Efforts to understand and model the current and future behavior of the global
phosphorus (P) cycle are limited by the availability of global data on rates of soil P processes, as well as their environmental controls. Here, we present a novel isotope pool dilution approach using 33P labeling of live and sterile soils, which allows for high-quality data on gross fluxes of soil inorganic P (Pi) sorption and desorption, as well as of gross fluxes of organic P mineralization and microbial Pi uptake to be obtained. At the same time, net immobilization of 33Pi by soil microbes and abiotic
sorption can be easily derived and partitioned. Compared with other approaches, we used short incubation times (up to 48 h), avoiding tracer
remineralization, which was confirmed by the separation of organic P and Pi using isobutanol fractionation. This approach is also suitable for strongly weathered and P-impoverished soils, as the sensitivity is increased by the extraction of exchangeable bioavailable Pi (Olsen Pi; 0.5 M NaHCO3)
followed by Pi measurement using the malachite green assay. Biotic
processes were corrected for desorption/sorption processes using adequate
sterile abiotic controls that exhibited negligible microbial and
extracellular phosphatase activities. Gross rates were calculated using
analytical solutions of tracer kinetics, which also allowed for the study of gross
soil P dynamics under non-steady-state conditions. Finally, we present major
environmental controls of gross P-cycle processes that were measured for
three P-poor tropical forest and three P-rich temperate grassland soils.
Introduction
Phosphorus (P) is a major limiting nutrient to terrestrial primary
production, particularly in highly weathered soils such as those found in the tropics. Globally, increasing imbalances between nitrogen (N) and P inputs
(i.e., increasing N : P stoichiometry of inputs) caused by human activities and
land-use changes through increased emissions of reactive N are suggested to
lead to progressive P limitation of terrestrial ecosystems, and the first signs thereof have already been identified (Penuelas et al., 2013). A decrease in the
relative P availability might have strong repercussions on future nutrient
limitations of natural ecosystems, on food production and on carbon (C)
sequestration (Penuelas et al., 2013; Penuelas et al., 2012; Yang et al.,
2013). Efforts to understand and model the current and future global P cycle
and its coupling to the global C and N cycles have been intensified, but are
strongly limited by the availability of global data on soil gross P processes
and their environmental controls (Reed et al., 2015). Therefore, large
investments in new projects, experiments and models have recently been
undertaken to advance our understanding of the terrestrial P cycle,
and to fill data gaps, e.g., IMBALANCE-P (http://imbalancep-erc.creaf.cat, last access: 2 August 2019) and
NGEE-Tropics (http://ngee-tropics.lbl.gov, last access: 2 August 2019).
Schematic representation of (a) major fluxes of soil P processes
controlling the availability of inorganic P (Pi) in soils, and of (b) the isotope pool dilution principle showing influxes of unlabeled Pi
(31P) into the available Pi pool labeled by a spike in
33Pi, and efflux of Pi in the ratio of 33P:31P as
present in the target pool. Biotic and abiotic processes of influx and efflux
are presented. (c) This causes a decline in the specific activity of Pi,
i.e., 33Pi:31Pi declines over time in sterile soils
(abiotic processes only) and live soils (biotic plus abiotic processes),
allowing for the biotic contributions to overall gross fluxes to be derived.
TPi represents total soil Pi, and TPo represents total organic
P; TPi includes occluded and fixed P as well as primary mineral P, and
TPo includes occluded Po in aggregates. Avail. refers to available.
Pi desorption includes Pi dissolution from minerals, and Pi
sorption includes Pi precipitation.
Soil Pi availability is governed by transfers of
exchangeable P between pools (immobilized/fixed P and occluded P), by the slow release of Pi from mineral P via the weathering of primary minerals and by the
mineralization of organic P (Po) (Fig. 1, this study; Bünemann, 2015;Turner
et al., 2007). In strongly weathered soils, primary mineral P pools are
depleted, and the largest fraction of P is found in occluded and fixed pools,
as well as in Po (Vitousek and Farrington, 1997; Yang and Post, 2011).
Phosphorus limitation in such soils is further aggravated by their high P
sorption potentials caused by high contents of Fe–Al (hydr)oxides
(Goldberg and Sposito, 1985). Most of the immediate P needs of
plants (and microbes) in natural and agricultural systems are supplied by
Po mineralization, catalyzed by extracellular phosphatases that are
released by soil microbes and plant roots (Richardson and Simpson,
2011), as well as by abiotic Pi desorption. Soil microbes and plant
roots can promote the release of P from primary and secondary minerals by
accelerating mineral dissolution and Pi desorption, via the exudation of (phyto)siderophores and organic acids (Mander et al., 2012; Ryan et al.,
2001).
Comparison of traditional isotope exchange kinetic kinetic (IEK) experiments
and the novel isotope pool dilution (IPD) approach to measure organic P
mineralization.
Factor and approachIsotope exchange (IEK)Isotope pool dilution (IPD)Tracer addition and incubation period33P, 32P; several time points across several days to weeks and months33P, (32P); two time points at 4 and 24 hMeasured P poolWater-extractable PiBicarbonate-extractable Pi and PoAbiotic controlsAbiotic controls measured in batch experiment with live soil: 100 min Pi exchange experiment in soil suspension 1 : 10 (soil : water), ±HgCl2 or sodium azide; microbial contributions in short-term experiment often not accounted forDuplicate autoclaving for abiotic controls to kill microbial biomass and extracellular enzymes; treatment of abiotic controls similar to live soils in terms of tracer addition, incubation period and extractionMicrobial processes inabiotic controlsMicrobial biomass active in abiotic controls if no microbicide added, extracellular phosphatases fully active (causing organic P mineralization in abiotic controls)Microbial biomass and phosphatases deactivated by autoclaving (no/almost no P mineralization occurring in abiotic controls)Preincubation of soilsto equilibrate to moisture and temperatureYes (to constant respiration – equilibrium conditionsnecessary)Yes (not necessary)Change in soil structure and P availabilityNo (if no microbicide is added)Potentially yes, as autoclaving might increase available P by death of microbial biomass and soil structure might change by autoclavingNumerical solution for Po mineralizationIsotopically exchangeable P within t minutes, E(t), derived as the inverse of the relative specific activity of phosphate in soil solution (water extractable Pi) over time in live soils. E′(t) derived for abiotic controls extrapolated from 100 min to length of full experiment, graphical solution of corrected data following Fardeau (1993). Differences in E′(t) and E(t) estimate gross Po mineralizationCalculation of IPD influx rates based on mass/isotope balance equations derived by Kirkham and Bartholomew (1954) for tracer: tracee experiments. Gross Po mineralization calculated as difference of IPD influx rates of live soils minus abiotic controls
Soil P cycling processes such as soil Pi sorption/desorption fluxes and gross Po mineralization rates, as well as the size of the exchangeable soil Pi pool have been measured by isotope exchange kinetic (IEK) techniques using 32P or 33P. These techniques are based on recurrent measurements of radiotracer recovery and Pi concentration in soil water extracts (Di et al., 1997; Frossard et al., 2011;Bünemann, 2015; Table 1, this study). A variety of IEK procedures and protocols are in use, and optimizations in methodology have been called for, particularly for Po mineralization
(Bünemann, 2015). Only over the last decade have common,
accepted protocols been adopted, and these protocols are currently used to measure soil
P processes following Oehl et al. (2001b). In this IEK
approach, abiotic sorption/desorption processes from
an isotopically exchangeable Pi pool are measured over a short time
period in short-term batch experiments (100 min, 1 : 10 (w:v) soil : water slurry, ± microbicides). This assumes that no microbial tracer uptake (blocked by
microbicides) and no organic P mineralization occurs, and that soils are in a
steady state, i.e., do not show changes in Pi concentration (Table 1). In such
short-term IEK experiments the decrease in radioactivity (radiotracer
recovery) in soil water is described by a power function:
r(t)/R=r1min/Rxt-n,
where R is the added radioactivity, and r(t) is the radioactivity recovered at any time t in soil water extracts. The parameters r1min/R and n (slope of the
regression indicating speed of isotopic exchange) are derived from the
log–log regression of r(t) versus time. This is based on steady-state
assumptions, i.e., that Pi concentration in soil water extracts (CP)
is constant. In some soils an extended version of this equation needs to be
applied:
r(t)/R=mx(t+m1/n)-n+rinf/R.
Here, rinf/R is the maximum possible dilution of the added radiotracer,
approximated as the ratio of CP to total inorganic P in soils. n and m
are derived from non-linear fitting procedures. Assuming that the tracer and
tracee behave similarly in the system, the specific activity of Pi in the soil solution should reflect the specific activity of isotopically
exchangeable P – termed the E value (in mg P kg-1 soil).
E(t)=CP/(r(t)/R).
Isotopic dilution, E′(t), is further measured over the full length of a moist soil incubation experiment lasting for several days to weeks,
constituting the total amount of exchangeable Pi or isotope dilution
caused by concurrent biological processes (Po mineralization) and
physicochemical processes. Short-term exchange kinetics are then extrapolated
over the full time period of the moist soil incubation, E(t)
(Fardeau et al., 1991). The difference between E′(t) and the
extrapolated E(t) value then provides the measure of gross Po
mineralization.
The isotope pool dilution approach (IPD) of Kirkham and Bartholomew (1954) was developed as a general tracer approach to measure gross rates of
soil element cycle processes, but was most frequently applied to nitrogen
cycling processes such as organic N mineralization and nitrification
(Booth et al., 2005). However, the IPD approach can also be
transferred to measure gross rates of P cycle processes (Di et al., 2000).
