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Development of a phosphatase activity assay using excised plant roots.

Introduction

The decline of soil fertility in the tropics and subtropics is a major hurdle towards sustainable food production and ensuring food security (Stoorvogel and Smaling 1998; Hartemink 2003; Hartemink 2006). Furthermore, considering that economically accessible phosphorus (P) fertiliser sources are expected to become exhausted over the next 50-100 years (Cordell et al. 2009), it is increasingly necessary to understand and improve the utilisation of other P sources to maintain food and fibre production into the future. A major alternative source of P is found in waste waters and solids. Another source is resident soil organic P, which becomes plant-available via root and microorganism processes.

Vast regions of tropical soils are highly weathered and contain low concentrations of bioavailable inorganic phosphate (Clinebell et al. 1995) but often contain considerable amounts of organic P compounds. Organic P has been noted as an important P source in maintaining P bioavailability in agricultural and natural resource landscapes (Johnson et al. 2003; Condron and Ticssen 2005). As much as 80% of total soil P can be in an organic form (Richardson 1994).

The nutritional significance of soil organic P, while recognised by many as being a long-term, slow-release P source (Vitousek and Sanford 1986; Zhang 1991; Tanner et al. 1998; Vitousek et al. 2010; Malik et al. 2012), is often not considered in evaluations of soil fertility (Schachtman et al. 1998; Baggie et al. 2004; Bolland and Guthridge 2007; Saifuzzaman et al. 2011). Organic phosphates bound within soil organic matter must be mineralised before being available for plant uptake (Tate and Salcedo 1988; Rojo et al. 1990; Hernandez et al. 1996; Bunemann et al. 2011). Rhizosphere soil has been shown to have up to eight times more phosphatase activity than bulk soil (Tarafdar and Jungk 1987; Fox and Comerford 1992), illustrating the role of enzyme-mediated organic-P cycling. Although the literature suggests possible direct uptake of dissolved organic P by phytoplankton and cyanobacteria (Bentzen and Taylor 1991; Cotner and Wetzel 1992; Bjorkman and Karl 1994; Glibert et al. 2004; Sebastian et al. 2012), this has not been demonstrated for higher plants. Hence, of special interest to the evaluation of soil fertility is the bioavailability of soil organic P mediated by an array of enzymes associated with roots and soil microorganisms.

The earliest attempts to determine phosphatase activity in plant roots were made with sterile hydroponic plants or germinating sterile seed cultures (Rose 1912; Rogers et al. 1941; McLachlan 1980; Gilbert et al. 1999) requiring specialised laboratory conditions. These were followed by pot-based experiments (Tarafder and Classen 1988; Helal 1990). Other methods included incubating commercially available enzymes with radiation-treated soil (Bishop et al. 1994), isotopic dilution (Oehl et al. 2001; Biinemann et al. 2007) and excised roots as a source of native phosphatases (MacFall et al. 1991; Antibus et al. 1997). All techniques have shortcomings, including the use of radioactive materials (Bishop et al. 1994; Oehl et al. 2001; Bunemann et al. 2007), long periods of incubation (Tarafder and Classen 1988; Helal 1990; Firsching and Classen 1996; Oehl et al. 2001), sterile conditions (Rose 1912; Pierre and Parker 1927; Rogers et al. 1941; McLachlan 1980; Tarafder and Classen 1988; Helal 1990; Bishop et al. 1994; Gilbert et al. 1999; Oehl et al. 2001; Bunemann et al. 2007), time invested in developing test plants (Tarafder and Classen 1988; Helal 1990), requirement of commercially available phosphatases (Bishop et al. 1994) and an inability to determine true phosphatase activity due to the fate of the inorganic P once mineralised (Tarafder and Classen 1988; Helal 1990).

Given the methodological limitations noted above, a simple and rapid method to assess the activity of phosphatase of field-grown roots and their ability to mineralise soil organic P is needed. This would provide a way to assess and classify organic P in terms of its bioavailability under various soil conditions and the characteristics of plant species or groups of species. Investigating greenhouse and field-grown roots implies that one is investigating the root and the biofilm of microorganisms associated with the root (Richardson and Simpson 2011). Throughout the present paper, that association should be assumed when the term 'root' is used.

