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Enzymatic Hydrolysis of Organic Phosphates Adsorbed on Mineral Surfaces Rickard Olsson,† Reiner Giesler,‡ John S. Loring,§ and Per Persson*,† †
Department of Chemistry, Umea University, SE-901 87 Umea, Sweden ‡ Climate Impacts Research Centre, Department of Ecology and Environmental Science, Umea University, SE-981 07 Abisko, Sweden § Pacific Northwest National Laboratory, Richland, Washington 99352, United States
bS Supporting Information ABSTRACT: Esters of phosphoric acid constitute a sizable fraction of the total phosphorus supply in the environment and thus play an important role in the global phosphorus cycle. Enzymatic hydrolysis of these esters to produce orthophosphate is often a required reaction preceding phosphorus uptake by plants and microorganisms. Generally, adsorption to environmental particles is assumed to limit this process. Here we show, however, that the rate of enzymatic hydrolysis of glucose-1-phosphate (G1P) adsorbed on goethite by acid phosphatase (AcPase) can be of the same order of magnitude as in aqueous solution. The surface process releases carbon to the solution whereas orthophosphate remains adsorbed on goethite. This hydrolysis reaction is strictly an interfacial process governed by the properties of the interface. A high surface concentration of substrate mediates the formation of a catalytically active layer of AcPase, and although adsorption likely reduces the catalytic efficiency of the enzyme, this reduction is almost balanced by the fact that enzyme and substrate are concentrated at the mineral surfaces. Our results suggest that mineral surfaces with appropriate surface properties can be very effective in concentrating substrates and enzymes thereby creating microchemical environments of high enzymatic activity. Hence, also strongly adsorbed molecules in soils and aquatic environments may be subjected to biodegradation by extracellular enzymes.
’ INTRODUCTION Phosphorus is one of the major nutrients needed to sustain life and is critical for food production. The demand is expected to increase over the next 50 years whereas the supply from workable phosphate rock mines is decreasing, resulting in increasing value.1 In this context, exploration of how soil phosphorus is utilized by plants or microorganisms and the mechanisms behind it is becoming increasingly important. Phosphorus occurs in nature almost exclusively as inorganic or organic phosphates.2,3 In the latter class the phosphate group is mainly incorporated into organic molecules via ester bonds. A unique property of phosphates compared to most other essential macronutrients is the unusually large reactivity toward solid particles in the environment, especially those containing Al, Mn, and Fe.2,4 6 Thus, reactions at environmental interfaces govern the fate of phosphorus and greatly influence biomass production in ecosystems.7 In this respect key processes are the transfer of phosphorus from adsorbed, solid, or organically bonded states into bioavailable forms, and these processes occur primarily at various interfaces. Processes leading to the mobilization of phosphorus may have adverse effects on for instance water quality. Organic phosphate is an umbrella term for a diverse class of phosphorus compounds including inositol phosphates, sugar phosphates, phospholipids, nucleic acids, and phosphoproteins.3,8 10 r 2011 American Chemical Society
In some environments these can constitute as much as 80% of the total phosphorus pool.11 A property in common to almost every organic phosphate is that hydrolysis of the ester bond is a necessary step for biouptake of phosphorus by plants and microorganisms.3,12 The hydrolysis is mediated by extracellular enzymes such as phosphatases and phytases13 15 but has also been shown to occur abiotically on certain mineral surfaces.16,17 Since both extracellular enzymes and organic phosphates are known to adsorb rapidly and strongly onto environmental particles, interfacial processes play a crucial role in the enzymatically driven reactions as well.18 21 A number of previous studies have shown that enzyme adsorption to soil constituents can have a large effect on their phosphate hydrolyzing activity, and in this respect pH and particle composition have been shown to be important parameters.14,15 In contrast, there are relatively few investigations that explore enzyme-catalyzed hydrolysis of adsorbed organic phosphates, which is a situation likely to occur in environmental systems.15 Mineral surfaces have been indicated to protect certain adsorbed Received: August 15, 2011 Accepted: November 21, 2011 Revised: November 15, 2011 Published: November 21, 2011 285