It then also relies on the labeling of the Pi pool with 33P or 32P and on subsequent time-resolved measurements of concentrations and specific activities of Pi (Table 1, Fig. 1b). However, in contrast to IEK techniques, changes in Pi concentrations and specific activities are then solved by mass balance equations developed specifically for gross rate calculations based on tracer studies (Kirkham and Bartholomew, 1954). In the following we list the criteria that have to be met by the IPD method to correctly determine gross rates of soil Po mineralization and soil Pi sorption/desorption (Di et al., 2000; Murphy et al., 2003; Kirkham and Bartholomew, 1954).
The tracer (32Pi or 33Pi) and tracee (unlabeled
31Pi) behave identically and are well mixed. This is given for the
different isotopes of P as long as radiotracer solution is homogeneously
distributed in the soil and sufficient time is provided for isotope
equilibration between added radiotracer and the native Pi pool.
The influx into the target (Pi) pool (i.e., the product of Po mineralization) has to be unlabeled (i.e., no tracer remineralization), in
order for it to dilute the tracer: tracee ratio over time (Fig. 1b, c).
Tracer remineralization via microbial tracer assimilation, mortality and
subsequent remineralization of labeled Po would result in an
underestimation of Po mineralization, but can be avoided by short
incubation times (1–2 d).
Abiotic release of Pi from a non-extractable pool (Pi desorption) causes an influx of unlabeled Pi into the target pool, resulting in an overestimation of the biotic process, Po mineralization, and has to be determined in parallel abiotic incubations of sterile soils. However, adequate abiotic controls with no contribution of biological processes have remained a major obstacle in measuring soil P dynamics with radiotracers,
both in IEK and IPD experiments. Procedures in earlier studies ranged from
short-term assays with no inhibitor addition as often performed in IEK assays
(Spohn et al., 2013; Oehl et al., 2001b), to amendments of HgCl2,
sodium azide, toluene or chloroform, and gamma irradiation or repeated
autoclaving (Kellogg et al., 2006; Bünemann, 2015; Bünemann et al.,
2007; Oehl et al., 2001b; Achat et al., 2010).
The soil extraction should target the bioavailable exchangeable Pi
pool. Pi in soil solution undergoes rapid equilibration with easily
adsorbed Pi. An incomplete extraction of this pool causes an
underestimation of Po mineralization rates, due to desorption from this pool, causing an influx of unlabeled tracer (and unlabeled Pi) into the target pool, and thus violates assumption no. 2 of IPD assays. The
commonly used soil water extractions target only a small fraction of this
target pool, whereas standard soil P extractants, such as Olsen, Mehlich 3 or
Bray 1, extract a larger fraction (Kleinman et al.,
2001) and, therefore, are suggested to be better suited to extract the
rapidly exchanging Pi pool (Kellogg et al., 2006).
The efflux from the isotopically labeled pool (i.e., microbial Pi
immobilization and Pi sorption into a non-extractable pool) occurs at
the ratio of tracer: tracee as present in the Pi pool at any specific
time, with no discrimination between native Pi and added radiotracer
(Fig. 1b). A short preincubation time is therefore needed to allow for
full mixing and isotopic equilibration of tracer and tracee (see point
no. 1).
Changes in specific activity need to be measured specifically in the target
pool, i.e., in extractable Pi for measurements of gross rates of
Po mineralization and Pi sorption/desorption. However, most current
approaches do not separate extractable Pi and Po but measure
radioactivity in unfractionated extracts, including radiolabeled Po
formed during the incubation, leading to an eventual overestimation of
Po mineralization.
The rates of Pi influx (Po mineralization, abiotic Pi release)
and Pi efflux (biotic and abiotic Pi immobilization) need to be
constant over the duration of incubation: (i) the initial phase of fast
immobilization by sorption, microbial uptake and isotopic equilibration of
radiotracer is excluded from calculations of gross rates, and (ii) incubation
takes place within a suitable time frame to avoid microbial turnover and
33Po remineralization (see point no. 2). The minimum two time
points necessary to measure concentration and specific activity of Pi
for the IPD calculations should therefore lie in between the initial phase
and the start of remineralization, but it is recommendable to test more time
points in the beginning to test the time linearity of IPD rates for specific soil
types.
Mooshammer et al. (2012) adopted such a protocol for
measurements of gross Po mineralization in decomposing plant litter,
following the knowledge of IPD processes based on 15N additions to study
gross rates of soil N cycling (Hart et al., 1994; Murphy et al., 2003; Wanek
et al., 2010; Braun et al., 2018). However, in plant litter P sorption and the
abiotic release of Pi from sorbed P pools do not interfere.
Consequently, the litter protocol cannot be directly transferred to soil
studies. In the present study we developed an IPD protocol to assess soil P
dynamics, based on the previous work for litter by
Mooshammer et al. (2012) and soils by Kellogg et al. (2006). The protocol is based on IPD
theory (Kirkham and Bartholomew, 1954; Di et al., 2000) applied to parallel
incubations of live and sterile soil with 33Pi tracer addition.
Gross rates of Pi sorption (abiotic immobilization) and Pi
desorption are determined in sterile soils, and allow for the correction of gross
Po mineralization and microbial Pi immobilization rates in live
soils. We used bicarbonate extractions to target the bioavailable
exchangeable Pi pool. To avoid tracer remineralization, we used short
incubation periods (up to 2 d). To confirm that no significant amount of
33Po was formed during incubation, Pi was also separated from
Po based on isobutanol fractionation (Jayachandran
et al., 1992). Pi concentrations were measured based on the
phosphomolybdate blue protocol. At very low Pi concentrations, e.g., in tropical soils, which are below the detection limit of the phosphomolybdate
blue method, Pi was determined by parallel measurements of Pi in
bicarbonate extracts using the more sensitive malachite green assay
(D'Angelo et al., 2001; Ohno and Zibilske, 1991). The protocol was tested
rigorously with two different soils, and then applied to six soils in total
(three tropical forest and three temperate grassland soils) to explore
environmental controls on gross soil P dynamics.
Soil characterization of three temperate grassland soils (soil 2, 4
and 6) and three tropical lowland forest soils (soil 3, 5 and 7).
ParameterUnitTemperate soils Tropical soils 246357Soil pH (10 mM CaCl2)6.306.256.804.154.154.10Clay(%)16.814.12.764.1219.626.2Silt(%)59.224.440.688.072.870.1Sand(%)24.061.456.67.927.613.74Total organic C(mg g-1 soil dw)48.3126.760.326.430.828.5Total N(mg g-1 soil dw)3.355.032.322.172.572.27Total P (TP)(mg g-1 soil dw)0.820.440.510.140.170.09Total organic P (TPo)(mg g-1 soil dw)0.400.250.110.090.130.07Soil Pi(µg g-1 soil dw)15.14.235.590.560.490.37TPo of TP(%)49.156.522.364.275.776.4Soil C : N14.425.226.012.112.012.5Soil C : TPo121507548293237406Soil N : TPo8.420.121.124.119.832.5Phosphatase(nmol MUF g-1 soil dw h-1)256316233139616982346Materials and methodsSoil materials and basic characterization
Soils (0–15 cm depth) were collected in summer 2015 from three temperate
grassland sites in Austria and in spring 2015 from three tropical lowland
forest sites in Costa Rica (Table 2). The grassland soils were extensively
managed meadows, collected in Lower Austria (48∘13–20′ N,
16∘12–17′ E) in the vicinity of Vienna, at elevations between 170
and 320 m. The tropical forest soils were collected along a topographic
gradient (ridge–slope–valley bottom) in wet evergreen old-growth forests in
southwestern Costa Rica close to the Piedras Blancas National Park (8∘41′ N,
83∘12′ W, 110–250 m a.s.l.). Soils were sieved to 2 mm and stored
in an air-dried state. Soil pH was measured in a 1 : 5 (w:v) mixture of air-dried soil in water after 60 min of equilibration using an ISFET electrode
(Sentron SI600 pH meter). Soil texture was quantified using a miniaturized
pipette/sieving protocol for 2–4 g air-dried soils (Miller and Miller,
1987), using 4 % sodium metaphosphate as a dispersant. Soil total C and
total soil N content were determined after grinding oven-dried soil in a ball
mill, using an elemental analyzer (EA 1110, CE Instruments, Thermo
Scientific). Temperate grassland soils were treated with 2 M HCl to remove
carbonates, re-dried, ground and then analyzed using an elemental analyzer for soil organic C. Total soil P and total soil Pi were measured after 0.5 M H2SO4 extraction of ignited soils (5 h at 450 ∘C in a
muffle furnace; O'Halloran and Cade-Menun,
2008) and of untreated soils, respectively, using the malachite green method
(Ohno and Zibilske, 1991; D'Angelo et al., 2001). Total organic P was
estimated by calculating the difference between total soil P and total soil
Pi. We must, however, submit that ignition methods tend to overestimate
soil organic P in highly weathered tropical soils (Condron
et al., 1990).
Schematic overview of the final isotope pool dilution (IPD)
procedure. MG refers to the malachite green procedure, and MR refers to the Murphy–Riley procedure to measure Pi concentrations; LSC denotes liquid scintillation counting to measure radioactivity in extracts. Isobutanol fractionation separates dissolved Pi from Po and thereby allows for highly specific measurements of concentrations and 33P activities in Pi, without interference from 33Po. Direct acidification of bicarbonate extracts measures dissolved Pi using malachite green, but LSC quantifies the sum of 33Pi and 33Po; however, the formation of the latter (33Po) turned out to be insignificant.
