SOILSOILSOILSOIL2199-398XCopernicus PublicationsGöttingen, Germany10.5194/soil-2-601-2016Long-term elevation of temperature affects organic N turnover and associated N2O emissions in a permanent grassland soilJansen-WillemsAnne B.anne.jansen@teagasc.ieanne.willems@bot2.bio.uni-giessen.deLaniganGary J.CloughTimothy J.AndresenLouise C.https://orcid.org/0000-0002-4890-889XMüllerChristophTeagasc Johnstown Castle, Wexford, Co. Wexford, IrelandInstitute for Plant Ecology, JLU Giessen, Heinrich-Buff-Ring 26–32, 35390 Giessen, GermanyDepartment of Soil and Physical Sciences, Faculty of Agriculture and Life Sciences, Lincoln University, 7647 Lincoln, New ZealandDepartment of Earth Science, University of Gothenburg, Gothenburg, SwedenSchool of Biology and Environmental Science, University College Dublin, Dublin, IrelandAnne B. Jansen-Willems (anne.jansen@teagasc.ie, anne.willems@bot2.bio.uni-giessen.de)30November20162460161430May201621June201630September201622October2016This work is licensed under a Creative Commons Attribution 3.0 Unported License. To view a copy of this license, visit http://creativecommons.org/licenses/by/3.0/This article is available from https://soil.copernicus.org/articles/2/601/2016/soil-2-601-2016.htmlThe full text article is available as a PDF file from https://soil.copernicus.org/articles/2/601/2016/soil-2-601-2016.pdf
Over the last century an increase in mean soil surface temperature has been
observed, and it is predicted to increase further in the future. In order to
evaluate the legacy effects of increased temperature on both nitrogen (N)
transformation rates in the soil and nitrous oxide (N2O) emissions, an
incubation experiment and modelling approaches were combined. Based on
previous observations that gross N transformations in soils are affected by
long-term elevated-temperature treatments we hypothesized that any associated
effects on gaseous N emissions (e.g. N2O) can be confirmed by a change
in the relative emission rates from various pathways. Soils were taken from a
long-term in situ warming experiment on temperate permanent grassland. In
this experiment the soil temperature was elevated by 0 (control), 1, 2 or
3 ∘C (four replicates per treatment) using IR (infrared) lamps over a period of 6
years. The soil was subsequently incubated under common conditions
(20 ∘C and 50 % humidity) and labelled as NO315NH4
Gly, 15NO3NH4 Gly or NO3NH415N-Gly. Soil
extractions and N2O emissions were analysed using a 15N tracing
model and source-partitioning model. Both total inorganic N
(NO3-+ NH4+) and NO3- contents were higher in soil
subjected to the +2 and +3 ∘C temperature elevations (pre- and
post-incubation). Analyses of N transformations using a 15N tracing
model showed that, following incubation, gross organic (but not inorganic) N
transformation rates decreased in response to the prior soil warming
treatment. This was also reflected in reduced N2O emissions associated
with organic N oxidation and denitrification. Furthermore, a newly developed
source-partitioning model showed the importance of oxidation of organic N as
a source of N2O. In conclusion, long-term soil warming can cause a legacy
effect which diminishes organic N turnover and the release of N2O from
organic N and denitrification.
N2O production via four processes (nitrification, denitrification,
co-denitrification and oxidation of organic N). Three uniformly distributed
pools were considered. These pools were ammonium (NH4+) with
a 15N atom fraction of an, nitrate (NO3-) with a
15N atom fraction of ad and organic N with a 15N atom
fraction of ao (= 0.003663). The N2O produced via co-denitrification
consists of one N atom from the nitrate pool and one from the organic N pool.
Introduction
Globally, managed pastures were estimated to occupy 34.7 million km2
in 2000, and this area is projected to increase by a further
13.4 % by 2050 (Tilman et al., 2001). Concomitantly, the Earth's mean
surface temperature has increased by 0.6 ∘C in the past century
with surface temperatures expected to increase by a further
1.5–4.5 ∘C resulting from a doubling of the atmospheric carbon
dioxide (CO2) concentration (IPCC, 2013). Agricultural
soils play a central role in the global carbon (C) and nitrogen (N) cycles
(French et al., 2009), and C–N interactions are to a large extent
affected by temperature (Luo, 2007). Thus, research into the effect of
elevated soil temperatures is essential to better understand biogeochemical
N cycling in grassland ecosystems.
Previous research generally showed an increase in both net (Peterjohn et
al., 1994; Rustad et al., 2001; Norby and Luo, 2004; Butler et al., 2012;
Bai et al., 2013; Björsne et al., 2014; X. Z. Zhang et al., 2015) and gross
(Larsen et al., 2011; Björsne et al., 2014) N mineralization under
elevated soil temperatures. However, not all studies found this effect
(Emmett et al., 2004; Niboyet et al., 2011; Andresen et al., 2015). An
effect on N immobilization or nitrification was generally not observed
(Emmett et al., 2004; Barnard et al., 2005; Andresen et al., 2010;
Niboyet et al., 2011; Bai et al., 2013; Björsne et al., 2014). In their meta-analyses, Dijkstra et al. (2010) and Bai et al. (2013) identified increases in inorganic N under elevated soil temperatures.
Most of this inorganic N increase occurred as nitrate (NO3-)
(Dijkstra et al., 2010). Peterjohn et al. (1994) also found that
average monthly ammonium (NH4+) concentrations increased in a
mineral soil under forest; however, daily average concentrations did not
differ. In the same study, no differences in NO3- concentrations
were observed, and the amount of extractable NO3- was very small.