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phosphate esters from enzymatic hydrolysis,15 but it is still an open question whether this is a general finding. Moreover, the mechanism of this process is largely unknown in the case where the enzyme is able to access adsorbed substrates. A central question is whether the enzyme is acting only on the soluble fraction that is replenished through substrate desorption or if the hydrolysis reaction occurs at the interface between the aqueous solution and the solid particles. In other words, is the overall rate determined by substrate desorption kinetics and solution diffusion or does enzyme adsorption also play a role in the reaction mechanism? In the present work we address these questions by studying a system consisting of an acid phosphatase (AcPase), glucose1-phosphate (G1P), and goethite (α-FeOOH) particles. The main objectives are to investigate if, how, and under what conditions a strongly adsorbed organic phosphate may be subjected to enzymatic hydrolysis. The enzyme is a broad-spectrum phosphatase similar to what has been used in previous studies,22,23 and the substrate is a representative sugar monoester that has been detected in soil samples at substantial concentrations.24,25 The mineral was chosen, as it is a common and well-characterized soil component with a high affinity for both inorganic and organic phosphates. 4,20,21 We combine wet-chemical analysis with a simultaneous infrared and potentiometric titration (SIPT) method that allows us to follow the interfacial reactions under in situ conditions. This approach provides molecular-level spectroscopic information in real-time of enzymatic hydrolysis of phosphate ester bonds at the water mineral interface.
diluted, and NaCl was added to give a Na(Cl) concentration of 50 mM. The goethite used in the present study was from the same batch as the one in our previous study on G1P goethite interactions.28 Enzymatic Hydrolysis of G1P in the Absence of Goethite. Volumes of a G1P stock solution were added to 40.0 mL of 50 mM NaCl, giving G1P concentrations of 1.2 mM and 0.6 mM. These G1P concentrations corresponded to the G1P amounts added in the simultaneous infrared and potentiometric titrations described below. The pH was adjusted to 5.0 and was kept constant with an automated and computer-controlled buret system. A sample was collected, and AcPase corresponding to 0.06 units/mL was added. Samples were then regularly collected to monitor the glucose level as a function of time. All samples were filtered through a 0.2 μm filter, and 1.0 mM NaOH was added to inhibit the enzyme. The glucose amounts in the samples were immediately determined by ion chromatography using a Dionex system (ICS-3000) with a CarboPac PA 20 guard column followed by a CarboPac PA 20 column, and a 20 mM NaOH eluent. Infrared Spectroscopy. The infrared spectra were recorded in attenuated total reflectance (ATR) mode using an experimental setup for a simultaneous infrared and potentiometric titration (SIPT) that has been described in detail by Loring et al.;29 a schematic description is provided in Supporting Information Figure S1. A goethite suspension was pumped peristaltically in a closed loop through fluoroelastomer (Chemsure Gore Industries) and PTFE tubing from a thermostatted (25 ( 0.05 °C) titration vessel to a flow-through ATR cell. The flow-through attachment was custom built of inert materials (e.g., Pyrex glass, PEEK, PTFE) and mounted on a single-reflection ZnSe 45° ATR accessory (FastIR, Harrick Scientific). A goethite overlayer was deposited onto the ZnSe crystal by evaporating 0.7 mL of a mineral suspension (ca. 2.5 g/L) onto the crystal at 75 °C for 2.5 h under N2-atmosphere. The ATR cell was placed inside an evacuated (5 mbar) infrared spectrometer (Bruker IFS66 v/s in a thermostatted room 25 ( 0.15 °C) equipped with a deuterated triglycine sulfate (DTGS) detector and a water-cooled Globar source. A volume of 80.0 mL of a 12 g/L goethite suspension was pipetted into the titration vessel and pumped over the overlayer. The final goethite concentration, after addition of G1P and enzyme, and the final pH adjustment, was 10 g/L. The pH was first adjusted to 9.4 and kept constant with an automated and computer-controlled buret system. A background spectrum of 4096 scans was collected of the overlayer and the goethite suspension, at a resolution of 4 cm 