Wetland Soils Soil Phosphorus Forms along a Strong Nutrient … · 2012-06-18 · Soil Phosphorus Forms along a Strong Nutrient Gradient in a Tropical Ombrotrophic Wetland Wetland

Soil Science Society of America Journal

Soil Sci. Soc. Am. J. doi:10.2136/sssaj2011.0365 Received 28 Oct. 2011. *Corresponding author ([email protected]). © Soil Science Society of America, 5585 Guilford Rd., Madison WI 53711 USA All rights reserved. No part of this periodical may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Permission for printing and for reprinting the material contained herein has been obtained by the publisher.

Soil Phosphorus Forms along a Strong Nutrient Gradient in a Tropical Ombrotrophic Wetland

Wetland Soils

Phosphorus is a key element limiting ecosystem processes in freshwater wetlands (Daniel et al., 1998; Rejmánková, 2001). Because much of the P in wetland soils occurs in organic forms (Davelaar, 1993; Newman and

Robinson, 1999; Reddy et al., 2005), P availability is dependent on the cycling of P from biological material. Biologically derived P in soils, however, represents a va-riety of compounds that differ markedly in their behavior and bioavailability (Celi and Barberis, 2005; Condron et al., 2005). The nature of these P forms is the result of the dynamic interplay among biological inputs (Makarov et al., 2005), abiotic stabilization (Celi and Barberis, 2005), and biological modification (Cheesman et al., 2010b), all of which are influenced by edaphic, environmental, and biological factors. Information on the role these factors have in determining P composition in wetland soils is therefore critical for understanding the cycling and productivity of wetland systems, as well as in explaining future responses to perturbations.

Unlike quantitative extraction and determination of P pools based on chemi-cal stability, techniques such as solution 31P NMR spectroscopy allow the assess-ment of actual P forms present in environmental samples (Cade-Menun, 2005; McKelvie, 2005; Turner et al., 2005). This has yielded valuable insight into the na-

Alexander W. Cheesman*Wetland Biogeochemistry LaboratorySoil and Water Science Dep.Univ. of Florida106 Newell HallGainesville, FL 32611

and

Smithsonian Tropical Research InstituteApartado 0843-03092Balboa, Ancón, Republic of Panama

Benjamin L. TurnerSmithsonian Tropical Research Institute Apartado 0843-03092Balboa, AncónRepublic of Panama

K. Ramesh ReddyWetland Biogeochemistry Lab.Soil and Water Science Dep.Univ. of Florida106 Newell HallGainesville, FL 32611

Phosphorus cycling influences productivity and diversity in tropical wetlands, yet little is known about the forms of P found in the accreting organic matter of these ecosystems. We used alkaline (NaOH–ethylenediamine tetraacetic acid [EDTA]) extraction and solution 31P nuclear magnetic resonance (NMR) spectroscopy to characterize P in surface soils across a strong nutrient gradi-ent within a tropical ombrotrophic peat dome. From the interior bog plain to the marginal Raphia taedigera swamp, total soil P increased from 14.6 to 70.9 g m−3 and resin-extractable P from 0.1 to 30 mg kg−1. Phosphatase activity declined across the same transect (364–46 mmol methylumbellifer-one kg−1 min−1), indicating an increase in P availability toward the periph-ery of the wetland. Organic P identified by solution 31P NMR spectroscopy included phosphomonoesters (12–17%), phosphodiesters (10–14%), and phosphonates (up to 3.3% of total P). Inositol phosphates were not detected in these acidic peats. Inorganic P forms included orthophosphate (9–25% of total P), pyrophosphate (up to 3%), and long-chain polyphosphates; the latter occurred in concentrations (up to 24% of total soil P) considerably higher than previously found in wetland soils. The concentration of residual (unex-tractable) P was similar among sites (mean 280 mg kg−1), resulting in an increase in its proportion of the total soil P from 29% at the P-rich margins to 55% at the P-poor interior. This is the first information on the P composition of tropical wetland soils and provides a basis for further study of the cycling and contribution of P forms to the nutrition of plants and microorganisms.

Abbreviations: AEM, anion exchange membrane; EDTA, ethylenediamine tetraacetic acid; NMR, nuclear magnetic resonance.

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ture of P forms and their biogeochemical processing in wetlands. For example, previous work has highlighted the diverse range of biogenic P forms that can be present in wetland soils (Sundaresh-war et al., 2009), the importance of site conditions in determin-ing P composition (Cheesman et al., 2010b), and differences in P composition between wetland and terrestrial soils (Turner and Newman, 2005; Turner et al., 2006). Yet despite clear evidence that P availability influences both vegetation (Hagerthey et al., 2008; Sjögersten et al., 2011) and biogeochemical processes such as C fluxes (Wright et al., 2011) and N2 fixation (Šantrůčková et al., 2010), little is known about P forms and their associated cycling in tropical wetlands.