Soil pretreatment and assay of sterilization efficiency (abiotic controls)
Before starting the experiments, the soils were re-equilibrated from an
air-dried state by rewetting to 60 % water holding capacity for 6 d at
20 ∘C. Gravimetric soil water content and water holding capacity
were determined prior to the experiment. Soils were then either sterilized
twice, 48 and 2 h before the start of the IPD experiments, by autoclaving at 121 ∘C for 60 min (sterile soils), or were kept at 20 ∘C
(live soils, Fig. 2). Sterilization efficiency was checked based on soil
enzyme activity measurements. Fluorescein diacetate (FDA) hydrolysis in soils
was measured as a proxy of viable, active microbial biomass (Green et al.,
2006; Schnurer and Rosswall, 1982), and the activity of acid
phosphomonoesterases, which are extracellular enzymes involved in Po
mineralization, was determined using methylumbelliferyl (MUF) phosphate
(Sirova et al., 2013; Marx et al., 2001).
33P IPD assay
A schematic representation of the final IPD protocol can be found in Fig. 2. Duplicate soil aliquots (2 g fresh weight) of sterile and live soil were
each amended with 20 kBq 33Pi (dilution of orthophosphoric acid
phosphorus-33 radionuclide, 5 m Ci mL-1, i.e., 185 MBq mL-1 HCl-free
water at a specified date, Perkin NEZ080002MC). Between 0.15 and 0.2 mL of
33P-label solution was added to each sample (Fig. 2); the volume added
was adjusted for each soil type to obtain an optimal water content in each
soil (∼ 75 % water holding capacity). Soils were extracted
with 30 mL (temperate soils) or 15 mL (tropical soils) of 0.5 M NaHCO3
(pH 8.5) after 4 and 24 h of incubation for 30 min on a horizontal shaker and
filtered through ash-free cellulose filters. Lower extractant volumes in
tropical and other P-poor soils were used to reach higher Pi concentrations in
the bicarbonate extracts for better quantification.
Following this, isobutanol fractionation of the bicarbonate extracts was
performed, separating Pi (into the organic phase) from Po (into the acidic aqueous phase) allowing for the measurement of the kinetics and specific activity of the Pi pool without interference from Po (Kellogg et al., 2006; Mooshammer et al., 2012). Isobutanol partitioning enables 100 % recovery of Pi with no hydrolysis of Po
(Jayachandran et al., 1992). For isobutanol fractionation, each 1.5 mL of the soil extracts, standards and blanks was amended by sequential addition of 1.5 mL acidified molybdate, 3 mL deionized water and 3 mL isobutanol. The acidified molybdate reagent consists of 5 g ammonium molybdate tetrahydrate ((NH4)6Mo7O24⚫4H2O) dissolved in 0.1 L 2.3 M H2SO4 (stable at room temperature for at least 3
months) and causes strong CO2 outgassing from the bicarbonate extracts. After the addition of all reagents the vials were shaken overhead for 1 min and then rested for 10 min for phase separation. For later photometric quantification of Pi in the isobutanol phase, standards ranging from 320 to ∼ 1 µM Pi (1 : 2 dilution series) and blanks, both of the same matrix as soil extracts (i.e., 0.5 M NaHCO3), were prepared and underwent isobutanol fractionation along with the samples. 33P recovery standards were also prepared and processed via the isobutanol fractionation protocol, consisting of the same volume of extractant (15 or 30 mL) and 33P tracer activity as added to soils (Fig. 2).
Response of soil enzyme activities to autoclaving: percentage
inhibition of (a) fluorescein diacetate (FDA) hydrolysis as a proxy for the
inhibition of live, cell-bound microbial enzyme activity and of (b) MUF-phosphomonoesterase activity as a proxy for the inhibition of
extracellular enzyme activity. Temperate grassland soils (2, 4 and 6) and
tropical forest soils (3, 5 and 7) were tested. A two-way ANOVA was carried out to test for the factors soil, time (1, 24 and 48 h after second autoclaving
cycle, in open, gray and black bars, respectively) and their interaction. P
values are presented.
Pi in the isobutanol phase was quantified using the phosphomolybdate
blue color reaction according to Murphy and Riley (1962). Briefly,
each 1.5 mL of the upper organic phase was transferred to vials and amended
with 2.1 mL molybdate free reducing agent, consisting of 1.32 g ascorbic acid
dissolved in 250 mL antimony potassium tartrate (APT) solution (145.4 mg APT
in 0.5 M H2SO4). The APT solution is stable at room temperature for
> 4 weeks, whereas the molybdate free reducing agent has to be
prepared fresh daily. Thereafter, samples were shaken overhead for 1 min and
rested for 20 min for phase separation and color development. A volume of 250 µL of the blue isobutanol phase was then pipetted into a microtiter
plate, and the absorbance was read at 725 nm with a microplate photometer (Tecan
Infinite M200, Tecan Austria GmbH, Grödig, Austria).
In parallel to the phosphomolybdate blue reaction of Pi in the
isobutanol phase, Pi concentrations were also determined directly in
acidified bicarbonate extracts using the malachite green approach
(D'Angelo et al., 2001). This method is 4–10 times more
sensitive than the commonly used phosphomolybdate blue method and was chosen
to account for the expectedly low Pi concentrations of the tropical
soils. Standards for calibration of the malachite green method were prepared
in 0.5 M NaHCO3, ranging from 50 to 0.039 µM Pi.
Acidification of bicarbonate extracts and standards (blanks) was performed on
2.5 mL sample aliquots by adding 250 µL 2.75 M H2SO4 (Fig. 2). Of the acidified samples and standards, 200 µL was pipetted into
a microtiter plate, and 40 µL of malachite green reagent A was added and
incubated for 10 min. Then, 40 µL of reagent B was added, and the absorbance
was read after 45 min at 610 nm with a microplate reader. Reagent A was
prepared by pipetting 50 mL deionized water into an amber 0.1 L glass bottle,
adding 16.8 mL concentrated H2SO4, and stirring and dissolving 1.76 g
ammonium heptamolybdate tetrahydrate
((NH4)6Mo7O24⚫4H2O). Reagent B was
prepared by heating 0.25 L of distilled H2O to 80 ∘C in an
amber 0.5 L glass bottle, dissolving 0.875 g PVA (polyvinyl alcohol, MW = 72 000 g mol-1) whilst continuously stirring, cooling to room temperature and
finally dissolving 87 mg malachite green oxalate in this solution. Both
reagents are stable for > 6 months at room temperature.
Radioactivity (33P activity) was measured in 0.25 mL aliquots of
acidified bicarbonate extracts and in 0.4 mL aliquots of the isobutanol
phase, after the addition of each 4 mL scintillation cocktail (Ultima Gold,
Perkin Elmer), by liquid scintillation counting (Tri-Carb 1600 TR, Packard,
Perkin Elmer) (Fig. 2).
Experiments
Time kinetics: high-resolution time kinetics of tracer and tracee dynamics
(33Pi, 31Pi) were measured in two soils (temperate
grassland, soil 4; tropical forest, soil 3; Table 2). After tracer addition
to live and sterile soils in triplicates, IPD assays were stopped by
extraction with 0.5 M NaHCO3 after 0, 1, 2, 4, 8, 24 and 48 h. Time
point 0 was assessed by adding the tracer solution and immediately extracting
the soils with 0.5 M NaHCO3.
Microbial 33P immobilization: the procedure outlined in Sect. 2.3 can
be combined with the direct determination of microbial P by extraction with
liquid chloroform-enriched salt solutions (Setia et al., 2012). Here, we
tested a sequential extraction–liquid chloroform extraction (sECE)
procedure. After 24 h of soil incubation in experiment (i), soil samples (2 g
fresh weight) were first extracted with 15 (soil 4) or 30 (soil 3) mL 0.5 M NaHCO3 for 30 min and centrifuged for 15 min at 10.000 g, before the
supernatant was decanted. The soil residue was then re-extracted with 15 (30) mL 0.5 M NaHCO3 containing 3 % (v:v) chloroform for 30 min and finally
filtered through ash-free cellulose filters. Volume corrections were applied
for extractant absorption by the soil pellet after centrifugation. Volume
corrections were calculated as soil wet weight after centrifugation minus
fresh weight weighed into each tube in grams, divided by the density of the
bicarbonate solution (in g mL-1), providing the carryover of extractant from
the first extraction (in mL).
Soil effects on tracer dynamics: live and sterile soils (2 g aliquots) of all
six soils (Table 2) were measured in triplicates for 33Pi activity
and Pi concentrations, and assays were stopped after 0, 4 and 24 h. Net
immobilization of 33P and gross process rates were calculated for the
time interval 4 to 24 h, and relationships between gross and net soil P
processes and soil physicochemical parameters were tested.
Calculations of abiotic and biotic net 33P immobilization
In addition to the measurement of gross rates, abiotic net 33P
immobilization (net soil Pi fixation) and biotic net 33P
immobilization (net soil microbial Pi immobilization) were calculated
based on the determination of the recovery of added tracer in soil extracts
of live and autoclaved soils (see above) after 0, 1, 2, 4, 8, 24 and 48 h. Abiotic immobilization (in % added tracer) was estimated as 100 % minus the percent 33P recovery in autoclaved soils. Total
immobilization was estimated as 100 minus the percent 33P recovery in
live soils. Biotic immobilization was calculated as the difference between
total and abiotic immobilization. These data provide a rapid assessment of
the abiotic versus microbial sink strengths for Pi, but do not represent
gross rates.