The objective of this study was to develop a simple, rapid method as a step towards the goal indicated above. This study documents a phosphatase evaluation method using greenhouse-and field-grown excised roots as the source of native phosphatases to access their ability to mineralise various sources of organic P by following the fate of inorganic P generated by mineralisation. While fluorometric evaluation of the activity of phosphatase (Sinsabaugh et al. 2003) is a useful method, this research approached the problem of using the direct measurement of inorganic P so it would be applicable to all laboratories with the most basic equipment.

Materials and methods

The method was developed using several preliminary experiments identifying incubation time, inorganic P fate and Qio values. The applicability of the method was defined by its subsequent use to determine root phosphatase activity as a function of plant variety, plant species, plant stress and organic P source. The initial experimental conditions are outlined in the following section. This basic framework was maintained for all experiments, with significant changes mentioned for each experiment where necessary.

Quantitative determinations of mineralised organic P were accomplished by analysing inorganic P produced in the system by the method of Murphy and Riley (1962) at wavelength 880 nm with a detection limit of 0.004 mg [L.sup.-1]. By focusing on the inorganic P product, studies are not limited to a specified set of organic P sources, thus diversifying the range of substrates for experimentation.

Experiments 1a and 1b. Effect of incubation time on P mineralisation

Bahiagrass (Paspalum notatum cv. Tifton 9) grown under greenhouse conditions was used as the model plant for method development. Grasses were grown at 25 [+ or -] 2[degrees]C and 80% relative humidity in 9-cm-diameter and 10-cm-deep pockets within an 18-pocket tray. The plants were brought from the greenhouse to the laboratory, where roots ~4 6 cm (3-6 mg dry weight) were excised using scissors, carefully rinsed with deionised water, placed in 8-mL glass test tubes containing the various treatment solutions and capped. The glass tubes contained 7 mL of treatment solution. Para-nitrophenyl phosphate (PNPP, catalogue No. 71768-5G; Sigma Chemical Co., St. Louis, MO, USA) was used as the model organic P compound, and treatment solutions were prepared in a matrix of 0.1 m sodium acetate buffered at pH 5.2.

Ten replicates were used for each of four treatments, which were: (1) +PNPP, +root; (2) -PNPP, +root; (3) +PNPP, -root; and (4) -PNPP, -root. Treatment 1, containing an excised root segment in 20 mg [L.sup.-1] of PNPP solution, determined the increase of inorganic P from PNPP mineralisation by the root, and any leakage from the root. Treatment 2, containing roots in buffered solution without PNPP, determined the inorganic and organic P leakage from roots. Treatment 3, containing buffered 20 mg [L.sup.-1] of PNPP solution, determined the degradation of PNPP without roots. Treatment 4, containing only the buffer solution, acted as a check or control for potential background inorganic P.

All test tubes were capped and incubated at 28[degrees]C in a low-temperature incubator (3524 Precision; Thermo Electron Corporation, Fisher Scientific, Waltham, MA, USA) for different lengths of time. For each time interval, 10 replicates were taken from the incubator and the roots removed to obtain dry weight (forced-air oven at 70[degrees]C for 24 h). The solutions were transferred to scintillation vials, stored in the refrigerator at 4[degrees]C and analysed for inorganic P within 30 h and for total P the following day. Solutions were analysed for inorganic P by the method of Murphy and Riley (1962). Total P was determined by digesting 5mL of a treatment solution with 100 [micro]L concentrated sulfuric acid and 200 mg of ammonium persulfate for 30 min in an autoclave at 121[degrees]C (15-20 psi). This is based on EPA method 365.1-12 (EPA 1993); however, the amount of persulfate was increased to the indicated level, based on initial studies that showed it was required for the complete digestion of PNPP. The digestates were analysed for inorganic P as mentioned above. Organic P values were calculated by the difference between total P and inorganic P. The total P analysis was used to confirm the organic P concentration of the prepared organic P substrates and to mass balance P at the end of the study as a check of experimental technique. Organic P mineralised by roots was expressed as mg organic P mineralised [g.sup.-1] dry weight of root. Sampling times for Expt 1a were 1,3,6, 12, 18and24h, while those for Expt 1b were 2, 4, 8, 10, and 12 h. Expts 1a and 1b were conducted separately.

