ArticlePDF Available

Priming and microbial nutrient limitation in lowland tropical forest soils of contrasting fertility

Authors:

Abstract and Figures

Priming is an increase in soil organic carbon decomposition following input of labile organic carbon. In temperate soils where biological activity is limited commonly by nitrogen availability, priming is expected to occur through microbial acquisition of nitrogen from organic matter or stimulated activity of recalcitrant-carbon degrading microorganisms. However, these priming mechanisms have not yet been assessed in strongly weathered tropical forest soils where biological activity is often limited by the availability of phosphorus. We examined whether microbial nutrient limitation or community dynamics drive priming in three lowland tropical forest soils of contrasting fertility (‘low’, ‘mid’ and ‘high’) by applying C4-sucrose (alone or in combination with nutrients; nitrogen, phosphorus and potassium) and measuring (1) the δ13C-signatures in respired CO2 and in phospholipid fatty acid (PLFA) biomarkers, and (2) the activities of enzymes involved in nitrogen (N-acetyl β-glucosaminidase), phosphorus (phosphomonoesterase) and carbon (β-glucosidase, cellobiohydrolase, xylanase, phenol oxidase) acquisition from organic compounds. Priming was constrained in part by nutrient availability, because priming was greater when sucrose was added alone compared to when added with nutrients. However, the greatest priming with sucrose addition alone was detected in the medium fertility soil. Priming occurred in parallel with stimulated activity of phosphomonoesterase and phenol oxidase (but not N-acetyl β-glucosaminidase); when sucrose was added with nutrients there were lower activities of phosphomonoesterase and phenol oxidase. There was no evidence according to PLFA δ13C-incorporation that priming was caused by specific groups of recalcitrant-carbon degrading microorganisms. We conclude that priming occurred in the intermediate fertility soil following microbial mineralization of organic nutrients (phosphorus in particular) and suggest that priming was constrained in the high fertility soil by high nutrient availability and in the low fertility soil by the low concentration of soil organic matter amenable to priming. This first study of priming mechanisms in tropical forest soils indicates that input of labile carbon can result in priming by microbial mineralization of organic nutrients, which has important implications for understanding the fate of organic carbon in tropical forest soils.
Content may be subject to copyright.
Priming and microbial nutrient limitation in lowland
tropical forest soils of contrasting fertility
Andrew T. Nottingham Benjamin L. Turner
Paul M. Chamberlain Andrew W. Stott
Edmund V. J. Tanner
Received: 24 August 2010 / Accepted: 2 August 2011 / Published online: 31 August 2011
ÓSpringer Science+Business Media B.V. 2011
Abstract Priming is an increase in soil organic
carbon decomposition following input of labile organic
carbon. In temperate soils where biological activity is
limited commonly by nitrogen availability, priming is
expected to occur through microbial acquisition of
nitrogen from organic matter or stimulated activity of
recalcitrant-carbon degrading microorganisms. How-
ever, these priming mechanisms have not yet been
assessed in strongly weathered tropical forest soils
where biological activity is often limited by the
availability of phosphorus. We examined whether
microbial nutrient limitation or community dynamics
drive priming in three lowland tropical forest soils of
contrasting fertility (‘low’, ‘mid’ and ‘high’) by
applying C
4
-sucrose (alone or in combination with
nutrients; nitrogen, phosphorus and potassium) and
measuring (1) the d
13
C-signatures in respired CO
2
and
in phospholipid fatty acid (PLFA) biomarkers, and (2)
the activities of enzymes involved in nitrogen (N-
acetyl b-glucosaminidase), phosphorus (phospho-
monoesterase) and carbon (b-glucosidase, cellobiohy-
drolase, xylanase, phenol oxidase) acquisition from
organic compounds. Priming was constrained in part
by nutrient availability, because priming was greater
when sucrose was added alone compared to when
added with nutrients. However, the greatest priming
with sucrose addition alone was detected in the
medium fertility soil. Priming occurred in parallel
with stimulated activity of phosphomonoesterase and
phenol oxidase (but not N-acetyl b-glucosaminidase);
when sucrose was added with nutrients there were
lower activities of phosphomonoesterase and phenol
oxidase. There was no evidence according to PLFA
d
13
C-incorporation that priming was caused by spe-
cific groups of recalcitrant-carbon degrading microor-
ganisms. We conclude that priming occurred in the
intermediate fertility soil following microbial miner-
alization of organic nutrients (phosphorus in particu-
lar) and suggest that priming was constrained in the
high fertility soil by high nutrient availability and in the
low fertility soil by the low concentration of soil
organic matter amenable to priming. This first study of
priming mechanisms in tropical forest soils indicates
that input of labile carbon can result in priming by
microbial mineralization of organic nutrients, which
has important implications for understanding the fate
of organic carbon in tropical forest soils.
A. T. Nottingham E. V. J. Tanner
Department of Plant Sciences, University of Cambridge,
Downing Street, Cambridge CB2 3EA, UK
A. T. Nottingham (&)
School of Geosciences, University of Edinburgh,
Drummond Street, Edinburgh EH8 9XP, UK
e-mail: anotting@staffmail.ed.ac.uk
B. L. Turner
Smithsonian Tropical Research Institute, Apartado
0843-03092, Balboa, Ancon, Republic of Panama
P. M. Chamberlain A. W. Stott
Centre of Ecology and Hydrology, Lancaster
Environment Centre, Lancaster LA1 4AP, UK
123
Biogeochemistry (2012) 111:219–237
DOI 10.1007/s10533-011-9637-4
Keywords Carbon dioxide Phenol oxidase
Phospholipid fatty acids Phosphomonoesterase
Priming effect Soil carbon
Introduction
Inputs of labile organic compounds to soil can
stimulate microbial mineralization of pre-existing
organic matter through ‘priming effects’ (Fontaine
et al. 2004). Priming effects have the potential to
feedback positively on climate change by contribut-
ing an additional source of atmospheric CO
2
. A net
input of atmospheric CO
2
would arise if environ-
mental change increases plant productivity and labile
carbon (C) inputs to soils, and these inputs are then
exceeded by soil CO
2
emissions due to priming.
Despite the large potential for priming to affect the
global C cycle they remain poorly understood,
especially in tropical forest soils. Tropical forests
contain 30% of global soil C (Jobbagy and Jackson
2000) and may already be subject to increased inputs
of labile C due to increased aboveground production
resulting from environmental change (Phillips et al.
2008), yet no studies have investigated priming
mechanisms in these ecosystems. Furthermore, prim-
ing mechanisms in tropical soils may differ markedly
to temperate soils because they are generally more
strongly weathered, with consequences for the nutri-
ent limitation to soil microorganisms (Walker and
Syers 1976; Cleveland et al. 2006).
Two mechanisms have been hypothesized to lead
to priming. First, priming may occur when microor-
ganisms become nutrient-limited during the degrada-
tion of new labile C and co-metabolize pre-existing
soil organic matter to meet their nutrient demands,
leading to soil C being mineralized and released as
‘primed’ CO
2
(Blagodatskaya and Kuzyakov 2008).
Supporting evidence for this mechanism is provided
by laboratory experiments on temperate soils, which
measured increased activity of organic nitrogen (N)-
degrading enzymes during priming (Asmar et al.
1994) and a reduction in priming when N was added
with labile C, due to a switch from soil C to added
labile C as the preferred substrate (‘preferential
substrate utilization’; Hagedorn et al. 2003; Blagod-
atskaya et al. 2007).
Second, priming may occur when microorganisms
that specialize in the degradation of stable soil C are
able to compete with other microorganisms and
utilize part of the new labile C. This mechanism is
thought to occur in nutrient-poor soils where slow-
growing soil C-specialist microorganisms are able to
gain a competitive advantage over fast-growing labile
C-specialist microorganisms. According to this the-
ory, the soil C-specialists increase in abundance by
utilizing some of the new labile C and persist to
degrade stable soil C when the labile C becomes
diminished (Fontaine et al. 2003). Supporting evi-
dence for this mechanism is provided by studies that
measured prolonged priming after supply of added
labile C is exhausted (Fontaine et al. 2004), and
priming alongside an increased dominance of the
microbial community by fungi (Carney et al. 2007).
A predictive understanding of priming remains
elusive in part because few studies have measured
changes in microbial community composition and
biochemical activity during priming (Blagodatskaya
and Kuzyakov 2008). Priming due to co-metabolism of
organic matter by nutrient-limited microorganisms can
be identified by an increase in the microbial synthesis of
extracellular enzymes to liberate nutrients from organic
matter (Kuzyakov et al. 2000; Schimel and Weintraub
2003; Blagodatskaya and Kuzyakov 2008). The
enzyme N-acetyl b-glucosaminidase is mostly present
in soils in extracellular forms (Parham and Deng 2000)
and can be produced by microorganisms in response to
N-deficiency to acquire N from organic matter (Sin-
sabaugh and Moorhead 1994; Olander and Vitousek
2000; Muruganandama et al. 2009). Phosphomonoes-
terase is released by microorganisms in response to
phosphorus (P) deficiency to hydrolyze ester bonds
between C and P in organic matter and liberate
orthophosphate (Quiquampoix and Mousain 2005).
Activity of these enzymes can indicate microbial
mineralization of nutrients from organic matter due to
N or P demand (Olander and Vitousek 2000;Allison
et al. 2007; Sinsabaugh et al. 2008) and consequently
may provide a useful indication of nutrient limitation
during priming. Priming due to increased abundance of
recalcitrant-C degrading microorganisms can be iden-
tified by a shift in microbial community composition
(e.g. increase in fungal:bacterial ratios; Carney et al.
2007), prolonged priming following the exhaustion of
the added substrate, and greater priming in low fertility
soils (Fontaine et al. 2003,2004).
220 Biogeochemistry (2012) 111:219–237
123
Priming has been suggested as the cause of reduc-
tions in soil C in temperate forest grown under
experimentally elevated CO
2
, despite increased inputs
of plant-C to soils (Carney et al. 2007; Langley et al.
2009). The parallel loss of soil C and increase in N
mineralization in the study by Langley et al. (2009)
suggests that priming occurred due to microbial
‘co-metabolism’ of organic matter to acquire N
(Kuzyakov et al. 2000), which is often considered the
limiting nutrient in temperate forests. In contrast,
studies performed in tropical forests suggest that
decomposition is limited by the availability of P
(Hobbie and Vitousek 2000; Kaspari et al. 2008). Many
tropical forest soils are strongly weathered and, while
they have abundant available N (Martinelli et al. 1999),
contain little available P due to its occlusion within
secondary minerals and immobilization within organic
matter (Walker and Syers 1976; Vitousek and Sanford
1986). Although the generalization is unlikely to hold
true for all tropical forests due to varying rates of rock
weathering and dust deposition (Porder and Hilley
2011), P-limitation of microbial C metabolism has
been demonstrated for numerous tropical forests where
experimental P-addition increased microbial mineral-
ization of dissolved organic matter (Cleveland et al.
2006) and increased soil CO
2
efflux (Cleveland and
Townsend 2006), and where elevated P concentration
in litter increased decomposition rates (Hobbie and
Vitousek 2000). Another study concluded P-limitation
of microbial metabolism of C in a tropical soil when
CO
2
efflux increased more rapidly following an
addition of C with P compared to C with N (Gnan-
kambary et al. 2008).