Calculations of gross rates of soil P dynamics
Calculation of gross IPD rates followed the mass balance equations of
Kirkham and Bartholomew (1954), as later applied by others for soil
gross P fluxes (Kellogg et al., 2006; Mooshammer et al., 2012). In these
gross P-flux studies abiotic processes were not corrected for; therefore, Pi influx
rates represent the sum of biotic (organic P mineralization) and
abiotic (desorption) processes, the latter of which do not play a significant
role in decomposing litter that is devoid of soil minerals
(Mooshammer et al., 2012). However, to calculate gross
Po mineralization for soils, gross rates of Pi desorption have to
be corrected for in live soils. In the present study, this abiotic correction
was performed by applying IPD calculations for influx (GI – gross influx;
Eq. 1) for sterile soils (abiotic influx by Pi desorption) and live
soils (total Pi influx), and taking the difference as the biotic influx
(i.e., gross Po mineralization). The same procedure was performed for
tracer efflux (GE – gross efflux; Eq. 2), calculating gross
immobilization fluxes for live soils (total Pi efflux) and sterile soils
(Pi sorption), with the difference providing gross rates of microbial Pi
immobilization. Both abiotic corrections are based on the assumption that
abiotic sorption/desorption processes are not affected by autoclaving, i.e.,
that these processes act similarly in sterile and in live soils.
1Gross influx is calculated asGI=Ct2-Ct1t2-t1×lnSAt1SAt2lnCt2Ct12and gross efflux is calculated asGE=Ct1-Ct2t2-t1×1+lnSAt2SAt1lnCt2Ct1,
where t1 and t2 represent incubation time (4 and 24 h; in days),
C represents the soil Pi concentration (in µg Pi g-1 soil dw), SA represents the specific activity (in Bq µg-1Pi)
and LN is the natural logarithm. Thus, gross rates are in micrograms of soil inorganic phosphorus per gram soil dry weight per day (µg Pi g-1 soil dw d-1). Net organic P mineralization rates can
easily be derived by subtracting gross microbial Pi uptake from gross
Po mineralization rates.
Due to the relatively rapid decline in 33P activity by radioactive
decay, all data were decay-corrected back to the start of each experiment,
i.e., the time point of tracer addition to the soil. This was done according
to Eq. (3):
Nt0=Nte-λt,
where Nt0 is the decay-corrected 33P activity in a sample (in Bq), Nt is the measured 33P activity at time of liquid scintillation counting, t is time (in days) elapsed between tracer addition and 33P activity measurement, e=2.71828 and λ is the decay constant of 33P (0.0273539).
Statistics
Regressions were performed in Sigmaplot 13.0 (Systat Software, Inc.) and
group differences were tested using one-way and two-way ANOVA tests followed by a Tukey HSD test in Statgraphics Centurion XVIII (Statpoint Technologies,
Inc.). Variance homogeneity was tested using a Levene test and if necessary data
were log, square root or rank transformed to meet assumptions of
homoscedasticity and normal distribution.
ResultsSoil characterization
Temperate grassland soils had a pH between 6.3 and 6.8, with a silt loam to
sandy loam texture (Table 2). Soil organic C contents ranged between 48 and
127 mg C g-1 soil dw, soil N from 2.3 to 5.0 mg N g-1 soil dw and soil total P from 0.44 to 0.82 mg P g-1 soil dw. Tropical forest soils had a pH between 4.1
and 4.2, and soil texture varied between silt, silty loam and sandy loam. Soil
organic C contents were lower at 26 to 31 mg C g-1 soil dw, soil N ranged from
2.2 to 2.6 mg N g-1 soil dw, and soil total P ranged from 0.09 to 0.17 mg P g-1 soil dw.
Organic P comprised a larger fraction of total P in tropical forest soils
(64 %–76 %) than in temperate grassland soils (22 %–57 %). Extractable soil
Pi was higher in temperate grasslands (4.2–13.1 µg P g-1 soil dw) compared with tropical forest soils (0.07–0.13 µg P g-1 soil dw). Acid phosphomonoesterase activities of tropical
forest soils (1396–2346 nmol MUF released g-1 soil dw h-1)
markedly exceeded those in temperate grasslands (233–256 nmol MUF released g-1 soil dw h-1).
Abiotic controls: soil sterilization efficiency
A separation of biotic and abiotic processes is based on the comparison of
gross rates using the IPD assay in live versus autoclaved soils, where the
latter should not exhibit any microbial activity (no FDA hydrolysis activity)
and no extracellular enzyme activities (no MUF-phosphatase activity), in
order to serve as abiotic controls. An incomplete inhibition of extracellular
phosphatase activities would lead to an underestimation of biological
processes and, therefore, of gross Po mineralization. Our results show
that two consecutive treatments of the soils by autoclaving, with a 48 h
incubation in between, effectively reduced microbial metabolic activity as
shown by the reduction in soil FDA hydrolysis by 90 % in soil 4 and by
97 %–99 % in all other soils (Fig. 3). Autoclaved soils did not show any
increase in soil microbial activity during the 2 d of incubation. On the
contrary, the inhibition of FDA hydrolysis even increased from 1 h (all
soil average: 94 %) towards 24 and 48 h after sterilization (average:
97 %–99 %). The inhibition of extracellular acid phosphatase activity was
almost complete in tropical soils (95 %–97 %) and strongly reduced in
temperate soils (79 %–80 %). Similar to FDA hydrolysis, the extent of
inhibition of phosphatase activity increased from day 0 (average: 86 %) to
day 1 and 2 (average: 88 %–89 %, Fig. 3). However, autoclaving increased
available Pi1.86±0.32-fold (mean ± 1 SD) in temperate
soils and 1.65±0.36-fold in the tropical soils (Fig. S1).
Relationship between (a)33P recoveries as measured directly
in acidified bicarbonate extracts and after isobutanol fractionation,
relative to the added tracer amount, and between (b)Pi concentrations
measured using the malachite green method in acidified bicarbonate extracts and
after isobutanol fractionation following the phosphomolybdate blue approach.
(c) Comparison of specific activities (SA) of Pi measured in acidified
bicarbonate extracts and after isobutanol fractionation. Regression in (c) is
only for temperate grassland soils (closed circles); for tropical forest
soils (open circles), Pi concentrations were close to the detection limit
of the phosphomolybdate method, impairing calculations of SA for isobutanol
fractionation. Linear regressions are given (slopes and intercepts ±1 SD).
Comparison of isobutanol fractionation and direct measurements of Pi and 33P activity
Soil Pi concentrations measured using the malachite green method directly
in acidified bicarbonate extracts were compared to those measured after
isobutanol fractionation by phosphomolybdate blue reaction, including both
live and sterile soils. Both approaches yielded similar soil Pi
concentrations, and the relationship showed no bias (slope =0.979±0.033, mean ±1 SE), with a coefficient of determination of 0.92 (Fig. 4b). The malachite green method is much more sensitive and therefore produced
more reliable results for the low-P soils from the three tropical forests.
Moreover, the relationship between 33P recoveries by isobutanol
fractionation and by direct measurements in acidified bicarbonate extracts
had a slope less than 1 (slope =0.875±0.010; Fig. 4a), indicating no
significant formation of 33Po during soil incubations. We also
found no 33Po formation in other soils using the same measurement
protocols, e.g., from the Jena biodiversity experiment (82 plots of temperate
grassland varying in soil texture and plant biodiversity,
slope =0.891±0.017) and from French Guiana (24 soils from two primary
forest regions, with soils sampled across topographic gradients,
slope =1.043±0.020) (same regression types as in Fig. 3a; data not
shown). The specific activities of Pi were indistinguishable between
both approaches for temperate soils (slope =0.977±0.064, R2=0.93, P<0.0001; Fig. 4c) but varied strongly for the tropical
soils, where soil Pi measurements in the isobutanol fraction were at or
below the limit of detection of the phosphomolybdate blue method. Specific
activities of Pi were initially higher in live than in sterile soils
(Fig. 4c). This was caused by the addition of the same amount of radiotracer
to both, sterile and live soils, but autoclaving caused a flush of Pi
from lysed soil microbes, which effectively lowered the specific activities
of Pi in sterile soils.