Experiment 2. Evaluation of inorganic P uptake by excised roots during incubation

Since the method was based on measuring mineralised P, it was necessary to determine whether significant quantities of inorganic P were being absorbed by the excised roots during the incubation time, or whether net inorganic P leakage from excised roots dominated. To determine possible inorganic P uptake, an external inorganic P source was added. Treatment solutions of 3.9 mg [L.sup.-1] of inorganic P were prepared using ammonium dihydrogen phosphate (ADP) in 0.1M sodium acetate buffer at pH 5.2. As with the previous experiment, buffer solutions at pH 5.2 with no added inorganic P were used as controls. The treatments in this experiment were: (7) +ADP, +root; (2) -ADP, +root; (3) +ADP, -root; and (4) -ADP, -root. Treatment 1, containing excised root in 3.9 mg [L.sup.-1] of ADP inorganic P solution, determined net inorganic P uptake by roots. Treatment 2, containing roots in the blank buffer solution, determined inorganic and organic P leakage from roots. Treatment 3, containing buffered 3.9 mg [L.sup.-1] of inorganic P without roots, was used to track possible changes in the experimental matrix without roots. Treatment 4, containing only the buffer solution, acted as a control for potential background inorganic P. This experiment was run for 4h at 28[degrees]C then analysed for inorganic P and organic P as described above. The 4-h time-frame was based on results from Expts 1a and 1b.

Experiments 3a-3d. [Q.sub.10] values for phosphatase activity

The temperature dependence of root phosphatase activity was measured to help determine whether organic P mineralisation measured in our experiments was due to enzyme activity. This was accomplished via the interpretation of [Q.sub.10] values calculated from four separately conducted experiments. The experimental setup was the same as described above in Expt 1, except that roots were subjected to both, 28[degrees]C and 38[degrees]C. Each experiment represents a range of times and required different sets of roots from the same plants, as they were conducted over 60 days. The time-frames of the four experiments were: 1,4 and 6 h for Expt 3a; 1,2 and 4 h for Expt 3b; 1 and 4 h for Expt 3c; and 1,2,4 and 6h for Expt 3d. The [Q.sub.10], was calculated by the equation:

[Q.sub.10] = ([R.sub.2]/[R.sub.1])[conjunction](10/(38 - 28)) (1)

where [R.sub.2] and [R.sub.1] are the measured reaction rates at 38[degrees]C and 28[degrees]C, respectively.

Experiment 4. Phosphatase activity as a function of bahiagrass variety and stress

Based on the experimentation above, method development from Expts 1-3 is summarised in Table 1, and was applied to roots of bahiagrass cvv. Tifton 9 and Argentine, to determine the effect of variety and stress on bahiagrass phosphatase activity. The treatments used in this experiment were: (7) BArg; (2) BT9; (3) BT9 +chlorotic; (4) BT9 + chlorotic + stunt; and (5) BT9 + Brown. Treatment 1 contained roots of cv. Argentine grown in 9-cm-diameter and 10-cm-dcep pots (BArg). Treatment 2 contained roots of cv. Tifton 9 grown in 9-cm-diameter and 10-cm-deep pots (BT9). Treatment 3 contained roots from cv. Tifton 9 with chlorotic leaves grown in seedling flat cells 2.54 cm long, wide and deep, with one grass stalk per cell (BT9 + chlorotic). Treatment 4 contained roots from cv. Tifton 9 with chlorotic and stunted leaves grown in seedling flat cells 2.54 cm long, wide and deep, with four or five grass stalks per cell. Treatment 5 contained brown roots from treatment 4. Active white roots were used for all other treatments. Brown roots used in treatment 5 were older sections of regular roots.