Another meta-analysis showed no effect of soil warming on total soil N,
NH4+ or NO3- in a Tibetan grassland (X. Z. Zhang et al.,
2015). Other studies also found no effect of soil warming on total soil N
(Bai et al., 2013) and inorganic N (Larsen et al., 2011).
N mineralization follows a step-wise sequence of protein depolymerization by
extracellular activity to oligomers (e.g. peptides) and monomers (e.g. amino
acids) and then uptake by microorganisms before mineralization to
NH4+ (Schimel and Bennett, 2004). Hence, the production of peptides
and amino acids as well as the mineralization of amino acids, affects the main
fluxes regulating gross N mineralization. Amino acids have a short residence
time in the soil due to either rapid assimilation by soil microbes or
mineralization, which occurs within a few hours (Farrell et al., 2014).
In heathland and grassland soils, no effect of soil warming on the amino acid
concentration has been observed (Chen et al., 2014; Andresen et al., 2015).
Nitrous oxide (N2O), a potent greenhouse gas with a global warming
potential of 298 on a 100-year basis, can be produced by several processes,
such as nitrification, partial denitrification, co-denitrification and the
oxidation of organic matter (Butterbach-Bahl et al., 2013; J. Zhang et
al., 2015) (Fig. 1). Laughlin and Stevens (2002) confirmed the
importance of co-denitrification for N2 production, a process that may
comprise 25 % of the total N balance in pastures (Selbie et al., 2015).
Müller et al. (2014) found that, for the same grassland soil as used in
this study, co-denitrification contributed 17.6 % of the total N2O
production. N2O emissions following fertilization with ammonium
nitrate (NH4NO3) may be greater than from urea fertilizer because of the
greater susceptibility to denitrification (Harrison and Webb, 2001). The
amount and form of N inputs primarily govern N2O emissions, with further
impacts resulting from climatic factors, such as temperature and
precipitation, and soil factors, such as C availability and microbial
community structure (Harrison and Webb, 2001; Müller et al., 2003;
Stark and Richards, 2008; Laughlin et al., 2009; Li and Lang, 2014).
However, the impact of elevated soil temperature on N2O production, in
semi-natural grasslands is unclear (Peterjohn et al., 1994; Bijoor et
al., 2008; Larsen et al., 2011). Furthermore, there has been very limited
research into the effect of elevated soil temperature on the different
N2O production processes. Maag and Vinther (1996) observed a
decrease in nitrification-associated N2O emissions and an increase in
denitrification-associated N2O with increasing soil temperature. It has
been suggested that this was due to the creation of anoxic conditions and the
associated depletion of oxygen following the increase in microbial
respiration with higher soil temperatures (Castaldi, 2000). Prolonged
elevated soil temperatures, on the other hand, could also lead to changes in
the microbial community (Avrahami and Conrad, 2003; French et al., 2009).
Several methods, such as source partitioning, have been used to quantify the
contributions of individual N pools to N2O emissions (Stange et al.,
2009, 2013; Rütting et al., 2010; Zhang et al., 2011; Zhu et al., 2011;
Müller et al., 2014). However, one of the assumptions of
the source-partitioning method is the absence of hybrid reactions such as
co-denitrification (J. Zhang et al., 2015). Because of the potential
importance of co-denitrification for the N2O production, it should not
be omitted from the analysis of N2O sources. Currently, only one
technique is available to identify several processes including a hybrid
reaction, which is a full 15N tracing approach (Müller et al.,
2014). This approach, however, requires data on NO2-,
NO3- and NH4+ pool sizes and measurements at multiple time
points. Furthermore, it requires at least multiple days of running the model
to be able to distinguish the different processes. A straightforward method
partitioning N2O fluxes into several pathways including a hybrid
reaction, which does not rely on measurements of NO2- and data at
multiple time points, would therefore be very beneficial.
The objectives of this study were to quantify the legacy effects of 6 years of elevated temperature (via IR heaters) on soil N cycling dynamics,
including (1) net and gross N transformation rates in the soil, (2) N2O
fluxes immediately after fertilization and (3) the processes responsible for
these N2O fluxes. Net and gross transformation rates were determined
using an extended version of a basic 15N tracing model described by
Müller et al. (2007). Since the publication of this basic model in 2007,
more than 50 peer-reviewed papers have been published, where the basic model
or modifications of the basic model have been used, demonstrating the robustness of the approach in various soils, ecosystems and climatic
conditions. To determine the processes involved in N2O production, a
new source-partitioning method was developed to allow the identification of
hybrid reactions. This source-partitioning method is a newly developed
method and not a modification of the 15N tracing model. To identify
the legacy effect of different in situ temperature treatments on the
internal N transformation processes, soil incubations were carried out under
identical moisture and temperature conditions in the laboratory. Based on
previous observations that gross N transformations in soils are affected by
long-term elevated-temperature treatments, we hypothesized that any
associated effects on gaseous N emissions (e.g. N2O) can be confirmed
by a change in the relative emission rates from various pathways. Thus, the
newly developed source-partitioning method would be helpful to confirm such a change.
Material and methodSite description and field treatment
The 100 m2 site was established on a permanent grassland of the
“Environmental Monitoring and Climate Impact Research Station Linden” in
Germany (50∘31.6′ N, 8∘41.7′ E). A full description of
the site can be found in Jansen-Willems et al. (2016). Briefly,
the site has been managed as a meadow with two cuts per year and fertilized
with 50–80 kg N ha-1 yr-1 for the last 3 decades. Since
1995, the N fertilizer input has been reduced to 40 kg N ha-1 yr-1,
as KAS (calcium–ammonium–nitrate). The mean annual temperature and precipitation
were 9.5 ∘C and 560 mm (observation period: 1995–2014), respectively.