Sensitivity of the IPD assay
The sensitivity of this assay is greatly improved relative to traditional
methods by using a combination of bicarbonate extractions and malachite green
Pi measurements. The detection limit of the IPD approach was 0.12 µg P g-1 soil dw d-1, based on 3 times the standard
deviation of gross Po mineralization, measured for the three tropical
soils (each measured in triplicate), and was, therefore, fully suitable across all
soil types tested so far. However, the precision suffers from IPD equations
that combine uncertainties from four measurements, two Pi concentrations
and two radioactivity measurements for the two time points in live as well
sterile soils. The coefficients of variation (CV) ranged between 1.0 % and
22.1 % (average 10.0 %) for Pi concentration across temperate and
tropical soils, and between 1.5 % and 22.1 % (average 9.6 %) for SA, the
two major input variables into the IPD equation. CVs increased towards lower
Pi concentrations and higher SA values, i.e., closer to the detection
limit of the malachite green method. The CVs might be reduced by working with
larger soil aliquots (increase from 2 to 5 or 10 g soil fresh weight) and by
duplicate measurements of all samples. Purely methodological CVs were lower,
at about 2.5 % and 0.9 % for Pi measurements using malachite green in the
range from 3 to 12 and 12 to 120 µM, respectively, and 0.8 % for liquid
scintillation counting. Therefore, much of the shown variability derived from
differences between biological soil replicates. However, the variability
found here compares well with CVs published for soil Pi concentrations
of 2 %–10 % (Bünemann et al., 2007) and 20 %–25 %
(Bünemann et al., 2012), and CVs for measured E values that are calculated from Pi concentrations and 33P
recoveries analogous to SA values ranging between 6 % and 16 %
(Bünemann et al., 2007), 8 % and 19 %
(Bünemann et al., 2012) and 9 % and 10 %
(Randriamanantsoa et al., 2015) across a range of cultivated
and non-cultivated soils from temperate to tropical regions. These variations
naturally propagate into higher errors in the measured rates of soil P
cycling and increase the limit of detection and the limit of quantification
of the various methods.
Test for linearity of change in 33P recoveries (a, b) and in
specific activities of Pi(c, d) over time, for a temperate grassland
soil (a, c) and a tropical forest soil (b, d). Data presented are for
33P measured directly in bicarbonate extracts of live soils (closed
circles) and sterile soils (open circles), and are shown in a
logarithmic manner (LN) on the y axes. Regression lines follow exponential decay which in
this linear–LN plot appears as a straight line; dashed lines represent
sterile soils, and solid lines represent live soils. Regressions were calculated for the
time intervals from 2 to 24 h (tropical soil) and from 4 to 48 h (temperate
soil).
Time kinetics
During the first hour of the incubation, we found a rapid drop in 33P recovery and in the SA of Pi (Fig. 4), while soil Pi concentrations increased slightly (Fig. S1). Thereafter, a dynamic equilibrium between added 33P tracer and the soil Pi pool was reached and concentrations of extractable Pi remained constant. A plot of ln(33P recovery) versus time for both live and sterile soils showed that the consumption of 33P occurred linearly between 4 and 48 h in the temperate soil and between 2 and
24 h in the tropical soil (Fig. 5). Similarly, the plot of ln(SA of Pi) versus time showed a linear relationship from 4 to 48 h in the temperate soil and for 2 to 48 h in the tropical soil, particularly in live soils (Fig. 5), with constant dilution of the isotopic signature of the pool over time. The regressions became insignificant in the sterile tropical soil, as 33P recovery and SA declined more slowly. The data clearly show that abiotic 33P processes (i.e., decreases in 33P recovery and SA of Pi over time in sterile soils) occurred, particularly in the temperate soil, and that this occurred over a prolonged period of time. More importantly, the dynamics of the abiotic 33P processes changed over time: rapid abiotic immobilization during the initial 0–4 h was followed by a period of slower but linear tracer immobilization.
Net 33P immobilization by abiotic and biotic processes
Abiotic net 33P immobilization (net soil P fixation) increased markedly from 0 to 48 h in the grassland soil (17 % to 58 % of added tracer), while it reached 83 % almost instantaneously in tropical soil and further increased to 90 % after 48 h (Fig. 6a). Similar patterns were found across all six soils, with significantly higher abiotic net immobilization in tropical than in temperate soils, increasing in both with time from 0 to 4 and 24 h (Fig. 6c). Biotic (microbial) net 33P immobilization ranged from 3 % to 8 % in the tropical soil and 8 to 17 % in the temperate soil in the time kinetics experiment, with a significant increase in the temperate but not in the tropical soil (Fig. 6b). Similarly, biotic net 33P immobilization was
low but increased with time in all three tropical soils (3 % to 6 %), while
it was significantly higher in temperate soils but increased (soil 6) or
decreased (soil 2 and 4) with time (Fig. 6d). Microbial immobilization was
very fast, with almost instantaneous 33P uptake by microbes (sampling at
0 h), ranging between 3 % (tropical soils) and 15 %–38 % (temperate soils).
Given the strong changes in both abiotic and biotic net 33P
immobilization, we suggest that it is best to measure them after 24 h (up to
48 h).
Sequential extraction–liquid chloroform-extraction (sECE) was applied to
directly follow net 33P uptake by microbes, whereas biotic net 33P immobilization was estimated indirectly as the difference in net 33P immobilization by live and sterile soils. In the two measured soils, sECE estimates of microbial net 33P uptake were higher than the microbial net 33P immobilization estimates (temperate soil was 24.6 % versus 16.0 %, and tropical soil was 16.8 % versus 7.5 %, for direct and indirect estimates, respectively). This indicates incomplete extraction of exchangeable Pi prior to microbial lysis with chloroform and re-extraction.
Net immobilization rates of 33Pi by abiotic processes
(sorption; a, c) measured in sterile soils and biotic processes (microbial
uptake; b, d) measured in live soils of a temperate grassland (soil 4) and a
tropical forest (soil 3) after 0, 1, 2, 4, 8, 24 and 48 h (a, b) and for
six soils measured after 0, 4 and 24 h (c, d). Temperate grassland soils
(2, 4 and 6) and tropical forest soils (3, 5 and 7) were investigated in (c) and (d).
Curvilinear regressions following the function “exponential rise to
maximum” were performed on the data in (a, b). Statistical analyses of data
in (c, d) were undertaken using a two-way ANOVA for the factors soil and time (0, 4 and
24 h after tracer addition), and the interaction between factors.
33P pool dilution rates of abiotic and biotic processes
We calculated gross Pi influx and efflux rates for live and sterile
soils. The calculated rates of sterile soils provide estimates of gross rates of
soil Pi sorption and desorption, and the difference between live and
sterile soils give the biotic influx (gross Po mineralization) and
efflux (gross microbial Pi uptake). Gross Po mineralization
significantly differed between soils, with two out of three temperate soils
(0.48 to 2.03 µg P g-1 dw d-1) exhibiting higher rates than
two out of three tropical soils (0.08 to 0.15 µg P g-1 dw d-1) (Fig. 7a). Gross rates of Pi sorption in temperate soils (2.06
to 6.14 µg P g-1 dw d-1) were higher than in tropical soils (0.15 to 0.32 µg P g-1 dw d-1), and a similar trend was found for gross rates of microbial Pi uptake (temperate soils: 0.44 to 1.13 µg P g-1 dw d-1; tropical soils: 0.05 to 0.12 µg P g-1 dw d-1; Fig. 7b). Gross rates of soil Pi desorption were significantly higher in temperate soils (1.44–3.63 µg P g-1 dw d-1) than in tropical soils (0.04–0.14 µg P g-1 d-1, Fig. 7a). The relative contribution of Po mineralization to total Pi release into the soil Pi pool ranged between 25.0 % and 73.8 %,
with two tropical P-poor soils showing the highest contributions (Fig. 7c).
Contributions of biological processes to gross Pi immobilization did not differ between soils (ranged from 11.5 % to 34.9 %).
Gross influx rates into the available soil Pi pool (a) and
gross efflux rates from this pool (b) measured by 33P isotope pool
dilution for six soils over the time period from 4 to 24 h and assessed in
sterile and live soils. Abiotic and biotic process rates are indicated by
open and closed bars, respectively. Temperate grassland soils (2, 4 and 6) and
tropical forest soils (3, 5 and 7) were studied. Presented are means ±1 SD
of triplicate live and sterile soils per time point and soil type. A one-way
ANOVA was performed on transformed data, as indicated in parentheses. Different
lowercase letters indicate significant differences between soils for abiotic
processes (open bars), uppercase letters indicate significant differences between soils for biological processes (black
bars).
Relationship between selected soil physicochemical parameters, net
abiotic and microbial immobilization fluxes, gross Pi influx rates by
biological processes (gross Po mineralization) and abiotic processes
(gross Pi desorption), and gross Pi efflux rates by biological
processes (gross microbial Pi uptake) and abiotic processes (gross
Pi sorption). Regression lines are for linear or power function fits,
and P and R2 values for these are shown. Open circles (∘)
depict tropical forest soils, and closed circles (•) depict temperate grassland
soils. Units are provided in Table 2 for soil physicochemical parameters and
phosphomonoesterase, and are the percentage (%) of added tracer for net processes, and micrograms of phosphorus per gram soil dry weight per day (µg P g-1 soil dw d-1) for gross process rates.
Physicochemical and biological controls on soil Pi processes
Gross Po mineralization was strongly positively correlated with total
soil P (R2=0.87, P<0.01; Fig. 8a) and with total as well as
extractable soil Pi concentration (R2>0.83, P<0.05; Fig. 8b), but not with soil organic P or its contribution to soil total P, nor with soil organic C, soil texture or soil acid phosphatase activity (Table S1). Gross abiotic Pi release rates via desorption and dissolution were strongly positively related to total soil P and bicarbonate soil Pi (R2=0.97 and 0.98, respectively, both P<0.001; Fig. 8c and Table S1), but not to other parameters such as soil pH, soil texture and soil organic C content. Gross Pi sorption rates exceeded gross Pi desorption rates approximately 2-fold, but both were strongly related (R2=0.99, P<0.001; Fig. 8e). Gross Pi sorption rates were strongly positively related to soil total P (R2=0.96, P<0.001;
Fig. 8d), soil total Pi (R2=0.88, P<0.05; Table S1) and
bicarbonate soil Pi (R2=0.99, P<0.001; Table S1), but
not to soil pH, soil organic C, nor clay content or soil texture.