Tropical peat domes are “self-emergent” organic wetlands within the humid tropics (Semeniuk and Semeniuk, 1997). Their upper surface shows a pronounced convex morphology leading to (or resulting from) their hydrologic isolation and an ombrotrophic hydrologic regime (Anderson, 1983; Andriesse, 1988; Belyea and Baird, 2006). They are systems of environ-mental, social, cultural, and economic importance, providing numerous direct ecosystem services to local populations (Cen-tral American Commission for Environment and Development, 2002; Ellison, 2004) as well as representing substantial and dy-namic pools in the global C cycle (Maltby and Immirzi, 1993; Yu et al., 2010). Often associated with the swamps of maritime Southeast Asia (Anderson, 1983; Anderson and Muller, 1975), significant peat deposits also occur throughout the Caribbean coastal plain (Ellison, 2004; Phillips and Bustin, 1996). Al-though these remain poorly studied in comparison with their Asian counterparts, there are strong similarities, including a vis-ible soil–vegetation catena across the convex surface (Phillips et al., 1997) and distinct gradients in nutrient availability (Sjöger-sten et al., 2011; Troxler, 2007).

We used solution 31P NMR spectroscopy to assess the functional forms of P present within the surface soils of a tropi-cal ombrotrophic wetland. By using a natural soil P gradient and range of vegetation types while controlling for more general site conditions, we aimed to derive novel information on the influ-ence of nutrient status on the chemical nature of soil P found in tropical wetlands. Such data are critical to our understanding of P cycling in wetlands, particularly given the extent to which an-thropogenic P enrichment affects such ecosystems (Cheesman et al., 2010b; McDowell, 2009; Turner et al., 2006).

METHODsstudy site

The Changuinola peat deposit is an approximately 80-km2 portion of the internationally recognized San San Pond Sak wet-land in Bocas del Toro province, northwest Panama (Ramsar Site 611, www.ramsar.org). Palynological evidence (Cohen et al., 1989; Phillips and Bustin, 1996; Phillips et al., 1997) sug-gests that Caribbean coastal systems such as the Changuinola deposit developed by different mechanisms and contain distinct community types compared with the well-studied peat domes of Southeast Asia (Anderson and Muller, 1975). They retain a

visible soil–vegetation catena, however, with current communi-ties ranging from a monodominant Raphia taedigera Mart. palm swamp at the periphery, through mixed and monodominant Campnosperma panamense Standl. swamp forest, to a central “bog plain” community dominated by herbaceous species and stunted trees (Phillips et al., 1997; Sjögersten et al., 2011).

samplingWe established nine sampling sites at 300-m intervals along

a linear transect running perpendicularly from a bordering canal toward the geodesic center of the peat dome (Fig. 1; Table 1) (Co-hen et al., 1989; Sjögersten et al., 2011; Troxler, 2007). Sample Site 9 could not be located in the very center of the bog plain due to practical constraints but was considered to be just inside the central vegetation zone (Myrica–Cyrilla bog plain). In September 2007, three replicate peat samples were collected from within 20 m of each site using a sharpened metal cutting head on a ridged polycarbonate tube. Each replicate was an amalgamated sample of three surface cores (7.5-cm diameter, 10-cm depth) collected from within 2 m of each other. Samples were immediately cooled (~10°C) and returned to the laboratory, where they were stored at 4°C until processing within 72 h of collection. This processing involved homogenization and the removal by hand of recogniz-able roots (>1-mm diameter) and lignified structures (seeds, twigs, etc.). Due to the high concentration of fine roots from herbaceous vegetation at Sites 8 and 9, however, a substantial amount of root biomass may have remained within these samples. The samples were subsequently split, with half being air dried (~22°C for 10 d) to constant weight and the remainder stored at 4°C in sealed bags (subsequently referred to as fresh sample). Air-dried samples were ground in a ball mill using tungsten carbide vessels and stored in airtight containers under ambient laboratory conditions until analysis. Additional soil was collected from Site 9 in November 2007 for 31P NMR spectroscopy, for which the total P was not significantly different from the September sample at this site (t-test: P > 0.05).

soil PropertiesSoil moisture was determined by gravimetric loss after dry-

ing at 105°C for 24 h and used to estimate the soil bulk density. Soil pH was determined on fresh soil using a 1:20 soil (oven-dry weight)/water ratio and a glass electrode. Total elemental concen-trations were determined on dried and ground samples. Total soil C and N were determined by combustion and gas chromatogra-phy using a Flash EA1112 (Thermo Scientific), while total P was determined by H2O2 + H2SO4 digestion (Parkinson and Allen, 1975) and detection by inductively coupled plasma–optical emis-sion spectrometry (Optima 2100, PerkinElmer).

