An essential nutrient element

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               Phosphorus (P) is an essential nutrient element for the growth of living organisms including plants, animals and microorganisms. It is a component of the adenosine triphosphate (ATP) that drives most energy-requiring biochemical processes, deoxyribonucleic acid (DNA) that is the seat of genetic inheritance, ribonucleic acid (RNA) that directs protein synthesis in plants and animals, phospholipids that compose cellular membranes and intermediate compounds of respiration and photosynthesis (Brady and Weil, 1996; Taiz and Zeiger, 1998; Fuentes et al., 2006).For most plant species, it is the second most abundantly required nutrient element after nitrogen and the total phosphorus content of healthy leaf tissue range between 0.2 and 0.4% of the dry matter (Brady and Weil, 2002).

               Phosphorus in soil comes from both pedogenic and anthropogenic sources (Bolan et al., 2005). In spite of its wide distribution in nature, P is a limited resource (Adnan et al., 2003; Shimamura et al., 2003) and is deficient in most soils with respect to its availability to plants (Vassilev et al., 2001). The P problem in soil fertility is threefold. First, the total P level of soils is low, ranging from 200 to 2000 kg P per hectare-furrow slice (HFS), with an average of about 1000 kg P per HFS. Second, the P compounds commonly present in soils are mostly unavailable for plant uptake, often because they are highly insoluble. Third, when soluble sources of P, such as those in fertilizers and manures are added to soils, they are fixed and only 10 to 15 percent of the P added through fertilizers is likely to be taken up by plants in the year of application (Brady and Weil, 2002). Hence, low P bioavailability limits crop production under most soil conditions.

               The research work carried out so far either at national or international level to improve P nutrition of crop plants has been mainly confined to exploring the physical and chemical processes related to improving P availability in soil. Microbial and biochemical processes which are the key elements behind almost all biochemical transformations leading to nutrients bioavailability in soil have in general been less explored. Microbial biomass is the most labile fraction of soil organic matter, and plays a vital role in sustainability of soil fertility and the functioning of soil ecosystem (Jenkinson and Ladd, 1981; Smith and Paul, 1990; Khan and Joergensen, 2006). The magnitude of microbial biomass pool directly affects the nutrients flux and their bioavailability in soil.

               Studies on the dynamics of microbial biomass, microbial P and enzymes activities such as dehydrogenase and alkaline phosphatase in relation to different P fractions in soil are important for understanding the role and contribution of different microbial/ biochemical parameters to P bioavailability in soils. The proposed research work will therefore be conducted to achieve the following objectives:

  1. To study different phosphorus fractions in soils of Potohar and their relationship with soil physical, chemical, and microbial parameters.
  2. To evaluate dynamics of different P fractions in soil in response to various organic amendments and their relationship to soil microbial parameters.


               The literature pertaining to the proposed research is reviewed as under:

               Stewart and Tiessen (1987) studied dynamics of soil organic phosphorus. They reported that microbial uptake of P and its subsequent release and redistribution play a central role in the soil organic P cycle. Interactions with soil minerals and stabilization of organic matter and associated P in organo-mineral complexes determine the persistence and build up of organic P through soil development, in different ecosystems and management conditions. They concluded that an understanding of organic P turnover in soils will be highly helpful in assessment of P fertility of many agricultural and native systems.

               Lee et al. (1990) studied the influence of microbial activity in mobilizing P, maintaining it in a plant-available state, and preventing its fixation, and the effect of N and biocides on these processes in a highly weathered Ultisol. Exchangeable aluminum and soil moisture were also determined, since they interact with microbes and soil P. They found that increased microbial activity reduced sorption of dissolved and organic P by soil, maintained inorganic P in soluble and labile pools, increased microbial P, decreased mineral P, increased exchangeable Al, and water retention. Additions of N and biocides had variable effects probably due to complex interactions between N, degrading biocides and microbial populations.