Experiment 5. Phosphatase activity of roots as a function of organic P source and plant species

Ths e method described in Table 1, using a 4-h incubation time, was applied to the excised roots of field-grown plants exposed to different forms of organic P. A completely randomised 3x3 (species x organic P form) factorial design with six replicates was used. In addition, bahiagrass roots from greenhouse-grown plants were evaluated in the same manner with different forms of organic P. The experiment with grasses from the greenhouse was performed as described in Expt 1. Roots from field-grown plants were excised from three different crop species: bahiagrass (cv. Pensacola, in the bloom stage just before the seed maturity stage), peanut (Arachis hypogaea cv. Georgia 06G, in the early reproductive stage), and cotton (Gossypium hirsutum cvv. Delta and Pineland 1050, in the early reproductive stage). The three organic P sources were PNPP, glucose-1-phosphate (G1P) (Catalog No. 56401-20-8; Fisher Science, Hanover Park, IL, USA) and phytic acid (Catalog No. P0109-10G; Sigma Chemical Co.) at a concentration of 20 mg PI. 1. Roots from all three species were collected from the same soil, which was an Ultisol with kaolinitic clay (Orangeburg soil series) and a sandy clay-loam surface texture (Soil Survey Staff 1999). The average Mchlich 1 and Mehlich 3 extractable P contents of this soil were 39.4 and 60.2 mg [kg.sup.-1].

Root samples were collected from the field early in the morning when the soil was moist. First, the surrounding soil was loosened with a spade and the plant was carefully uprooted. The entire root system was separated from the shoots with pruning shears, deposited with adhering soil in ziploc bags, and placed on ice in a cooler. Samples were immediately taken to the laboratory where the roots were cleaned of soil, rinsed with deionised water, excised, and placed in the glass test tubes as described in Table 1. All field roots were brown in colour.

Calculation of organic P mineralisation

The information from the preceding experiments was used to calculate the amount of organic P mineralised [g.sup.-1] root dry weight. The equation used to determine organic P mineralisation for all experiments was:

Net P mineralised = [P.sub.i] - [P.sub.il] - [P.sub.ia] (2)

where [P.sub.i] is the increase in inorganic P in the (+organic P, +root) treatment, [P.sub.il], is the inorganic P leaked by excised roots in the (-organic P, +root) treatment, and [P.sub.ia] is the inorganic P obtained through abiotic mineralisation of organic P in the (+organic P, -root) treatment. The control had insignificant inorganic P in all experiments and did not need to be included in the equation.

Statistical analyses

Normality and goodness-of-fit tests were conducted on the data from each experiment. Where the data were not normally distributed, log-transformations were used to stabilise the variance and analyses were conducted on the transformed data. Log-transformed data were back-transformed for the Figures. Phosphatase activity was calculated from Eqn 2 for Expts 1, 4 and 5. Inorganic P uptake was calculated for Expt 2, and Eqn 1 was used to calculate Q10 values for Expt 3. Treatment effects for each experiment were determined by analysis of variance (ANOVA) based on a randomised, replicate design using the General Linear Model routine in Statistica (StatSoft Inc. 2011). Treatment main effects or interactions were further evaluated using a post-hoc mean comparison employing a Duncan's test with a significance level of P=0.05. Where appropriate, least-square log means from the statistical analyses were back-transformed using the equation:

z = exp(y x 2.302 + 0.5[[sigma].sup.2] x 5.301)

where z is the back-transformed mean from the log mean, y is the log mean and [[sigma].sup.2] is the variance of the log mean.

Results

Experiments 1a and 1b. Effect of incubation time on P mineralisation

Phosphatase activity, determined as a measure of organic P mineralised [g.sup.1] root dry weight, showed two distinct mineralisation patterns (Fig. 1). Mineralisation over the course of both the 24 h and 12 h experiments followed a linear relationship (y = 6.19x + 0.62, [R.sup.2] = 0.95) up to and including the first 6h. At 6-8 h the mineralisation rate dropped by 44-72% then increased linearly with different slopes (p = 2.05x + 3.38, [R.sup.2] = 0.94; and y = 1.17x + 1.25, [R.sup.2] = 0.99) (Fig. 1). The greatest amount of PNPP mineralised before the shift was 35.8 mg [g.sup.-1] at 6 h and 31.1 mg [g.sup.-1] at 4 h for the 24-h and 12-h experiments, respectively.