The site was divided into 16 plots: four rows of four plots. The
16 plots were, according to a Latin square design, assigned to one of four
treatments. From 28 January 2008, the soil temperature of each plot,
measured at 5 cm depth, was elevated by 0, 1 (mean 0.8 standard error (SE) 0.02),
2 (mean 1.9 SE 0.03) or 3 (mean 2.6 SE 0.03) ∘C
above ambient temperature, using infrared heaters. The use of heaters will
also affect the soil moisture content. The temperature treatments (including
any moisture effect) are referred to as Tcontrol, T1, T2 and
T3, respectively. The infrared heaters were installed at different
heights to create the different temperature elevations (Jansen-Willems et al., 2016).
Incubation, labelling and extraction
On the day the heaters were turned off, all soil within a circular area of
318 cm2 directly underneath each infrared lamp was excavated to 7.5 cm
for the tracing experiment. A small subsample of each plot was dried at
70 ∘C for 48 h, ground and analysed by a CNH Macro Elemental
Analyser (Hanau, Germany) for total N content. A subsample of the soil for
each plot was dried at 105 ∘C for 24 h to determine the soil
gravimetric water content. The remaining field-moist soil was kept at
4 ∘C (for less than 60 h) until further analysis whereupon the
soil from each field plot was sieved through a 10 mm sieve to homogenize it
and to remove roots. Incubations were carried out in 750 mL jars (WECK GmbH
u. Co. KG, Wehr, Germany). Thirteen jars per field plot were prepared each
with an average of 67 (SD 8.4) g dry soil per jar (except for plots 3, 5,
7, 11 and 14, where only 10 jars were prepared due to lack of soil). All
jars were closed with glass lids that were fitted with septa to allow for
gas sampling. During gas flux analyses the jars were sealed using a clamp
and a rubber ring between the jar and the lid. At other times a gap was left
between the jar and the lid to allow air exchange while minimizing water
loss. Two days after soil sampling (day -55), all jars were put in a dark
climate chamber at 20 ∘C and 50 % humidity and incubated for
55 days prior to 15N labelling (day 0).
Soil gravimetric moisture data were used to determine the exact amount of
dry soil in each jar and to calculate the amount of water to be added to
ensure the same soil water content in each jar. On day -53 the soil moisture
in each jar was adjusted to a water-filled pore space (WFPS) of 64 %. On
day -43 and -5 the jars were watered to replenish the water lost due to evaporation.
For the 15N tracing study three different labels were used,
NO315NH4 Gly, 15NO3NH4 Gly and
NO3NH415N-Gly (at 60, 60 and 99 atm % 15N, respectively). All solutions contained 50 µg NO3-N, 50 µg NH4-N
and 30 µg Gly-N g-1 soil. On day 0, the label was added to each jar using a needle with side ports to inject the
solution into the soil to minimize disturbance while providing an equal
distribution in the soil (Müller et al., 2007). For each field plot,
jars were set up for four soil extractions, at day 0, 1, 3 and 6, and three labels, except for plot 3, 5, 7, 11 and 14, where, due
to the lack of soil, no NO3NH415N-Gly label addition was possible.
The soil in each jar was extracted with 2 M KCl using the blending procedure
of Stevens and Laughlin (1995). The 15N enrichments of
NO3- and NH4+ in the extracts were determined by
converting NO3- and NH4+ into N2O following the
procedures by Stevens and Laughlin (1994) for determination of the 15N
enrichment in NO3- and Laughlin et al. (1997) for the 15N
enrichment in NH4+. The extraction of soil prior to 15N
addition took place on day -2. The other extractions took place at
0.11 days (±0.004), 1.02 days (±0.001), 2.95 days (±0.001) and
5.93 days (±0.001) after 15N substrate addition and are hereafter
referred to as day 0, 1, 3 and 6.
Gas sampling
Gas samples were taken from 43 different jars (one jar per 15N label) for each plot. During the pre-incubation gas samples were taken 1, 46 and
48 days before label addition. After labelling, gas samples were taken
immediately prior to soil extractions.
Gas samples were taken using a 60 mL syringe (Ecoject Plus, Gelnhausen,
Germany). At time zero (t0) 15 gas samples were taken from 15 different
jars. Then at time 1 (t1) a gas sample was taken through the rubber
septum. At both t0 and t1 the syringe was flushed twice with
headspace gas to ensure a representative sample was taken. The times between
t0 and t1 during each of the seven different gas samplings (three
before label addition and four immediately prior to extraction) were 120–129,
120, 180, 233, 240, 235 and 214 min, respectively. Gas samples
were analysed within 24 h after sampling using a gas chromatograph (GC; Bruker) equipped with
an electron capture detector (ECD) for N2O analysis. An average of the
concentrations measured in the 15 samples was used as the t0
concentration for all 43 jars. Fluxes were based on the parts per million and time
difference between t0 and t1. They were calculated using the
constant gas law, with ambient pressure, and temperature was assumed to be
20 ∘C (the temperature of the incubation room). The fluxes were
then converted to a per dry gram basis.