               Maguire et al. (2000) conducted a study to identify the effect of biosolids applications on the forms and release potential of P in agricultural soils. They collected samples from eight farms with a history of biosolids amendments, selecting fields that had setback areas (where biosolids applications were not permitted) to allow comparison of amended and unamended soils and analyzed them for P fractions (soluble P, Al-P, Fe-P, reluctant soluble P, and Ca-P; their sum equals total P), sequentially desorbable P (Fe-strip), oxalate P, Al and Fe, Mehlich-1 P, and the degree of P saturation. Results showed that following a N-based biosolids nutrient management plan could significantly increase total P (from 403 to 738 mg kg-1) and initially desorbable P (from 32 to 61 mg kg-1). The main soil components associated with P retention (Alox and Feox) also tended to be increased by biosolids amendment and this may help mitigate P release. Biosolids amendment significantly increased Fe-P (from 137 to 311 mg kg-1), probably due to Fe added to biosolids during production, and there was also a strong trend for higher Al-P where biosolids had been applied. Desorbable P was initially greatest from biosolids sites, but with increasing extractions, the release converged towards that from the setback areas. Mehlich-1 P and Pox were good predictors of desorbable P release, as measured by one and five sequential extractions with Fe-strips. Desorbable P, by both one and five Fe-strip extractions, was more closely correlated with Al-P than Fe-P, especially in setback areas, indicating that Al-P is probably the most important source of desorbable P independent of biosolids amendment.

               Qualls and Richardson (2000) studied the influence of P additions on the elevation of microbial biomass P in the soil. In order to isolate the effects of P enrichment, they placed bags containing cattail (Typha domengensis Crantz) and sawgrass (Cladium jamaicense Pers.) litter into two sets of experimental channels into which controlled inputs of five different phosphate concentrations were added continuously. After one year of incubation, litter was analyzed for C, P, N, Cu, Ca, and K content. Loss of C at the end of one year increased linearly with increasing average PO4 content in the channels with a similar slope for both species of litter. Immobilization caused an absolute increase in P content of the litter up to approximately nine fold across the range of water P concentrations, while immobilization of N, Ca, and K did not vary with water P concentrations. The microbial biomass P was up to nine times higher in the surface soil of the most enriched channel compared with the control, but this elevation in concentration was restricted to the upper 12 cm of soil.

               Cleveland et al. (2002) tested the effects of P availability on the decomposition of multiple forms of C, including dissolved organic carbon and soil organic carbon (SOC) using natural gradients in P fertility created by soils of varying age underlying tropical rain forests in southwestern Costa Rica, combined with direct manipulations of carbon (C) and P supply. Results from a combination of laboratory and field experiments suggested that C decomposition in old, highly weathered oxisol soils is strongly constrained by P availability. In addition, P additions to these soils (no C added) further revealed that microbial utilization of at least labile fractions of SOC was also P limited. This was regarded to be the first direct evidence of P limitation of microbial processes in tropical rain forest soil. They suggested that P limitation of microbial decomposition might have profound implications for C cycling in moist tropical forests, including their potential response to increasing atmospheric carbon dioxide.

               Kabba and Aulakh (2004) conducted an experiment to examine the effect of climatic conditions and crop residue quality on N, P, and S mineralization in soils with contrasting P status. They observed the effect of three temperatures (15 °C, 30 °C, and 45 °C) and two moisture regimes (60% and 90% water-filled pore space (WFPS)) on the mineralization-immobilization of N, P, and S from groundnut (Arachis hypogea) and rapeseed (Brassica napus) residues (4 t ha-1) in two soils with contrasting P fertility. Crop residue mineralization was differentially affected by incubation temperature, soil aeration status, and residue quality. Only the application of groundnut residues (low C: nutrient ratios) resulted in a positive net N and P mineralization within 30 days of incubation, while net N and P immobilization was observed with rapeseed residues. The initial P content influenced the mineralization of N and P, which was significantly higher in the soil with a high initial P fertility (18 mg P (kg soil)-1) than in soil with low P status (8 mg P (kg soil)-1).