Inorganic and organic P leakage from roots comprised 1-3% and 5-10% of mineralised organic P (Fig. 2). Organic P mineralisation in the absence of roots was 0.0005% of the total P mineralised, therefore an insignificant contributor to inorganic P in these studies.

Experiment 2. Evaluation of inorganic P uptake by excised roots during incubation

Underestimation of phosphatase activity through potential root uptake of inorganic P was tested over 4 h using a solution of 3.9 mg [L.sup.-1] of orthophosphate. The final P concentration of 4.1 mg [L.sup.-1] (Fig. 3) indicated no net uptake of inorganic P, but rather a net inorganic P exudation, consistent with the results of Expt 1.

Experiments 3a-3d. [Q.sub.10] values for phosphatase activity

The technique was further confirmed by determining [Q.sub.10] values of the root-microbe systems (Fig. 4). The calculated [Q.sub.10] values of phosphatase activity from our root-microbe system were used to verify that the mineralisation of organic P was indeed a function of root activity and not physical reactions, such as cell wall degradation, leaching of inorganic P from roots or abiotic organic P mineralisation. [Q.sub.10] values ranged from 1.52 to 1.95 with a mean 1.7 and a 95% confidence interval of 1.37-2.01.

Experiment 4. Phosphatase activity as a function of bahiagrass variety and plant stress

There was no significant difference in mean root phosphatase activities of bahiagrass cultivars (Argentine v. Tifton 9) (Fig. 5). Phosphatase activities of both cultivars were similar to that of chlorotic bahia-T9. Phosphorus mineralisation rates were highest with the bahiagrass that exhibited stunted growth, along with chlorotic leaves (12.5 mg [g.sup.-]), while brown bahiagrass roots demonstrated the least P mineralisation potential (2.1 mg [g.sup.-]).

Experiment 5. Phosphatase activity of roots as a function of organic P source and plant species

Three different sources of organic P (PNPP, G1P, phytate) were used to evaluate the phosphatase activity of bahiagrass roots grown under greenhouse conditions (Fig. 6) and field-grown roots of three crop species (Fig. 7). The organic P mineralised by roots from bahiagrass grown in a greenhouse was 6.2, 1.5 and 0.5mg [g.sup.-1] for PNPP, G1P and phytate, respectively. The organic P mineralised using field-grown roots followed the same trend as the greenhouse-grown bahiagrass (i.e. PNPP > G1P > phytate). It is important to note that while roots collected from the greenhouse were actively growing white roots, those collected from the field for all three species were entirely brown. The phosphatase mineralisation values of field-grown brown roots of bahiagrass were consistent with the greenhouse brown roots of bahiagrass in Expt 4. With respect to plant species, excised roots of peanut mineralised significantly more organic P than did bahiagrass roots, and both species mineralised more organic P than cotton roots, regardless of the organic P source.

Discussion

Effect of incubation time on P mineralisation

The potential organic P mineralisation rate was linear through 6h of incubation. Beyond 6h the mineralisation potential dropped significantly in both independent experiments. The reason for this drop in mineralisation potential is not clear but it may be due to [O.sub.2] availability. The focus of this study was to determine a suitable time-frame for measuring phosphatase activity, and we concluded that an appropriate incubation time for this method was 1-4 h. Therefore, an incubation time of up to 4 h was chosen for subsequent experiments to ensure adequate inorganic P concentrations for calculating phosphatase activity with precision and accuracy.

Determination of net inorganic and organic P contributions from excised root leakage, necessary to avoid possible underestimation or overestimation of P mineralisation (Fig. 2), were incorporated into the experimental protocol. This approach was generally lacking in other studies using excised roots (MacFall et al. 1991; Antibus et al. 1997), and it was shown by these data to be a necessary component. Considering the rate of exudation of both organic and inorganic P from roots, the 1-4 h period appeared adequate to account for these influences.