1. All spectra presented were ratioed against this background spectrum, thus removing contributions from the spectrometer and the ATR cell as well as the ionic medium and the pure goethite film at pH 9.4. No additional subtraction was performed to isolate the G1P bands. To isolate the protein amide bands as a result of possible enzyme adsorption, the spectrum at t0 (i.e., the spectrum at AcPase addition) was subtracted from the subsequent spectra using a subtraction factor of 1. A volume of G1P solution was then added to reach a total concentration of 1.36 or 0.69 μmol G1P/m2 of goethite. The pH was adjusted to 4.5, 5.0, or 5.5, and this was done comparatively fast to avoid base-catalyzed hydrolysis of G1P; the target pH values were reached in approximately 5 min. Sample absorbance spectra (512 scans) were collected as a function of time to follow the adsorption of G1P, and the adsorption reaction was assumed to be equilibrated after approximately 2.5 h. The change in peak
’ EXPERIMENTAL SECTION Chemicals, Solutions, and Suspensions. Deionized (MilliQ Plus) and boiled water was used to prepare all solutions and goethite suspensions. NaCl (Merck, p.a.) dried at 180 °C was used to prepare a background electrolyte concentration of 50 mM Na(Cl). pH adjustments were made with NaOH (50 mM) and HCl (50 mM). A 25 mM G1P stock solution was prepared by dissolving a weighed amount of G7000 (Sigma-Aldrich, 98%), and the concentration was verified by an acid base titration using standardized HCl and NaOH. The G1P and the 50 mM Na(Cl) solutions were sterilized by filtration through a 0.2 μm Sarstedt filter (Filtropur S 0.2). This procedure together with the fact that experiments were only analyzed during the first 10 h minimized effects from possible bacterial contamination. The G1P solution was kept frozen until use. An enzyme solution was prepared by dissolving a weighed amount of phosphatase, acid from potato, P1146 (Sigma). The isoelectric point of the enzyme was ca. 4.6 as determined by a Malvern Zetasizer Nano ZS, and a 6 mg/mL solution was indicated to contain both monomers and oligomers. The enzymatic activity was determined according to an assay based on Bergmeyer et al.,26 in which 1 unit of the enzyme hydrolyzes 1.0 mmol of p-nitrophenyl phosphate per minute at pH 4.8 at 37 °C. The synthesis and characterization of goethite (α-FeOOH) have been described previously.27 Briefly, goethite was prepared in polyethylene bottles by adding 1.8 L of 2.5 M KOH (EKA, p.a.) to 10 L of 0.15 M Fe(NO3)3 (Merck, p.a.) at a rate of 10 mL/min. The precipitates were aged for 96 h at 60 °C and dialyzed for 2 weeks. The resulting particles were identified to be goethite by X-ray powder diffraction, and the specific surface area was determined to 86.9 m2/g using N2 BET analysis. The pH of point of zero charge of the goethite particles was 9.4. The suspension was 286
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intensities in consecutive spectra, collected 8 min apart, was then 0.5% or less of the total peak intensities. After equilibration, AcPase corresponding to 0.06 units/mL was added. Again, spectra (512 scans) were recorded as a function of time. To quantitatively follow the hydrolytic process, samples of the goethite suspension were collected. First, the samples were filtered through a 0.2 μm filter, and then 1.0 mM NaOH was added to inhibit the enzyme. Finally, the glucose amounts in the samples were immediately determined by ion chromatography as described above. The IR spectrum of AcPase adsorbed on goethite in the absence of G1P at pH 5.0 was collected with the same ATR FTIR cell as used for the SIPT measurements described above. The sample was prepared in batch mode at the same enzyme and goethite concentrations as in the SIPT experiments. Prior to analysis, the sample was centrifuged and the wet paste was applied directly onto the ATR crystal. Also the spectrum of the supernatant was recorded, and this was subtracted from the paste spectrum to isolate the spectral features of AcPase adsorbed on goethite. To collect spectra of aqueous AcPase, 17.4 mg of the enzyme was dissolved in 5.00 mL of 50 mM NaCl. The pH of the AcPase solution was adjusted to 5.0. Background spectra of the empty ATR cell were recorded, before spectra of the AcPase solution and a 50 mM NaCl solution at pH 5.0 were collected. Spectra were an average of 512 scans collected at a resolution of 4 cm 1. To isolate the spectrum of the AcPase, the spectrum of the 50 mM NaCl solution was subtracted from that of the AcPase solution.