               Saleque et al. (2004) conducted the experiment to evaluate the effect of different nutrient management in wetland rice on the changes of soil P fraction at different depths. Soil samples from five depths (0-5, 5-10, 10-15, 15-30, and 30-50 cm) were collected from a long-term experimental field. The field received six treatments for 10 year: absolute control with no fertilizer applied (T1), one-third of recommended fertilizer doses (T2), two-thirds of recommended fertilizer doses (T3), full doses of recommended fertilizers (T4), T2 + 5 Mg cow dung (CD) and 2.5 Mg ash ha-1 (T5), and T3 + 5 Mg CD and 2.5 Mg ash ha-1 (T6). The apparent balance of P compared with the initial P status after 10 years varied from -115 kg ha-1 under T1 to 348 kg ha-1 under T6. The P fractionation study was conducted over the treatments and soil depth. Treatment and depth had no significant effect on solution P. Larger concentrations of NaHCO3 soluble P, NaOH extracted inorganic P (Pi), and acid P were observed under treatments with organic fertilizers (T5 and T6) than with other treatments at 0 to 5, 5 to 10, and 10 to 15cm depths. The concentrations of NaHCO3-P, NaOH-Pi and acid P fractions were lowest under T1 and T2 treatments. At 15 to 30 cm or lower soil depths, none of the P fractions were affected by treatments. The change in NaOH organic P (Po) and residual P (extracted with HNO3 + HClO4) with soil depth was not significant, and the differences in these P fractions under the tested P treatments were not large.

               Kaur et al. (2005) conducted studies to compare the soils receiving organic manures with and without chemical fertilizers for the last 7 years with pearl millet-wheat cropping sequence for soil chemical and biological properties. The application of farmyard manure, poultry manure and sugarcane filter cake alone or in combination with chemical fertilizers improved the soil organic C, total N, P, and K status. The increase in soil microbial biomass C and N was observed in soils receiving organic manures only or with the combined application of organic manures and chemical fertilizers compared to soils receiving chemical fertilizers only. Basal and glucose-induced respiration, potentially mineralizable N, and arginine ammonification were higher in soils amended with organic manures with or without chemical fertilizers, indicating that more active microflora is associated with organic and integrated system using organic manures and chemical fertilizers together which is important for nutrient cycling.

               Khan and Joergensen (2006) conducted a study to analyze the amounts of microbial-biomass C, biomass N, and biomass P in 11 rain-fed arable soils of the Potohar plateau, Pakistan, in relation to the element-specific total storage compartment, i.e., soil organic C, total N, and total P. Average contents of soil organic C, total N, and total P were 3.9, 0.32, and 0.61 mg per g soil, respectively. Less than 1 percent of total P was extractable with 0.5 M NaHCO3. Mean contents of microbial biomass C, biomass N, and biomass P were 118.4, 12.0, and 3.9 µg per g soil, respectively. Values of microbial biomass C, biomass N, biomass P, soil organic C, and total N were all highly significantly interrelated. The mean crop yield level was closely connected with all soil organic matter and microbial biomass-related properties. The fraction of NaHCO3-extractable P was also closely related to soil organic matter, soil microbial biomass, and crop yield level. This revealed the overwhelming importance of biological processes for P turnover in alkaline soils.

               Smith et al. (2006) conducted an experiment to determine the effect of different stages of sewage sludge treatment on phosphorus (P) dynamics in amended soils using samples of undigested liquid (UL), anaerobically digested liquid (AD) and dewatered anaerobically digested (DC) sludge. Sludges were taken from three points in the same treatment stream and applied to a sandy loam soil in field-based mesocosms at 4, 8 and 16 t ha-1 dry solids. Mesocosms were sown with perennial ryegrass (Lolium perenne cv. Melle), and the sward was harvested after 35 and 70 days to determine yield and foliar P concentration. Soils were also sampled during this period to measure P transformations and the activities of acid phosphomonoesterase and phosphodiesterase. Data showed that the AD amended soils had the greatest plant-available and foliar P content up to the second harvest, but the UL amended soils had the greatest enzyme activity. Characterization of control and 16 t ha-1 soils and sludge using solution 31P nuclear magnetic resonance (NMR) spectroscopy after NaOH-EDTA extraction revealed that P was predominantly in the inorganic pool in all three sludge samples, with the highest proportion (of the total extracted P) as inorganic P in the anaerobically digested liquid sludge. After sludge incorporation, P was immobilized to organic species. The majority of organic P was in monoester-P forms, while the remainder of organic P (diester P and phosphonate P) was more susceptible to transformations through time and showed variation with sludge type.



               The study will consist of three experiments, details of which are described below:

  1. Study-1:
  2.                Study of phosphorus fractions in Pothwar soils and their relationship with soil microbial and biochemical parameters.