We asked whether priming effects occur due to (1)
co-metabolism of soil organic matter due to microbial
nutrient limitation, and/or (2) increased abundance of a
specific group of soil C-degrading microorganisms. To
investigate this, we assessed priming effects following
sucrose additions with and without mineral nutrients to
soils of contrasting fertility from three different
lowland tropical forests in the Republic of Panama.
Materials and methods
Soils
Soils were collected from three 1 ha lowland tropical
forest plots from the Center for Tropical Forest Science
(CTFS) network in the Republic of Panama (Pyke et al.
2001; Turner and Engelbrecht 2011). We selected sites
at Rio Paja (‘low-fertility’—plot 26 in Turner and
Engelbrecht 2011), Pipeline Road (‘mid-fertility’—plot
15 in Turner and Engelbrecht 2011)and Campo Chagres
(‘high-fertility’)due to their marked variationin nutrient
status; total C, N and P increased in the order: Rio
Paja \Pipeline Road \Campo Chagres (Table 1;
plotted in context to other CTFS and RAINFOR forest
sites in Fig. 1). Detailed information on the P compo-
sition of the soils is reported elsewhere (Turner and
Engelbrecht 2011). Soils at Rio Paja are derived from
fine grained rhyolitic tuff, soils at Pipeline Road are
derived from marine sedimentary parent material of the
Gatuncillo Formation, and soils at Campo Chagres are
derived from calcareous sandstone of the Alajuela
Formation. For each site, surface soil (0–10 cm) was
collected from 30 random locations using a 2.5 cm
diameter soil corer. Within 24 h of collection, soils were
returned to the laboratory and visible stones and roots
were removed by hand. Soils were then mixed, sieved
(\2 mm) to isolate the fine earth fraction, weighed
(500 g air-dry weight) into PVC containers (16 cm
diameter, 10 cm height), adjusted to 60% water holding
capacity and pre-conditioned at 22°C for 10 days prior
to experimental treatments. The temperature and mois-
ture content (by daily watering) were then maintained
throughout the experiment.
Experimental design
Treatments were a sugar-cane sucrose solution and
a combined nutrient solution (N in NH
4?
and NO
3-
,
P and K in KH
2
PO
4
) applied as follows: control (no
addition; CTL), nutrient addition (NPK), sucrose
addition (C) and sucrose and nutrient addition (CNPK).
Additions of sucrose and nutrients on the basis of dry
soil were: 4000 mg C kg
-1
, 100 mg N kg
-1
, 100 mg P
kg
-1
, and 130 mg K kg
-1
. Additions of C were within
the range of annual input of C to tropical forest soils.
For example, nearby tropical forest soils in Panama
receive 470–650 g litterfall-C ha
-1
year
-1
(Wieder
and Wright 1995; Kaspari et al. 2008), equivalent to
2300–3300 mg C kg
-1
soil year
-1
from litterfall alone
(assuming that the majority is mineralized within the
top 20 cm of soil with a bulk density of 1 g cm
-3
).
Treatments were allocated according to a random-
ized block design with four replicates of each
treatment and four controls and stratified according
Biogeochemistry (2012) 111:219–237 221
123
to untreated soil CO
2
efflux. Sucrose and nutrients
were applied once in 20 ml solutions (20 ml C
solution and/or 20 ml NPK solution) during daily
watering of soils. Measurements of CO
2
efflux and
d
13
C values in CO
2
efflux were made on 10 occasions
for the low-fertility soil (days 0, 1, 2, 3, 4, 6, 8, 10,
14, 18); seven occasions for the mid-fertility soil
(days 0, 1, 2, 3, 4, 6, 8); and nine occasions for the
high-fertility soil (days 0, 1, 2, 3, 4, 6, 8, 10, 12).
Measurements for the low-fertility soil continued for
18 days because of the unexpectedly long time it took
for the added sucrose to decompose and measure-
ments for the mid-fertility soil discontinued at 8 days
because of equipment problems.
Soil samples (0–2 cm depth) were taken 3 days
after treatments were imposed and analyzed for
extractable nutrients, microbial nutrients, enzyme
activities, and PLFAs. We sampled soil 3 days after
treatments were imposed to quantify chemical and
biological properties during priming effects, rather
than during the peak of priming, which could only be
determined retrospectively. Soils were extracted for
nutrients on the day of sampling to minimize the
rapid changes in nutrients that can occur during
storage (Turner and Romero 2009).
Soil CO
2
efflux and d
13
C determination
Soil CO
2
efflux was measured using a Li-8100 soil
respiration system (infra red gas analyzer; Li-Cor,
Lincoln, Nebraska, USA). To sample soil CO
2
efflux
for determination of d
13
C we used 4-l static chambers
constructed from thick PVC and fitted over each
container to give an air tight seal (Bertolini et al.
2006). Air samples were collected during a near-
linear increase in chamber CO
2
concentration, char-
acterized in preliminary tests at different rates of soil
CO
2
efflux, with the first sample extracted 3 min after
chamber placement and the final sample extracted at
2–3 times ambient CO
2
. Samples for determination of
d
13
C values were collected in 12 ml exetainers, wax-
sealed, shipped to Lancaster, UK and analyzed within
approximately 3 months. The d
13
C values of CO
2
samples were determined by isotope ratio mass
spectrometry using a Micromass TraceGas Pre-con-
centrator coupled to an Isoprime isotope ratio mass
spectrometer (Micromass, Wythenshawe, UK). A
further 20 ml of soil chamber air was extracted and
analyzed for CO
2
concentration using a gas chro-
matograph (Shimadzu GC-14B, Columbia, MD,
USA) equipped with an electron capture detector
(Loftfield et al. 1997) which was calibrated with four
Fig. 1 Tropical forest sites in Panama (including sites from
this study: LOW, MID, HIGH; and other CTFS sites: C1–C16)
and across the Amazon (RAINFOR sites: R1–R11), distributed
according to soil fertility as determined by total concentration
of nitrogen and phosphorus (measured at 0–10 cm depth).
CTFS site descriptions are given in Turner and Engelbrecht
(2011): LOW (Rio Paja P26), MID (Pipeline P15), HIGH
(Campo Chagres), C1 (Rio Paja P25), C2 (Pipeline P09), C3
(Sherman P02), C4 (Pipeline P08), C5 (Buena Vista P12), C6
(Santa Rita P32), C7 (Albrook), C8 (Pipeline P17), C9
(Mocambo), C10 (Cerro Torre), C11 (Las Cruces P27), C12
(Buena Vista P13), C13 (Sherman P01), C14 (BCNM P18),
C15 (Gamboa P24), C16 (Cerro Galera). RAINFOR site
descriptions are given in Quesada et al. (2011): R1 (SUC-02),
R2 (CUZ-03), R3 (HCC-21), R4 (CHO-01), R5 (BOG-02),
R6 (ELD-12), R7 (CAX-02), R8 (TAP-04), R9 (SIN-01),
R10 (JUR-01), R11 (MAN-12)
Table 1 Site locations, total soil carbon, total nutrients, pH
and soil texture for untreated soils
Low-fertility Mid-fertility High-fertility
C(gkg
-1
) 28.1 40.1 104.0
N(gkg
-1
) 2.1 3.5 8.9
P (mg kg
-1
) 71 200 696
C/N 13.4 11.5 11.7
C/P 396 201 149
N/P 30 18 13
pH 4.4 6.3 6.8
Sand (%) 4 38 32
Silt (%) 62 27 21
Clay (%) 33 36 47
Soils were designated: ‘low-fertility’ (Rio Paja), ‘mid-fertility’
(Pipeline Road) and ‘high fertility’ (Campo Chagres). Data are
single analyses of pooled sub-samples; soil texture data are
from Turner and Engelbrecht (2011)
222 Biogeochemistry (2012) 111:219–237
123
standard gases (360, 706, 1505 and 5012 ppm CO
2
Deuste Steininger GmbH, Mu
¨hlhausen, Germany).
Soil nutrients, enzymes and microbial biomass
Soil inorganic N (in NO
3-
and NH
4?
) and microbial
C and N were determined by K
2
SO
4
extraction.
Microbial C and N were determined as the difference
between chloroform-fumigated and unfumigated soil
samples following a 24 h fumigation period (Vance
et al. 1987) and corrected for efficiency of the
extraction procedure with k-factors of 0.45 for C (Wu
et al. 1990) and 0.54 for N (Joergensen and Mueller
1996). Total C and N in the extracts were determined
by combustion and gas chromatography using a
Thermo-Electron Flash 1112 Elemental Analyzer
(CE Elantech, Lakewood, NJ). Readily-exchangeable
phosphate (extractable P) and microbial P were
determined by extraction with anion-exchange mem-
branes and hexanol fumigation based on the method
described by Kouno et al. (1995). Phosphate was
recovered from anion-exchange membranes by shak-
ing for 1 h in 50 ml of 0.25 M H
2
SO
4
, with detection
in the acid solution by automated molybdate color-
imetry using a Lachat Quickchem 8500 (Hach Ltd,
Loveland, CO, USA). Extractable P was determined
by P recovered from unfumigated samples and
microbial P was calculated as the difference between
the fumigated and unfumigated samples.
Five enzymes involved in C and nutrient cycling
were measured using microplate fluorimetric assays
with 200 lM methylumbelliferone (MU)-linked sub-
strates (Marx et al. 2001): b-glucosidase (degradation
of labile C), cellobiohydrolase (degradation of cellu-
lose), N-acetyl b-glucosaminidase (degradation of
N-glycosidic bonds), phosphomonoesterase (degrada-
tion of monoester-linked simple organic phosphates)
and xylanase (degradation of hemicellulose). Soil
samples were collected from soil containers to 2 cm
depth, stored at 3°C and assayed within 3 days, which
does not appear to greatly alter observed activities in
tropical forest soils (Turner and Romero 2010). On the
day of the assay, 2 g soil (dry weight basis) was added
to 200 ml 1 mM NaN
3
solution and dispersed by
stirring on a magnetic stir plate. After 5 min and while
stirring, 50 ll aliquots of soil suspension were
removed using an 8-channel pipette and dispensed
into a 96-well microplate containing 50 ll modified
universal buffer solution (Tabatabai 1994) adjusted to
soil pH. Each microplate included assay wells (soil
solution plus 100 ll MU substrate), blank wells (soil
solution plus 100 ll of 1 mM NaN
3
) and quench wells
(soil solution plus 100 ll MU standard). A further
control plate was prepared with MU substrates and
standards with no soil solution to determine quenching
by soil solution in assay plates. There were eight
analytical replicate wells for each assay. Microplates
were incubated at 22°C for either 1 h (b-glucosidase,
N-acetyl b-glucosaminidase, phosphomonoesterase)
or 4 h (cellobiohydrolase, xylanase). Following incu-
bation, 50 ll of 0.5 M NaOH was added to terminate
the reaction and plates were immediately analyzed on a
Fluostar Optima spectrofluorometer (BMG Labtech,
Offenburg, Germany) with excitation at 360 nm and
emission at 450 nm.
A further enzyme, phenol oxidase (degradation of
phenolic compounds), was measured using 5 mM L-
dihydroxyphenyalanine (L-DOPA) as substrate (e.g.
Waldrop and Firestone 2004a). Briefly, 1 g soil
(oven-dry basis) was added to 100 ml of 5 mM
bicarbonate buffer and mixed well; 100 llof5mM
L-DOPA solution and 100 ll of soil solution were
then added to a 96-well plate. Control plates were
made using 100 ll of 5 mM bicarbonate buffer and
100 ll aliquots of soil solution. There were 16
analytical replicates and controls per soil sample.