For the 15N abundance of N2O, a 30 mL sample was taken at t1
and transferred to a 12 mL Exetainer® vial
(Labco Ltd, High Wycombe, Buckinghamshire, UK). The over-pressurized sample
vials were returned to ambient pressure immediately before analyses of
stable isotopes. This was performed using a double-ended needle fixed
vertically in a clamp stand with the ventral needle submerged 3–4 mm in a
beaker of water and the gas sample held upside down and pushed onto the
dorsal needle. The excess pressure in the sample vial was thus released
causing the water to bubble until the pressure inside the vial had
equilibrated with the ambient atmospheric pressure. Cessation of bubbling
implied equal pressure had been reached. The 15N enrichments of
15N2O and 15N2 were determined using an automated
isotope ratio mass spectrometer (Sercon Ltd 20-20), as described by
Stevens et al. (1993), interfaced to a TGII cryofocusing unit (Sercon
Ltd 20-20). The detection limit for atom % 15N of a 50 ppm N2O
standard gas was 0.00003 (n= 10); SD was 0.00009 atom % 15N.
Respective values for a 0.4 ppm N2O standard were higher (0.00084 (n= 10); SD 0.003).
15N tracing model
The 15N tracing analysis tool described by Müller et al. (2007)
was used to quantify gross soil N transformations. In the current study, the
only changes to the original model were the addition of an amino acid
(glycine) pool and the transformations to and from this pool. The model
(Fig. 2) considered 7 N pools and 13 N transformations. The N pools
were NH4+, NO3-, amino acid glycine (AA), labile (Nlab)
and recalcitrant (Nrec) organic N, adsorbed
ammonium (NH4ads+), and stored nitrate (NO3sto-).
Description of N transformations and average gross N fluxes per
treatment (diagram shown in Fig. 2). Standard deviation in brackets.
K stands for kinetics, where 0 implies the use of zero-order and 1 the use of
first-order kinetics in the model. The p is the p value of the one-way
ANOVA, with ns (non-significant) if p> 0.1 (p value in bold if
< 0.05). For the Holm–Šídák pairwise comparisons, t indicates a tendency to be
different from control (p< 0.10).
Transformation KAverage gross flux (µg N g soil-1 day-1) pTcontrolT1T2T3MNrecMineralization of Nrec to NH4+ or AA03.18(1.95)5.42(2.50)0.91(0.73)1.35(0.90)0.040INH4NrecImmobilization of NH4+ to Nrec116.12(9.23)13.43(6.92)17.45(6.53)4.72(3.65)nsMNlabMineralization of Nlab to NH4+ or AA135.86(16.49)28.01(8.92)36.14(10.17)35.43(8.78)nsINH4NlabImmobilization of NH4+ to Nlab130.59(19.34)22.28(14.65)30.54(8.82)29.59(19.78)nsONrecOxidation of Nrec to NO3-03.64(0.96)1.99(1.31)2.02(0.56)2.92(1.34)nsINO3Immobilization of NO3- to Nrec15.64(2.74)2.15(1.31)4.57(2.62)4.97(3.10)nsONH4Oxidation of NH4+ to NO3-115.40(2.30)11.64(1.65)14.21(1.92)15.26(2.58)nsDNO3Dissimilatory NO3- reduction to NH4+00.18(0.05)0.24(0.12)0.36(0.12)0.14(0.10)nsANH4Adsorption of NH4+134.26(19.67)20.41(19.61)23.64(11.50)15.81(12.84)nsRNH4aRelease of adsorbed NH4+133.22(21.43)20.51(12.33)24.77(6.15)16.41(9.07)nsANO3Adsorption of NO3-128.08(14.18)55.23(37.72)82.39(58.45)62.99(47.75)nsRNO3sRelease of stored NO3-123.70(10.48)53.23(10.63)78.49(36.84)59.96(22.29)0.096MAAMineralization of AA to NH4+132.21(7.67)17.40(4.32)27.29(9.52)15.32(3.63)t0.045
15N tracing model for analyses of gross soil N transformation
rates. Abbreviations of the transformations are explained in Table 1. The pools
are explained in Sect. 2.4.
The initial NO3- and NH4+ pool sizes were determined by
extrapolating the first two extraction times back to time zero. The initial
AA pool size was set to 30 µg N g-1 soil, corresponding to the
application of glycine (Gly). The initial NH4ads+ and
NOsto3- were based on the difference between the added and
initial N (Müller et al., 2004). The initial pool sizes for organic N
(Nrec and Nlab) were based on previous field measurements.
However, these organic N values were not critical because for Nrec,
zero-order kinetics were used (independent of initial pool size), and for
Nlab, the quick turnover time ensures that a small pool will be
governed quickly by the dynamics of the in- and out-flowing rates.
The N transformations are described in Table 1. The N transformations were
calculated based on zero- or first-order kinetics (Table 1). Whether
Nlab and Nrec were transformed into AA or NH4+ was
determined by two factors: one for MNlab and one for MNrec.
These factors determine the fraction of the MNlab or MNrec flowing into
the AA pool, with the remainder entering the NH4+ pool. For each
temperature treatment the kinetic parameters and the two split factors were
simultaneously optimized by minimizing the misfit between the modelled and
measured NH4+ and NO3+ concentrations and their
respective 15N enrichments (Müller et al., 2004). For treatment
T2 the measurements of the 15N Gly label were not included in the
optimization because only one replicate was available for this label.
A Markov chain Monte Carlo Metropolis algorithm (MCMC-MA) was used for the
optimization, which practices a random walk technique to find global minima
(Müller et al., 2007). The uncertainties (standard deviation) of the
observations were taken into account by the optimization routine. The
MCMC-MA routine was programmed in MatLab-Simulink (Mathworks Inc) as
described in Müller et al. (2007). The most suitable parameter set
was determined using the Akaike information criterion (AIC). Gross and net
nitrification and gross and net mineralization were calculated using
Eqs. (1)–(4), in which SF stands for split factor. The combined standard
deviation was calculated by ((SD rate 1)2+ (SD rate 2)2+ …)0.5,
in which the SD of MNx. SFMNx is the SD
of MNx multiplied by the SF.