                   In this study, representative soil samples from 12-15 prominent soil series of the Pothwar plateau will be collected from the agricultural fields. The soil samples of 1.5 kg will be taken at 0-15 cm depth from 4 different locations of each of the selected site. The samples will be properly labeled, packed in polyethylene bags, brought to the laboratory. The field moist samples will be hand picked to remove stones, larger plant residues and soil animals (earth worms etc.), passed through a 2-mm sieve, mixed thoroughly and stored in polyethylene bags at 5 °C prior to biological analysis.

                   A portion of each soil sample will be air-dried, ground to powder form and analyzed for selected physico-chemical properties such as particle size analysis, EC, pH, CEC, CaCO3, organic C, total N, total P, P fractionation and water soluble cations (Na, K, Ca, Mg) and anions (CO3, HCO3, Cl, SO4).The soil samples prepared and stored for biological analysis will be equilibrated to room temperature, incubated at 30 °C for 7 days after moisture adjustment to 50 percent of their water holding capacity (WHC) and analyzed for microbial biomass C, biomass N, biomass P, soil respiration, and the activities of enzymes like dehydrogenase and alkaline phosphatase. The data obtained will be analysed statistically to evaluate the relationship of different P fractions with soil physical, chemical, microbial and biochemical properties.

  3. Study-II:
  4.                Dynamics of phosphorus fractions in soils amended with organic manures and their relationship with soil microbial and biochemical parameters.

                   In this study, two soils deficient in plant available phosphorus with variable physico-chemical properties such as clay content, pH, or organic C will be selected on the basis of the results of study-I. The soils will be adjusted to 50 percent of their water holding capacity and incubated at 30 °C for 7 days prior to amendment addition. The treatments will include: 1) Control; 2) Farmyard manure (FYM); 3) Poultry litter (PL) and 4) Biogenic waste compost (BWC), each applied to 600 g (oven dry basis) soil separately at the rate of 1 percent of the oven dry soil weight. All the treatments will be quadruplicated according to completely randomized design (CRD).

                   After amendment addition, soil samples will be transferred to 2 litre capacity incubation jars and incubated at 30 °C for a period of 72 days. The CO2 evolved will be absorbed into 2M NaOH solution in 100 ml beakers. The NaOH solution will be changed after 1, 2, 3, 5, 7, 10, and 14 days and thereafter weekly. Soil samples of 50 g oven dry weight will be taken at 0, 14, 28, 56, and 72 days of incubation for the determination of microbial biomass C, biomass N, biomass P, and 0.5M NaHCO3-extractable P. Different P fractions and activities of enzymes like dehydrogenase and alkaline phosphatase will be measured in samples collected at 0 and 72 days of incubation.

  5. Study-III:
  6.                Relationship between microbial biomass, enzyme activities and P availability in soils amended with organic manures under wheat crop.

                   A greenhouse experiment will be conducted in completely randomized design (CRD) to evaluate the effect of organic amendments on the relationship between soil microbial biomass, enzyme activities and P availability in soil under wheat crop. For this purpose, two soils used in study-II will be collected, passed through 2-mm sieve and amended with organic manures. The treatments will include: 1) Control; 2) Farmyard manure (FYM); 3) Poultry litter (PL) and 4) Biogenic compost (BC), each applied to 5 kg (oven dry basis) soil separately at the rate of 1 percent of the oven dry soil weight. All the treatments will be quadruplicated and the moisture contents will be adjusted to field capacity gravimetrically.

                   Seeds of wheat will be sown after 2 weeks of organic amendments addition. Soil samples will be collected at 0, 14, 28, 42 and 56 days after the sowing of seeds and analyzed for microbial biomass C, biomass N, biomass P, enzyme activities like dehydrogenase and alkaline phosphatase, and 0.5M NaHCO3 extractable P. After 56 days, plants will be harvested and data on plant growth parameters such as plant height, oven dry weight etc. will be recorded. Plant samples will be washed properly with distilled water, oven dried at 60 oC, ground in Wiley Mill and analyzed for important macro and micronutrients and phosphorus uptake will be calculated.


  1. Soil Analysis:
  2. Particle size analysis:
  3.                To 40 g of soil sample, 40 ml of 1 % sodium hexametaphosphate and 150 ml of distilled water will be added and the suspension will be kept over night. After stirring for ten minutes, the contents will be shifted to 1000 ml capacity cylinder and reading will be recorded with the soil hydrometer. Soil textural class will be determined by using ISSS triangle (Gee and Bauder, 1986).

