Combined effects of carbon, nitrogen and phosphorus on CH4 production and denitrification in
wetland sediments
Sang Yoon Kim1, Annelies J. Veraart1, Marion Meima-Franke1 and Paul L.E. Bodelier1*
1Netherlands Institute of Ecology (NIOO-KNAW), department of microbial ecology, Wageningen, the
Netherlands
*Corresponding author: Paul L.E. Bodelier
Phone: +31 (0)317473485
Fax: +31 (0)317473675
E-mail address: [email protected]
Paper type: Regular article
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1. Introduction
Methane (CH4) is the second most potent greenhouse gas in the atmosphere after carbon
dioxide (CO2) and has 34 times the global-warming potential of CO2 over a 100-year horizon (IPCC,
2013). Global atmospheric CH4 concentration has increased from pre-industrial level of 0.715 ppm to
1.824 ppm in 2013 (WMO, 2014). Wetlands, including rice paddies, are the largest natural source of
CH4 to the atmosphere, accounting for approximately 139 – 343 Tg CH4 yr-1. These ecosystems
contribute 32 – 47% to the total global CH4 emissions (Denman, 2007). CH4 is produced by a complex
microbial group which degrades organic matter by anaerobic methanogenesis (Conrad et al., 2007).
Methanogenesis is mediated by acetoclastic, hydrogenotrophic, and methylotrophic methanogens that
belong to the Euryarchaeota (Liu and Whitman, 2008).
Methanogenesis can be regulated by various factors including temperature (Conrad, 2002;
Glissman et al., 2004; Inglett and Inglett, 2013), pH (Wang et al., 1993; Ye et al., 2012), substrate
availability (O'Connor et al., 2010), and availability of electron acceptors (D'Angelo and Reddy, 1999).
Although a number of studies have been carried out to identify the main factors which control CH4
dynamics from wetlands, the effect of nutrients on CH4 dynamics is poorly understood. Many studies
point at nitrogen (N) as an important variable influencing CH4 cycles in wetland ecosystems (Bodelier,
2011). Effects of N addition can act directly on the methanogenic community, but may have indirect
effects, by stimulating the bacteria capable of denitrification. Denitrifiers and methanogens compete for
organic carbon (C), and furthermore denitrification produces toxic intermediates (NO, NO2, and N2O)
which can inhibit methanogenic archaea in wetland sediments, thereby reducing CH4 production (Roy
and Conrad, 1999). Although denitrification mitigates eutrophication effects in wetlands by reducing
availability of reactive nitrogen, incomplete denitrification also contributes to the emission of N 2O,
which is an even more potent greenhouse gas than CH4, with 300 times the global warming potential
of CO2 (IPCC, 2013).
Besides N, also phosphorus (P) can be an important regulating factor of methanogenesis. For
example, in Dutch drainage ditches, the water column PO43- concentration was found to be an
important predictor of CH4 emissions (Schrier-Uijl et al., 2011). Not only is P one of the most important
nutrients influencing the microbial activity, including decomposition processes (Cleveland et al., 2002),
but higher P concentrations also elevate microbial growth rates (Makino et al., 2003; Sterner and
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Elser, 2002). Recently, Medvedeff et al. (2014) suggested that P addition stimulated methanogenic
activity in P-limited calcareous subtropical wetland soil, increasing the activity of methanogens directly
or indirectly via fermentative bacteria that produce methanogenic substrates.
Anthropogenic impacts on natural ecosystems have been increasing and largely influencing
the balance of essential nutrients such as C, N and P for decades (Howarth and Marino, 2006; Wang
et al., 2014). These changes in nutrient availability can impact the growth and activity of methanogens
and denitrifiers, and may significantly influence CH4 cycling. However, little is known about the
combined effects of C, N and P on CH4 cycling, and their impact on the interaction between
methanogenesis and denitrification remains unclear in wetland ecosystems.
