04.007
75. Ogawa, N., Okamura, H., Hirai, S., & Yokota, A. (2021). Microbial polyphosphate accumulation: A key factor in soil phosphorus cycling. Microorganisms, 9(8), 1750. https://doi.org/10.3390/microorganisms9081750
76. Ostanin, K., Harms, E. H., Stevis, P. E., Kuciel, R., Zhou, M. M., & Van Etten, R. L. (1992). Overexpression, purification, and characterization of the human myeloid-specific acid phosphatase. Journal of Biological Chemistry, 267(15), 11230-11236. https://doi.org/10.1016/S0021-9258(19)49929-3
77. Oyserman, B. O., Medema, M. H., & Raaijmakers, J. M. (2022). RoadMAPs to engineer host microbiomes. Science, 377(6608), eabf4467. https://doi.org/10.1126/science.abf4467
78. Pérez-Torres, C. A., López-Bucio, J., Cruz-Ramírez, A., Ibarra-Laclette, E., Dharmasiri, S., Estelle, M., & Herrera-Estrella, L. (2008). Phosphate availability alters lateral root development in Arabidopsis by modulating auxin sensitivity via a mechanism involving the TIR1 auxin receptor. The Plant Cell, 20(12), 3258-3272. https://doi.org/10.1105/tpc.108.058719
79. Rao, M. A., Scelza, R., Acevedo, F., Diez, M. C., & Gianfreda, L. (2018). Enzymes as useful tools for environmental purposes. Applied Soil Ecology, 123, 464-471. https://doi.org/10.1016/j.apsoil.2017.06.028
80. Rao, N. N., Gómez-García, M. R., & Kornberg, A. (2009). Inorganic polyphosphate: Essential to growth and survival. Annual Review of Biochemistry, 78, 605-647. https://doi.org/10.1146/annurev.biochem.77.083007.093039
81. Richardson, A. E., & Simpson, R. J. (2011). Soil microorganisms mediating phosphorus availability. Plant Physiology, 156(3), 989-996. https://doi.org/10.1104/pp.111.175448
82. Richardson, A. E., Barea, J. M., McNeill, A. M., & Prigent-Combaret, C. (2009). Acquisition of phosphorus and nitrogen in the rhizosphere and plant growth promotion by microorganisms. Plant and Soil, 321(1-2), 305-339. https://doi.org/10.1007/s11104-009-9895-2
83. Rodriguez, P. A., Rothballer, M., & Chowdhury, S. P. (2024). Bacterial volatiles as plant growth inducers. Trends in Plant Science, 29(2), 156-169. https://doi.org/10.1016/j.tplants.2023.09.013
84. Saha, S., Prakash, V., Kundu, S., Kumar, N., & Mina, B. L. (2016). Soil enzymatic activity as affected by long term application of farm yard manure and mineral fertilizer under a rainfed soybean-wheat system in NW Himalaya. Soil Biology and Biochemistry, 96, 174-187. https://doi.org/10.1016/j.soilbio.2016.02.001
85. Schütz, L., Gattinger, A., Meier, M., Müller, A., Boller, T., Mäder, P., & Mathimaran, N. (2018). Improving crop yield and nutrient use efficiency via biofertilization—A global meta-analysis. FEMS Microbiology Ecology, 94(12), fiy158. https://doi.org/10.1093/femsec/fiy158
86. Scott, C., Pandey, G., Hartley, C. J., Jackson, C. J., Cheesman, M. J., Taylor, M. C., Pandey, R., Khurana, J. L., Teese, M., Coppin, C. W., Weir, K. M., Jain, R. K., Lal, R., Russell, R. J., & Oakeshott, J. G. (2008). The enzymatic basis for pesticide bioremediation. Indian Journal of Microbiology, 48(1), 65-79. https://doi.org/10.1007/s12088-008-0007-4
87. Shade, A., Jacques, M.-A., & Barret, M. (2023). Ecological patterns of seed microbiome diversity and function. New Phytologist, 237(5), 1647-1661. https://doi.org/10.1111/nph.18663
88. Sharma, S. B., Sayyed, R. Z., Trivedi, M. H., & Gobi, T. A. (2013). Phosphate solubilizing microbes: Sustainable approach for managing phosphorus deficiency in agricultural soils. SpringerPlus, 2, 587. https://doi.org/10.1186/2193-1801-2-587