’ RESULTS AND DISCUSSION Enzymatic Hydrolysis of G1P in Solution and on α-FeOOH Surfaces. Macroscopic Results. The enzymatic hydrolysis ex-
periments were designed to facilitate direct comparison between hydrolysis of dissolved G1P and G1P goethite surface complexes. Thus, the reaction was studied in the absence and presence of goethite as a function of total concentration of G1P and time at a fixed AcPase concentration. The G1P concentrations were chosen so that in the goethite experiments >99.7% of the ligand was adsorbed (corresponds to a maximum G1P concentration of 4.1 μM in solution). Furthermore, the experiments were carried out in absence of buffers typically used in enzymatic studies, and instead pH was controlled via an automatic buret titrator. Thus, potential buffer surface interactions that can have a substantial effect on the interfacial processes were excluded.30 The broad-spectrum AcPase studied herein hydrolyzed G1P in aqueous solution (Figure 1A). Under the current experimental conditions 25% to 35% of the total G1P concentration was converted into glucose and orthophosphate within 10 h of reaction time. At both G1P concentrations, the reaction approximately followed first-order kinetics and the rate constants obtained were similar (Supporting Information Figure S2). The rate of hydrolysis of the G1P goethite complexes was strongly dependent on the total G1P concentrations and thus the surface coverage (Figure 1B and 1C); note that at enzyme addition, essentially all G1P was adsorbed to the goethite particles and that the hydrolysis in absence of enzyme under these conditions was negligible.28 At high surface coverage (1.36 μmol/m2), the rate of release of glucose into solution was on the same order of magnitude as the hydrolysis in aqueous solution; at pH 5, the first-order rate constant decreases from 4.6 10 4 min 1 in solution to 3.2 10 4 min 1 at the interface (Supporting Information Figures S2 and S3).
Figure 1. Enzyme-catalyzed release of glucose from glucose-1-phosphate in aqueous solution (A) and adsorbed on goethite (B and C) as a function of time. C is a blow-up of the results obtained at total G1P concentrations of 0.64 and 0.92 mM. In the goethite experiments, the G1P concentrations of 0.64, 0.92, and 1.23 mM correspond to 0.69, 1.00, and 1.36 μmol/m2, respectively. The experiments were conducted in 50 mM NaCl at 25 ( 0.05 °C.