Plates were analyzed on a Fluostar Optima spectro-
fluorometer (BMG Labtech, Offenburg, Germany),
with phenol oxidase activity calculated as the
increase in absorbance at 450 nm over 1 h.
Phospholipid fatty acids (PLFA)
Phospholipid fatty acids were extracted from freeze
dried soils using the method of Crossman et al.
(2004). Following sampling (3 days after treatments
were imposed), soils were stored at -35°C for
3–6 months and then freeze-dried approximately
1 month prior to PLFA extraction. PLFA fingerprints
were identified and quantified by gas chromatography
(GC). The d
13
C values of PLFA were determined
using gas-chromatography-combustion-isotope ratio
mass spectrometry (GC-C-IRMS). All analytical
conditions are described in Chamberlain et al.
(2006). Molecular structures of PLFA are described
using standard nomenclature: the first number refers
to the total number of C-atoms and the number
after the colon refers to the number of double bonds.
Biogeochemistry (2012) 111:219–237 223
123
A number following a x is the location of the first
double bond relative to the aliphatic end of the
molecule. Notations ‘Me’, ‘OH’ and ‘cy’ are,
respectively, methyl, hydroxy, cyclopropane groups
and notations ‘i’ and ‘a’ respectively are iso- and
anteiso-branched fatty acids. PLFA biomarkers were
grouped as Gram-positive bacterial (15:0, i15:0,
a15:0, i16:0, i17:0, a17:0, 7Me17:0), Gram-negative
bacterial (16:1x5, 16:1x7, 17:1x8, 7,9cy17:0,
18:1x7, 7,8cy19:0, 19:1), fungal (18:2x6, 18:1x9)
and non-specific saturated (14:0, 16:0, 18:0; Frost-
ega
˚rd and Ba
˚a
˚th 1993; Zelles 1999).
Calculations and statistics
The percent of respired CO
2
originating from added
sucrose was calculated according to:
%C
sucrosederived ¼dCdT
dCdL

100 ð1Þ
where d
C
is the d
13
C value of the respired CO
2
from
control soils, d
T
is the d
13
C value in respired CO
2
from treated soils and d
L
is the d
13
C value of the
sucrose C (e.g. Balesdent et al. 1988). The increase in
soil C mineralization (primed CO
2
) was then calcu-
lated from the increase in soil respiration in treat-
ments relative to controls minus the contribution of
sucrose C (Kuzyakov et al. 2000). We also used a
variation of Eq. 1to calculate the mass of C within
individual PLFAs at day three. The proportion of
sucrose-derived C within each PLFA was calculated
using Eq. 1, where d
C
is the d
13
C value of a specific
PLFA from untreated control soils, d
T
is the d
13
C
value for the same PLFA from treated soils and d
L
is
the d
13
C value of the sucrose (e.g. Nottingham et al.
2009). Real priming effects (extra soil-derived CO
2
from soil organic matter) were distinguished from
apparent priming effects (extra soil-derived CO
2
from
substitution of soil-labeled with sucrose-labeled C in
microbial biomass) by comparing total primed CO
2
-C
with microbial C and by examining changes of soil-
labeled C within PLFA biomarkers. For each parti-
tioned C-source we calculated standard errors from
summed respective variances. Cumulative evolution
of CO
2
-C was calculated by integrating the area
beneath the CO
2
efflux versus time curve.
We used repeated measures ANOVA to study the
variation in total soil CO
2
efflux and partitioned
components of soil CO
2
efflux with time between
three treatments and a control. For all other analyses
one-way ANOVAs were used with either soil type or
treatment as the factor and respective response
variables were: cumulative CO
2
evolution, PLFA
concentration, quantity of sucrose-derived C or soil-
derived C within PLFAs, concentration of PLFA
biomarker groups (Gram-positive, Gram-negative,
fungal) and fungal/bacterial ratios (according to the
ratio of 18:2x6,9/bactPLFAs; Bardgett et al. 1996).
Pair-wise comparisons were made using post-hoc
Tukey HSD analysis and significant differences were
determined at PB0.05. Treatment effects on the
concentrations of soil and sucrose-derived C within
PLFA were further examined using Principal Com-
ponents Analysis (PCA) to construct new variables
from multivariate PLFA data sets. Prior to analysis,
data were tested for normality using a Ryan-Joiner
test and non-normal data were log-transformed. All
statistical analyses were performed using Minitab
(version 15, Minitab Inc., PA, USA).
Results
Total, sucrose and soil-derived soil CO
2
efflux
Additions of C and CNPK led to significant increases
in total soil CO
2
efflux for all soils (P\0.001;
Fig. 2). For CNPK compared to C additions, the peak
in total soil CO
2
efflux was higher and occurred
earlier for low-fertility and mid-fertility soils but was
higher and occurred at the same time for the high-
fertility soil (Fig. 2). Following C additions, total soil
CO
2
efflux peaked after 10, 3 and 2 days and
following CNPK additions after 3, 1, and 2 days for
low-, mid- and high-fertility soils respectively
(Fig. 2). For CNPK compared to C additions, the
cumulative evolution of total C over 8 days was
significantly higher for all soils (P\0.001; Fig. 3).
Addition of NPK alone had no significant effect on
total soil CO
2
efflux for any soil (P[0.90; data not
shown).
Priming, identified by a significant increase in soil-
derived CO
2
efflux in treated relative to control soils,
occurred following C additions for all soils (for all
comparisons, P\0.05; Fig. 2). The peak in primed
CO
2
efflux following C additions occurred over
0–10 days for low-fertility, 0–4 days for mid-fertility
and 0–3 days for high-fertility soils, and was prior to
224 Biogeochemistry (2012) 111:219–237
123
the peak in sucrose-derived CO
2
for all soils (Fig. 2).
The cumulative evolution of primed CO
2
-C over
8 days following C additions was higher for soils in
the order low-fertility \high-fertility \mid-fertility
(the portion of soil-derived CO
2
-C in excess of basal
CO
2
-C in Fig. 3). For CNPK compared to C
additions, primed CO
2
efflux was significantly lower
for all soils (Figs. 2,3).
The peak in sucrose-derived CO
2
efflux was higher
for all soils and occurred earlier for low-fertility and
Fig. 2 Total CO
2
-C efflux (lgCO
2
-C g
-1
soil h
-1
) following
treatments was partitioned into basal soil-derived C, sucrose-
derived C, primed soil-derived C for low-fertility (a,b), mid-
fertility (c,d) and high-fertility (e,f) soils following ?C and ?
CNPK additions. Data are means ±1 standard error (n =4)
Biogeochemistry (2012) 111:219–237 225
123
mid-fertility soils following CNPK compared to C
additions (Fig. 2). Cumulative evolution of sucrose-C
8 days following the addition of 4 mg sucrose-C g
-1
was higher for soils in the order low-fertility\high-
fertility \mid-fertility, with 0.50, 0.72 and 1.00 mg
CO
2
-C g
-1
respired respectively. Following CNPK
additions, cumulative evolution of sucrose-derived C
was higher for soils in the order high-fertility\low-
fertility \mid-fertility, with respiration of 0.97, 1.18,
1.42 mg CO
2
-C g
-1
, respectively; significantly higher
compared to C additions for all soils (P\0.01; Fig. 3).
Sucrose-derived CO
2
-C evolution following addition of
CNPK compared to C alone was proportionally higher
for soils in the order high-fertility (35%) \mid-fertility
(46%) \low-fertility (136%) (Fig. 3).
Soil nutrients
Concentrations of NO
3-
and extractable P in untreated
soils were higher in the order low-fertility \mid-fertil-
ity \high-fertility, with significant differences between
all soil types (P\0.05; Table 2). There was an excep-
tion to this pattern for NH
4?
, which was higher in soils in
the order low-fertility =high-fertility \mid-fertility,
with significantly higher concentrations of NH
4?
in
mid-fertility compared to other soils (P\0.05).
The conce ntrations of NO
3-
,NH
4?
and extractable P
were generally lower in soil where sucrose-C was added
alone compared to all other treatments, with the
exception of extractable P in the low-fertility soil, in
which concentrations in untreated soils were already
very low (Table 2). Following NPK and CNPK addi-
tions, there were significantly higher concentrations of
NO
3-
,NH
4?
and extractable P for all soils (P\0.01)
except NH
4?
following CNPK addition in the low-
fertility soil (P=0.28), relative to controls (Table 2).
Soil pH was significantly different in some treated
soils relative to controls: higher for low-fertility
following CNPK additions, higher for the high-fertility
soil following C and CNPK additions, and lower for the
low-fertility soil following NPK additions (Table 2).
Microbial biomass
The concentrations of microbial C and microbial N in
untreated soils increased in the order low-fertil-
ity \mid-fertility \high-fertility, with significant
differences between all soils (P\0.05; Fig. 4); while
microbial P concentrations increased in the order:
low-fertility =mid-fertility \high-fertility with sig-
nificantly higher concentrations of microbial P in the
high-fertility soil compared to other soils (for all
comparisons, P\0.05). It was notable that, although
the low-fertility soil had a concentration of extractable
P four-fold lower than the mid-fertility soil (0.7 cf.
2.9 mg kg
-1
), there was no significant difference in
microbial P between the two soils (P=0.20).
Microbial C was significantly higher in CNPK
compared to C treated soils for low- and high-fertility
(P\0.05), but not mid-fertility soils (P[0.05;
Fig. 4). Microbial N was significantly higher in the
CNPK treated low-fertility soil, but was significantly
lower in CNPK treated high-fertility and NPK treated
mid-fertility soils relative to untreated controls (for all
comparisons, P\0.05; Fig. 4). Microbial P was
significantly higher in all CNPK treated soils relative
to untreated controls (for all comparisons, P\0.05;
Fig. 4). The ratio of microbial C:N was significantly
higher in CNPK treated high-fertility soils (P\0.05)
and there was a trend of higher ratios of microbial
C:P in all C treated soils, relative to controls.
Enzyme activities
There were many significantly lower enzyme activ-
ities in treated soils relative to control soils. In
Fig. 3 Cumulative sucrose- and soil-derived CO
2
-C evolution
over 8 days following treatments. Each pot received 4 mg
sucrose-C g
-1
soil. Primed CO
2
-C is the portion of soil-derived
CO
2
-C in excess of basal CO
2
-C; basal CO
2
-C, represented by
the dashed reference lines through the soil-derived compo-
nents, was constant for each soil and assumed equal to total
CO
2
-C in control (CTL) treatments. Significant differences in
the evolution of sucrose- and soil-derived CO
2
-C between
treatments for each soil type are highlighted by asterisks
(PB0.05). Data are means ±1 standard error (n =4)
226 Biogeochemistry (2012) 111:219–237
123
particular, the low-fertility soil had lower enzyme
activities following NPK addition (b-glucosidase,
N-acetyl b-glucosaminidase, phosphomonoesterase,
P\0.001); following C addition (b-glucosidase,
cellobiohydrolase, P\0.01; phosphomonoesterase,
xylanase, P\0.001) and following CNPK addition
(phosphomonoesterase, xylanase P\0.001) com-
pared to controls (Table 3). In the mid-fertility soil
there was significantly lower xylanase activity fol-
lowing C addition (P\0.05) and in the high-fertility
soil significantly lower cellobiohydrolase and xylan-
ase activity following C addition (P\0.05) and
b-glucosidase activity following CNPK addition
(P\0.05), compared to controls.