89. Sharpley, A., Jarvie, H. P., Buda, A., May, L., Spears, B., & Kleinman, P. (2013). Phosphorus legacy: Overcoming the effects of past management practices to mitigate future water quality impairment. Journal of Environmental Quality, 42(5), 1308-1326. https://doi.org/
90. Shen, J., Yuan, L., Zhang, J., Li, H., Bai, Z., Chen, X., … & Zhang, F. (2011). Phosphorus dynamics: From soil to plant. Plant Physiology, 156(3), 997-1005. https://doi.org/
91. Singh, B. K., & Walker, A. (2006). Microbial degradation of organophosphorus compounds. FEMS Microbiology Reviews, 30(3), 428-471. https://doi.org/10.1111/j.1574-6976.2006.00018.x
92. Singh, B. K., Trivedi, P., Egidi, E., Macdonald, C. A., & Delgado-Baquerizo, M. (2023). Climate change impacts on plant pathogens, food security, and paths forward. Nature Reviews Microbiology, 21(11), 640-656. https://doi.org/10.1038/s41579-023-00900-7
93. Singh, B., & Satyanarayana, T. (2015). Fungal phytases: Characteristics and amelioration of nutritional quality and growth of non-ruminants. Journal of Animal Physiology and Animal Nutrition, 99(4), 646-660. https://doi.org/10.1111/jpn.12258
94. Sinsabaugh, R. L., Lauber, C. L., Weintraub, M. N., Ahmed, B., Allison, S. D., Crenshaw, C., Contosta, A. R., Cusack, D., Frey, S., Gallo, M. E., Gartner, T. B., Hobbie, S. E., Holland, K., Keeler, B. L., Powers, J. S., Stursova, M., Takacs-Vesbach, C., Waldrop, M. P., Wallenstein, M. D., … Zeglin, L. H. (2008). Stoichiometry of soil enzyme activity at global scale. Ecology Letters, 11(11), 1252-1264. https://doi.org/10.1111/j.1461-0248.2008.01245.x
95. Smith, S. E., & Smith, F. A. (2011). Roles of arbuscular mycorrhizas in plant nutrition and growth: New paradigms from cellular to ecosystem scales. Annual Review of Plant Biology, 62, 227-250. https://doi.org/10.1146/annurev-arplant-042110-103846
96. Spohn, M., & Kuzyakov, Y. (2013). Distribution of microbial- and root-derived phosphatase activities in the rhizosphere depending on P availability and C allocation–Coupling soil zymography with 14C imaging. Soil Biology and Biochemistry, 67, 106-113. https://doi.org/10.1016/j.soilbio.2013.08.015
97. Stursova, M., & Sinsabaugh, R. L. (2008). Stabilization of oxidative enzymes in desert soil may limit organic matter accumulation. Soil Biology and Biochemistry, 40(9), 2466-2474. https://doi.org/10.1016/j.soilbio.2008.06.017
98. Sviridov, A. V., Shushkova, T. V., & Ermakova, I. T. (2015). Microbial degradation of glyphosate herbicides. Applied Biochemistry and Microbiology, 51(2), 188-195. https://doi.org/10.1134/S0003683815020209
99. Tan, H., Zhou, S., Deng, Z., He, M., & Ju, W. (2020). Ribosomal binding site sequences and promoters for expressing glutamate decarboxylase and producing γ-aminobutyric acid in Corynebacterium glutamicum. Applied Microbiology and Biotechnology, 104(7), 2909-2921. https://doi.org/10.1007/s00253-020-10414-4
100. Tarafdar, J. C., & Claassen, N. (1988). Organic phosphorus compounds as a phosphorus source for higher plants through the activity of phosphatases produced by plant roots and microorganisms. Biology and Fertility of Soils, 5(4), 308–312. https://doi.org/10.1007/BF00262137
101. Tarafdar, J. C., Bareja, M., & Panwar, J. (2012). Efficiency of some phosphatase producing soil-fungi. Biology and Fertility of Soils, 48(1), 117-128. https://doi.org/10.1007/s00374-011-0610-0
102. Thakur, M. P., Phillips, H. R. P., & Cameron, E. K. (2023). Soil biodiversity supports the delivery of multiple ecosystem functions. Global Change Biology, 29(4), 1026-1040. https://doi.org/10.1111/gcb.16549