Phosphorus CharacterizationAnion Exchange Membranes

Readily extractable and microbial P were operationally deter-mined using anion exchange membranes (AEMs; BDH Prolabo 551642S, VWR International) using a method based on Kouno

6 Soil Science Society of America Journal

et al. (1995) and Myers et al. (2005), with modifications described in Cheesman et al. (2010c). The difference in phosphate between nonfumigated and hexanol-fumigated samples was attributed to microbial P, and we did not correct values for unrecovered biomass ( Jenkinson et al., 2004). Extracted P in nonfumigated samples was assumed to represent exchangeable inorganic phosphate, although

some organic and condensed inorganic P forms might also be in-cluded (Cheesman et al., 2010c).

solution Phosphorus-31 NMR spectroscopySoil P composition was determined by standard alka-

line extraction and solution 31P NMR spectroscopy (Cade-

Fig. 1. Overview of study transect and sampling sites within the Changuinola peat deposit, san san Pond sak, northwest Panama. Access route originates at a canal cut in 1908 and approximates previous leveling transects of the site (Cohen et al., 1989; Phillips and Bustin, 1996). Elevation increase from site 1 to site 9 is approximately 4 m.

Table 1. soil characteristics from sampling stations across the Changuinola ombrotrophic peat dome.

siteVegetation

plot† pH Bulk densityTotal elements

C N P

Mg m−3 ————————— g kg−1 ————————— mg kg−1

1 1 3.6 ± 0.21‡ 0.069 ± 0.006 ab§ 498 ± 9.9 a 29 ± 0.5 a 1028 ± 43 a

2 3.8 ± 0.12 0.064 ± 0.009 ab 508 ± 7.3 ab 29 ± 1.0 a 1014 ± 51 a

3 3.9 ± 0.05 0.060 ± 0.008 abc 515 ± 6.7 ab 28 ± 0.7 ab 956 ± 69 a

4 2 3.7 ± 0.03 0.078 ± 0.012 a 535 ± 9.7 ab 26 ± 1.3 abc 655 ± 81 bc

5 3.9 ± 0.08 0.064 ± 0.007 ab 506 ± 6.2 ab 28 ± 0.2 ab 710 ± 21 b

6 3 3.6 ± 0.29 0.056 ± 0.005 bc 507 ± 4.2 ab 25 ± 0.5 bcd 659 ± 31 bc

7 4 3.7 ± 0.19 0.040 ± 0.005 cd 458 ± 22.7 c 23 ± 1.9 cd 672 ± 81 b

8 3.8 ± 0.34 0.050 ± 0.004 bcd 500 ± 3.1 a 19 ± 1.9 e 388 ± 23 d

9 5 3.6 ± 0.27 0.033 ± 0.005 d 417 ± 5.3 d 22 ± 1.7 de 442 ± 58 cd

† Permanent vegetation sampling plot established by Sjögersten et al. (2011).‡ Mean ± one standard deviation (n = 3).§ Superscript letters denote homogenous subsets determined by post-hoc Tukey’s honestly significant difference test.


Menun and Preston, 1996; Turner et al., 2007). The extraction (0.25 mol L−1 NaOH and 50 mmol L−1 EDTA) was applied to air-dried soils with shaking for 4 h, as well as fumigated and nonfumigated fresh soils after application of the AEM strips (Cheesman et al., 2010a) to assess the potential contribution of live microbial cells to the P composition of the air-dried soils. All samples were extracted in a 1:30 soil (oven-dry weight)/solu-tion ratio at ambient room temperature (~22°C) before being centrifuged at 7000 rpm (maximum relative centrifugal force ~7500 ´ g) (Sorvall RC6, SLA 1500 Rotor, Thermo Fisher Scientific) for 20 min and the supernatant decanted. Field rep-licates were combined on an equal-volume basis (15 mL), mixed with 1 mL of an internal standard (methylenediphosphonic acid [MDP], 50 mg P L−1), frozen (−80°C), and lyophilized to await resuspension and NMR spectroscopy. A second, independent subsample was analyzed for total P by a double acid digest us-ing concentrated H2SO4 and HNO3 (Rowland and Haygarth, 1997), with P detection by automated molybdate colorimetry.

Spectra were acquired using a Bruker Avance 500 Console with a Magnex 11.75 T/51-mm magnet, using a 10-mm BBO probe. Lyophilized samples (~300 mg) were resuspended in 2.7 mL of resuspension fluid (1 mol L−1 NaOH and 0.1 mol L−1 EDTA) and 0.3 mL D2O in a 15-mL centrifuge tube. Samples were vortexed for 1 min, filtered through a prewashed 1-mm GF-B in-line syringe filter, and loaded into a 10-mm NMR tube (1008-UP-7 Norell). Spectra were acquired at a stabilized 25°C with a 30° tip angle (calibrated on samples spiked with addition-al orthophosphate), a zgig pulse program (Berger and Siegmar, 2004), 0.4-s acquisition time (100-ppm spectral width), and a 2-s pulse (d1) delay. Results presented here are of ~40,000 scans accumulated as four sequential experiments with free induction decay signals summed post-acquisition by Bruker proprietary software. Additional qualitative spectra from alkaline extrac-tions after pre-extraction with AEM strips were acquired using a 5-mm NMR probe and a Bruker Avance 500-MHz spectrometer using a 45° pulse, a 1-s d1 delay, and a zgig pulse program.