There were also several significantly higher enzyme
activities in treated soils relative to control soils. In the
low-fertility soil, phenol oxidase was higher following
C addition (P\0.05). In the mid-fertility soil, phos-
phomonoesterase and phenol oxidase were higher
following C addition (P\0.05) and N-acetyl b-
glucosaminidase was higher following CNPK addition
(P=0.05). In the high-fertility soil there were signif-
icantly higher activities for b-glucosidase following
NPK addition (P\0.05), N-acetyl b-glucosaminidase
following CNPK addition (P=0.05), and phospho-
monoesterase following C additions (P\0.001).
PLFA concentration and carbon incorporation
Total concentration of PLFA was similarly high in
low- and high-fertility soils; both were significantly
higher than the mid-fertility soil (P\0.001;
Table 4). Fungal:bacterial ratios increased in soils
in the order mid-fertility (0.08) \high-fertility
(0.12) \low-fertility (0.16). Gram-positive bacteria
PLFA concentrations increased in soils in the order
low-fertility \mid-fertility \high-fertility, whereas
Gram-negative bacteria PLFA and fungal PLFA
concentrations increased in the order mid-fertil-
ity \high-fertility \low-fertility. There were no
significant differences in total PLFA concentration
following any treatments; the only microbial group
that was significantly different following treatments
was the higher concentration of fungal PLFA bio-
markers in the low-fertility soil following CNPK
additions (P\0.001; Fig. 5; Table 4). Accordingly,
fungal:bacterial ratios were not significantly different
following treatments among soils except for low-
fertility soil following CNPK additions, where they
increased from 0.16 to 0.52 (P\0.001; Table 4).
Following C addition, there was significant sucrose-
derived C incorporation in the low-fertility soil for
a saturated (18:0) biomarker; mid-fertility soil for
Table 2 Nutrients and pH in surface soils measured 3 days following treatments
Low-fertility Mid-fertility High-fertility
NO
3-
mg N kg
-1
CTL 1.8 ±0.4 8.0 ±0.4 22.3 ±0.7
?NPK 31.7 ±1.9* 47.8 ±1.3* 91.1 ±3.0*
?C 0.2 ±0.1 1.0 ±0.1* 0.1 ±0.1*
?CNPK 22.2 ±2.2* 23.6 ±07* 45.3 ±1.1*
NH
4?
mg N kg
-1
CTL 0.3 ±0.1 1.7 ±0.3 0.3 ±0.1
?NPK 45.0 ±2.2* 26.5 ±1.2* 83.9 ±3.4*
?C 0.1 ±0.1 0.3 ±0.1 0.2 ±0.1
?CNPK 1.1 ±0.7 7.0 ±0.6* 14.5 ±2.3*
Extractable P mg P kg
-1
CTL 0.7 ±0.1 2.9 ±0.1 4.6 ±0.2
?NPK 234.3 ±24.5* 102.2 ±11.4* 461.9 ±33.0*
?C 0.7 ±0.1 0.6 ±0.2 0.9 ±0.2
?CNPK 179.3 ±14.9* 92.9 ±22.8* 256.0 ±20.1*
pH CTL 4.4 ±0.1 6.3 ±0.1 6.8 ±0.1
?NPK 4.2 ±0.1 5.6 ±0.1* 6.8 ±0.1
?C 4.5 ±0.1 6.1 ±0.1 7.1 ±0.1*
?CNPK 4.7 ±0.1* 6.3 ±0.1 7.0 ±0.1*
Significant differences between treatments and controls are highlighted by asterisks (where PB0.05). Data are means ±1 standard
error (n =4)
Biogeochemistry (2012) 111:219–237 227
123
Gram-positive (i15:0, 7Me17:0), Gram-negative
(16:1x7, 18:1x7), saturated (16:0, 18:0) and fungal
(18:1x9) biomarkers; and high-fertility soil for Gram-
positive (a15:0, i16:0, i17:0), Gram-negative (16:1x7)
saturated (16:0) and fungal (18:2x6, 18:1x9) biomark-
ers (for all comparisons, P\0.001; Fig. 5). Following
CNPK addition, sucrose-derived C was significantly
incorporated into PLFA in the low-fertility soil for
Gram-negative (16:1x7, 19:1), saturated (16:0, 18:0)
and fungal (18:1x6, 18:1x9) biomarkers with major
incorporation into fungal biomarkers (P\0.001;
Fig. 5). In mid- and high-fertility soils CNPK additions
led to significant sucrose-derived C incorporation in
broad ranges of biomarkers: in Gram-positive (i15:0,
a15:0, i16:0), Gram-negative (16:1x7, 18:1x7), satu-
rated (16:0, 18:0) and fungal (18:2x6, 18:1x9) bio-
markers in the mid-fertility soil; and in Gram-positive
(i15:0, a15:0, i16:0, i17:0), Gram-negative (18:1x7),
saturated (16:0) and fungal (18:2x6, 18:1x9) biomark-
ers in the high-fertility soil (P\0.001; Fig. 5).
Soil-derived C concentrations in PLFA for treated
soils compared to controls were only significantly
different following CNPK additions. There was
significantly less soil-derived C in Gram-negative
Fig. 4 Microbial carbon (a), nitrogen (b) and phosphorus (c);
and the ratios of carbon/nitrogen (d) and carbon/phosphorus
(e) in surface soils measured 3 days following treatments.
Significant differences (PB0.05) are shown by asterisks
between ?CNPK and ?C treated soils. Data are means ±1
standard error (n =4)
228 Biogeochemistry (2012) 111:219–237
123
Table 3 Enzyme activities in soils three days following treatments
N-cycling P-cycling C-cycling
N-acetyl glucosaminidase
(nmol MU min
-1
g
-1
)
Phosphomonoesterase
(nmol MU min
-1
g
-1
)
b-Glucosidase
(nmol MU
min
-1
g
-1
)
Cellobiohydrolase
(nmol MU
min
-1
g
-1
)
Xylanase
(nmol MU
min
-1
g
-1
)
Phenol oxidase
(mg h
-1
g
-1
)
Low-fertility
CTL 3.42 ±0.16 40.12 ±1.05 3.05 ±0.17 0.31 ±0.01 1.43 ±0.01 0.60 ±0.08
?NPK 2.02 ±0.21* 26.79 ±0.58* 1.72 ±0.10* 0.33 ±0.02 1.27 ±0.05 0.77 ±0.08
?C 2.76 ±0.19 33.35 ±1.14* 1.93 ±0.08* 0.26 ±0.03* 1.00 ±0.13* 0.90 ±0.02*
?CNPK 3.26 ±0.08 31.45 ±0.81* 2.69 ±0.02 0.31 ±0.05 0.96 ±0.11* 0.86 ±0.03
Mid-fertility
CTL 1.82 ±0.12 7.80 ±0.45 1.03 ±0.08 0.17 ±0.01 0.52 ±0.09 1.62 ±0.03
?NPK 1.75 ±0.07 7.25 ±0.27 1.01 ±0.07 0.18 ±0.01 0.49 ±0.03 1.51 ±0.03
?C 1.76 ±0.25 12.71 ±0.79* 0.78 ±0.05 0.15 ±0.02 0.28 ±0.02* 1.99 ±0.06*
?CNPK 2.29 ±0.18* 6.56 ±0.36 0.98 ±0.05 0.17 ±0.01 0.34 ±0.01 1.82 ±0.14
High-fertility
CTL 1.43 ±0.14 6.32 ±0.27 2.59 ±0.07 0.45 ±0.01 0.85 ±0.06 1.01 ±0.08
?NPK 2.02 ±0.25 6.84 ±0.37 2.99 ±0.15* 0.43 ±0.02 0.80 ±0.02 1.10 ±0.10
?C 1.95 ±0.09 12.28 ±0.36* 2.29 ±0.06 0.34 ±0.04* 0.65 ±0.02* 1.11 ±0.04
?CNPK 2.16 ±0.19* 7.79 ±0.58 2.15 ±0.06* 0.39 ±0.02 0.71 ±0.08 1.18 ±0.05
Significant differences between treatments and controls are highlighted by asterisks (PB0.05). Data are means ±1 standard error (n =4)
Biogeochemistry (2012) 111:219–237 229
123
(17:1x8) and fungal (18:1x9) biomarkers in the low-
fertility soil and significantly less soil-derived C in a
fungal (18:2x6) biomarker in the high-fertility soil
(P\0.001). There was significantly higher soil-
derived C in a Gram-negative biomarker (16:1x7) in
both mid- and high-fertility soils following CNPK
addition (P\0.001).
The relatively large treatment and soil-fertility
effects on sucrose-derived C within PLFA compared
to the smaller effects on soil-derived C within PLFA
were summarized by multivariate analyses (Fig. 6).
The concentration of soil-derived C within PLFA
varied widely among treatments and soils (Fig. 6a).
In contrast, for the concentration of sucrose-derived
C within PLFA (Fig. 6b) there was a clear separation
along the X-axis according to treatment (explaining
34% of variation) and along the Y-axis according to
soil-fertility (explaining 28% of variation).
Discussion
Priming and soil fertility
The largest release of additional soil-derived (i.e.
primed) CO
2
following sucrose-C addition was for soil
from the site of intermediate fertility, suggesting that
priming and soil fertility were not linearly correlated.
Microbial metabolism of sucrose-C, on the other hand,
was limited by soil fertility, because respiration of
sucrose-derived CO
2
was proportionally higher when
added in combination with nutrients for soils in the
order of decreasing fertility. Therefore, factors other
than fertility constrained microbial metabolism of soil
C through priming in the low-fertility soil. We
hypothesize that priming was limited by both nutrient
deficiency to microorganisms (Blagodatskaya et al.
2007) and the concentration and ‘lability’ of soil
organic matter (Kuzyakov and Bol 2006). Thus,
sucrose-C additions did not lead to significant priming
in the high fertility soil because microbial nutrient
limitation was insufficient, while in the low fertility
soil priming was limited because the pool of soil C
amenable to priming was insufficient. Priming may
also have been influenced by soil pH and mineralogy.
Soil pH can affect priming due to the adsorption of
enzymes on soil surfaces, which occurs when soil pH is
lower than the isoelectric point of the enzyme (Qui-
quampoix 2000); clay content can affect priming due to
the stabilization of soil organic C within clay-mineral
complexes, which can protect it from priming (Ras-
mussen et al. 2007). In the low-fertility soil where
priming was lowest, both clay content and pH were
relatively low, suggesting that pH rather than
Table 4 PLFA concentration within microbial groups 3 days following treatments
Total
(lgCg
-1
soil)
Bacterial
(lgCg
-1
soil)
Gram-positive
bacterial
(lgCg
-1
soil)
Gram-negative
bacterial
(lgCg
-1
soil)
Fungal
(lgCg
-1
soil)
Saturated
(lgCg
-1
soil)
Fung/Bact
Low-fertility
CTL 62.9 ±10.1 42.1 ±7.1 14.7 ±1.8 27.4 ±5.5 6.9 ±1.1 13.9 ±2.1 0.16 ±0.01
?C 55.7 ±2.5 37.2 ±1.8 12.9 ±0.5 24.3 ±1.3 6.2 ±0.4 12.2 ±0.4 0.2 ±0.0
?CNPK 63.7 ±3.7 34.3 ±0.9 10.9 ±0.2 23.5 ±0.8 18.0 ±2.7* 11.4 ±1.9* 0.52 ±0.09*
Mid- fertility
CTL 46.9 ±3.0 35.2 ±5.5 18.1 ±3.0 17.1 ±2.6 2.9 ±0.4 8.8 ±1.3 0.08 ±0.01
?C 39.1 ±4.7 28.2 ±3.6 13.6 ±1.8 14.7 ±2.0 2.4 ±0.2 8.5 ±1.0 0.1 ±0.0
?CNPK 47.3 ±1.8 34.8 ±0.7 15.8 ±0.6 19.1 ±1.3 2.6 ±0.1 9.8 ±1.3 0.07 ±0.01
High-fertility
CTL 63.9 ±32 46.6 ±2.7 22.4 ±1.7 24.2 ±1.6 5.6 ±0.3 11.6 ±0.8 0.12 ±0.01
?C 69.0 ±4.9 50.1 ±3.9 23.6 ±2.3 26.5 ±1.7 5.6 ±0.4 13.3 ±0.7 0.1 ±0.0
?CNPK 71.2 ±5.8 55.8 ±5.1 27.3 ±2.7 28.5 ±2.5 6.9 ±0.6 8.5 ±1.9 0.12 ±0.01
Fungal/bacterial ratios are the ratio of 18:2x6,9:bacterial PLFAs. Significant differences between treatments and controls are highlighted
by asterisks (PB0.05). Data are means with 1 SE (n =4)
230 Biogeochemistry (2012) 111:219–237
123
mineralogy may have had some influence on priming.