Evaluation of inorganic P uptake by excised roots during incubation

Determination of inorganic P uptake by excised roots is important to ensure that a measurable amount of P mineralised by root phosphatases is not absorbed by the roots, which can lead to an underestimation of mineralisation. Regardless of the method used to determine organic P mineralisation, the fate of inorganic P is always uncertain unless it is specifically tested. This step was lacking in previous root studies. There was no measureable net uptake of inorganic P by excised bahiagrass roots. However, there was an increase in inorganic P concentrations (0.2 mg [L.sup.-1]), indicating a net leaching of inorganic P by roots. The findings of this experiment were consistent with results of Expt 1, which showed the same relative amount of inorganic P leaching. The lack of measureable inorganic P uptake by incubated, excised roots eliminated the need to consider P absorption within the 4-h incubation time in the subsequent experiments.

[Q.sub.10] values for phosphatase activity

Chemical reactions have [Q.sub.10] values between 2 and 3, in contrast to physical processes with [Q.sub.10] values only slightly greater than 1 (Salisbury and Ross 1969; Graham and Patterson 1982). The Ql0 values presented here are supporting evidence that the reactions were associated with temperature-dependent biochemical processes, rather than physical processes. Abiotically driven P mineralisation was insignificant and therefore not likely to require monitoring in future use of this excised root method under conditions similar to that used in these experiments.

Phosphatase activity as a function of bahiagrass variety and plant stress

Although there was no statistical difference between bahiagrass cultivars, there were significant differences based upon root morphology and plant stress. Brown roots showed significantly lower phosphatase activity than white roots (which represented the other treatments), while roots from chlorotic-stunted Tifton 9 bahiagrass showed the greatest phosphatase activity among all the treatments. Phosphatase activity has been regarded as a soil quality indicator (Amador et al. 1997), and abiotic stresses such as drought (Sharma el al. 2004; Sun et al. 2010), salinity (Sharma et al. 2004; Bybordi and Ebrahimian 2011) and nutrient deficiency (McLachlan 1980) have increased the phosphatase activity. Restricted root growth has been shown to reduce plant photosynthesis (Poorter et al. 2012), and reduced photosynthetic conditions have been shown to increase phosphatase activity (Espeland and Wetzel 2001). A heightened nutrient-acquiring mechanism is the probable cause of high phosphatase activity of the chloroticstunted bahiagrass. These results show that the excised root method can respond to differences in root physiology and management conditions.

Phosphatase activity of roots as a function of organic P source and plant species

Para-nitrophenylphosphate was the organic P compound most susceptible to mineralisation. Studies have shown higher mineralisability of PNPP than of most other organic P compounds (Juma and Tabatabai 1988). Although not naturally occurring, easy spectrometric quantification of generated PNP (para-nitrophenol) has made PNPP the organic P form of choice in assessing phosphatase activity of roots. However, in order to eventually investigate mineralisability of soil organic P, PNPP cannot be used as an indicator since it does not naturally occur in soils. Nonetheless, the technique of tracking inorganic P is possible regardless of the organic P in question and can be accomplished in the most basic laboratory conditions.

Phosphate diesters (mainly nucleic acids and phospholipids) are considered the most readily mineralisable forms of organic P in soil, followed by mono-esters such as inositol phosphate (Bowman and Cole 1978; Feng et al. 2003). This explains why phytate tends to accumulate in soils relative to the diesters (Feng et al. 2003). Our data demonstrated that phytic acid mineralised to a much lower degree than the other organic P compounds. However, Feng et al. (2003) also noted that when phytate was provided as the source of P in amounts similar to soluble inorganic P, phytate provided a similar contribution to net plant P uptake. Another possible reason for low general mineralisability of organic P by field roots could be that these roots were collected during seed-set, when the internal processes of annual species are oriented towards seed production rather than acquiring nutrients. If this is so, then this indicates another venue where this method can be used to indicate plant condition.