Spectra interpretation was performed using wxNUTS ver-sion 1.0.1 for Microsoft Windows (Acorn NMR Inc.). Spectra were referenced and integrated against the internal standard (MDP) set at d = 17.46 ppm after comparison to an externally held 85% H3PO4 set at d = 0 ppm. Spectra were integrated over set spectral windows corresponding to known P bonding classes (Turner et al., 2003b), with spectral deconvolution applied to the region between 8 and 3 ppm to separate orthophosphate from phosphomonoesters and −3 to −5 ppm to separate pyro-phosphate and polyphosphate end residues.

Hydrolytic Enzyme AssaysTo assess biological investment in P acquisition from organ-

ic compounds, the activities of two hydrolytic enzymes involved in the P cycle (phosphomonoesterase and phosphodiesterase) were determined using fluorogenic substrates based on a stan-dard microplate assay (Marx et al., 2001) as described in Turner and Romero (2010). For each sample, soil suspensions were

prepared in a 1:100 soil/water ratio (containing 1 mmol L−1 NaN3 to prevent microbial activity). The soil suspension (50 mL) was then pipetted into wells on a micro-well plate (eight wells per substrate) containing 100 mL of 200 mmol L−1 substrate [4-methylumbelliferyl phosphate or bis(4-methylumbelliferyl) phosphate] and 50 mL of 200 mmol L−1 NaOAc–HOAc buffer adjusted to pH 4.0 (the approximate mean pH of the peat). Plates were incubated for 30 min at 26°C to approximate the daytime soil temperature in the Changuinola peat deposit. The reaction was terminated by adding 50 mL of 0.5 mol L−1 NaOH (final solution pH >11) and the plates were read immediately on a FLUOstar Optima multidetection plate reader (BMG Labtech), with excitation at 360 nm and emission at 460 nm. Control plates containing substrate, buffer, and 1 mmol L−1 NaN3 (no soil suspension) were prepared and analyzed immediately before and after the analysis of the soil samples to account for initial fluorescence as well as pH-induced instability of the substrates. Each soil had corresponding blanks, methylumbelliferone (MU) standards, and soil-specific quench standards. All enzyme activi-ties are expressed as micromoles MU per kilogram of dry soil per minute (mmol MU kg−1 min−1).

Data AnalysisAll statistical tests were performed in SPSS for Win-

dows version 17.0.0 statistical software (SPSS Inc.). Data were checked for normality by application of a Shapiro–Wilk test and visual inspection. Natural logarithms were used for sta-tistical analysis if the assumption of normality was improved. Differences between basic biogeochemical characteristics were explored via a simple analysis of variance using a univariate gen-eral linear model (GLM), with site number as the fixed factor. Post-hoc analysis (Tukey’s honestly significant difference) was used to identify significant homogeneous subsets. The relation-ship between hydrolytic enzyme activity and total soil P was ex-plored by use of the SPSS curve estimation of an inverse func-tion. Site biogeochemical characteristics were averaged among samples (n = 3) and analyzed for correlation (Pearson’s r) with P forms identified by 31P NMR.

REsuLTssoil Biogeochemical Properties

Surface peat from across the wetland transect was very acidic (pH 3.7 ± 0.4) and of low bulk density, ranging from 0.03 g cm−3 in the central bog plain to 0.08 g cm−3 in forested portions of the transect (Table 1). Total C concentrations ranged between 417 and 535 g kg−1, with significant differences among sites (P < 0.001) but with no clearly discernible pattern.

Total N and P concentrations declined (P < 0.001) from the relatively nutrient-rich Site 1 (total N = 29 g kg−1, total P = 1.0 g kg−1) to oligotrophic Site 9 (total N = 22 g kg−1, total P = 0.4 g kg−1), with total soil P showing the same gradient as that observed by Sjögersten et al. (2011) (1.0–0.4 g kg−1). For both N and P, post-hoc analysis (Tukey’s honestly significant difference) showed a similar pattern of four homogeneous subsets of sites


across the gradient, with proximate sites showing no significant differences with each other. When nutrient concentrations were expressed on a volumetric basis, the low bulk densities in the cen-tral region amplified the gradients in N and P across the transect (Fig. 2A and 2B), with a significant (P < 0.001) fivefold increase in total P (14.6 to 70.9 g m−3) and almost threefold increase in to-tal N (0.73 to 2.01 kg m−3). Molar N/P ratios ranged from 62 at Site 1 to 111 at Site 9 (Fig. 2C), while the molar C/P ratio ranged from 1252 at Site 1 to 3335 at Site 8 (data not shown).

Phosphorus BiogeochemistryConcentrations of readily exchangeable P extracted by AEM

strips were <3 mg kg−1 for Sites 5 to 9 but up to 30 mg kg−1 (2.8% of total P) at other sites (Table 2). Phosphorus released by hexanol fumigation differed significantly (P < 0.05) among sites, represent-ing between 18 and 38% of soil total P, even without the use of a correction factor to account for unrecovered biomass (Bunemann et al., 2008; Jenkinson et al., 2004). There was a highly significant negative correlation (Pearson’s r = −0.55, P < 0.01) between total soil P and the proportion of total P released by fumigation.