The following combined rates were calculated.
Grossnitrification:ONrec+ONH4Netnitrification:ONrec+ONH4-INO3-DNO3Grossmineralization:MNlab⋅SFMNlab+MNrec⋅SFMNrec+MAANetmineralization:MNlab⋅SFMNlab+MNrec⋅SFMNrec+MAA-INH4Nrec-INH4Nlab-INO3
Determining the contribution of different processes to N2O flux
The N2O fluxes, from the soil labelled with NO315NH4 Gly
and 15NO3NH4 Gly, were separated into four different
processes. These were nitrification, denitrification, co-denitrification and
oxidation of organic matter. The N2O was assumed to be derived from
three uniformly distributed pools and based on initial substrate 15N
enrichments; isotopic discrimination was considered negligible for all four
processes. The pools and processes accounting for the N2O production
are shown in Fig. 1. The 15N content of the organic matter was
considered to be at natural abundance (0.3663 atom %). The N2O
produced via co-denitrification consists of one N atom from the
NO3- pool and one N atom from the organic N pool. The chance that
the N2O produced via nitrification, denitrification or oxidation of
organic N contains zero, one or two 15N enriched atoms can be described
by Eqs. (5), (6) and (7), respectively, where ax (the 15N fraction
of the pool) is an for nitrification, ad for denitrification and
ao for the oxidation of organic N; an, ad and ao are
explained in Fig. 1.
Chanceofzero15Natoms:1-ax2Chanceofone15Natom:21-axaxChanceoftwo15Natoms:ax2
The chance that the N2O produced via co-denitrification consists of
zero, one or two 15N enriched atoms is described by Eqs. (8), (9) and (10), respectively.
Chanceofzero15Natoms:1-ad1-a0Chanceofone15Natom:ad1-a0+a01-adChanceoftwo15Natoms:ada0
The chance that the N2O in the gas sample contains zero, one or two
15N atoms is described by Eqs. (11), (12) and (13), respectively, where
the subscripts “d”, “n” and “o” refer to the fractions of N2O produced by
denitrification, nitrification and oxidation of organic N, respectively. The
fraction of N2O produced by co-denitrification is 1-d-n-o as all of the
N2O produced was assumed to come from one of the four processes.
Chanceofzero15Natoms:n1-an2+d1-ad2+o1-ao2+(1-n-d-o)1-ad1-a0Chanceofone15Natom:2n1-anan+2d1-adad+2o1-aoao+(1-n-d-o)ad1-a0+a01-adChanceoftwo15Natoms:nan2+dad2+oan2+(1-n-d-o)ada0
The automated continuous-flow isotope ratio mass spectrometer enabled the
measurement of 45R (45I/44I) and 46R
(46I/44I), where xI is the ion currents at m/zx. The 45R
and 46R were corrected for the presence of 18O. This, therefore,
means that 45R is the fraction of N2O molecules containing one
15N atom divided by the fraction of N2O molecules containing zero
15N atoms, and 46R is the fraction of N2O molecules
containing two 15N atoms divided by the fraction of N2O molecules
containing zero 15N atoms. The expected fractions are described by
Eq. (11)–(13), where ao was set to 0.003663 and an and ad
were considered to be the 15N abundance of NH4+ and
NO3-, respectively, while n, d and o were quantified using the
fminsearchbnd function in MatLab (The MathWorks Inc, Natick, MA). For this the 45R,
46R, an and ad of soil labelled with NO315NH4
Gly and soil labelled with 15NO3NH4 Gly were used. The amount
of N2O produced via each process was calculated by multiplying the
average N2O flux from the jars labelled with NO315NH4
Gly and 15NO3NH4 Gly with the fractions of N2O produced
by the four different processes. This was carried out separately for each
plot and time step. Because of missing 15NH4 data, the different
processes were not distinguished for plot 1, time step 3. Total N2O flux
contributions were calculated using linear interpolations between time steps.
NH4-N content (a), NO3-N content (b) and
N2O emissions (c) at the extraction times. Time point 0 is the
time of label addition (15NH4NO3 Gly, NH415NO3 Gly or
NH4NO315N Gly). The ammonium and nitrate content at time point 0
is based on unlabelled soil. The N2O flux at time point 0 is based on the
average flux of the three gas samplings before label addition. The error bars are
the standard error of the mean; * shows a significant difference in
NH4-N from Tcontrol (p< 0.03), # shows a significant
difference in NO3-N from Tcontrol (p< 0.0001), and
Δ shows a significant difference in N2O flux from
Tcontrol (p< 0.05).
Modelled vs. measured data. The lines are modelled data, and the squares,
circles and triangles are the measured data points. Error bars are standard
deviations. Time is the time in days from the moment of label addition.
Statistical analyses
Treatment differences in total soil N were analysed with the non-parametric
Kruskal–Wallis test using IBM SPSS statistics (version 22) because one
sample per plot was taken, resulting in only four measurements per
treatment. The effect of treatment N2O fluxes (including different
processes) and inorganic N (NO3-+NH4+), NO3-
and NH4+ concentrations was analysed using the MIXED procedure in
SAS (Version 9.3, SAS institute). The N2O fluxes were transformed using
log(flux + 10). The N2O fluxes via the different processes were
transformed using flux1/4. A Tukey–Kramer adjustment was used to
correct for multiplicity effects in pairwise comparisons. Residual checks
were made to ensure that the assumptions of the analysis were met. The
effect of treatment on modelled N transformation rates was analysed using a
one-way ANOVA based on the averages and standard deviations in Matlab
(Version 2013b, The MathWorks Inc.). The pairwise comparisons were
calculated with the Holm–Šídák test in SigmaPlot (Version 11.0, Systat Software Inc.).