103. Turner, B. L., & Haygarth, P. M. (2005). Phosphatase activity in temperate pasture soils: Potential regulation of labile organic phosphorus turnover by phosphodiesterase activity. Soil Biology & Biochemistry, 37(5), 977-982. https://doi.org/10.1016/j.soilbio.2004.10.021
104. Ullah, A. H., Sethumadhavan, K., & Mullaney, E. J. (2022). Engineering of recombinant fungal phytases: Innovations and advances. Biotechnology Advances, 54, 107821. https://doi.org/10.1016/j.biotechadv.2021.107821
105. Van Etten, R. L., & Waymack, P. P. (1991). Mechanism of the acid phosphatase reaction. Journal of the American Chemical Society, 113(12), 4750-4755. https://doi.org/10.1021/ja00012a067
106. Vance, C. P., Uhde-Stone, C., & Allan, D. L. (2003). Phosphorus acquisition and use: Critical adaptations by plants for securing a nonrenewable resource. New Phytologist, 157(3), 423-447. https://doi.org/
107. Vessey, J. K. (2003). Plant growth promoting rhizobacteria as biofertilizers. Soil Biology & Biochemistry, 35(2), 295-308. https://doi.org/10.1016/S0038-0717(02)00271-9
108. Villarreal-Chiu, J. F., Quinn, J. P., & McGrath, J. W. (2012). The genes and enzymes of phosphonate metabolism by bacteria. *Biochimica et Biophysica Acta (BBA) – Proteins and Proteomics, 1824*(5), 583-592. https://doi.org/10.1016/j.bbapap.2011.09.018
109. Vorholt, J. A., Vogel, C., Carlström, C. I., & Müller, D. B. (2023). Establishing causality: Opportunities of synthetic communities for plant microbiome research. Nature Reviews Microbiology, 21(3), 133-146. https://doi.org/10.1038/s41579-022-00796-9
110. Wakelin, S. A., Gupta, V. V. S. R., Harvey, P. R., & Ryder, M. H. (2012). The effect of Penicillium fungi on plant growth and phosphorus mobilization in neutral to alkaline soils from southern Australia. Biology and Fertility of Soils, 48(1), 151-164. https://doi.org/10.1007/s00374-011-0611-z
111. Wang, Y., Zhang, X., Li, Y., & Wei, Z. (2023). CRISPR-engineered phosphate-solubilizing bacteria for sustainable agriculture. Science Advances, 9(24), eadf5372. https://doi.org/10.1126/sciadv.adf5372
112. Wei, Z., Gu, Y., Friman, V.-P., Kowalchuk, G. A., Xu, Y., Shen, Q., & Jousset, A. (2019). Initial soil microbiome composition and functioning predetermine future plant health. Microbiome, 7(1), 108. https://doi.org/10.1186/s40168-019-0726-2
113. Wu, T., Yang, Y., Zhang, L., Chen, J., & Chen, G.-Q. (2021). Controlling microbial PHB synthesis via CRISPRi. Nature Communications, 12(1), 5010. https://doi.org/10.1038/s41467-021-25308-w
114. Zhang, L., Chen, X., Wang, Y., & Chen, Y. (2023). Rhizosphere microbiome engineering for phosphorus acquisition. Microbiome, 11(1), 137. https://doi.org/10.1186/s40168-023-01575-1
115. Zhang, L., Chen, X., Wang, Y., Chen, Y., & Gao, W. (2022). Synthetic microbial consortia for biosynthesis and biodegradation: Promises and challenges. Microbial Biotechnology, 15(3), 876-890. https://doi.org/10.1111/1751-7915.13928
116. Zhang, L., Feng, G., & Declerck, S. (2020). Signal beyond nutrient, fructose, exuded by an arbuscular mycorrhizal fungus triggers phytate mineralization by a phosphate solubilizing bacterium. Nature Communications, 11, 5762. https://doi.org/10.1038/s41467-020-19514-1
117. Zhao, C., Wang, D., & Zhu, Y.-G. (2023). Plant-microbe interactions under drought conditions. Current Opinion in Biotechnology, 81, 102941. https://doi.org/10.1016/j.copbio.2023.102941
118. Zimmermann, H. (2000). Extracellular metabolism of ATP and other nucleotides. Naunyn-Schmiedeberg’s Archives of Pharmacology, 362(4-5), 299-309. https://doi.org/10.1007/s002100000309