Phosphatase activity (both phosphomonoesterase and phos-phodiesterase) increased from the nutrient-rich margin of the wetland to the oligotrophic interior, with central sites showing rates up to seven times that of the peripheral Site 1 (Fig. 3). Values differed significantly among sites (P < 0.001), with phosphatase activity showing a highly significant (P < 0.001) inverse relation-ship with total soil P (phosphomonoesterase, R2 = 0.55; phospho-diesterase, R2 = 0.74).

Phosphorus Recovery in sodium Hydroxide– EDTA Extraction

An average of 63% of total soil P was extracted in NaOH–EDTA from dried soils, with site averages ranging from 45 to 71% of total soil P. Extraction of P from dried soils was similar to that recovered from fresh soils by the combined use of AEM and NaOH–EDTA extraction after hexanol fumigation (Cheesman et al., 2010a). Although more P was recovered from dried soil (paired t-test, P < 0.001), the difference represented an average of only 0.3% of the total soil P, suggesting that alkaline extraction of air-dried soils recovers P in live microbial biomass (Turner et al., 2003a) and that unextractable residual P is a physically distinct pool. Extraction of air-dried soils for 16 h resulted in an aver-age increase in recovery of 4.1% of total soil P across all samples (paired t-test, P < 0.001). Given the known hydrolysis of vari-ous phosphodiesters in alkaline solution (Turner et al., 2003b), however, we used a standard 4-h extraction for detailed 31P NMR analysis. Replicate extractions using a 4-h extraction yielded a consistent recovery (±10% of total soil P) based on molybdate colorimetry of digested alkaline solutions (data not shown).

solution Phosphorus-31 NMR spectroscopyA diverse range of P forms was present in the soils, includ-

ing organic (phosphomonoesters, phosphodiesters, and phos-

Fig. 2. (A) Mass of total P, (B) mass of total N, and (C) N/P molar ratio from nine study sites (average ± sE, n = 3) across four different vegetation types found within the Changuinola peat dome. samples were taken along a transect between the periphery of the peat dome (site 1) to near the geodesic center (site 9), with general vegetation groupings based on “phasic communities” identified by Phillips et al. (1997). All attributes show significant overall differences among sites (P < 0.001), with letters above bars indicating homogenous subsets as derived from Tukey’s honestly significant difference post-hoc analysis.


phonates) and inorganic (orthophosphate, pyrophosphate, and long-chain polyphosphate) forms (Fig. 4; Table 3).

Phosphomonoester concentrations ranged from 45 to 174 mg P kg−1 (11.5–17.1% of total soil P) (Fig. 4; Table 3) and did not include peaks characteristic of myo-, scyllo-, neo-, or D- chiro-inositol hexakisphosphate (Fig. 5) (Turner and Richardson, 2004; Turner et al., 2003c; Turner et al., 2012). Concentrations of DNA ranged from 47 to 105 mg P kg−1, which represented between 8.7 and 13.3% of the total soil P. Phosphodiesters other than DNA were detected at low levels (0.6–3% of total soil P) but phosphodiesters such as phosphatidyl choline and RNA that are hydrolyzed in alkaline solution are likely to be underestimated (Makarov et al., 2002; McDowell and Stewart, 2005; Turner et al., 2003b). Phosphonates occurred at up to 31 mg P kg−1 and ranged from 1.7 to 3.3% of total soil P. These compounds were not de-tected at the low-P Sites 8 and 9, although this may be due to a low signal/noise ratio and an inability to resolve their presence, as opposed to a true absence.

Inorganic P identified by solution 31P NMR ranged be-tween 100 and 400 mg P kg−1 (23–39% of total soil P) and in-cluded both orthophosphate and polyphosphates. Orthophos-phate concentrations ranged from 29 to 258 mg P kg−1 (7–25% of total soil P) and pyrophosphate from 2 to 32 mg P kg−1 (up to 3% of total soil P). Longer chain polyphosphates were found in surprisingly high concentrations from all study sites, ranging from 69 to 165 mg P kg−1 (10–24% of total soil P). Concen-

Fig. 4. solution 31P NMR spectra showing a range of P forms present in surface soils from select sites across the study transect: A = phosphonates, B = methylenediphosphonic acid (MDP, internal standard d = 17.46 ppm), C = orthophosphate, D = phosphomonoesters, E = phosphodiesters, F = DNA, G = polyphosphate (end-chain residues), H = pyrophosphate, I = polyphosphate (mid-chain residues). spectra plotted using 15-Hz line broadening referenced and scaled using internal standard MDP. ‡Due to a lack of sample material, spectra were acquired on additional samples collected in November 2007.

Fig. 3. Enzyme activity from nine study sites (average ± sE, n = 3) across four different vegetation types found within the Changuinola peat dome. samples were taken along a transect between the periphery of the peat dome (site 1) to near the geodesic center (site 9), with general vegetation groupings based on “phasic communities” identified by Phillips et al. (1997). Both phosphomonoesterase and phosphodiesterase activities show significant difference among sites (P < 0.001), with letters above bars indicating homogeneous subsets as derived from Tukey’s honestly significant difference post-hoc analysis.