ResultsSoil nitrogen pool sizes
Total soil N content did not differ between soil warming treatments prior to
the incubation study. A significant interaction between treatment and time
affected soil NH4+ concentrations; thus, these results are
therefore given separately for each time step. No such interaction was found
for NO3- or total inorganic N (NO3-+NH4+)
concentrations. The total inorganic N content differed with temperature
treatment (p< 0.0001) (all pairwise comparisons were also
significant; p< 0.0001). The total inorganic N content was in the
order of T1<Tcontrol<T3<T2.
Soil NH4+ concentrations increased from 2 µ g N g-1
soil to between 28 and 54 µg N g-1 soil upon label addition, and
subsequently decreased over the next 5 days to ca. 9 µg N g-1
soil (Fig. 3a). Soil NH4+ concentrations did not differ as a
result of the soil warming treatments on either days 0 or 6. However, on day 1,
treatment T1 had a lower NH4+ concentration compared to
all other treatments (p< 0.029), while the soil NH4+
concentration in the T2 treatment was higher than in the Tcontrol
or T1 treatments (p< 0.001). Three days after label addition
the NH4+ concentration in the T1 treatment remained lower
compared to the T2 and T3 treatments (p< 0.001 and 0.044, respectively).
After the initial increase in NO3- due to label addition, the
NO3- concentrations continued to slowly increase over the
following 6 days (Fig. 3b). NO3- concentrations were
significantly different among the treatments (p< 0.001), with
differences also occurring with respect to the initial NO3-
concentrations prior to label addition (p< 0.001). The highest
NO3- concentrations occurred in the T2 treatment followed by
the T3 and Tcontrol, while the lowest NO3- concentration
was observed in the T1 treatment.
Soil N transformations
The modelled and observed concentrations and 15N enrichments were in
good agreement with R2> 0.97 for all runs (Fig. 4). The
gross rates of most N transformations did not differ as a result of the
previously imposed soil warming treatment (Table 1). However, the rates of
recalcitrant N mineralization were reduced under the T2 and T3
treatments (p= 0.040). Mineralization of amino acids also became slower
with increasing temperatures (p= 0.045). However, the overall gross
mineralization of organic N to NH4+ did not differ with the
previously imposed warming treatments. This was because the mineralization
of labile organic N was the major contributor to total mineralization, and
this rate was not significantly affected by previous warming (Table 2). Net
mineralization did not differ as a result of the previously imposed warming
treatments. Despite the fact that the release of stored NO3-
tended to increase with warming (p= 0.096) and also that cumulative
ONH4 and ONrec rates tended to be different (p= 0.095), no
significant effect on net nitrification could be observed (Table 2).
Cumulative N2O flux via four processes between 3 h and 6 days
after labelling. N2O fluxes based on average flux from soil labelled with
15NH4NO3 Gly or NH415NO3 Gly. The cumulative flux
per process is an average over the four plots per treatment. Error bars are
standard error of the mean (SEM). Percentages are the average percentage of
flux produces via each process; SEM between brackets; * indicates significantly
lower cumulative flux compared to the control (p< 0.05).
Gross mineralization (MinGross), net mineralization (MinNet),
gross nitrification (NitGross) and net nitrification (NitNet)
rate in µg N g soil-1 day-1. Including the contributions
from the different N pools for the gross transformations (italics), where
Nlab is a labile organic N pool, Nrec is a recalcitrant
organic N pool, NH4+ is the ammonium pool and NAA is the
amino acid Gly pool. t indicates one-way ANOVA tendency p< 0.1.
In response to N supply, N2O emissions immediately increased and
decreased thereafter (Fig. 3c). While treatments T2 and T3 had
lower N2O fluxes than the control treatment (p= 0.004 and p= 0.036,
respectively), no interaction between incubation time and treatment was
observed. The N2O fluxes from the T2 treatment were also lower
than those from the T1 treatment (p= 0.016). However, observed fluxes
from the T1 treatment did not differ from the control treatment and
N2O fluxes from the T2 treatment did not differ from the T3 treatment.
The newly developed partitioning model was successful to identify cumulative
N2O fluxes (Fig. 5) and N2O contribution at each extraction time
(Fig. 6) associated with nitrification, denitrification, co-denitrification
and the oxidation of organic N between 0.11 and 5.93 days after N addition.
The oxidation of organic N was the main source of N2O at all sampling
dates, comprising between 63 and 85 % of the total N2O flux (Fig. 5).
The percentage contribution made by organic N to N2O fluxes increased
over the sampling period, rising from a minimum of 40 % in the control
treatment to virtually 100 % across all treatments by day 6 (Fig. 6). The
fluxes from organic N oxidation were the highest in the control treatment,
followed by T1, and lowest for T2 and T3. Significant
differences were found between the control and the T2 and T3
treatment (p= 0.011 and p= 0.002, respectively) and between T1 and
T3 (p= 0.039). The amount of N2O produced via denitrification was
also the highest under the control treatment, followed by T1 and
T3. It was the lowest under T2. Compared to the control treatment,
denitrification contributed less to N2O under the T2 and T3
treatments (p< 0.0001 and p= 0.002, respectively). The
contribution of denitrification also differed between treatments T2 and
T1 (p= 0.004). Co-denitrification only contributed to the N2O
flux during the first day after substrate addition. The highest amount of
N2O produced via co-denitrification was found under the control
treatment, followed by T1. Under T2 and T3 treatments, the
contribution of co-denitrification was minor. However, these differences
were not significant. No significant differences were found in the amount of
N2O produced via nitrification.