However, our hypothesis that sucrose-C addition did
not induce significant priming because high nutrient
demand in this soil had already led to depletion of the
limited pool of C amenable to priming was supported
by the high enzyme activities and low organic C
Fig. 5 Concentration of sucrose-derived C and soil-derived C
within PLFA 3 days after ?C and ?CNPK and CTL
treatments for low-fertility (a,b), mid-fertility (c,d) and high-
fertility (e,f) soils. Significant incorporation of sucrose-derived
C and significant differences in soil-derived C between
treatments and controls are highlighted by asterisks (where
PB0.05). Data are means ±1 standard error (n=4)
Biogeochemistry (2012) 111:219–237 231
123
concentration in the untreated low-fertility soil—it has
the lowest C concentration of these three soils, and also
of a much wider set of soils in the region (Table 1;
Turner and Engelbrecht 2011). We suggest that soils at
this site undergo constant and intense priming during
the decomposition of plant residues, consistent with
descriptions of priming as a phenomena that preferen-
tially degrades labile fractions of soil organic matter
(Kuzyakov and Bol 2006) and constrains the accumu-
lation of C in surface soils (Fontaine et al. 2007).
Priming mechanisms
Our results suggest strongly that priming was due to
microbial demand for nutrients required to mineralize
the added sucrose-C. This was indicated by signifi-
cant reductions in priming when C was added in
combination with nutrients (100% for low-fertility,
33% for mid-fertility, 44% for high-fertility soils;
Fig. 3), the depletion of NO
3-
,NH
4?
and extractable
P (Table 3), the increases in microbial C:N and C:P
ratios (Fig. 4), and increases in phosphomonoesterase
activity (Table 3). It seems likely that when sucrose-
C was added alone, energy derived from its metab-
olism was used to degrade soil organic matter through
production of extracellular enzymes to liberate addi-
tional nutrients, which simultaneously made soil C
available for microbial metabolism (‘co-metabolism’;
Kuzyakov et al. 2000).
The rapid priming (the peak in primed-CO
2
occurred before the peak in sucrose derived-CO
2
)
due to microbial nutrient demand as indicated by our
data is consistent with some studies of temperate soils
(Zyakun and Dilly 2005; Blagodatskaya et al. 2007;
Nottingham et al. 2009), but at odds with others that
do not predict such rapid priming (Fontaine et al.
2004) or an increased production of extracellular
enzymes (De Nobili et al. 2001). This suggests that
different mechanisms may apply in different soils on
different time scales. For example, priming through
increased abundance and activity of slow-growing
recalcitrant-C degrading microorganisms that persist
after the added labile C is exhausted (Fontaine et al.
2004) is not consistent with our detection of short-
term priming, which was greatest in the soil of
intermediate fertility with no change in microbial
community composition. Priming by increased
growth of recalcitrant-C degrading microorganisms
may be more important on longer time scales
following addition of substrates that take longer to
decompose (e.g. the long-term depletion of soil C and
altered microbial community composition in Carney
et al. (2007)). Similarly inconsistent with our results
is ‘apparent priming’, either through turnover of soil
microbial biomass (Wu et al. 1993), or a small
increase in microbial mineralization of endo-cellular
C reserves with no change in decomposition of soil
organic matter (where microorganisms increase their
metabolic activity following a trace C input in
anticipation of a larger input in the near future; De
Nobili et al. 2001). These apparent priming mecha-
nisms are not consistent with the large release of soil-
derived C from the mid-fertility soil (360 mg CO
2
-C
g
-1
soil), which was 120% greater than the original
concentration of microbial C. Further evidence that
priming measured in our study was not ‘apparent’
Fig. 6 The effect of sucrose-C and nutrient additions on
concentration of soil-derived C and sucrose-derived C within
microbial community composition as revealed by principal
components analysis, showing asoil-derived C concentration
in PLFA and bsucrose-derived C concentration in PLFA. Data
are grouped into low-fertility (circles), mid-fertility (squares)
and high-fertility (triangles) soils following sucrose-C addition
(?C, grey symbols), sucrose and nutrient addition (?CNPK,
white symbols) and no addition controls (CTL, black symbols)
232 Biogeochemistry (2012) 111:219–237
123
was that despite the cumulative release of 260 mg
soil-derived CO
2
-C g
-1
soil over three days between
sucrose-C addition and extraction of soil for PLFA
analyses, there was no significant reduction in soil C
within PLFA biomarkers (Fig. 5), which would be
predicted if there was significant substitution of soil-
labeled C by sucrose-labeled C within microorgan-
isms (see Perelo and Munch 2005; Schneckenberger
et al. 2008; Nottingham et al. 2009). Thus, apparent
priming can only account for a minor part of the
short-term increase in the efflux of soil-derived CO
2
measured in our study.
Microbial carbon utilization: enzyme activity
and PLFA
Microbial priming of C from soil organic matter was
supported in part by changes in activities of enzymes
that degrade organic C. We measured higher activity of
phenol oxidase during priming in low- and mid-
fertility soils. This oxidative enzyme catalyzes the
degradation of phenolic C compounds, including
lignin and humus, and can limit the accumulation of
soil organic matter (Sinsabaugh 2010). Oxidative
enzymes have been linked to stimulated decomposi-
tion of recalcitrant-C in experimentally warmed soils
(Waldrop and Firestone 2004b) and soils beneath a
deciduous forest growing under elevated CO
2
through
priming effects (Carney et al. 2007). The lack of
increase in phenol oxidase activity when nutrients were
added to soils alongside C (Table 3), a result similar to
that found by Carreiro et al. (2000) following leaf-litter
and nutrient additions to a temperate deciduous forest
soil, suggests that its induction was in response to
nutrient-deficiency and associated with microbial
acquisition of nutrients from soil organic matter.
The cause of differences in the activities of
hydrolytic C-degrading enzymes during priming
was less clear. The addition of sucrose-C led to
lower activities of several hydrolytic enzymes that
catalyze the degradation of more labile forms of
organic C; b-glucosidase (in low- and high-fertility
soils), cellobiohydrolase (in low- and high-fertility
soils), and xylanase (in all soils). These patterns can
be explained by a depletion of labile forms of soil C
following priming; the sucrose-stimulated microor-
ganisms rapidly exhausted the organic-C forms that
are degraded by these enzymes. A similar pattern was
detected by Waldrop and Firestone (2004b) who
attributed a 50% reduction in activity of hydrolytic
cellulases and hemicellulases in experimentally
warmed soils to a depletion of labile soil C. The
generally lower activities may also be due to
microbial metabolism of the enzymes themselves as
a nutrient source following sucrose-C addition (e.g.
as a N source; Treseder and Vitousek 2001), or of the
soil organic matter to which these enzymes were
adsorbed.
The absence of reduction in soil-derived C within
PLFA during priming provided strong evidence that
the ‘primed’ CO
2
efflux originated from soil organic
matter rather than turnover of microbial biomass.
However, whether priming resulted from the activity of
specific microbial groups was inconclusive. The par-
allel stimulation of priming and activity of phenol
oxidase suggested that priming was induced by acti-
nomycetes or fungi, known producers of phenol
oxidase (Kirk and Farrell 1987), but this was not
reflected by stimulated incorporation of primed soil C
into their PLFA biomarkers (Fig. 5; the PLFA bio-
marker, 10Me18:0 for actinomycetes was not detected;
Frostega
˚rd and Ba
˚a
˚th 1993). We found a higher
concentration of soil-derived C within the Gram-
negative biomarker 16:1x7 (following CNPK addition
to mid- and high-fertility soils; Fig. 5), which has been
detected by other studies following substrate additions
to temperate soils (Waldrop and Firestone 2004a;
Nottingham et al. 2009). The soil C incorporated into
this bacterial biomarker may originate from primed
soil organic matter, but it was notable that in our study
there was no increase in soil C-incorporation into this
biomarker in soils when sucrose-C was added alone
and priming was greatest. Further studies using
different molecular techniques are required to resolve
whether: (i) specific microorganisms caused priming
and there was rapid release and re-assimilation of C
within microorganisms following death of microor-
ganisms to re-distribute soil-labeled C within PLFA
biomarkers, or (ii) priming resulted from the activity of
various functionally redundant microbial groups.
While there were no conclusive patterns in primed
soil-derived C within microbial groups, there were
clear patterns in sucrose-derived C (Fig. 6). Sucrose-
derived C was incorporated into similar bacterial
biomarkers (a15:0, 16:1x7) for mid- and high-
fertility soils, but for the low-fertility soil it was
preferentially incorporated into fungal biomarkers
(18:2x6, 18:1x9), which led to a large shift in
Biogeochemistry (2012) 111:219–237 233
123
microbial community composition in CNPK-treated
soils (Table 4). Fungal-based food webs are more
retentive of nutrients than bacterial-based foods webs
(Coleman et al. 1983), are associated with more
acidic soils (Alexander 1964; low-fertility soils had
pH of 4.4) and with later retrogressional and therefore
less fertile stages of soil development (Allison et al.
2007; Wardle et al. 2008). A recent experiment on a
soil devoid of organic matter similarly found that the
metabolism of added sucrose-C was predominantly
by fungi (Engelking et al. 2008). Thus, we hypoth-
esize that due to low pH and low concentration of
available nutrients (Table 3) nutrient cycling was
more retentive and dominated by fungal activity in
the low-fertility soil. Further evidence for this was the
high enzyme activity in the untreated low-fertility
soil (Table 4), an indication of high allocation of
microbial resources into nutrient acquisition.
Microbial nutrient limitation
and acquisition of nutrients
Priming most likely resulted from microbial utilization
of organic nutrients, in particular P, with evidence for
both increased demand and increased acquisition of P
by microorganisms during priming. The increased
acquisition of mineral nutrients during priming was
reflected by lower concentrations of mineral N and P in
sucrose-C treated soils (Table 2), although this was not
reflected as a significant increase in microbial N or P
(Fig. 4), which is not surprising given that concentra-
tions of microbial nutrients were up to an order of
magnitude higher than concentrations of mineral
nutrients. An increased microbial demand for P was
reflected by significantly higher phosphomonoesterase
activity for both mid- and high-fertility soils following
sucrose-C addition compared to controls, providing
evidence that there was increased microbial production
of phosphomonoesterase in response to P deficiency to
access additional P from organic sources. In contrast,
there was no change in N-acetyl b-glucosaminidase
activity (to release N) following sucrose-C addition
(Table 3), suggesting that N was less limiting than P or
that other enzymes we did not measure were involved
in N-acquisition (e.g. enzymes involved in the hydro-
lysis of amino acids; Acosta-Martinez and Tabatabai
2000). Our interpretation of up-regulated phospho-
monoesterase activity is supported by several studies
that have measured higher phosphomonoesterase
activity in response to greater P deficiency in soils
(Olander and Vitousek 2000; Allison et al. 2007;
Sinsabaugh et al. 2008).