Table 2. Phosphorus identified by anion exchange membrane (AEM) extraction of fresh soils at sampling stations across the Changuinola ombrotrophic peat dome.

site AEM-extractable P Fumigation-released P

——————— mg kg−1 ——————— % total soil P1 29.5 ± 9.6 a† 184 ± 40.5 abc 18 ± 4.6 a

2 17.6 ± 4.2 ab 232 ± 20.1 cd 23 ± 0.4 ab

3 27.6 ± 7.9 a 193 ± 19.4 bcd 20 ± 2.1 ab

4 5.4 ± 8.1 bc 138 ± 52.5 ab 21 ± 6.0 ab

5 0.1 ± 0.0 c 267 ± 24.2 d 38 ± 3.9 c

6 0.9 ± 0.7 c 190 ± 25.3 bcd 29 ± 2.7 bc

7 2.9 ± 2.1 bc 148 ± 12.4 ab 22 ± 3.6 ab

8 0.2 ± 0.0 c 110 ± 8.5 a 28 ± 2.5 abc

9 0.3 ± 0.2 c 151 ± 6.0 ab 35 ± 4.3 c

† Mean ± one standard deviation (n = 3); superscript letters denote homogenous subsets determined by post-hoc Tukey’s honestly significant difference test.


trations of polyphosphates determined in dried soils correlated (Pearson’s r = 0.80, P < 0.05) with fumigation-released P in fresh soils. Polyphosphate was also detected in alkaline extracts of fresh (undried) soil extracts (Fig. 6), indicating that their presence was not an artifact of fungal growth during sample air drying (Kou-kol et al., 2008). When extracts of fresh soils were analyzed by solution 31P NMR after application of AEM strips but without the use of hexanol as a biocide, polyphosphates were detected. When samples were pre-extracted with AEM strips and hexanol fumigation, however, polyphosphates were not detected or were found only in trace concentrations (Fig. 6).

The concentration of residual P (i.e., P not extracted or iden-tified by NMR analysis) ranged from 193 to 333 mg P kg−1 and was correlated with total P (Pearson’s r = 0.75, P < 0.05). When expressed as a proportion of the total P, residual P increased from 29% at the relatively enriched Site 1 to 55% at the oligotrophic Site 9 (Table 3), with values correlated negatively with total P (Pearson’s r = −0.87, P < 0.005).

The two forms of P with the largest absolute change in their contribution to the total P both correlated negatively (P < 0.01) with residual P (orthophosphate, Pearson’s r = −0.83; phospho-

monoesters, Pearson’s r = −0.80). In addition, there was a signifi-cant positive correlation (Pearson’s r = 0.79, P < 0.05) between soil total P and the ratio of P identified as phosphomonoesters and phosphodiesters, ranging from 1.40 at Site 2 to a low of 0.88 at Site 8.

DIsCussIONSurface peat at all sites was acidic and presumably derived

from the current standing vegetation. Given that mineral matter deposited from external sources can be considered minimal in an ombrotrophic system (Phillips et al., 1997), the significant dif-ferences in C concentrations may reflect differences in biosilica deposition from opal phytoliths and diatoms (Wüst et al., 2002) or variance in C decomposition and relative peat accretion rates across the transect (Craft and Richardson, 1993).

Elemental analysis confirmed previous studies in identify-ing the presence of a strong nutrient gradient across the distinct vegetation communities (Sjögersten et al., 2011; Troxler, 2007). Total P was at the high end of the range observed in other tropi-

Table 3. Phosphorus forms identified by solution 31P nuclear magnetic resonance (NMR) spectroscopy in soils from the Changuinola ombrotrophic peat dome. The concentration was determined by multiplying the proportion of spectral area by the total P determined by digest of alkaline extracts.

siteTotal

P Phosphonate Orthophosphate Phosphomonoesters DNAOther

Phosphodiesters† PyrophosphatePolyphosphate‡ Residual

PER MR

——————————————————————————————— mg P kg−1 soil (% of total P) ———————————————————————————————1 1028 27 (3) 258 (25) 167 (16) 105 (10) 29 (3) 32 (3) 34 (3) 76 (7) 301 (29)

2 1014 31 (3) 233 (23) 174 (17) 93 (9) 30 (3) 20 (2) 14 (1) 107 (11) 310 (31)

3 956 18 (2) 170 (18) 143 (15) 91 (10) 26 (3) 11 (1) 25 (3) 138 (14) 333 (35)

4 655 15 (2) 129 (20) 79 (12) 57 (9) 10 (1) 26 (4) 19 (3) 61 (9) 260 (40)

5 710 16 (2) 82 (12) 86 (12) 72 (10) 14 (2) 4 (1) 18 (3) 147 (21) 271 (38)

6 659 16 (2) 83 (13) 75 (11) 61 (9) 9 (1) 3 (<0.5) 9 (1) 135 (20) 268 (41)

7 672 11 (2) 76 (11) 89 (13) 90 (13) 10 (1) 3 (<0.5) 9 (1) 124 (19) 260 (39)

8 388 ND§ 29 (7) 45 (11) 47 (12) 3 (1) 2 (1) 4 (1) 65 (17) 193 (50)9¶ 578 ND 54 (9) 68 (12) 57 (10) 3 (1) 7 (1) ND 72 (12) 317 (55)

† Includes RNA and phospholipids not hydrolyzed by alkaline extraction and solution 31P NMR spectroscopy.‡ Polyphosphate ER = end-chain P residue; MR = mid-chain P residue.§ ND, not detected or trace.¶ Spectra acquired on samples collected in November 2007.