  4. Soil moisture:
  5.                Soil samples will be taken in metallic cans and weights will be recorded. The samples will be dried to constant weight at 105oC in the oven. Afterwards, the samples will be removed from the oven and weight will be recorded after cooling. Soil moisture will be determined by the following formula:

    Weight of wet soil - weight of oven dry soil

    Soil moisture = 100

                   Weight of oven dried soil

  6. Water holding capacity:
  7.                Water holding capacity will be determined by preparing the saturated soil paste. The paste will be then transferred to a porous Buckner funnel for the removal of surplus moisture and the water contents held by the soil will be determined gravimetrically (Anderson and Ingram, 1993).

  8. Electrical conductivity (EC):
  9.                Soil water suspension will be prepared using a soil to water ratio of 1:2.5. The contents will be allowed to equilibrate for 30 minutes and the electrical conductivity will be recorded using conductivity meter (Page et al., 1982).

  10. pH:
  11.                Soil water suspension will be prepared using a soil to water ratio of 1:2.5. The contents will be allowed to equilibrate for 30 minutes and the pH will be measured using a calibrated pH meter (Page et al., 1982).

  12. Total organic carbon:
  13.                Two gram soil will be taken in a 500 ml Erlenmeyer flask. Ten ml of 1 N potassium dichromate will be added into flask and the flask will be swirled to mix the contents. Then 20 ml of conc. sulphuric acid will be added in the soil suspension. Flask will be swirled for one minute and allowed to stand for 30 minutes. Then 200 ml of water, 10 ml of phosphoric acid and 1 ml of diphenylamine indicator will be added in the flask. The contents will be titrated against 0.5 N ferrous sulphate solution until colour changes from blue to red (Page et al., 1982).

  14. Total nitrogen:
  15.                Total nitrogen (TN) will be determined by colorimetric analysis of digested soil samples. To 0.2 g of soil sample in separate digestion tubes, 4.4 ml of digestion mixture containing selenium powder, lithium sulphate and hydrogen per oxide will be added and it will be digested for two hours at 360oC till solution is colourless, 50 ml of water will be added and mixed well. After cooling, it will be made up to 100 ml and mixed. After the settlement, the clear solution will be analysed for total nitrogen colorimetrically.

  16. Cation exchange capacity:
  17.                Air-dried soil and 33 ml of 1N sodium acetate trihydrate will be shaken in a centrifuge tube for 5 minutes. Then, solution will be centrifuged at 3000 rpm until the supernatant becomes clear. The supernatant will be discarded 4 times after decanting. Then, 33 ml ethanol will be added and centrifuged again for three times. Sodium from sample will be replaced with ammonium acetate solution and measured on the flame photometer.

    CEC (meq/100g) = meq/l Na (from calibration curve)*A/Wt*100/1000


                   A = total volume of the extract (ml).

                   Wt = weight of the air-dry soil (g).

  18. Calcium carbonate:
  19.                One gram soil will be taken in 250 ml flask. 10 ml of 1N HCl will be added, and contents will be heated at 50-60 0C and thereafter cooled. 50 ml of deionized water and 2-3 drops of phenolphthalein indicator will be added and titrated with 1N NaOH until faint pink color will be developed. Percentage of carbonate in soil will be calculated by using formula (Page et al., 1982).

    % CaCO3 = {(10x N HCl) - (R x N NaOH) x 0.05 X 100 / Wt (soil)

              N HCl = Normality of HCl

                R = Volume of NaOH

              N NaOH = Normality of NaOH

  20. Soluble Calcium and Magnesium
  21.                By titrating the saturation extract with 0.01 N EDTA (Versenate) solution in the presence of NH4Cl + NH4OH buffer solution using Eriochrome Black T (EBT) as indicator. The colour will change from wine red to blue (H.B.60, Method, 7 P.94).

  22. Soluble Sodium and Potassium:
  23.                The same extract will be analysed for Sodium and Potassium contents by Flame Photometer (Richards, 1954).