Priming and nutrient limitation in tropical forests
Our study provides further evidence that when soil
microorganisms have sufficient energy, the availability
of soluble soil nutrients regulates soil organic C
mineralization by priming (Fontaine et al. 2011).
Furthermore, our study suggests that the ultimate
control on microbial respiration in moist and aerobic
soils is the availability of labile C rather than nutrients,
and that nutrients only limit microbial respiration when
labile C is abundant (Fig. 2; NPK additions had no
significant effect on CO
2
efflux, data not shown).
While we are cautious in making ‘real world’ infer-
ences based on results from our laboratory study, the
implication that the controls on microbial sources of
soil CO
2
efflux shift between labile C and nutrients,
with priming occurring when there are large inputs of
labile C relative to nutrients, is supported by observa-
tions in tropical forests. Two studies hypothesized a
release of primed soil organic matter within soil CO
2
efflux following the addition of C to soils: Sayer et al.
(2007) to explain increased soil CO
2
efflux from soils
that received experimentally increased litterfall, and
Cleveland et al. (2010) to explain increased soil CO
2
efflux from soils that received more concentrated
dissolved organic matter following a drought treat-
ment. Two further studies in tropical forests where
nutrients were added without C further support our
interpretation that priming occurs when there are large
inputs of labile C relative to nutrients; both studies
provide evidence in support of suppressed priming in
the presence of excess nutrients. Koehler et al. (2009)
observed no or negative responses of soil CO
2
efflux
to N additions, while Cleveland and Townsend
(2006) observed little response of soil CO
2
efflux to
P addition, except during the dry-wet season transition
when litter decomposition rates were highest and
therefore the C limitation of microbial activity was
lower.
Priming resulting from nutrient limitation in
tropical forest soils, which can contain abundant
organic N (Hedin et al. 2005) and organic P (Turner
and Engelbrecht 2011; Dieter et al. 2010), would
have large consequences for the tropical forest C
balance under elevated CO
2
by increasing the pool of
234 Biogeochemistry (2012) 111:219–237
123
nutrients available for plant assimilation and desta-
bilizing soil organic C. In particular, we suggest that
P limitation of heterotrophic activity in tropical forest
soil (as demonstrated by Hobbie and Vitousek 2000;
Cleveland et al. 2006; Cleveland and Townsend
2006; Kaspari et al. 2008), is alleviated in the
presence of excess labile organic C to provide energy
for mineralization of organic P in parallel with
priming effects.
Conclusions
This is the first study to investigate priming effect
mechanisms in tropical forest soils. Priming resulted
from microbial nutrient limitation and appeared to be
stimulated by microbial acquisition of P from organic
matter. Priming appeared to be constrained in low
fertility soils by low concentrations of organic C, and
in high fertility soils by the abundance of available
nutrients. This provides a framework for understand-
ing the extent and distribution of priming effects in
lowland tropical forests, and suggests that nutrient
availability constrains soil C storage in such ecosys-
tems. However, a better understanding of contrasting
tropical forest soils and how their properties may
influence priming intensity on different time scales is
still required before this potentially important feed-
back mechanism can be included in C-cycle models.
Acknowledgments We thank Tania Romero, Dayana Agudo
and Dianne de la Cruz for laboratory support; Marie-Soleil
Turmel for field assistance; Beto Quesada for sharing data from
RAINFOR sites; and Marife Corre and Rodolfo Rojas for the
GC-analyses of CO
2
samples. For their insightful comments on
earlier drafts of this manuscript, we thank two anonymous
reviewers. This work was funded by a NERC grant (NER/S/A/
2004/12241A), Gonville and Caius College Cambridge, a
NERC LSMSF grant (allocation number SI-025), and a STRI
short term fellowship to ATN.
References
Acosta-Martinez V, Tabatabai MA (2000) Arylamidase activ-
ity of soils. Soil Sci Soc Am J 64:215–221
Alexander M (1964) Introduction to soil microbiology. Wiley,
New York
Allison VJ, Condron LM, Peltzer DA, Richardson SJ, Turner
BL (2007) Changes in enzyme activities and soil micro-
bial community composition along carbon and nutrient
gradients at the Franz Josef chronosequence, New Zea-
land. Soil Biol Biochem 39:1770–1781
Asmar F, Eiland F, Nielsen NE (1994) Effect of extracellular-
enzyme activities on solubilization rate of soil organic
nitrogen. Biol Fert Soils 17:32–38
Balesdent J, Wagner GH, Mariotti A (1988) Soil organic-matter
turnover in long-term field experiments as revealed by C-13
natural abundance. Soil Sci Soc Am J 52:118–124
Bardgett RD, Hobbs PJ, Frostega
˚rd A
˚(1996) Changes in soil
fungal:bacterial biomass ratios following reductions in the
intensity of management of an upland grassland. Biol Fert
Soils 22:261–264
Bertolini T, Inglima I, Rubino M, Marzaioli F, Lubritto C,
Subke JA, Peressotti A, Cotrufo MF (2006) Sampling
soil-derived CO
2
for analysis of isotopic composition: a
comparison of different techniques. Isot Environ Health
Stud 42:57–65
Blagodatskaya E, Kuzyakov Y (2008) Mechanisms of real and
apparent priming effects and their dependence on soil
microbial biomass and community structure: critical
review. Biol Fert Soils 45:115–131
Blagodatskaya EV, Blagodatsky SA, Anderson TH, Kuzyakov
Y (2007) Priming effects in chernozem induced by glu-
cose and N in relation to microbial growth strategies. Appl
Soil Ecol 37:95–105
Carney KM, Hungate BA, Drake BG, Megonigal JP (2007)
Altered soil microbial community at elevated CO
2
leads to
loss of soil carbon. Proc Natl Acad Sci USA 104:4990–4995
Carreiro MM, Sinsabaugh RL, Repert DA, Parkhurst DF
(2000) Microbial enzyme shifts explain litter decay
responses to simulated nitrogen deposition. Ecology 81:
2359–2365
Chamberlain PM, Bull ID, Black HIJ, Ineson P, Evershed RP
(2006) Collembola trophic preferences determined using
fatty acid distributions and compound-specific stable
carbon isotope values. Soil Biol Biochem 38:1275–1281
Cleveland CC, Townsend AR (2006) Nutrient additions to a
tropical rain forest drive substantial soil carbon dioxide
losses to the atmosphere. Proc Natl Acad Sci USA 103:
10316–10321
Cleveland CC, Reed SC, Townsend AR (2006) Nutrient reg-
ulation of organic matter decomposition in a tropical rain
forest. Ecology 87:492–503
Cleveland CC, Wieder WR, Reed SC, Townsend AR (2010)
Experimental drought in a tropical rain forest increases
soil carbon dioxide losses to the atmosphere. Ecology
91:2313–2323
Coleman DC, Reid CPP, Cole CV (1983) Biological strategies
of nutrient cycling in soil systems. Adv Ecol Res 13:1–55
Crossman ZM, Abraham F, Evershed RP (2004) Stable isotope
pulse-chasing and compound specific stable carbon iso-
tope analysis of phospholipid fatty acids to assess methane
oxidizing bacterial populations in landfill cover soils.
Environ Sci Technol 38:1359–1367
De Nobili M, Contin M, Mondini C, Brookes PC (2001) Soil
microbial biomass is triggered into activity by trace
amounts of substrate. Soil Biol Biochem 33:1163–1170
Dieter D, Elsenbeer H, Turner BL (2010) Phosphorus frac-
tionation in lowland tropical rainforest soils of central
Panama. Catena 82:118–125
Biogeochemistry (2012) 111:219–237 235
123
Engelking B, Flessa H, Joergensen RG (2008) Formation and
use of microbial residues after adding sugarcane sucrose
to a heated soil devoid of organic matter. Soil Biol Bio-
chem 40:97–105
Fontaine S, Mariotti A, Abbadie L (2003) The priming effect
of organic matter: a question of microbial competition?
Soil Biol Biochem 35:837–843
Fontaine S, Bardoux G, Abbadie L, Mariotti A (2004) Carbon
input to soil may decrease soil carbon content. Ecol Lett
7:314–320
Fontaine S, Barot S, Barre P, Bdioui N, Mary B, Rumpel C
(2007) Stability of organic carbon in deep soil layers
controlled by fresh carbon supply. Nature 450:277–280
Fontaine S, Henault C, Aamor A, Bdioui N, Bloor JMG, Maire
V, Mary B, Revaillot S, Maron PA (2011) Fungi mediate
long term sequestration of carbon and nitrogen in soil
through their priming effect. Soil Biol Biochem 43:86–96
Frostega
˚rd A, Ba
˚a
˚th E (1993) Shifts in the structure of soil
microbial communities in limed forests as revealed by
phospholipid fatty acid analysis. Soil Biol Biochem 25:
723–730
Gnankambary Z, Stedt U, Nyberg G, Hien V, Malmer A (2008)
Nitrogen and phosphorus limitation of soil microbial
respiration in two tropical agroforestry parklands in the
south-Sudanese zone of Burkina Faso: the effects of tree
canopy and fertilization. Soil Biol Biochem 40:350–359
Hagedorn F, Spinnler D, Siegwolf R (2003) Increased N
deposition retards mineralization of old soil organic
matter. Soil Biol Biochem 35:1683–1692
Hedin LO, Brookshire J, Menge DNL, Barron AR (2005) The
nitrogen paradox in tropical forest ecosystems. Annu Rev
Ecol Evol Syst 40:613–635
Hobbie SE, Vitousek PM (2000) Nutrient limitation of decom-
position in Hawaiian forests. Ecology 81:1867–1877
Jobbagy EG, Jackson RB (2000) The vertical distribution of
soil organic carbon and its relation to climate and vege-
tation. Ecol Appl 10:423–436
Joergensen RG, Mueller T (1996) The fumigation-extraction
method to estimate soil microbial biomass: calibration of
the K
EN
value. Soil Biol Biochem 28:33–37
Kaspari M, Garcia MN, Harms KE, Santana M, Wright SJ,
Yavitt JB (2008) Multiple nutrients limit litterfall and
decomposition in a tropical forest. Ecol Lett 11:35–43
Kirk TK, Farrell RL (1987) Enzymatic ‘combustion’’: the
microbial degradation of lignin. Annu Rev Microbiol
41:465–505
Kouno K, Tuchiya Y, Ando T (1995) Measurement of soil
microbial biomass phosphorus by an anion-exchange
membrane method. Soil Biol Biochem 27:1353–1357
Koehler B, Corre MD, Veldkamp E, Sueta JP (2009) Chronic
nitrogen addition causes a reduction in soil carbon dioxide
efflux during the high stem-growth period in a tropical
montane forest but no response from a tropical lowland
forest on a decadal time scale. Biogeosciences 6:
2973–2983
Kuzyakov Y, Bol R (2006) Sources and mechanisms of
priming effect induced in two grassland soils amended
with slurry and sugar. Soil Biol Biochem 38:747–758
Kuzyakov Y, Friedel JK, Stahr K (2000) Review of mecha-
nisms and quantification of priming effects. Soil Biol
Biochem 32:1485–1498
Langley JA, McKinley DC, Wolf AA, Hungate BA, Drake BG,
Megonigal JP (2009) Priming depletes soil carbon and
releases nitrogen in a scrub-oak ecosystem exposed to
elevated CO
2
. Soil Biol Biochem 41:54–60
Loftfield N, Flessa H, Augustin J, Beese F (1997) Automated
gas chromatographic system for rapid analysis of the
atmospheric trace gases methane, carbon dioxide, and
nitrous oxide. J Environ Qual 26:560–564
Martinelli LA, Piccolo MC, Townsend AR, Vitousek PM,
Cuevas E, McDowell W, Robertson GP, Santos OC,
Treseder K (1999) Nitrogen stable isotopic composition of
leaves and soil: tropical versus temperate forests. Bio-
geochemistry 46:45–65
Marx MC, Wood M, Jarvis SC (2001) A microplate fluori-
metric assay for the study of enzyme diversity in soils.