Fig. 5. Detail of solution 31P NMR spectra from site 7 soils. spectra plotted using 2-Hz line broadening and referenced using internal standard methylenediphosphonic acid (d = 17.46 ppm).

Fig. 6. solution 31P NMR spectra of soils from sites 1 and 9 after application of anion exchange membranes with (F) and without (NF) a hexanol fumigation step. spectra plotted using 15-Hz line broadening referenced and scaled using internal standard methylenediphosphonic acid (d = 17.46 ppm).


cal ombrotrophic systems, with sites in Kalimantan ranging from 272 to 373 mg kg−1 (Page et al., 1999) and the Peruvian low-land Amazonia ranging from 130 to 590 mg kg−1 (Lahteenoja et al., 2009). This may reflect the relatively young age and shal-low depth of the study peatland (Phillips et al., 1997) or periodic deposition of P during its formation from either volcanic ash or catastrophic flooding events (Cohen et al., 1989).

Membrane-extractable and total P increased significantly toward peripheral R. taedigera sites, yet the high total C/P molar ratios (range 1252–3335) and total N/P (62–111), in addition to high rates of phosphatase enzyme activity at all sites, suggest the potential for P limitation of vegetation and microbes within the soils of all sample sites (Cleveland and Liptzin, 2007). Phos-phomonoesterase activity at peripheral sites (Sites 1–3) were similar to those reported in a Malaysian forested peat swamp ( Jackson et al., 2009), suggesting the potential for high organic P turnover in tropical wetlands (Penton and Newman, 2008). The marked increase in phosphatase activity alongside a decreasing nutrient concentration toward the center of the dome indicates a clear increase in biological investment in the acquisition of P from organic forms (Sinsabaugh and Moorhead, 1994; Wright and Reddy, 2001).

Across all sites, a substantial proportion of the total P was held in a pool released by fumigation. Although it is likely that P from fine roots liberated by the action of hexanol contribut-ed to this pool (Sparling et al., 1985), fumigation without the use of a correction factor underestimates the microbial biomass (Brookes et al., 1982; Myers et al., 1999). It is therefore likely that a substantial proportion of P identified in the NMR spectra represents live microbial biomass. A similar conclusion can be drawn when comparing the extraction efficiency of the coupled AEM and alkaline extraction of fumigated fresh soil to that of air-dried soil. The large and significant increase in extraction ef-ficiency between nonfumigated fresh and air-dried soil is con-trary to reports of the influence of pretreatment on other peat-based soils. For example, direct alkaline extraction of fresh and air-dried soils resulted in a similar recovery of total P for three sites in the Florida Everglades, with the notable exception of soft-water floc samples (pH 5.8) (Turner et al., 2007). We interpret this as evidence of either a significant pH ´ pretreatment inter-action on the alkaline extraction efficiency or physical protection of P pools and microbial biomass in the highly fibrous peats of this study system.

The diversity of P forms identified by solution 31P NMR varied little among vegetation types or across the nutrient gradi-ent, yet the relative proportions of different compounds showed progressive alteration in relation to the basic biogeochemical gradient and microbial P. This study showed that, similar to other organic wetlands, a significant proportion of organic P oc-curred as phosphodiesters or potential phosphodiester hydroly-sis products (Turner and Newman, 2005). This is in contrast to terrestrial soils, in which up to 90% of organic P occurs as phos-phomonoesters (Condron et al., 2005), with isomers of inositol hexakisphosphate often forming a substantial proportion of the

total organic P (Turner, 2007; Turner et al., 2012). This distinc-tion has been attributed to differential stabilization in the or-ganic matter and redox conditions prevalent in wetlands (Turner and Newman, 2005; Turner et al., 2006), although the recent detection of the phosphomonoester myo-inositol hexakisphos-phate within anaerobic sediments (McDowell, 2009; Turner and Weckström, 2009) suggests a complex interplay between inputs and site-specific stabilization processes. The general distinction between terrestrial soils and wetlands (Turner and Newman, 2005) might also represent a difference in the proportion of to-tal soil P found within viable microbial biomass compared with the extracellular environment (Oberson and Joner, 2005), which seems likely here given the high proportion of total soil P found within the microbial biomass.