The slower rate of hydrolysis at pH 4.5 and 5.5 as compared to pH 5 (Figure 1B and Figure S3) indicates that the catalytic optimum at the interface is close to the reported optimum in solution of pH 4.8 (See Experimental Section). At low surface concentration (0.69 μmol/m2), the extent of hydrolysis was barely detectable and the first-order rate constant decreased by more than 1 order of magnitude to 1.3 10 5 min 1 (Supporting Information Figure S4). Although desorption of G1P from goethite has been shown to be slightly slower at the lower surface concentration, it cannot explain this large difference in the enzymatic hydrolysis of adsorbed G1P.28 It is therefore unlikely that AcPase hydrolyzes the soluble fraction of G1P in bulk solution only. Instead the results point toward a process where the mineral surface concentrates both the enzyme and substrate at the interface, and it is differences in interfacial properties that cause the markedly different hydrolysis rates. The occurrence of an interfacial enzymatic hydrolysis reaction was further corroborated by the rapid decrease in enzymatic activity of the supernatant, implying fast enzyme adsorption (see text in Supporting Information). Irrespective of G1P surface 287
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Figure 2. Infrared spectra collected as a function of time in the amide and G1P regions from the SIPT experiments of the reaction between AcPase and G1P goethite complexes at pH 5 and at 0.69 μmol G1P per m2 of goethite (A, B), 1.00 μmol G1P per m2 of goethite (C, D), 1.36 μmol G1P per m2 of goethite (E, F). All spectra in the amide region are plotted on the same scale and so are the G1P spectra. Note however the different ordinate scales of the two regions. Each data set was collected over a 10 h period containing approximately 75 spectra. The arrows indicate intensity changes as a function of time.
coverage, after 1 h of reaction time, the remaining enzymatic activity in the supernatant was below the detection limit of the method (i.e., below 1.3% of the initial activity). Enzymatic Hydrolysis of G1P on α-FeOOH Surfaces. Infrared Spectroscopic Results. The IR spectroscopic experiments were well suited to study the current system, as the main IR bands of G1P (900 1250 cm 1) are separated from the strong amide I and II bands (1500 1700 cm 1) characteristic of proteins.28,31 Hence, changes in adsorbed G1P and interactions between the enzyme and the goethite particles can be monitored simultaneously and in real-time with the SIPT method. In agreement with the quantitative hydrolysis results (Figure 1), the IR spectroscopic trends were dependent on the total concentrations of G1P and showed pronounced differences in both the G1P and amide regions (Figure 2). The results obtained at the highest G1P concentration, 1.36 μmol/m 2 , showed a pronounced decrease of the G1P band at 1140 cm 1 as a function time (Figure 2F), which is a direct consequence of hydrolysis of adsorbed G1P. This was accompanied by an equally pronounced increase of the amide bands at 1640 and 1547 cm 1, respectively (Figure 2E). The ratio between the 1140 cm 1 intensities after a reaction time t and t0 (i.e., at enzyme addition) provides a spectroscopic estimate of G1P hydrolysis, and this ratio was in good agreement with the glucose release determined from solution analysis (Figure 3).
Figure 3. The fraction of hydrolyzed G1P goethite complexes at pH 5 and 1.36 μmol G1P per m2 of goethite calculated from the measured glucose concentrations in solution (red squares), and from the main IR band at 1140 cm 1 of adsorbed G1P (blue diamonds). The amide band area is the integrated area between 1450 and 1700 cm 1 (green triangles).
The rate of hydrolysis of the G1P goethite complexes at 1.36 μmol/m2 can be compared to the enzyme adsorption observed by IR spectroscopy using the integrated amide IR bands as a measure. The curves in Figure 3 show that the accumulation of enzymes at the interface was initially faster than the rate of hydrolysis. This result is in agreement with two-dimensional correlation analysis of the spectroscopic data which shows that adsorption of AcPase precedes degradation of the G1P goethite complexes (supplementary text and Figure S5 in Supporting Information). A similar trend was also observed at 1.00 μmol/m2 288
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Figure 4. Normalized infrared spectra in the amide region collected at pH 5 and 1.00 μmol G1P per m2 of goethite (left) and 1.36 μmol G1P per m2 (right). For comparison, infrared spectra of AcPase adsorbed to goethite (top) and an aqueous solution of AcPase (middle) have been included in the left panel. The spectrum of the aqueous solution of AcPase is also included in the right panel (red spectrum drawn with thicker line width). The arrows indicate directions of shifts, and the double-headed arrow indicates band broadening as a function of time.