  24. Phosphorus Fractionation
  25.                Sequential Phosphorus Fractionation procedure will be adopted. Fraction of inorganic and organic P will be performed on each soil by a modified P fractionation scheme of Sui and Thompson (1999). Soil P fractions including Solution P, NaHCO3-P, NaOH-Pi-P, NaOH-Po-P, Acid P and Residual P will be measured in sequence as described below:

    1. Solution P, by shaking 1 g soil in 30 mL of 0.05 M CaCl2 for 16 h, centrifuging, filtering, and measuring P in the filtrate.
    2. NaHCO3-P, by shaking the residue from (1) in 30 mL of 0.5 M NaHCO3 for 16 h, centrifuging, filtering, and measuring P in the filtrate.
    3. NaOH-Pi-P, by shaking the residue from (2) in 30 mL of 0.1 M NaOH, centrifuging, filtering, and measuring P in the filtrate after acidifying 5 mL (with concentrated HCl) and centrifuging.
    4. NaOH-Po-P, by digesting 5 mL of the filtrate from (2) in 6 mL of concentrated H2SO4 for 1 h, cooling, adding 5 mL of H2O2, and reheating until the residue became white, determining P in the digest, and subtracting the NaOH-Pi-P from it (Hedley et al., 1982).
    5. Acid P, by shaking the residue from (3) in 30 mL of 1:1 mixture of 1 M HCl/1 M H2SO4, centrifuging, filtering, and measuring P in the filtrate.
    6. Residual P, by refluxing the soil residue from (5) in 6 mL of a 5:2 mixture of concentrated HNO3 and HClO4, and determining P from the digest (Hedley et al., 1982).

                   All P will be determined colorimetrically (Murphy and Riley, 1962) after neutralization when necessary with dilute HCl and NaOH and the neutral pH indicated by the light yellow color of the solution in the presence of P-nitrophenol indicator. Absorbance for P will be determined at a wavelength of 712 nm by spectrophotometer.

Analyses of Soil Microbial Biomass:

  1. Microbial biomass C (Cmic):
  2.                Microbial biomass C and biomass N will be estimated by fumigation-extraction (Brookes et al., 1985) in the 30 g samples removed from each of the incubation beakers. One portion of 10 g (on oven dry basis) moist soil will be fumigated for 24 h at 25 °C with ethanol-free CHCl3. Following fumigant removal, the sample will be extracted with 40 ml 0.5M K2SO4 by 30 min horizontal shaking at 200 rev min-1 and filtered through a folded filter paper (Whatman No. 40). The non-fumigated 10 g portion will be extracted similarly at the time when fumigation commenced. Organic C in the extracts will be measured as CO2 by infrared absorption after combustion at 760 °C using a Shimadzu automatic TOC analyzer (Shimadzu Corp. Japan). Microbial biomass C will be calculated as follows: Microbial biomass C = Ec / kEC, where Ec = (organic C extracted from fumigated soils) - (organic C extracted from non-fumigated soils) and KEC = 0.45 (Wu et al., 1990).

  3. Microbial biomass N (Nmic):
  4.                Total N in the extracts will be measured as NO2 after combustion at 760 °C using a Shimadzu-N chemoluminescence detector (Shimadzu Corp. Japan). Microbial biomass N will be calculated as follows: Microbial biomass N = EN / kEN, where EN = (total N extracted from fumigated soils) - (total N extracted from non-fumigated soils) and kEN = 0.54 (Brookes et al., 1985; Joergensen and Muller, 1996).

  5. Microbial biomass P (Pmic):
  6.                Soil microbial biomass P will be also measured by fumigation-extraction (Brookes et al., 1982) as described by Joergensen et al. (1995). Three portions equivalent to 5 g oven-dry soil will be taken from the 50 g soil sample used for measuring the basal respiration and each will be extracted with 100 ml of 0.5 M NaHCO3 (pH 8.5) after different pre-treatment. The first portion will be used for the fumigation treatment (see above), the second portion for the non-fumigation treatment, and the third portion for estimating P fixation by the addition of 25 µg P g-1 soil as KH2PO4 to the extractant. P will be analysed by a modified ammonium molybdate-ascorbic acid method as described by Joergensen et al. (1995). Microbial biomass P will be calculated as follows:

    Microbial biomass P = EP / kEP / recovery, where EP = (PO4-P extracted from fumigated soil) - (PO4-P extracted from non-fumigated soil), kEP = 0.40 (Brookes et al., 1982). Recovery was calculated as follows: 1- ((PO4-P extracted from non-fumigated and spiked soil) - (PO4-P extracted from non-fumigated soil)) / 25.