N2O flux divided into four processes at different time points after
fertilization. N2O fluxes based on average flux from soil labelled with
15NH4NO3 Gly or NH415NO3 Gly. The portrayed flux
per process is an average over the four plots per treatment. Error bars are
standard error of the mean. The scale of the y axis is different for each time point.
Discussion
Prior to incubation the inorganic N, as well as the NO3-
concentrations, were higher in the T2 and T3 treatments as a
result of the 6-year warming treatment. This suggests that a sustained
increase in temperature led to an increase in net mineralization and net
nitrification. This is in line with previous studies showing increases in
net mineralization in response to warming (Peterjohn et al., 1994; Rustad
et al., 2001; Norby and Luo, 2004; Bai et al., 2013; Björsne et al.,
2014; X. Z. Zhang et al., 2015). An increase in net nitrification in response to
soil warming, while less common, has also been shown (Barnard et al.,
2005; Bai et al., 2013; Björsne et al., 2014; X. Z. Zhang et al., 2015). Both
could be due to infield temperatures being more favourable for optimal
microbial activity. In agreement with previous research (Bai et al.,
2013; X. Z. Zhang et al., 2015), the total soil N pool did not differ among
warming treatments. This result may be due to the fact that the relative
sizes of the N pools differ: since the total soil N pool is significantly
larger than the inorganic N pool, it may take longer to register a change
(Galloway et al., 2008; Bai et al., 2013).
During incubation all soil was kept at 20 ∘C, regardless of the
in-field treatment to investigate any legacy impacts of sustained soil
warming on inherent soil N cycling. It has been suggested that changes in
the microbial community structure could alter the sensitivity of the
microbial community to temperature shifts (Balser et al., 2006). While
both net and gross mineralization rates did not differ as a result of the
previously imposed soil warming treatments, the mineralization of
recalcitrant N and mineralization of amino acids did differ. Lowest rates
were found under T2 (MNrec) and T3 (MNrec and MAA).
A similar effect to warming was found by Jamieson et al. (1998), who
reported decreased gross N mineralization rates in spring following winter
warming of soil. Adaptation of the microbial community, altering the
sensitivity to temperature shifts, could possibly provide an explanation why
no differences in net and gross mineralization and even decreases in
individual mineralization rates were found. However, no data were available
to test this hypothesis. Another possible explanation for the reduction in
mineralization rates could be a depletion of substrate due to the 6 years of elevated temperatures.
Previous research in heathland and grassland soils showed no significant
effect of warming on amino acid mineralization rates (Andresen et al.,
2015). The lower rates in the current study, however, could be due to a
change in amino acid oxidase activity (Vranova et al., 2013). Another
possible explanation for the lower amino acid mineralization rates could be
an increase in direct microbial assimilation of amino acids (Farrell et
al., 2014), since direct assimilation of glycine and larger amino acids is
well known (Barraclough, 1997; Andresen et al., 2009, 2011). Chen et al. (2015),
however, did not show an effect of warming on the microbial uptake
of amino acids. The fact that NH4+ immobilization rates were not
affected by previously imposed warming in the current study is in line with
previous research (Niboyet et al., 2011; Bai et al., 2013; Björsne et
al., 2014). It has been suggested that the depletion of labile C due to
warming might initiate a decrease in immobilization rates (Bai et al.,
2013). In the current experiment a labile carbon source (Gly) was added to
the soil, which could explain why no reduction in NH4+ immobilization was found.
Nitrous oxide emissions were highest shortly after label addition and
declined thereafter. Thus, initial higher rates from NH4+ and
NO3- were due to label addition. The higher absolute rate of
organic N oxidation at the start of the incubation did not come solely from
the Gly addition. If this had been the case, highest N2O 15N
enrichment would have been observed at the first measurement following
addition of the NO3NH415N-Gly label. However, for all
treatments the highest 15N enrichment of N2O was found in the
second measurement after label addition. The lower net rates of N2O
production at the end of incubation period could possibly have been caused
by N2O consumption; however, the consumption of pathway-specific
N2O emissions cannot be evaluated with the current model. However, as
WFPS was set to 64 %, it is unlikely that N2O consumption occurred,
as this would predominantly occur only under fully reductive conditions (but
see Goldberg and Gebauer, 2009, for an exception).
Oxidation of organic N was found to be the main source of N2O. The
production of N2O from an unlabelled organic source would most likely
follow a combined process of organic N oxidation via heterotrophic
nitrifiers to nitrite, followed by a reduction of nitrite to gaseous N
products (Butterbach-Bahl et al., 2013). This process, where oxidation
and reduction processes occur hand in hand, would be conceptually similar to
the nitrifier-denitrification process (Wrage et al., 2001). Most research,
however, does not take the oxidation of organic N into account as a possible
source of N2O (J. Zhang et al., 2015) even though recent studies
showed that this process contributed 54–85 % of N2O emissions in
pastures (Rütting et al., 2010; Müller et al., 2014). These
contributions are in line with the current study. Müller et al. (2014)
also showed that the fraction of N2O contributed via the
oxidation of organic N was lowest immediately following NH4NO3
addition and that this fraction increased to over 80 %, while the
contribution of denitrification decreased with time even though
NO3- concentrations increased. Because of the large contribution
of oxidation of organic N in N2O emissions, this pathway should not be
omitted in future research.