The aim of this study was to investigate effects and possible interactions of N and P additions
on CH4 production and denitrification in wetland sediments. More specifically we tested the following
hypotheses: (i) sole P addition stimulates CH4 production due to elevated microbial growth rates
including methanogens in wetland sediments, (ii) combined N and P addition enhance denitrification
rates which lead to increased substrate competition between methanogens and denitrifiers, repressing
CH4 production from wetland sediments. To this end incubation studies were performed with sediment
derived from agricultural ditches in the Netherlands following methane production, denitrification,
functional gene abundance and sediment physico-chemistry. Because effects of N and P are expected
to act through competition for carbon, experiments were carried out with and without added C.
2. Materials and methods
2.1. Preparation and incubation of sediment slurries
The sediment samples were collected from the top layer (0-5 cm) of a drainage ditch sediment
in the spring of 2014 (Nigtevecht, The Netherlands; coordinates: 52° 16’ 41.92” N, 05° 01’ 40.11” E).
The drainage ditches are common water management practices in the Netherlands. These ditches
serve as important waterways and efficiently provide water to agricultural fields. The sediment pH was
neutral (6.92) and had comparatively high organic C content (63 g kg-1) as well as CH4 production
potential (106.9 nmol g-1 d.w hr-1) with low nutrient contents in pore water (Table 1). The sediment and
pore water characteristics are presented in Table 1. Prior to use, sediment samples were passed
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through a stainless steel sieve (2 mm) by a wet-sieving method, and then stored at 4 °C until further
processing.
Anaerobic slurries were prepared (10 g fresh sediment and 13 ml amendment solution) in 150
ml serum bottles. The bottles were vigorously shaken by hand to homogenize the sediment slurries,
capped with sterilized butyl stoppers and then flushed with N2 for 1 hr. All bottles were incubated at
25 °C in the dark for 12 days on a shaker (120 rpm). In order to verify potential interacting effects of N,
P and C on CH4 production, we used a factorial design consisting of two C treatments (final conc. 0 or
10 mM C as CH3COOH), three P treatments (final conc. 0, 1, and 10 mM P as KH2PO4) and 3 N
treatments (final conc. 0, 2.5, and 5.0 mM N as KNO3), leading to a total of 18 combinations of C, N
and P. Each experiment was carried out in triplicate.The final concentrations of C, N and P, are
presented in Figure S1. N application level (0 - 5.0 mM, mean 2.5 mM) was chosen to resemble
agriculture fields that receive high N loads (< 10 mM as 200 kg N ha -1), as suggested by Roy and
Conrad (1999). In addition, P application level was determined by N to P ratios (0.25 - 5.0) based on
possible N and P input from fertilization by chemical fertilizer and manure applications which globally
ranged from 4.3 to 5.7 (Mean ca. 5) by estimating N and P balance for a century from 1950 to 2050
(Bouwman et al., 2013). C addition (10 mM) was chosen to achieve a ratio of 4 C: 1 N, which provides
sufficient C to the denitrifiers (Payne, 1981).
Note that although added nutrients were in the high range of drainage ditch water column
concentrations (Veraart, 2012), concentrations in the pore water, which are more relevant for this
study, frequently reach the mM range (Veraart et al., 2015), with our highest added concentrations
reflecting worst-case scenarios. Furthermore, nutrient concentrations are expected to increase with
fertilizer application and heavy agricultural runoff in wetland ecosystem after extreme weather events,
such as heavy rainfall, which due to global change is expected to occur more frequently in Western
Europe.
A control set of bottles (n = 3) was prepared to monitor the changes in headspace CH4
concentration during incubation and an additional control series (n = 3) was used for chemical analysis
of the pore water.