Soil Enzymes Analysis:

  1. Alkaline Phosphatase:
  2.                One gram soil will be mixed with 0.2ml of toluene, 4ml of MUB (modified universal buffer) of pH 11, 1ml of p-nitrophenyl phosphatase solution and flask will be placed in an incubator at 37°C for one hour. Then, 1ml of 0.5M CaCl2 and 4 ml of 0.5M NaOH will be added and soil suspension will be filtered through a Whatman no. 2v folded filter paper. Yellow colour intensity will be measured at 400nm wavelength by using a spectrophotometer (Alef and Nannipieri, 1995).

  3. Dehydrogenase:
  4.                Air-dried soil (20g) will be mixed with 0.2g of CaCO3 and 6g of this mixture will be placed in each of the three test tubes. After adding 1ml of 3% aqueous solution of TTC (Triphenyl Tetrazolium Chloride) and 2.5ml of distilled water, samples will be incubated at 37°C for 24 hours. Then, 10ml of methanol will be added and filtered after shaking. The red color intensity will be measured by using a spectrophotometer at a wavelength of 485nm (Alef and Nannipieri, 1995).

Plant Analyses:

Following plant analyses will be carried out:

  1. Total nitrogen and total phosphorus:
  2.                Total nitrogen (TN) and total phosphorus (TP) will be determined from the same digested plant samples; however, their colorimetric determinations will be carried out separately.

  3. Digestion for total nitrogen and phosphorus:
  4.                To 0 .2 g of ground plant material in separate digestion tubes, 4.4 ml of digestion mixture containing selenium powder, lithium sulphate and hydrogen per oxide will be added and it will be digested for two hours at 360oC till solution is colourless, 50 ml of water will be added and mixed well. After cooling, it will be made up to 100 ml and mixed. After the settlement, the clearer solution will be ready for further analysis for TN and TP colorimetrically.

  5. Colorimetric determination of total nitrogen:
  6.                To 0.1 ml of each standard and sample, 5 ml of reagent containing sodium salicylate, sodium citrate, sodium tartarate and sodium nitroprusside will be added. It will be mixed well and left for 15 minutes. Then 5 ml of reagent containing a solution of NaOH, water and sodium hypochlorite will be added to each test tube and left for one hour for full colour development. Absorbance of samples will be measured using spectrophotometer at 665 nm (Anderson and Ingram, 1993).

    Plant TN will be calculated by the following formula:

    TN % = C/W ´ 0.01

    Where C is corrected concentration (µg /ml) and W is weight of sample (g).

  7. Colorimetric determination of total phosphorus:
  8.                To 1 ml of each standard and sample in test tubes, 4 ml of ascorbic acid will be added. Then 3 ml of molybdate reagent containing ammonium molybdate, antimony sodium tartarate and sulphuric acid will be added, mixed well and left for one hour for full colour development. The absorbance of standards and samples will be read at 880 nm (Anderson and Ingram, 1993).

    TP will be calculated by the following formula:

    P in digest (%) = C/W ´ 0.1

    Where C is corrected concentration (µg /ml) and W is weight of sample (g).

  9. Micronutrients in plant material:
  10.                One gram of dried plant sample will be taken in 50 ml conical flask and will be kept for overnight after adding 5 ml concentrated nitric acid (HNO3) and 5 ml perchloric acid (HClO4). Next day 5 ml concentrated HNO3 will be added again and will be digested on the hot plate till material will become clear. After digestion the material will be cooled down and the volume will be made up to 50 ml with distilled water and stored in clean airtight bottles for the analysis of micronutrients (Fe, Cu, Mn, Zn) by atomic absorption spectrophotometer (Rashid, 1986).

  11. Statistical Analyses:
  12.                All the results will be presented as arithmetic means expressed on an oven dry basis. In study-I, the relationships between the different soil properties will be analysed by principal component analysis (PCA). In studies II and III, the significance of treatment effects will be tested either by a soil-specific one way analysis of variance (ANOVA) using Tukey/ Kramer HSD (honestly significant difference) test or by a two-way ANOVA using soils and organic sources/ substrates as independent factors and sampling date as repeated measures (Steel and Torrie.1980). All statistical analyses will be performed using StatView 5.0 (SAS Inst. Inc.).


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Anderson, J.M. and J.S.I. Ingram. 1993. Tropical Soil Biology and Fertility. CAB International, Wallingford, UK.

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