Soil Biol Biochem 33:1633–1640
Muruganandama S, Israel DW, Robarge WP (2009) Activities
of nitrogen-mineralization enzymes associated with soil
aggregate size fractions of three tillage systems. Soil Sci
Soc Am J 73:751–759
Nottingham AT, Griffiths H, Chamberlain PM, Stott AW,
Tanner EVJ (2009) Soil priming by sugar and leaf-litter
substrates: a link to microbial groups. Appl Soil Ecol
42:183–190
Olander LP, Vitousek PM (2000) Regulation of soil phospha-
tase and chitinase activity by N and P availability. Bio-
geochemistry 49:175–190
Parham JA, Deng SP (2000) Detection, quantification and
characterization of b-glucosaminidase activity in soil. Soil
Biol Biochem 32:1183–1190
Perelo LW, Munch JC (2005) Microbial immobilisation and
turnover of C-13 labelled substrates in two arable soils
under field and laboratory conditions. Soil Biol Biochem
37:2263–2272
Phillips OL, Lewis SL, Baker TR, Chao KJ, Higuchi N (2008)
The changing Amazon forest. Philos Trans R Soc Lond B
363:1819–1827
Porder S, Hilley GE (2011) Linking chronosequences with the
rest of the world: predicting soil phosphorus content in
denuding landscapes. Biogeochemistry 102:153–166
Pyke CR, Condit R, Aguilar S, Lao S (2001) Floristic com-
position across a climatic gradient in a neotropical low-
land forest. J Veg Sci 12:553–566
Quesada CA, Lloyd J, Anderson LO, Fyllas NM, Schwarz M,
Czimczik CI (2011) Soils of Amazonia with particular
reference to the RAINFOR sites. Biogeosciences 8:
1415–1440
Quiquampoix H (2000) Mechanisms of protein adsorption on
surfaces and consequences for extracellular enzyme
activity in soil. In: Bollag JM, Stotzky G (eds) Soil bio-
chemistry, vol 10. Marcel Dekker, New York, pp 171–206
Quiquampoix H, Mousain D (2005) Enzymatic hydrolysis of
organic phosphorus. In: Turner BL, Frossard E, Baldwin
DS (eds) Organic phosphorus in the environment. CAB
International, Wallingford, pp 89–121
Rasmussen C, Southard RJ, Horwath WR (2007) Soil miner-
alogy affects conifer forest soil carbon source utilization
and microbial priming. Soil Sci Soc Am J 71:1141–1150
Sayer EJ, Powers JS, Tanner EVJ (2007) Increased litterfall in
tropical forests boosts the transfer of soil CO
2
to the
atmosphere. PLoS One 2:e1299
236 Biogeochemistry (2012) 111:219–237
123
Schimel JP, Weintraub MN (2003) The implications of exo-
enzyme activity on microbial carbon and nitrogen limi-
tation in soil: a theoretical model. Soil Biol Biochem
35:549–563
Schneckenberger K, Demin D, Stahr K, Kuzyakov Y (2008)
Microbial utilization and mineralization of C-14 glucose
added in six orders of concentration to soil. Soil Biol
Biochem 40:1981–1988
Sinsabaugh RL (2010) Phenol oxidase, peroxidase and organic
matter dynamics of soil. Soil Biol Biochem 42:391–404
Sinsabaugh RL, Moorhead DL (1994) Resource-allocation to
extracellular enzyme-production—a model for nitrogen
and phosphorus control of litter decomposition. Soil Biol
Biochem 26:1305–1311
Sinsabaugh RL, Lauber CL, Weintraub MN, Ahmed B, Allison
SD, Crenshaw C, Contosta AR, Cusack D, Frey S, Gallo
ME, Gartner TB, Hobbie SE, Holland K, Keeler BL,
Powers JS, Stursova M, Takacs-Vesbach C, Waldrop MP,
Wallenstein MD, Zak DR, Zeglin LH (2008) Stoichiom-
etry of soil enzyme activity at global scale. Ecol Lett
11:1252–1264
Tabatabai MA (1994) Soil enzymes. In: Weaver R et al (eds)
Methods of soil analysis. Part 2. Microbiological and
biochemical properties. Soil Science Society of America,
Madison, pp 778–833
Treseder KK, Vitousek PM (2001) Effects of soil nutrient
availability on investment in acquisition of N and P in
Hawaiian rain forests. Ecology 82:946–954
Turner BL, Engelbrecht BMJ (2011) Soil organic phosphorus
in lowland tropical rain forests. Biogeochemistry 103:
295–315
Turner BL, Romero TE (2009) Short-term changes in
extractable inorganic nutrients during transport and stor-
age of tropical rain forest soils. Soil Sci Soc Am J 73:
1972–1979
Turner BL, Romero TE (2010) Stability of hydrolytic enzyme
activity and microbial phosphorus during storage of
tropical rain forest soils. Soil Biol Biochem 42:459–465
Vance ED, Brookes PC, Jenkinson DS (1987) An extraction
method for measuring soil microbial biomass-C. Soil Biol
Biochem 19:703–707
Vitousek PM, Sanford RL (1986) Nutrient cycling in moist
tropical forest. Annu Rev Ecol Syst 17:137–167
Waldrop MP, Firestone MK (2004a) Microbial community
utilization of recalcitrant and simple carbon compounds:
impact of oak-woodland plant communities. Oecologia
138:275–284
Waldrop MP, Firestone MK (2004b) Altered utilization pat-
terns of young and old soil C by microorganisms caused
by temperature shifts and N additions. Biogeochemistry
67:235–248
Walker TW, Syers JK (1976) Fate of phosphorus during ped-
ogenesis. Geoderma 15:1–19
Wardle DA, Walker LR, Bardgett RD (2008) Ecosystem
properties and forest decline in contrasting long-term
chronosequences. Science 305:509–513
Wieder RK, Wright SJ (1995) Tropical forest litter dynamics
and dry season irrigation on Barro-Colorado Island, Pan-
ama. Ecology 76:1971–1979
Wu J, Joergensen RG, Pommerening B, Chaussod R, Brookes
PC (1990) Measurement of soil microbial biomass C by
fumigation extraction—an automated procedure. Soil Biol
Biochem 22:1167–1169
Wu J, Brookes PC, Jenkinson DS (1993) Formation and
destruction of microbial biomass during the decomposi-
tion of glucose and ryegrass in soil. Soil Biol Biochem
25:1435–1441
Zelles L (1999) Fatty acid patterns of phospholipids and
lipopolysaccharides in the characterisation of microbial
communities in soil: a review. Biol Fert Soils 29:111–129
Zyakun AM, Dilly O (2005) Use of carbon isotope composition
for characterization of microbial activity in arable soils.
Appl Biochem Microbiol 41:512–520
Biogeochemistry (2012) 111:219–237 237
123
... Soil pH determines microbially mediated organic matter decomposition processes due to an optimal pH range for enzymes (Table S1) and thus for hydrolysis and oxidization of SOM, and energy investments by microorganisms to regulate the pH in their cells (Wang & Kuzyakov, 2023a) as well as the nutrient release into the soil solution (Nottingham et al., 2012;Pan et al., 2022; Figure 1). The nutrient availability controlled by these processes, in turn, strongly regulates microbial activity and plant productivity and thus the PE intensity. ...
... Nutrient availability controls plant growth and productivity and thus the input of rhizodeposits and litter into the soil (Prescott et al., 2020). Nutrient availability determines stoichiometric ratios (C:N:P:S) of the litter and consequently adaptation of microbial community structure, diversity, and activity, and thus microbial metabolic mechanisms and rates (Nottingham et al., 2012). Nutrient availability determines the competition intensity between roots and microorganisms and thus plant and microbial nutrient niches as well as microbial metabolisms and strategies (Wang et al., 2022a(Wang et al., , 2022b. ...
... For example, N limitation increased the root exudation rate by 60% in boreal pine forest soil as compared to N-rich soils (Högberg et al., 2010). The intensity of SOM priming induced by sucrose addition followed an order of slight nutrient (especially P) limitation > high nutrient availability > strong nutrient limitation (Nottingham et al., 2012). This is confirmed by the coincidence of the highest PE intensities (Figure 5a) and moderate nutrient limitation ( Figure 1) and alkaline soils (Figure 1), decreases PE intensity because of low microbial activity and low content of SOM amenable to priming (Nottingham et al., 2012). ...
Article
Full-text available
Priming of soil organic matter (SOM) decomposition by microorganisms is a key phenomenon of global carbon (C) cycling. Soil pH is a main factor defining priming effects (PEs) because it (i) controls microbial community composition and activities, including enzyme activities, (ii) defines SOM stabilization and destabilization mechanisms, and (iii) regulates intensities of many biogeochemical processes. In this critical review, we focus on prerequisites and mechanisms of PE depending on pH and assess the global change consequences for PE. The highest PEs were common in soils with pH between 5.5 and 7.5, whereas low molecular weight organic compounds triggered PE mainly in slightly acidic soils. Positive PEs up to 20 times of SOM decomposition before C input were common at pH around 6.5. Negative PEs were common at soil pH below 4.5 or above 7 reflecting a suboptimal environment for microorganisms and specific SOM stabilization mechanisms at low and high pH. Short‐term soil acidification (in rhizosphere, after fertilizer application) affects PE by: mineral‐SOM complexation, SOM oxidation by iron reduction, enzymatic depolymerization, and pH‐dependent changes in nutrient availability. Biological processes of microbial metabolism shift over the short‐term, whereas long‐term microbial community adaptations to slow acidification are common. The nitrogen fertilization induced soil acidification and land use intensification strongly decrease pH and thus boost the PE. Concluding, soil pH is one of the strongest but up to now disregarded factors of PE, defining SOM decomposition through short‐term metabolic adaptation of microbial groups and long‐term shift of microbial communities.
... Total and specific PLFA biomarker abundance were consistent between microbial communities under both plant types. Both peat types are dominated by Gram positive and Gram negative bacteria consistent with measurements made in other forest soils in Panama 10,37,[63][64][65] . Previous microbial community characterisations of peat from Changuinola have noted a dominance of Acidobacteria (a phylum of Gram negative bacteria) 43 , a finding also reported for other tropical 66 and temperate peats 67 . ...