2.2. Measurements of CH4 production and chemical parameters
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Methane concentrations were measured every day. Before sampling the headspace and pore
water, bottles were shaken vigorously to homogenize the sediment slurries. Gas samples (200 µl)
were taken using a gas-tight pressure-lock syringe flushed with N2. The CH4 in the headspace was
measured using an Ultra GC gas chromatograph (Interscience, the Netherlands) equipped with Rt-Q-
Bond (30 m, 0.32 mm, ID) capillary column and a flame ionization detector (FID). The temperature of
the column, injector and detector were adjusted to 80, 150, and 250 °C, respectively. Helium and H2
were used as the carrier and burning gases, respectively. Pore water samples (1 ml) were taken with
sterile disposable syringes equipped with long needles flushed with N2 to prevent O2 leakage that
might influence N mineralization processes such as ammonification and nitrification in the system. The
pore water was transferred to Eppendorf tubes (2 ml) and centrifuged for 15 min at 15,000 × g at 4 °C.
The supernatant was collected and stored at −20 °C until further analysis. Nutrient contents (NH4+,
NO3- and NO2
-) in the sediment were determined using an auto analyzer (QuAAtro, Seal analytical Inc.,
Beun de Ronde, Abcoude, The Netherlands). The pH in the slurries was measured directly after
addition of the amendment solutions and after incubation.
2.3. Denitrification potential
After the methanogenic incubations, denitrification potential measurements were conducted
using the acetylene inhibition method (Philippot et al., 2013; Qin et al., 2012). Briefly, 5g of fresh slurry
from the bottles after incubation was transferred to new bottles (150 ml) and 7.5 ml of a solution
containing KNO3 (1 mM) and glucose (1 mM) was added. The bottles were sealed and capped with
sterilized butyl stoppers and then flushed with N2 for 10 min. Acetylene was added (10% v/v of the
headspace) and the bottles were incubated at 25 °C on a shaker (120 rpm). Nitrous oxide (N2O) in the
headspace was measured after 24 hrs. of incubation using a TRACE 1300 (Thermo Scientific, the
Netherlands) equipped with HS-Q (1m, 2.0 mm, ID) packed column fitted with an electron capture
detector (ECD). The temperature of the column and injector was 90 °C and the detector was set at
350 °C. N2 gas was used as the carrier.
2.4. Quantification of mcrA and nirS gene copy numbers in sediment slurries
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After incubation, DNA was extracted from 0.10 g of freeze-dried sediment following a slightly
modified protocol based on the FastDNA spin kit for soil (MP Biomedicals, Solon, OH) previously
described in detail in Wang et al. (2012). Nucleic acids were routinely quantified using a NanoDrop
1000 spectrophotometer (Thermo) before quantitative PCR (qPCR). The copy numbers of the mcrA
gene, encoding the methyl coenzyme-M reductase were used as proxy for methanogenic abundance
and the copy numbers of the nirS gene, encoding the cytochrome cd1 nitrite reductase were used as
proxy for denitrifier abundance. Primer sets of mlas/mcrA-rev were used for mcrA (Steinberg and
Regan, 2008) and nirScd3af/nirSR3cd for nirS (Throbäck et al., 2004). Real-time qPCR was
performed in a Rotor-Gene Q real-time PCR cycler (Qiagen, the Netherlands). Briefly, qPCR reaction
(total volume 20 µl) for mcrA gene contained 10 µl of reaction mixtures 2X SensiFAST SYBR
(BIOLINE, the Netherlands), 3.5 µl of forward primers (4 pmol µl-1), 3.5 µl reverse primers (5 pmol µl-1),
1 µl Bovine Serum Albumin (5 mg ml-1; Invitrogen, the Netherlands), and 2 µl diluted template DNA (2
ng µl-1). Amplification was carried out as follows: for the mcrA gene, initial denaturation at 95 °C for 3
min, followed by 45 cycles of denaturation at 95 °C for 10 sec, annealing at 60 C for 10 sec, and
extension at 72 °C for 25 sec. qPCR reaction (total volume 20 µl) for nirS gene contained 10 µl of
reaction mixtures 2X SensiFAST SYBR (BIOLINE, the Netherlands), 2 µl of forward primers (5 pmol µl -
1), 2 µl reverse primers (5 pmol µl-1), 1 µl Bovine Serum Albumin (5 mg ml-1; Invitrogen, the
Netherlands), and 2.5 µl diluted template DNA (2 ng µl -1). Amplification was carried out as follows: for
the nirS gene, initial denaturation at 95 °C for 3 min, followed by 40 cycles of denaturation at 95 °C for
10 sec, annealing at 60 °C for 10 sec, and extension at 72 °C for 20 sec and 86 °C for 5 sec. In each
qPCR, the fluorescence signal was obtained at 72 °C after each cycle, and melt curves were obtained
from 70 °C to 99 °C (1 °C temperature by rising). Amplicon specificity was determined from the melt
curve. To avoid the inhibitory effects of substances co-extracted with the DNA, amplification of serial
dilutions was carried out for the slurry samples in each treatment.