although at much lower rates (Supporting Information Figure S6). Hence, the build-up of a catalytically active surface layer of AcPase is necessary for hydrolysis of the G1P goethite complexes. At total concentrations of 0.69 and 1.00 μmol/m2, only small changes in the G1P intensities were observed (Figure 2B and 2D). The growth of the G1P bands at the lowest concentration showed that the system was not in complete equilibrium before enzyme addition although bulk solution measurement indicated complete adsorption.28 The slow reaction time is explained by the experimental procedure leading to an initial uneven surface distribution of G1P between the goethite particles in the system, which is followed by a slow redistribution process (for further details, see Figure S1). At 1.00 μmol/m2 the redistribution is almost balanced by the small extent of hydrolysis (Figure 1C) as indicated by the practically constant G1P band intensities (Figure 2C). The macroscopic solution experiments discussed above indicated a rapid enzyme adsorption irrespective of G1P concentration but in contrast to the results at 1.36 μmol G1P/m2 no or only weak amide bands were detected at 0.69 and 1.00 μmol G1P/m2, respectively (Figure 2A,C,E). These seemingly conflicting results were explained by the presence of at least two fractions of enzymes displaying different adsorption modes. At low G1P concentrations, the enzyme adsorbed rapidly onto particles close to the point of addition in a nonactive form and did not distribute evenly over the particles in the system whereas at high G1P concentration AcPase was more labile, leading to an even distribution and appearance of amide bands in the IR spectra. The detection of the labile fraction coincided with the increase in hydrolysis of adsorbed G1P molecules (Figure 1B,C, and 2C,E), suggesting that this fraction was also catalytically active. The existence of different enzyme adsorption modes was further corroborated by analysis of the time-dependent features of the normalized amide bands (Figure 4). On pure goethite and initially at a G1P concentration of 1.00 μmol/m2, the amide I band was broadened and blue-shifted as compared to aqueous AcPase, indicating structural distortions of the enzyme. Also the amide II band was very broad whereas the shift was not pronounced. With increasing time the spectra shifted toward the band positions in the spectrum of the aqueous species, although the bandwidths of amide bands remained substantially greater (Figure 4). In contrast, at coverage 1.36 μmol G1P/m2, the initial spectra closely resembled that of aqueous AcPase, and maxima and widths of both amide bands were very similar to aqueous AcPase (Figure 4). However, increasing enzyme adsorption as a function of time caused significant band broadening, indicating surface-induced distortions. It is important to note that the changes of the amide bands
Figure 5. (a) Infrared difference spectrum calculated by subtracting the spectrum at t = 0 from that at t = 10 h (G1Ptot = 1.36 μmol/m2 and pH = 5). The subtraction factor was adjusted to minimize contribution from G1P remaining at the surface. (b) Infrared spectrum of orthophosphate adsorbed onto goethite at pH 5 and 0.5 μmol/m2.
at high and low G1P concentration are different, which implies that the nature of the enzyme distortions also is different. The analysis of the amide bands together with the previously presented data suggest that on pure goethite or at low G1P coverage the positively charged goethite surface provides sites for formation of enzyme-surface complexes that are characterized by structural changes of the three-dimensional AcPase structure as compared to the solution species. These species display fast adsorption and slow desorption rates. The complexes are possibly mediated by hydrogen bonding between the enzyme and surface functional groups. At high G1P coverage the enzyme adsorption mode changes. We ascribe this to blocking of the high-affinity surface sites by G1P, and to the glucose end of adsorbed G1P facing the solution, creating more organic-like surface properties presumably with increasing