Article
Full-text available
Tropical peatlands are carbon-dense ecosystems that are significant sources of atmospheric methane (CH4). Recent work has demonstrated the importance of trees as an emission pathway for CH4 from the peat to the atmosphere. However, there remain questions over the processes of CH4 production in these systems and how they relate to substrate supply. Principally, these questions relate to the relative contribution of recent photosynthetically fixed carbon, released as root exudates, versus carbon substrate supply from the slowly decomposing peat matrix to CH4 emissions within these ecosystems. Here, we examined the role of root inputs in regulating CH4 production inferred from soil emissions using a combination of in situ tree girdling, in situ¹³C natural abundance labelling via stem injections, and a ¹³CO2 labelling of transplanted plants of two contrasting plant functional types, a broadleaved evergreen tree, and a canopy palm. Girdling of broadleaved evergreen trees reduced CH4 fluxes by up to 67%. Stem injections of trees and palms with a natural abundance label resulted in significant isotopic enrichment of CH4 fluxes, reinforcing the link between root carbon inputs and peat CH4 fluxes. Ex situ¹³CO2 labelling of plants resulted in significant ¹³C enrichment of peat CH4 fluxes. Taken together, our results demonstrate for the first time that plant root exudates make a substantial contribution to CH4 production in tropical peatlands.
... For example, warming can increase the desorption of "mineral-associated" C and extracellular enzymes that form an otherwise stable pool, sorbed to the surfaces of mineral particles (Wallenstein et al., 2011;Fanin et al., 2022), and that may form a substantial pool in clay-rich tropical soils (Kirsten et al., 2021). The pattern could also result from biotic processes, where increased C release from decomposing leaflitter can accelerate enzyme synthesis for soil organic matter mineralization ("priming"), a phenomenon observed in a nearby field experiment where leaf-litter inputs were experimentally increased (Sayer et al., 2011;Tanner et al., 2024) and in another experiment using BCI soils following the addition of 13 C-labeled substrates (Nottingham et al., 2012). It is therefore possible that both abiotic and biotic processes contributed to accelerated enzyme activities and, in turn, organic matter mineralization under warming. ...
Chapter
Full-text available
p dir="ltr">Climate warming poses detrimental but poorly understood consequences for tropical forest biodiversity and carbon storage, especially through its impact on soils. To address this uncertainty, the Soil Warming Experiment in Lowland Tropical Rainforest (SWELTR) was initiated on Barro Colorado Island in Panama; one of a few emerging experiments designed to understand the effects of a warmer climate on the tropical land-surface. In this chapter, I describe results following up to three years of experimental whole-profile soil warming, showing high sensitivity of soil organic matter degradation and an unexpectedly large release of carbon dioxide (CO2 ) to the atmosphere. I consider the biogeochemical mechanisms that may be contributing to this CO2 emission and, finally, the research priorities to understand the longer-term and wider-scale implications of warming on tropical forest soils. Better understanding of these feedbacks is vital for developing mitigation strategies to conserve the biodiversity and carbon storage of tropical forests in a warming world. </p
... The addition of relatively labile carbon in litter (i.e., easily used for microbial metabolism and growth) also appears to have stimulated decomposition of existing soil carbon in what is called a "priming effect" (Sayer et al., 2011). A laboratory study showed that the priming effect of adding a labile carbon substrate to soils from Panamanian forests was greatest when the availability of other nutrients was moderate, and a broad group of microbes increased decomposition of existing soil organic matter to mine for additional nutrients (Nottingham et al., 2012). Overall, these data suggest that even large increases in aboveground litterfall (in this case a doubling) led to relatively modest increases in soil carbon stocks, and only in surface soils. ...
Chapter
Full-text available
p dir="ltr">This chapter presents soil carbon dynamics in the Barro Colorado Nature Monument (BCNM) and surrounding lowland Panamanian forests. I first review principles in soil carbon science for tropical forests, including soil carbon stocks, fractions, turnover times, inputs, and losses. I then review research on patterns and mechanisms of spatial and temporal variability in soil carbon dynamics, including among-site, seasonal, and depth-related variation as well as findings of experimental studies. Soil carbon stocks vary strongly among sites with soil weathering status, declining with fertility and increasing with clay content within the BCNM and in central Panama more generally. Rainfall seasonality drives seasonal variation in soil carbon inputs and losses. Experimental irrigation, nutrient addition, and litter manipulation studies have provided insight into the role of moisture, nutrient availability, and plant inputs in controlling soil carbon dynamics. Finally, I suggest future research directions, including constraining turnover times for different soil carbon fractions across depths. </p
... 29,30 Additionally, there can be changes in the direction and magnitude of the PE on SOC mineralisation because of the input of organic substrate that is accompanied by changes in soil nutrients or the composition of the microbial community over time, 23,31,32 such as switching from a negative to a positive PE rate on native SOC, 33 and an increase or decrease in the magnitude of the positive PE rate. 34 To validate the reliability of our method, we compared the results with those obtained using traditional GC/IRMS. A highly significant positive linear relationship (r 2 > 0.996; p < 0.001; Figure 7) covering almost the full incubation period indicated that CRDS-SCGIM produced results that were consistent with those of the traditional method. ...
Article
Full-text available
Rationale: Soil microbial heterotrophic C‐CO 2 respiration is important for C cycling. Soil CO 2 differentiation and quantification are vital for understanding soil C cycling and CO 2 emission mitigation. Presently, soil microbial respiration (SR) quantification models are based on native soil organic matter (SOM) and require consistent monitoring of δ ¹³ C and CO 2 . Methods: We present a new apparatus for achieving in situ soil static chamber incubation and simultaneous CO 2 and δ ¹³ C monitoring by cavity ring‐down spectroscopy (CRDS) coupled with a soil culture and gas introduction module (SCGIM) with multi‐channel. After a meticulous five‐point inter‐calibration, the repeatability of CO 2 and δ ¹³ C values by using CRDS‐SCGIM were determined, and compared with those obtained using gas chromatography (GC) and isotope ratio mass spectrometry (IRMS), respectively. We examined the method regarding quantifying SR with various concentrations and enrichment of glucose and then applied it to investigate the responses of SR to the addition of different exogenous organic materials (glucose and rice residues) into paddy soils during a 21‐day incubation. Results: The CRDS‐SCGIM CO 2 and δ ¹³ C measurements were conducted with high precision (< 1.0 µmol/mol and 1‰, respectively). The optimal sampling interval and the amount added were not exceeded 4 h and 200 mg C/100 g dry soil in a 1 L incubation bottle, respectively; the ¹³ C‐enrichment of 3%–7% was appropriate. The total SR rates observed were 0.6–4.2 µL/h/g and the exogenous organic materials induced ‐49%–28% of priming effects in native SOM mineralisation. Conclusions: Our results show that CRDS‐SCGIM is a method suitable for the quantification of soil microbial CO 2 respiration, requiring less extensive lab resources than GC/IRMS.
... Thus, various studies have added powders or solutions containing compounds found in exudates to sieved soil in an attempt to explore e.g. the mechanisms behind rhizosphere priming effects (Basiliko et al., 2012;Nottingham et al., 2012;Koranda et al., 2013;Wild et al., 2014;Girkin et al., 2018;Jilling et al., 2021) or changes in the microbial community (Shi et al., 2011;Mau et al., 2015;Papp et al., 2020). In contrast to root exudation which creates localized high C concentrations with strong concentration gradients (Kuzyakov and Razavi, 2019;Vetterlein et al., 2020), the substrate addition results in uniform distribution of moderately enhanced C availability throughout the whole soil volume, which may affect microbial responses to substrate input considerably. ...
... Changes in the chemical composition of DOM can have an influence on soil microbial characteristics, potentially leading to alterations in the availability and fractions of soil P. In general, the decomposition and mineralization of soil organic matter, caused by the input of fresh organic matter to the soil, has a substantial effect on the accessibility of P (Nottingham et al., 2012). Anderson et al. (1974) suggested that organic P has a tendency to adsorb onto soil minerals, thus making it difficult for microorganisms to utilize. ...
Article
Full-text available
The chemodiversity of exogenous dissolved organic matter (DOM) is significantly increased during the composting process, yet its influence on soil phosphorus (P) transformation is uncertain. A 60‐day incubation experiment was conducted to assess the availability and fractions of P in lime concretion black soil in response to exogenous DOM input, with three treatments: control (CK), soil with DOM from 30 days decomposed straw (DOM1), and soil with DOM from 90 days decomposed straw (DOM2). The results demonstrated that the addition of DOM caused an increase in the content of available P in the soil, particularly the addition of DOM1 with more protein‐like substances and a lower degree of humification. Compared to the control, DOM1 and DOM2 treatments showed an increase in available P content of 17.3%‐54.3% and 0.6%‐18.2%, respectively. However, the response of the soil's physicochemical and biological properties varied considerably. DOM1 treatment caused a marked decrease in soil pH compared to DOM2 treatment, which then led to the release of Ca ²⁺ and Mg ²⁺ and improved the solubility of inactive P. Moreover, DOM1 treatment was found to significantly raise the content of microbial biomass P (MBP), particularly at 60 days. The Mantel test and random forest analysis revealed that the strongest correlation was between MBP and available P. Furthermore, DOM1 treatment had a stimulating effect on the growth of phosphate‐solubilizing microorganisms, including Arthrobacter , Penicillium and Mortierella . This investigation could provide insights into how the chemodiversity of exogenous DOM affects the activation of legacy‐P in soil, thus enhancing the soil P supply capacity in agricultural systems.
Article
Full-text available
Tropical ecosystems face escalating global change. These shifts can disrupt tropical forests' carbon (C) balance and impact root dynamics. Since roots perform essential functions such as resource acquisition and tissue protection, root responses can inform about the strategies and vulnerabilities of ecosystems facing present and future global changes. However, root trait dynamics are poorly understood, especially in tropical ecosystems. We analyzed existing research on tropical root responses to key global change drivers: warming, drought, flooding, cyclones, nitrogen (N) deposition, elevated (e) CO 2 , and fires. Based on tree species‐ and community‐level literature, we obtained 266 root trait observations from 93 studies across 24 tropical countries. We found differences in the proportion of root responsiveness to global change among different global change drivers but not among root categories. In particular, we observed that tropical root systems responded to warming and eCO 2 by increasing root biomass in species‐scale studies. Drought increased the root: shoot ratio with no change in root biomass, indicating a decline in aboveground biomass. Despite N deposition being the most studied global change driver, it had some of the most variable effects on root characteristics, with few predictable responses. Episodic disturbances such as cyclones, fires, and flooding consistently resulted in a change in root trait expressions, with cyclones and fires increasing root production, potentially due to shifts in plant community and nutrient inputs, while flooding changed plant regulatory metabolisms due to low oxygen conditions. The data available to date clearly show that tropical forest root characteristics and dynamics are responding to global change, although in ways that are not always predictable. This synthesis indicates the need for replicated studies across root characteristics at species and community scales under different global change factors.
Article
Full-text available
The tropical forests of the Amazon Basin occur on a wide variety of different soil types reflecting a rich diversity of geologic origins and geomorphic processes. We here review the existing literature about the main soil groups of Amazonia, describing their genesis, geographical patterns and principal chemical, physical and morphologic characteristics. Original data is also presented, with profiles of exchangeable cations, carbon and particle size fraction illustrated for the principal soil types; also emphasizing the high diversity existing within the main soil groups when possible. Maps of geographic distribution of soils occurring under forest vegetation are also introduced, and to contextualize soils into an evolutionary framework, a scheme of soil development is presented having as its basis a chemical weathering index. We identify a continuum of soil evolution in Amazonia with soil properties varying predictably along this pedogenetic gradient.