2.5. Statistical analysis
Statistical analyses were conducted using R studio software (ver.2.6.0). Determination of
differences between parameters was performed through three-way analysis of variance (ANOVA)
including N, P and C additions and their interactions.
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3. Results
3.1. Effects of C, N and P on CH4 production and potential denitrification
3.1.1 CH4 production and its lag phase
C, N, P and their interaction all significantly affected CH4 production and its lag phase (Figures
1 and 2, Table 2, three-way ANOVA, see Table 3). Overall, we observed most CH4 in the headspace
of samples where no N was added, of which samples with added C had the highest final CH4
concentrations (113.2 µM CH4). When C was added, effects of N and P clearly interacted: the lowest
CH4 production (1.14 µM CH4) was found in the samples where the highest P (10 mM) and N (5 mM)
were added. Combined C and P addition at 10 mM increased the lag phase of CH4 production,
especially in combination with N addition (Figure 1 and Table 2).
3.1.2. Denitrification potential
Denitrification potential was on average 28.2 nmol N g -1 d.w-1, and showed interacting effects
of C, N and P (Figure 2, three-way ANOVA, see Table 3). When no P was added, addition of N and C
moderately stimulated denitrification. With 1 mM P added, C addition moderately enhanced
denitrification, but for the treatment without C the effect of N is not clear. Interestingly, a very different
pattern can be seen for the highest P addition (10 mM), where there seems to be an additive inhibitory
effect of N and C addition: the lowest denitrification potential (8.28 nmol N g-1 d.w-1) was found when P,
N and C were added.
3.2. Changes in biogeochemical properties
3.2.1. Changes of NO3-, NO2
-, and NH4+ in pore water
NO3- was consumed within 48 hr in sediment slurries without P addition, but decreased slower
in slurries where P was added, irrespective of C addition. In sediment slurries not amended with N and
C, no NO2- was detected during the incubation period. NO2
- was only observed in the sediment to
which both C and N were added, and was higher in the slurries with added P (10 mM). For example,
there was a NO2- accumulation, up to 0.27 mM in the slurries where C, N and P was added (C, N, and
P: 10, 5.0, and 10 mM). The NO2- peak occurs when NO3
- levels are still decreasing, and vanishes with
disappearing NO3-. Interestingly, although without P addition, only small fluctuations in NH4
+ occurred
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in all treatments, when C, N and P were added in combination, an increase in NH 4+
above the baseline
variation can be observed (especially at N 2.5 mM), which later decreases again. Overall, NH4+
concentrations were maintained at a higher level in P-added slurries (0.036 - 0.103 mM on average)
than in slurries where no P was added (0.020 - 0.034 mM on average) (Figure 3).
3.2.2. Changes of pH
C addition decreased the pH of the sediment slurries, but during incubation pH in these
slurries neutralized again (Figure S2). P addition also acidified the slurries, with lower pH in P10
slurries (ca. 6.30) than in P0 and P1 slurries (ca. 6.73), and these effects remained after incubation.
3.2.3. mcrA and nirS gene copy numbers
The abundances of the mcrA gene and nirS gene were determined by qPCR after slurry
incubation (Figure 4). The mcrA gene ranged from 6.78 × 107 to 1.10 × 108 and nirS gene copy
numbers ranged from 5.21 × 106 to 6.62 × 107 copies g-1 dry sediment. Unexpectedly, the copy
numbers of mcrA genes were higher in samples without C addition but increased with P addition
except N0 treatment. The effect of N addition on mcrA gene copy was not observed under C addition.