hydrophobic character and with different hydrogen bonding properties. This results in an even distribution of enzyme between the particles and enzyme surface complexes where the enzyme remains in structural conformations that are catalytically active. A uniform surface distribution also increases the likelihood of a close proximity between the enzyme and the substrate. This conceptual model involving at least two different fractions of surface enzyme nicely unify the quantitative results obtained from bulk solution with the infrared spectroscopic observations. It should be pointed out, however, that neither the model nor the actual experimental data contain information on where at the interface the actual enzymatic hydrolysis occurs. We believe though that it is likely that G1P desorbs from the surface site to interact with the catalytically active site of the interfacial enzyme. 289
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Environmental Science & Technology A final piece of information with respect to the complete reaction scheme was provided by analysis of the G1P bands at the beginning and end of the experiment. Figure 5a shows the spectral features of a surface species that increases in importance as the hydrolysis reaction progresses. This species is responsible for the observed intensity increase of IR bands in Figure 2F. The bands at 1042 and 1095 cm 1 were also detected in the twodimensional correlation analysis (Figure S5), and the signs of the synchronous and asynchronous signals indicate that these bands grow in with time but the growth is delayed as compared to the amide bands. Furthermore, the bands characterizing this new species are practically identical to those of orthophosphate adsorbed onto goethite. Accordingly, the interfacial hydrolysis of G1P separates the products into two fractions; glucose is released into solution whereas orthophosphate readsorbs to goethite. The conversion of adsorbed G1P into adsorbed orthophosphate will likely have an effect on the surface charge, which may influence the properties of the interfacial enzyme and could be a contributing factor to the changes in the amide bands observed at the G1P concentration of 1.36 μmol/m2 (Figure 4). The aggregative results have shown that the properties of the mineral surfaces and their coatings are essential determinants for the interfacial enzymatic activities. This strong dependence on interfacial properties offers an explanation to the reported variations in enzyme activities in the presence of mineral surfaces.14,15,22 Under optimal conditions, mineral surfaces can be very efficient in concentrating enzymes and substrates, thereby increasing the proximity between the reacting species. This will create local environments at interfaces displaying high enzymatic activity despite the fact that the concentrations of substrates and enzymes in bulk solutions are very low. Our results have bearing on the role of interactions at mineral surfaces with respect to stabilization of organic molecules in soils,32 and they offer a mechanistic explanation for how phosphorus and carbon can be mobilized via enzyme activity despite strong surface interactions. Our results may thus shed light on previous findings showing that microbial stimulation can mobilize old carbon or adsorbed organic P in mineral soils.33,34
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’ REFERENCES (1) Cordell, D.; Drangert, J.-O.; White, S. The story of phosphorus: Global food security and food for thought. Global Environ. Change 2009, 19 (2), 292–305. (2) Hinsinger, P. Bioavailability of soil inorganic P in the rhizosphere as affected by root-induced chemical changes: a review. Plant Soil 2001, 237, 173–195. (3) Paytan, A.; McLaughlin, K. The oceanic phosphorus cycle. Chem. Rev. 2007, 107 (2), 563–576. (4) Hiemstra, T.; VanRiemsdijk, W. H. A surface structural approach to ion adsorption: The charge distribution (CD) model. J. Colloid Interface Sci. 1996, 179 (2), 488–508. (5) Khare, N.; Hesterberg, D.; Martin, J. D. XANES investigation of phosphate sorption in single and binary systems of iron and aluminum oxide minerals. Environ. Sci. Technol. 