Phosphatase activity of excised root showed significant differences between species in the order peanut > bahiagrass > cotton. To our knowledge, rhizosphere properties of bahiagrass have not previously been investigated; therefore, these data represent the first evaluation of bahiagrass phosphatase mineralisation potential. Cotton reportedly has limited root phosphatase activity and appears to lack the ability to mobilise non-labile organic P sources (Wang et al. 2008). Peanut, on the other hand, is known for its ability to efficiently utilise organic P, but there is uncertainty over the mechanisms (Shibata and Yano 2003). Our experiments suggest that root phosphatases are a contributing factor.

Bahiagrass field roots were brown, and the overall phosphatase activity values were lower than measured with white roots from greenhouse-grown bahiagrass. Interestingly, the values for field roots were very similar to values recorded from greenhouse-derived brown roots, further supporting the method's sensitivity to root physiological and/or morphological differences. New roots are white in colour but change to light brown within weeks or months, becoming darker with age (Comas et al. 2000; Wells and Eissenstat 2001). Several physiological changes have been identified during the transition of white roots to brown roots, including death of epidermal and cortex cells, resulting in limited permeability to water and ions. Declines in nutrient uptake (Comas et al. 2000; Bouma et al. 2001; Wells and Eissenstat 2002), respiration rate and dehydrogenase activity (Comas et al. 2000; Voider et al. 2005) have been associated with root browning. Our experiments showed a significant decrease in phosphatase activity with root browning.

Conclusions

The excised root technique developed for determining phosphatase activity is quick, effective, accurate, reproducible and cost-effective. It was linear over a period of 6h, allowing for quantitative measurement of root phosphatase activity. This method does not require expensive laboratory equipment, radioactive isotopes, sterile conditions or long incubation periods, and it is not restricted to PNPP as the sole organic-P source for testing. This technique is sensitive enough to respond to different plant species, root physiological conditions, and management practices. Important considerations in the use of this method are inorganic and organic P leaching from excised roots. The ability to detect potential uptake of mineralised P into the roots results in more accurate phosphatase activity values. The potential ability to determine utilisation of organic P can contribute to better nutrient management. It is important to remember that this method determines potential root phosphatase activity; organic P mineralisability in soil is constrained by several factors apart from phosphatase availability, such as soil moisture, soil tortuosity, availability of organic P sources. Potential phosphatase activity can be used to evaluate a cropping system's potential to utilise organic P. We intend to use this method in future studies to define bioavailable soil organic P in a range of environmental matrices.

http://dx.doi.org/10.1071/SR13198

Acknowledgements

We thank Dr S. George and Dr C. M. Bliss for their support. We also acknowledge Kelly O'Brian and Ron Bolton for their assistance in the greenhouse and field.

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Jaya Das (A,B,C), Nicholas Comerford (A), David Wright (A), Jim Marois (A), and Cheryl Mackowiak (A)

(A) North Florida Research and Education Center, University of Florida, 155 Research Road, Quincy, FL 32351-5677, USA.

(B) Present address: Wright State University, 268 Brehm Lab, 3640 Colonel Glenn Highway, Dayton, OH 45435, USA.

(C) Corresponding author. Email: das.29@wright.edu

Table 1. Proposed excised root method for determining potential
phosphatase activity

Step 1.   For each replication use treatments of (1) +roots,
            +organic P; (2) +roots, -organic P; (3) -roots, -organic
            P. The treatment (-roots, +organic P) is not used because
            results showed that organic P was not significantly
            abiotically mineralised
Step 2.   Excise roots 4-6 cm in length
Step 3.   Immediately place roots in test tubes containing 7 mL of
            solution and stopper
Step 4.   Place test tube in incubator for 1-4h (depending on
            investigator's objective and precision to measure
            inorganic P)
Step 5.   After incubation, remove roots from test tube and measure
            inorganic P in solution
Step 6.   Dry roots to a constant weight and express P mineralised
            on root dry-weight basis. Could also be done on a root
            length and surface-area basis with proper measurements
            of the roots
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Author:Das, Jaya; Comerford, Nicholas; Wright, David; Marois, Jim; Mackowiak, Cheryl
Publication:Soil Research
Article Type:Report
Date:Mar 1, 2014
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