Abiotic net Pi immobilization was most strongly, negatively related to soil pH (R2=0.95, P<0.001; Fig. 8l) and weakly, negatively related to soil
Pi sorption (R2=0.59, P=0.073; Fig. 8j). Gross
microbial Pi uptake rates were directly proportional to microbial
biomass P measured by sECE (R2=0.95, P<0.01; Fig. 8g), and
positively related to net microbial Pi immobilization (R2=0.85,
P<0.01; Fig. 8i). We found a negative curvilinear relationship
between net immobilization rates by sorption and microbes (R2=0.97,
P<0.001; Fig. 8f).
Discussion
About a decade ago, Kellogg et al. (2006) compared two
IEK techniques with an IPD approach, identifying several biases of the
different approaches and making recommendations for further development. The
authors recommended the IPD approach with soil extraction using 0.5 M sodium
bicarbonate, as it is best suited for any potential soil type. However, this
approach is currently underused and has had issues with abiotic controls. IPD
methods are state-of-the-art to measure gross processes of soil N cycling
(Murphy et al., 2003), but have rarely been applied to
soil P cycling processes (Mooshammer et al., 2012; Di et al., 2000; Kellogg
et al., 2006). Here, we present a novel and versatile approach to derive
quantitative estimates of soil P-cycling processes that drive soil P
availability in low- to high-P soils. The approach quantifies gross rates of
soil Po mineralization and the abiotic release of Pi from
non-extractable soil Pi pools (Pi desorption and dissolution), both of which cause a gross influx of Pi into the soil available Pi pool. Furthermore, gross rates of Pi immobilization by soil sorption and precipitation and by microbial uptake processes are derived from the same data by calculating the efflux from the soil Pi pool in sterile soils (abiotic) and in live minus sterile soils (biotic processes), respectively.
In contrast to many earlier IEK assays, the IPD approach presented here is
based on real isotope pool dilution theory (Kirkham and
Bartholomew, 1954), and not on curvilinear extrapolation of E values (Table 1). Moreover, IEK assays of Po mineralization necessitate steady-state conditions (constant Pi and microbial biomass P pools, and constant rates of isotope exchange and respiration) to allow for the extrapolation of short-term exchange processes to the full length of the moist soil incubation experiments. IPD approaches can accommodate non-steady-state conditions as caused by flush effects and disturbances
(Mooshammer et al., 2017) or as induced by the addition of organic matter. The equations to estimate IPD rates can easily be solved for soils where target pool concentrations increase (net mineralization) or decrease (net immobilization) over time and where microbial biomass P changes (Kirkham and Bartholomew, 1954), and do not necessitate constant pool sizes as has been incorrectly suggested in previous studies (Di et al., 2000; Randhawa et al., 2005).
Soil sterilization
33P IPD experiments in soils differ from the more common 15N IPD variants for gross N processes (Murphy et al., 2003),
as the persistence of abiotic P processes over time (Figs. 5, 6) needs
to be accounted for via the use of sterile soils. Our data clearly show that
the dynamics of abiotic 33P processes change over time. Therefore, the IPD rates in the sterile soils need to be measured over the same time period and under similar environmental conditions as in the live soils. It is likely insufficient to extrapolate from short-term (100 min) batch incubations run under very different conditions to correct for abiotic processes in the respective live moist soil incubations over weeks.
Bünemann et al. (2007) indicated that batch
incubations (1 : 10 (w:v) soil : water suspensions) have higher water-soluble and isotopically exchangeable Pi concentrations (measured as extractable Pi and as E values) and tend to have higher tracer recoveries (measured as r/R, i.e., water-soluble 33Pi recovered relative to total
33Pi added) compared with moist soil incubations. Therefore, incubation conditions should also match between live and sterile soils.
We chose autoclaving as the sterilization procedure as other procedures only
reduce or eliminate microbial activity (gamma irradiation, azide, mercuric
chloride, toluene or chloroform treatment) but do not curtail extracellular
enzyme activities (Blankinship et al., 2014; Wolf et al., 1989; Tiwari et
al., 1988; Oehl et al., 2001b). Given that Po mineralization is mediated by extracellular phosphatases, previous isotope experiments using short-term batch experiments with or without microbicides or γ-irradiation did not inhibit phosphatases and, therefore, did not allow for the separation of abiotic and biotic processes of Pi release in soils. While the application of phosphatase inhibitors might be another viable option, we are only aware of one study testing this; the application of silver nanoparticles to soils showed a general inhibitory effect on soil enzymes (Shin et al., 2012). Previous tests in our laboratory with two commercial phosphatase inhibitor cocktails (Sigma-Aldrich) at 10-fold the recommended final concentration did not significantly decrease IPD rates in two soils (data not shown), indicating an insufficient inhibition of extracellular phosphatases. However, more rigorous tests of soil enzyme activities with synthetic substrates (e.g., MUF-Pi) and of P mineralization based on 33P IPD using increasing concentrations and different types of commercial phosphatase
inhibitor cocktails might clarify whether this approach is viable or not.
In contrast, autoclaving soils twice was highly efficient in suppressing
biological activities, and those soils had no or very low extracellular
enzyme activity and no residual microbial metabolic activity. Previous
studies have shown (almost) total inhibition of hydrolytic enzyme activities
(including phosphomonoesterases) by autoclaving, in a wide range of arable,
grassland and forest soils (Serrasolsas et al., 2008; Kedi et al.,
2013; Blankinship et al., 2014; Tiwari et al., 1988). Other studies have
demonstrated successful killing of bacterial and fungal cells in soils by
autoclaving (Carter et al., 2007; Blankinship et al., 2014; Serrasolsas and
Khanna, 1995b; Alphei and Scheu, 1993). Most importantly, the final step in
Po mineralization is catalyzed by phosphomonoesterases, which were fully inactivated by autoclaving in all soils tested so far.
It must be noted that autoclaving could potentially alter the physicochemical
properties of soils, thereby affecting abiotic sorption/desorption kinetics.
Despite this, in previous studies autoclaving up to two times and steam
sterilization did not affect the cation exchange capacity, nor did it impact base saturation, soil surface area, contents of total organic carbon and total
nitrogen, and it only slightly affected soil pH (Wolf et al., 1989; Tanaka et al.,
2003; Serrasolsas and Khanna, 1995b). Autoclaving might, however, weaken soil
aggregates and therefore increase the number of sites accessible for
sorption/desorption processes that were previously hidden in aggregates.
However, we did not find clear support for or against this in the literature
as autoclaving only weakly affected soil aggregate size distribution, causing
a 0.5 % to 1 % increase in clay-sized aggregates compared with silt-sized aggregates (Berns et al., 2008). In contrast, aggregate
stability and aggregation increased upon autoclaving in two other studies
(Lotrario et al., 1995; Salonius et al., 1967). The effects of autoclaving on
soil aggregation and soil P dynamics could be tested by measuring P-process
rates on intact aggregates < 2 mm and after destroying them by
ultrasonication or grinding. In our study autoclaving caused a pulse of
labile P into the available soil P pool due to the lysis of microbial biomass
(Fig. S1), as has also been demonstrated for P and N by Serrasolsas and
Khanna (1995a, b). Soil Pi concentrations increased significantly in the autoclaved soils studied here, but only by an average of 1.86-fold in the
three temperate soils and 1.65-fold in the three tropical forest soils,
which was in the range found by others, e.g., 1.3- to 1.6-fold
(Skipper and Westermann, 1973) and 1.5- to 1.6-fold
(Anderson and Magdoff, 2005), but lower than reported
elsewhere, e.g., 2.6- to 11-fold (Serrasolsas and Khanna, 1995a).
Autoclaving was also demonstrated to increase the tracer recovery (r/R) and
decrease the velocity of its decline over time, as expected due to loss of
microbial biomass (Bünemann et al., 2007). Therefore, autoclaving slightly affects the soil Pi pool and most likely has minor effects on its abiotic sorption/desorption dynamics, whereas it inhibits biological reactions. Nonetheless, the effects of microbial lysis on Pi sorption/desorption could be tested in sterile soils by adding increasing concentrations of non-labeled Pi alongside the 33Pi tracer and then could be corrected for in future 33P-IPD experiments. As stated earlier, changes in the Pi concentration caused by autoclaving can easily be accounted for in IPD approaches, as long as abiotic process rates remain unaffected by the treatment. However, the estimation of the contribution of abiotic and biotic processes is based on calculating the difference in P fluxes between sterile and non-sterile soils. This assumes that biotic and abiotic fluxes are additive while there is potential that both processes compete for available Pi. In this case we would overestimate abiotic process rates in autoclaved soils, due to lack of competition by biotic processes. This could effectively cause an underestimation of biotic processes, i.e., organic P mineralization and microbial Pi uptake. To date we have no approach at hand to cope with this potential bias. Thus, overall, there is the potential for method improvement, particularly in terms of using abiotic controls circumventing autoclaving (e.g., bactericide combined with phosphomonoesterase inhibitors) or correcting for autoclave-induced changes in aggregation and in soil Pi content.