2005, 39, 2152–2160. (6) Yao, W.; Millero, F. J. Adsorption of phosphate on manganese dioxide in seawater. Environ. Sci. Technol. 1996, 30, 536–541. (7) Vance, C. P.; Uhde-Stone, C.; Allan, D. L. Phosphorus acquisition and use: critical adaptations by plants for securing a nonrenewable resource. New Phytol. 2003, 157, 423–447. (8) Benitez-Nelson, C. R. The biogeochemical cycling of phosphorus in marine systems. Earth Sci. Rev. 2000, 51, 109–135. (9) Turner, B. L.; McKelvie, I. D.; Haygarth, P. M. Characterisation of water-extractable soil organic phosphorus by phosphatase hydrolysis. Soil Biol. Biochem. 2002, 34, 27–35. (10) Ingall, E. D.; Schroeder, P. A.; Berner, R. A. The nature of organic phosphorus in marine sediments: New insights from 31P NMR. Geochim. Cosmochim. Acta 1990, 54, 2617–2620. (11) Paul, E. A.; Clark, F. E. Soil Microbiology and Biochemistry, 2nd ed.; Academic Press: New York, 1996. (12) Raghothama, K. G.; Karthikeyan, A. S. Phosphate acquisition. Plant Soil 2005, 274 (1 2), 37–49. (13) Luo, H.; Benner, R.; Long, R. A.; Hu, J. Subcellular localization of marine bacterial alkaline phosphatases. Proc. Natl. Acad. Sci. U.S.A. 2009, 106 (50), 21219–23. (14) Leprince, F.; Quiquampoix, H. Extracellular enzyme activity in soil: effect of pH and ionic strength on the interaction with montmorillonite of two acid phosphatases secreted by the ectomycorrhizal fungus Hebetomu cylindrosporum. Eur. J. Soil Sci. 1996, 47, 511–522. (15) Giaveno, C.; Celi, L.; Richardson, A. E.; Simpson, R. J.; Barberis, E. Interaction of phytases with minerals and availability of substrate affect the hydrolysis of inositol phosphates. Soil Biol. Biochem. 2010, 42 (3), 491–498. (16) Baldwin, D. S.; Beattie, J. K.; Coleman, L. M.; Jones, D. R. Phosphate ester hydrolysis facilitated by mineral phases. Environ. Sci. Technol. 1995, 29 (6), 1706–1709. (17) Baldwin, D. S.; Beattie, A. K.; Coleman, L. M. Hydrolysis of an organophosphate ester by manganese dioxide. Environ. Sci. Technol. 2001, 35 (4), 713–716. (18) Norde, W. Adsorption of proteins from solution at the solidliquid interface. Adv. Colloid Interface Sci. 1986, 25 (4), 267–340. (19) Helassa, N.; Quiquampoix, H.; Noinville, S.; Szponarski, W.; Staunton, S. Adsorption and desorption of monomeric Bt (Bacillus thuringiensis) Cry1Aa toxin on montmorillonite and kaolinite. Soil Biol. Biochem. 2009, 41 (3), 498–504. (20) Celi, L.; Presta, M.; Ajmore-Marsan, F.; Barberis, E. Effects of pH and electrolytes on inositol hexaphosphate interaction with goethite. Soil Sci. Soc. Am. J. 2001, 65 (3), 753–760. (21) Ognalaga, M.; Frossard, E.; Thomas, F. Glucose-1-phosphate and myoinositol hexaphosphate adsorption mechanisms on goethite. Soil Sci. Soc. of Am. J. 1994, 58 (2), 332–337. (22) Rao, M. A.; Violante, A.; Gianfreda, L. Interaction of acid phosphatase with clays, organic molecules and organo-mineral complexes: kinetics and stability. Soil Biol. Biochem. 2000, 32 (7), 1007–1014. (23) Rosas, A.; Delaluzmora, M.; Jara, A.; Lopez, R.; Rao, M.; Gianfreda, L. Catalytic behaviour of acid phosphatase immobilized on
’ ASSOCIATED CONTENT
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Supporting Information. A schematic description of the setup (Figure S1), evaluation of rate constants (Figures S2 S4), description of determination of enzyme activity in supernatants (text), description of two-dimensional correlation analysis (text and Figure S5), and comparison between AcPase adsorption and rate of hydrolysis at 1.0 μmol G1P/m2 (Figure S6). This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*Tel: +46 90 786 5573. Fax: +46 90 786 7655. E-mail: per.persson@ chem.umu.se.
’ ACKNOWLEDGMENT Prof. Pernilla Wittung-Stafshede is gratefully acknowledged for helpful comments and suggestions. The Swedish Research Council and the Kempe Foundation funded this work. One of us (P.P.) acknowledges financial support from the Wenner-Gren Foundations and the Blaustein Visiting Professorship Fund of the School of Earth Sciences, Stanford University. 290
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