In particular, N addition gradually increased nirS gene copy numbers in slurries with added P or C, but
copy numbers decreased when P and C were both added. We found a weak correlation but significant
negative exponential relations between CH4 production and mcrA gene copy numbers (R2=0.164,
P=0.051). A positive exponential correlation between nirS gene copy numbers and denitrification
potential (R2=0.288, P<0.01) was also observed.
3.3. Relationship between CH4 production and denitrification potential
P additions (P1 and P10) without added N showed similar patterns as the control without
added P, and did not influence CH4 production and denitrification potential, irrespective of C addition
(Figure 5). N additions without added P decreased CH4 production but increased denitrification
potential in sediments with and without added C. Combined N and P addition showed additive effects
on CH4 production and denitrification potential. However, the processes had a different response to C
addition: when C was added, CH4 production and denitrification were more strongly inhibited at high
N:P ratios (2.5 - 5.0) than at low N:P ratios (0.25 - 0.5).
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4. Discussion
We hypothesized that sole and combined effects of N and P on methanogenesis and
denitrification in wetlands sediments act through competition for C. Experiments were carried out with
and without added C to measure CH4 production potential and potential denitrification as well as
biogeochemical properties during slurry incubations. The present study shows combined effects of N
and P on CH4 and N cycles in wetland sediments, and indicates that these effects may indeed occur
through competition for C - albeit in a more complex interaction than initially hypothesized - which will
be discussed in the following sections.
4.1. Effects of C, N and P on CH4 production and denitrification
Our study showed that CH4 production was significantly affected by C addition in sediment
slurries. It is well-known that acetate can be directly utilized by acetogenic methanogens to produce
CH4 under anaerobic conditions (Conrad, 2007). This stimulation was not reflected in the mcrA gene
abundance, for which we have no definite explanation other than referring to other studies which also
found no relation between gene abundance and methane production (Cadillo‐Quiroz et al., 2006;
Galand et al., 2003). Possibly, acetate and phosphate have formed acetylphosphate, which may have
affected enzyme functioning in the metabolic pathways and qPCR process.
N additions inhibited CH4 production (Figure 2). Its effect increased with increasing N
concentration. The inhibitory effect of N compounds on the methanogenic microbial community can be
caused by denitrification intermediates (NO, NO2-) that are toxic for methanogenic archaea, or by
competition between denitrifiers and methanogens for acetate (Kluber and Conrad, 1998; Roy and
Conrad, 1999). NO2- was only produced in excess when C and N were added, and this effect was
much stronger under high P (Figure 3). This result indicates that the increased C/N ratio influenced the
first denitrification steps and that the N:P ratio can also affect NO 2- accumulation. The accumulation of
NO2- stops when NO3
- levels decrease, which is also expected from theoretical denitrification
energetics (van de Leemput et al., 2011).
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Roy and Conrad (1999) speculated that competition for substrate between denitrifiers and
methanogens is not the main mechanism of suppression of methanogenesis in rice soil. However, we
observed inhibitory effects of N addition on CH4 production and lag phase in slurries where no C was
added (Figure 2). In these slurries, no NO2- accumulation was observed (Figure 3), which indicates
that the inhibitory effect is mainly due to the competition for C between methanogens and other
microorganisms.
Our study showed that CH4 production was not directly influenced by P addition in sediment
slurries, irrespective of C addition. However, P addition slowed down NO3- uptake by denitrifiers, which
can increase C availability for methanogens by decreasing competition. Therefore, we expected CH4
production to be increased during the delayed NO3- uptake. Different to our expectation, CH4
production was significantly lower when N and P were added, irrespective of C addition (Figure 2). It
appears that low N:P ratios decreased denitrification. A negative correlation between P and
denitrification was also observed in the sediment of C-poor shallow lakes (Veraart, 2012). On the other
hand, seasonal positive correlations between total P and denitrification were observed in a meta-
analysis of denitrification across aquatic ecosystems (Piña-Ochoa and Álvarez-Cobelas, 2006).