Soil Pi extraction using bicarbonate
Similar to 15N IPD assays, where salt extractions are employed to target
the available inorganic or organic N pool (Murphy et al., 2003; Wanek et
al., 2010; Hu et al., 2017), we focused on the potentially bioavailable,
salt-extractable Pi pool that more suitably reflects the plant- (and microbial)
accessible amount of soil Pi (Fardeau et al., 1988; Olsen et
al., 1954; Horta and Torrent, 2007) than the water extractable Pi pool, which is mostly assessed with soil IEK methods. The applied 0.5 M NaHCO3 extraction (pH 8.5, Olsen P) promotes the displacement of Pi (and the extraction of labile Po), particularly from Al–Fe (hydr)oxides and soil
organic matter, by competition of bicarbonate anions with Pi. The
underlying process is an increase of the negative charge on surfaces and a
decrease of the concentration and activity of Ca2+ and Al3+,
thereby increasing P solubility in acid to alkaline soils (Horta and
Torrent, 2007; Schoenau and O'Halloran, 2008; Demaria et al., 2005). Several
studies compared soil P tests such as Bray III, resin P and Olsen P to soil
water Pi and plant P uptake in order to assess how well they reflect the available Pi pool. These studies demonstrated that soil tests like bicarbonate extractions (Olsen P), resin P and DGT (diffusive gradients in thin films technique) closely resembled the SA values of Pi extracted by water or 10 mM CaSO4 or from plants (Six et al., 2012; Fardeau et al., 1988; Demaria et al., 2005). Others further showed that isotopically exchangeable Pi in soil water extracts (E values) and those extracted by plant roots in plant growth experiments (L values) were also strongly related (Bühler et al., 2003; Frossard et al., 1994). Bicarbonate extracted 8- to 22-fold greater amounts of exchangeable Pi compared with water, and SA of Pi in bicarbonate extracts reached 66 %–90 % of the SA values measured in soil water extracts (Demaria et al., 2005). IPD approaches require fast extractions to quickly terminate the assay after 4 and 24 h, which renders water extractions (generally 16 h), resin P (16 h) and DGT (up to 48 h in low P soils; Six et al., 2012) impossible.
Bicarbonate extractions only take 30–60 min and therefore represent a viable alternative. Moreover, it makes the IPD assay 8-fold more
sensitive on average, as a greater amount of exchangeable Pi is extracted by
bicarbonate than with water (Kleinman et al., 2001).
Underestimation of this labile Pi pool – even if specific activities
thereof are correctly measured – also causes underestimation of IPD rates,
given that Pi concentrations linearly affect IPD rates according to IPD Eqs. (1) and (2) above.
Microbial P dynamics
We observed very fast microbial Pi immobilization in live soils (within minutes; extraction started directly after tracer addition), causing net immobilization of 33P by 3 %–38 %. Similar results were reported within 1.5 to 4 h by others, ranging from 6 % to 37 % (Bünemann et al., 2012; Kellogg et al., 2006). This has two major repercussions. (i) Rapid uptake might cause microbial Pi assimilation and efflux or exudation of 33Po metabolites without microbial death and turnover. However, the comparison between specific activities and 33P recoveries of the direct measurement and after isobutanol fractionation (see below, and Fig. 3) showed that no significant release of microbial 33Po occurred during the 24 and 48 h incubations. The short extraction times used in this study also decrease the likelihood of significant hydrolysis of Po compounds. (ii) Rapid microbial 33Pi uptake clearly rules out the use of Po
mineralization assays that measure abiotic IEK in short-term batch
experiments (100 min) without the addition of a microbicide or without prior
sterilization and then extrapolate these “abiotic” process rates to the
full experimental duration.
Microbial Pi uptake can be derived indirectly as the difference in
33P recovery between live and sterile soils (Fig. 5, this study), more directly by sECE (this study), or by parallel water or bicarbonate extraction with and without the addition of liquid chloroform or hexane (measuring resin strip or extractable Pi), or by chloroform fumigation extraction (Bünemann et al., 2012; Oberson et al., 2001; Oehl et al., 2001a; Spohn and Kuzyakov, 2013). Microbial net 33P immobilization measured by direct sECE was higher relative to the difference in 33P immobilized in live minus sterile soils, pointing towards (i) overestimation of microbial net 33P immobilization by sECE due to incomplete extraction of nonmicrobial 33Pi by one-time bicarbonate extraction prior to sECE, or (ii) overestimation of abiotic sorption processes by autoclaving. In favor of (i),
repeated extractions of soils with Bray 1 showed that soils
continued to release P at lower rates in subsequent extractions after readily
extractable P was removed by the first extraction (Serrasolsas et al.,
2008; Messiga et al., 2014). Repeated extractions with bicarbonate also showed that the first extraction only removed 67 %–78 % of the 33Pi that was extractable with three consecutive extractions (D. Wasner, unpublished data, 2017). In favor of (ii), Kellogg et al. (2006) found
higher net 33P immobilization or sorption in sterile than in live soils. This was interpreted as a lack of microbial competition for P in
sterile soils. However, we found a weak positive relationship (R=0.749,
P=0.087; Table S1) between gross microbial Pi uptake and gross Pi sorption. This opposes the idea of strong competition between sorption and microbial uptake on the basis of gross process measurements. Another possible mechanism underlying (ii) could be changes in soil structure and reactive surfaces enhancing soil P sorption. Delineation of the causes could be performed by a comparison of sECE with liquid chloroform–fumigation
extraction (CFE), i.e., parallel assessments of microbial 33P uptake,
using a comparison of 33P in bicarbonate versus bicarbonate + liquid chloroform or bicarbonate + liquid hexane extracts. Given the continued extraction of Pi from exchangeable Pi pools in serial extraction tests, parallel determination of microbial P and 33P by CFE is recommended compared with sequential extractions by sECE.
Comparison of isobutanol fractionation with direct measurements of Pi and 33P activity
We showed that 33P IPD assays can be performed specifically on the
Pi pool using isobutanol fractionation in high-P soils. However, due to low production or persistence of 33Po, results closely conformed with measurements run without Pi-Po fractionation by malachite green and direct 33Ptotal estimates. This was ascertained for forest soils from French Guiana and Costa Rica, and for grassland soils from Austria and Germany (data not shown for French Guiana and Germany). Isobutanol fractionation has previously been applied in radiotracer studies on P dynamics in soils (Kellogg et al., 2006) and
litter (Mooshammer et al., 2012) to ascertain the separation of Pi from any possible radiolabeled Po contaminant,
although without comparison to SA in unfractionated bicarbonate extracts.
Oehl et al. (2001a) also applied isobutanol fractionation to
water extracts of fumigated and control soils, demonstrating that with long
extraction times (16 h) 33Pi activities in water extracts with and without isobutanol fractionation were comparable. It was suggested that
33Po possibly released during fumigation was cleaved by soil
phosphatases during extraction. This may not apply for short-term extractions
(e.g., 0.5 M NaHCO3 for 30 min, as used in this study) where hydrolysis by phosphatases would not necessarily occur due to short contact times. Measurements of 33P isotope pool dilution in soils based on bicarbonate extracts can therefore be interchangeably be performed by (i) direct
measurements of 33Ptot and Pi in acidified bicarbonate
extracts and after (ii) isobutanol fractionation on 33Pi and
Pi. However, this needs to be validated for other types of soil, and may change significantly after longer incubation periods (weeks), when microbial 33Pi uptake, assimilation and turnover causes the release of 33Po into the soil. The shortcut of performing direct measurements of Pi concentration and 33P in acidified bicarbonate extracts comes along with a 4- to 10-fold greater sensitivity of the malachite green assay relative to phosphomolybdate blue measurements of soil Pi. Another option to increase the measurement sensitivity for Pi (and possibly also for 33Pi) for strongly sorbing low-P soils has been adopted by Randriamanantsoa et al. (2013), and is based on the concentration of the phosphomolybdate blue complex from a large volume of extract into a smaller volume of hexane, with subsequent phase separation (Murphy and Riley, 1962). This allows for the quantification limits of Pi to be decreased 66-fold compared with the classical Murphy–Riley protocol, and 14-fold compared with the malachite green procedure (Randriamanantsoa et al.,
2013), but involves the handling of large volumes of radiolabeled extracts.
Time kinetics
During the first few minutes, equilibration between the tracer and tracee was not
achieved, which was indicated by the enhanced extractability of the added tracer
(33Pi) relative to more strongly bonded native tracee (soil
exchangeable Pi). The fast process of equilibration caused very rapid
declines in the SA of Pi during the first few minutes. Thereafter, microbial uptake and soil P fixation caused a rapid drawdown of extractable
33Pi and a further decrease in the SA of soil Pi, while soil Pi concentrations did not change after the initial phase of tracer–tracee equilibration (Fig. 4). These processes slowed down within the first 1–2 h but did not cease, and declines in 33P recoveries and in the SA of Pi occurred throughout the incubation period, in sterile as well as in live soils. Thereafter, time kinetics of IPD were relatively constant between 4 and 24 h for both temperate and tropical soils, as shown by the linearity of the relationship in a plot of ln(SA of Pi) versus time. This linear relationship is conceptually different from the plot of log(recovery, r/R) versus log(time) in short-term IEK batch experiments, which provides the parameter “n”, i.e., the slope or the rate of decline in tracer recovery due to sorption over time (Bünemann, 2015). Based on constant IPD rates in the abovementioned time interval we advise running 33P pool dilution experiments for an incubation period of 4 to 24 h. This time frame is well within the linear range, as it lies after the rapid abiotic equilibration, and is long enough to allow significant pool dilution to occur for sensitive measurements of organic P mineralization. Longer incubation times are not recommended due to the risk of 33Po
release from dying microbes, potentially causing a 33Pi reflux
via remineralization, violating a major assumption of IPD theory.