Phosphonates, while not present in the highly studied cal-careous or soft-water peatlands of South Florida (Turner and Newman, 2005; Turner et al., 2006), have been detected in acidic northern hemisphere blanket bogs and subarctic tundra (Bed-rock et al., 1994; Turner et al., 2004). It has been proposed that phosphonates are either more prevalent within the microbial biomass of these systems (Ternan et al., 1998) or that phospho-nates experience greater extracellular stability under cold, acidic, and saturated conditions (Condron et al., 2005). The presence of phosphonates in the tropical wetland studied here demonstrates that they may also be found in substantial quantities in acidic peats with a range of total P concentrations and a high soil tem-perature (26°C).

A large proportion of total P occurred as inorganic polyphos-phates from all sites, including those where P availability is expected to be low and potentially limiting to both plant productivity and microbial activity. Polyphosphates play an integral role in archeal, prokaroyotic, and eukaryotic cells (Kornberg et al., 1999) and are often associated with luxury microbial P uptake (i.e., high-P envi-ronments) (Hupfer et al., 2007; Khoshmanesh et al., 2002), includ-ing activated sludge processing (Reichert and Wehrli, 2007). Poly-phosphates have also been detected in a range of natural wetland and aquatic systems, however, including oligotrophic lake sediments (Ahlgren et al., 2006; Hupfer et al., 2004), Carolina bays (Sundar-eshwar et al., 2009), the humic-acid fraction of reseeded peatlands (Bedrock et al., 1994), and subarctic tundra (Turner et al., 2004). In this study, the large concentrations of inorganic polyphosphates determined by solution 31P NMR spectroscopy of dried soil extracts correlated strongly with fumigation-released (microbial) P in fresh soils, as well as appearing to be released by hexanol fumigation of fresh soils (Fig. 6). Therefore, polyphosphate measured in this wet-land probably represent intracellular stores associated with P homeo-stasis or microbe sporulation (Brown and Kornberg, 2004) under conditions of fluctuating redox (Davelaar, 1993), P storage associ-ated with arbuscular or saprotrophic fungi (Koukol et al., 2008), or a generalized microbial metabolic response to environmental stress or nutrient deficiency within an ombrotrophic environment (Seuffer-held et al., 2008). It should also be noted that polyphosphates, al-though stable under the alkaline extraction conditions, are catalyti-cally degraded by the presence of divalent cations (Harold, 1966).


The addition of EDTA improves the recovery of polyphosphates (Cade-Menun and Preston, 1996) and reduces Fe-induced hydro-lysis in alkaline extracts (Hupfer et al., 1995), yet some researchers interested in their role in lacustrine and marine sediments have ad-opted a pre-extraction with either buffered dithionite or EDTA to further preserve polyphosphates (Ahlgren et al., 2007; Hupfer et al., 2004; Reitzel et al., 2007). Therefore, the presence of large quantities of polyphosphates identified within this study when not using a pre-extraction step may be the result of intrinsically low divalent cation concentrations and a reduced degradation rate compared with other wetland extractions.

In ombrotrophic wetlands, the vast majority of organic mat-ter in surface soils is autochthonously derived. Phosphorus inputs within this organic matter consist of a variety of forms dependent on the nature and structure of the biotic community (Harrison, 1987; Makarov et al., 2005), yet microbial modification of P forms within the detrital material of organic wetland soils is strongly in-fluenced by nutrient availability (Cheesman et al., 2010b). Given the rapid nature of microbially mediated processes (Oberson and Joner, 2005), especially in a tropical setting, it is likely that soil P forms represent a balance among primary eukaryotic inputs mod-erated by prokaryotic processing, as well as forms present within the live biomass (i.e., microbial cells, roots, and the microfauna). The reduction in P availability toward the central portion of the wetland suggests an increase in the role of organic P cycling, which corresponds with a pronounced decrease in the relative propor-tions of total soil P identified as orthophosphate and phospho-monoesters, as well as in the ratio of phosphomonoesters to phos-phodiesters. The absolute concentration of residual P not identi-fied by alkaline extraction and 31P NMR, however, was relatively consistent across the pronounced gradient in total P. This resulted in a gradient in the proportion of total P found as residual P, po-tentially reflecting an inherently stable pool that comes to domi-nate soil P as turnover rates increase and other standing pools are reduced. Further work is need to establish whether this pattern represents differences in extracellular stabilized P or changes in the constitutive components of the microbial biomass (Makarov et al., 2005) that represent a major fraction of the soil P within this tropi-cal ombrotrophic wetland.

ACKNOWLEDGMENTsWe thank G. Jacome and P. Gondola of the Smithsonian Tropical Research Institute, Bocas del Toro Research Station, for logistical support and Dr. Jim Rocca, Dr. Alex Blumenfeld, and Tania Romero for analytical support. The project was supported by a grant from the USDA–CREES National Research Initiative (no. 2004-35107-14918) and the External User Program of the National High Magnetic Field Laboratory administered through the Advanced Magnetic Resonance Imaging and Spectroscopy (AMRIS) facility of the McKnight Brain Institute, University of Florida.

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Wetland Soils Soil Phosphorus Forms along a Strong Nutrient … · 2012-06-18 · Soil Phosphorus Forms along a Strong Nutrient Gradient in a Tropical Ombrotrophic Wetland Wetland