A decrease in N2O produced via denitrification was found in soil
previously subjected to higher-temperature treatments. This could be due to
a decrease in the rate of denitrification. However, though complete
denitrification was likely not a dominant process in these aerobic soils, it
is also possible that under treatment T2 and T3 more of the
NO3- underwent complete denitrification, forming N2 as
opposed to N2O. This highlights the importance of the gaseous N
stoichiometries, in particular the N2 / N2O ratio. Stevens and
Laughlin (2001) reported N2 / N2O ratios in a fine loamy grassland
soil of 2.2 and 0.5 from control and combined slurry plus NO3-
fertilizer treatments, respectively. However, Clough et al. (1998)
showed that ratios can vary between 6.2 and 33.2 following 15N-labelled
urine application to ryegrass (Lolium perenne)–white clover (Trifolium repens) pasture on four different
soils (silt loam, sandy loam, peat and clay soils). Unfortunately, due to
methodological restrictions we were not able to detect significant N2
fluxes, as they were < 4 g N2-N ha-1 day-1 (Stevens and Laughlin, 1998).
The adaptation of microorganisms to long-term elevated-temperature treatments might also provide an explanation for the decrease in N2O emissions
during the incubation with soil previously subjected to increasing soil
warming temperatures (Avrahami and Conrad, 2003; French et al., 2009;
Pritchard, 2011). Enhanced NO3- concentrations in the T2 and
T3 treatments, at the end of the field experiment, also suggest an in
situ reduction of denitrification and/or co-denitrification. A possible
explanation for the in situ reduction of denitrification could be the altered
field soil moisture content. While during the incubation, soil moisture was
purposely kept constant (WFPS of 64 %); in the field, however, moisture
conditions were affected by the heating treatment, leading to generally
drier, and thus more aerated, conditions in the heated plots
(Jansen-Willems et al., 2016). Under low WFPS, nitrification is
predominantly responsible for N2O efflux (Bollmann and Conrad, 1998;
Bateman and Baggs, 2005). This may be a consequence of altered soil moisture
or changes in soil texture and physical soil structure. The reduction of
NO3- (denitrification) takes place under more anoxic to anaerobic
conditions (Smith, 1997) because under aerobic conditions, denitrifiers
reduce O2 rather that NO3- (Arah, 1997). Any reduction in
soil moisture could therefore lead to a decrease in the in situ denitrification rate.
Co-denitrification was observed to be significant in Tcontrol and
T1 shortly after N addition. Rates were comparable with those from true
denitrification. Co-denitrification is a co-metabolic process which uses
inorganic and organic N compounds concurrently and converts them to the same
end products as in denitrification. Gases produced via this process are a
hybrid N–N species where one atom of N comes from NO2- and the
other one from a co-metabolized compound (Spott et al., 2011). The
conditions for increased co-denitrification are still not fully understood,
but the presence of fungi along with adequate amino acid pools appears to
enhance losses via this pathway (Laughlin and Stevens, 2002; Spott et al., 2011).
Laughlin and Stevens (2002) found that fungi dominated denitrification
and co-denitrification in grassland soils. It has been suggested that
warming could increase the relative contribution of fungi to the soil
microbial community (Zhang et al., 2005; Pritchard, 2011). Most fungi
lack N2O reductase, resulting in N2O as the final denitrification
product (Saggar et al., 2013). It can therefore be expected that warming
would lead to an increase in N2O produced via denitrification and
co-denitrification. However, the opposite was found in the current
experiment although the changes in co-denitrification were not significant.
The reduced co-denitrification and total denitrification rates seem to
indicate a reduction in fungal-mediated N processes under elevated
temperatures in these soils. Further research is required to elucidate the
effect of increased temperatures on N processes mediated by fungi.
Conclusion
Sustained increases in soil temperatures over 6 years (between 2 and
3 ∘C) led to an increase in both total inorganic soil N and
NO3- pools. Subsequent analyses of gross N transformations,
during an incubation of these soils under common temperature and moisture
conditions to study the legacy effect of increased temperatures, revealed
that mineralization of amino acids (glycine) and recalcitrant organic N
decreased with previously imposed elevated temperatures. This decrease in
mineralization was also correlated with a decrease in N2O emissions
from organic N turnover. However, elevated temperature did not cause a
significant change in relative N2O emissions from the different
pathways as hypothesized, but it led to an absolute decrease in N2O
emission rates. A new, easy to use, source-partitioning method was developed
to determine the contribution of four different pathways to N2O
emissions. Emissions of N2O in the first 6 days after fertilization
were decreased for soils previously subjected to higher temperatures as a
consequence of a reduction in the rates of denitrification and the oxidation
of organic N. For all treatments, oxidation of organic N was the main
contributor to N2O emissions and should therefore in future research
not be omitted as a possible source of N2O.
Data availability
The data will be made available via the following database:
http://www.face2face.center.
Acknowledgements
This study was funded by the LOEWE excellence programme FACE2FACE,
AGRI-I (RSF 10/SC/716) and the Walsh fellowship programme. The study was also
associated with the German Science foundation research unit DASIM (DFG 2337).
The funding was used in experimental design, data collection and analyses, and writing the report. The views expressed in this paper are those of the
authors and do not necessarily represent the views of collaborators, authors'
institutions or the funding agencies. The authors want to gratefully
acknowledge the assistance of Christian Eckhardt, Andre Gorenflo, Cecile Guillet,
Lisa Heimann, Bram Jansen, Birte Lenz, Gerhard Mayer, Gerald Moser,
Manjula Premaratne, David Rex, Sonja Schimmelpfennig, Jochen Senkbeil,
Nicol Strasilla and Till Strohbusch.
Edited by: S. Billings
Reviewed by: two anonymous referees
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