Comparison of Po mineralization rates with published values
The detection limit of the IPD approach was 0.12 µg P g-1 soil dw d-1. In comparison, the detection limits for gross Po
mineralization by the IEK approach were 0.20 µg P g-1 soil dw d-1 using the modified protocol including the hexane concentration of
phosphomolybdate blue for tropical soils (Randriamanantsoa et
al., 2015) and 0.6–2.6 µg P g-1 soil dw d-1 using the traditional IEK approach on temperate soils (Bünemann et al.,
2007). Values of gross Po mineralization measured via IPD in this study ranged between 0.08 and 0.15 µg P g-1 soil dw d-1 in tropical
forest soils and 0.48–2.03 µg P g-1 soil dw d-1 in temperate grassland soils and were, therefore, well within the range of those compiled for
IEK measurements by Bünemann (2015) for 14 different
soils, including temperate arable, grassland and forest soils (0.1–12.6 µg P g-1 soil dw d-1) and one tropical arable soil (0.8 µg P g-1 soil dw d-1). To date, the highest gross Po
mineralization rates have been reported for decomposing beech litter, i.e.,
22.5–86.3 µg P g-1 soil litter dw d-1
(Mooshammer et al., 2012). A direct comparison of the
present IPD and the IEK approaches on the same soils might help to clarify
how far the approaches really deviate or converge in their gross Po
mineralization rate estimates.
Physicochemical and biological controls on soil Pi processes
We found that gross Po mineralization was strongly positively correlated with total soil P but not to soil organic P, soil organic C, soil texture or soil acid phosphatase activity. This indicates that gross Po mineralization might be driven by total P rather than by soil enzyme activity, and that total soil Po does not represent the Po fraction accessible to soil phosphatases well. A few studies have demonstrated positive correlations between gross Po mineralization and soil Po
(Lopez-Hernandez et al., 1998) or litter Po (or its
inverse C : P; Mooshammer et al., 2012). However,
Wyngaard et al. (2016) did not find this relationship
between gross Po mineralization and total soil Po; however, they did note a relation with the Po content of the coarse soil fraction, which points in a similar direction to our results. Moreover, Po mineralization might rather be controlled by soil phosphodiesterases targeting DNA, RNA, teichoic
acids and phospholipids, than by phosphomonoesterases that are responsible
for the final extracellular dephosphorylation of Po. In contrast to our results, positive relationships were found between gross Po
mineralization and phosphomonoesterase activities in two studies (Spohn et
al., 2013; Oehl et al., 2004), although not across studies
(Bünemann, 2015). Thus, a larger set of soils varying in soil
pH, texture and mineralogy might provide better insights into the
controls of soil Po mineralization, such as effects by extracellular
phosphatase activity (phosphomonoesterases and phosphodiesterases), and the
availability, stabilization and accessibility of organic P in soils, among
others. Moreover, high Pi availability (i.e., bicarbonate Pi)
strongly suppressed phosphomonoesterase activity in soils, causing a negative
correlation between the enzyme activity and extractable Pi. In contrast, extractable Pi was positively related to gross Po mineralization, indicating that high-Pi conditions suppressed phosphatase production but not Po mineralization across these soils. This was also found as a positive correlation between gross Po mineralization and water-extractable Pi by others (Schneider et
al., 2017).
The contribution of gross Po mineralization to total Pi supply including Pi desorption from exchangeable Pi pools and dissolution ranged between 25 % and 74 %, with a trend towards larger contributions in low-P tropical soils (35 %–74 %) compared with temperate soils (25 %–51 %). This clearly demonstrates that biological processes contribute importantly to the Pi supply in soils, particularly in low-P soils, as also pointed out
by Bünemann (2015). In low-P forest soils biological
processes were shown to dominate over physicochemical processes, while in
P-rich forest soils abiotic processes controlled gross Pi supply rates (Bünemann et al., 2016). It was also found that the
contributions of microbial processes decreased with soil depth, whereas in deep soils diffusive fluxes (i.e., gross Pi desorption) dominated the soil
Pi supply due to low total Po contents relative to total P
(Achat et al., 2012, 2013).
Gross abiotic Pi release rates via desorption and dissolution were strongly positively related to total soil P and bicarbonate Pi, but not to other parameters such as soil pH, soil texture and soil organic C
content. In contrast to the weak effects of soil pH and texture on gross soil
Pi supply, soil mineralogy and particularly oxalate-extractable Fe and Al as proxies for Fe-Al (hydr)oxides play a major role in controlling abiotic dynamics of phosphate ions in soils, across the full range from acidic to alkaline soils (Achat et al., 2016). Fe-Al (hydr)oxides provide large positively-charged surface areas in weathered soils that are highly reactive to phosphate ions, more so than clay minerals such as kaolinite, illite and others (Hinsinger, 2001; Regelink et al., 2015). Therefore, soil mineralogy might provide further interesting insights into the controls of abiotic processes as demonstrated by (Achat et al., 2011, 2016), but can also affect Po mineralization via strong effects on the sorption strength of organic matter and of Po compounds. Moreover, the positive relations of Pi availability and Pi desorption with soil organic C contents reported elsewhere has been explained by competitive sorption of Pi and SOC or DOC to reactive surfaces such as positively charged
metal (hydr)oxides (Regelink et al., 2015; Achat et al., 2016).
Gross Pi sorption rates exceeded gross Pi desorption rates
approximately 2-fold but both were strongly related, indicating close and
rapid cycling of available Pi through sorption/desorption processes. The
observed rates indicate that soils immobilized more Pi then they
mobilized by abiotic processes, causing an intermediate drawdown of
available Pi pools. Two processes work against this drawdown of Pi in soils, i.e., Po mineralization and microbial P release via turnover and lysis. Moreover, plants (and microbes) might also desorb this sorbed Pi by the release of phytosiderophores and organic acids and thereby replenish Pi and reinject it into the organic P cycle. Similar to the soil C-N cycle, we might also expect an active “bank mechanism” regulating nutrient and C sequestration in soils (Fontaine et
al., 2011). At high nutrient availability priming effects are low, allowing
the sequestration of nutrients and SOC buildup. At low nutrient availability
microbes (and plants) release nutrients from SOM and from mineral surfaces
stimulated by root exudates, effectively mining inorganic and organic P
stored in soils.
The strong positive relationship between gross Pi sorption rates and
soil total P, soil total Pi and bicarbonate soil Pi, and the lack of a relationship with soil pH, soil organic C, clay content and soil texture highlights again that specific soil minerals, particularly metal (hydr)oxides and to a lesser extent clay minerals such kaolinite, which are factors not fully captured by soil pH and soil texture alone, are responsible for Pi sorption in soils (Regelink et al., 2015). In the IEK
experiments it was found that the rate of abiotic Pi depletion from soil solution via sorption was positively related to Al-Fe (hydr)oxide content and negatively related to soil organic C divided by the Al and Fe oxide content (Achat et al., 2016; Tran et al., 1988). The strong negative relation between abiotic net Pi immobilization and soil pH reconfirms that strongly weathered, acid tropical soils have a higher P sorption and fixation capacity than temperate soils.
Finally, gross microbial Pi uptake rates were directly proportional to microbial biomass P measured by sECE. We also found greater Pi
immobilization potentials through sorptive reactions (28 %–92 %) than through biological sinks (5 %–37 %) in the soils studied here. The importance of the rapid net uptake of tracer by soil microbes has also been demonstrated by other studies (e.g., Bünemann et al., 2012).
However, the presented IPD approach allowed for the estimation of
gross rates of microbial Pi uptake in addition to net microbial Pi immobilization for the first time. Gross rates of microbial uptake were calculated from the IPD approach, not necessitating the application of any extraction factor to calculate microbial biomass P from chloroform-labile P (kEP-factor). The
use of extraction factors becomes necessary when studying net Pi uptake over prolonged time periods in tracer experiments and for the correction of net Po mineralization rates (Bünemann, 2015; Bünemann et al., 2007).
Conclusion and outlook
The combination of this IPD assay with advanced numerical modeling
approaches, as applied by Müller and Bünemann (2014), might further enhance the precision of estimates of simultaneously
occurring soil P-cycle processes and thereby advance the knowledge of major
controls of the transformations and fluxes of this important nutrient in
terrestrial ecosystems. There is an ever-increasing need for high-quality data
on soil P processes, even more so to calibrate terrestrial biogeochemical
models and incorporate nutrient controls on plant productivity in global
models. This IPD approach may provide highly important quantitative data to
implement soil P-cycling processes into global biogeochemical models. This
will further enhance our current understanding of nutrient controls on the
global terrestrial C cycle and improve our capabilities to predict future
changes by increasing discrepancies in N and P inputs into the terrestrial
biosphere.
Data availability
The data of the different experiments are freely available
upon request from the corresponding author.
The supplement related to this article is available online at: https://doi.org/10.5194/bg-16-3047-2019-supplement.
Author contributions
The project was conceived and supervised by WW. DZ, JP
and DW performed the measurements and data evaluation. WW wrote the
paper with contributions from all coauthors.
Competing interests
The authors declare that they have no conflict of interest.
Acknowledgements
We are indebted to the Isotope Laboratory managers for
access and training (Virginie Canoine, Markus Schmid).
Review statement
This paper was edited by Sébastien Fontaine and reviewed
by two anonymous referees.
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