However, P-release from the sediment and denitrification occur under similar anoxic conditions,
confounding ecosystem observations, and making negative correlations between P and denitrification
even more interesting. The potential inhibitory effect of P on denitrification may be direct or indirect.
Potential direct effects are not well understood, but may be through P-sensitivity of the denitrifiers due
to adaptation to the low prevailing P conditions in this sediment. Indirect effects through enhanced
competition for C and N with other bacterial populations are also possible. We used copy numbers of
nirS, reflecting the denitrifier community containing the cytochrome cd1 nitrite reductase, as a proxy for
denitrifier abundance. This may give an incomplete picture, because part of the denitrifiers will use
nirK rather than nirS, and mere gene presence does not directly reflect functional activity. Nonetheless
abundance of nirS positively correlated with potential denitrification (Figure S3). Copy numbers of nirS
were also highest when both P and N were present, which may hint at an N and P co-limitation of the
denitrifying community. However, much like mcrA copy numbers, this effect of N and P on nirS copy
numbers was only observed when no C was added.
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4.2. Competition between denitrification, DNRA and methanogenesis
C addition may have shifted the competitive balance of different nitrate respiring and C
degrading communities. Possibly, communities carrying out DNRA (dissimilatory nitrate reduction to
ammonium) may have been better competitors for acetate under N and P addition, suppressing the
abundance of nirS-denitrifiers as well as methanogenic activity. DNRA-communities compete with
denitrifiers for C and N, as these pathways occur under similar conditions (Francis et al., 2007; Scott
et al., 2008; Tiedje, 1988; Yin et al., 2002). The C:N ratio is a key determinant of the competitive
outcome between both processes, favoring DNRA at high C availability (Smith, 1982), in line with our
findings of lower denitrification under combined N, P and C addition. Our ‘DNRA-competition’-
hypothesis is further supported by the observed increase in NH4+ concentrations in slurries with
combined N, P and application (Figure 2). However, we estimate that about 10% of the NO3- was
converted to NH4+ - rather than gaseous nitrogen - in these slurries, leaving part of the difference in
denitrification potential unexplained. A closed mass balance or isotope tracer approach would be
optimal to accurately trace N-conversions.
Interestingly, combined C and P addition seems to enhance initial ammonification and DNRA,
suggesting that these steps of the N cycle are P limited in this sediment. Clearly, further studies are
needed to test P-limitation of ammonification, DNRA, and denitrification and consequently its
implications for CH4 cycling. Isotope tracing methods based on single cells (Krause et al., 2014) such
as stable isotope tracers and nano-SIMS, in combination with bacterial community analyses to identify
shifts between dominant populations and N-respiring pathways (Kraft et al., 2014) will be promising
ways to study this.
5. Conclusions
We have summarized the main findings of this study in a conceptual scheme depicted in
Figure 6. The effects of N, P and C additions and the interaction between methanogenesis and
denitrification turned out to be more complicated than hypothesized. Instead of simply stimulating
growth of methanogens or denitrifiers by N or P additions we observed an interactive effect of C, N
and P which may be explained by a modulation of electron flow towards DNRA, consuming electron
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donors in the process. Despite the fact that this hypothesis still has to be verified experimentally it is
safe to conclude that P might play an important modulating role in carbon degradation and C-N cycle
interactions in wetland sediments with possible consequences for greenhouse gas emissions from
these ecosystems.
Acknowledgements
This work was supported a grant (823.001.008) of the Netherlands Organisation for Scientific
Research (NWO). This work was supported by the National Research Foundation of Korea (NRF)
grant funded by the Korea government (MSIP) (No. NRF-2013R1A2A2A07068946). We thank Anne
Steenbergh for help with the statistical analysis. This publication is publication no. xxxx of the
Netherlands Institute of Ecology.
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Table 1. Characteristics of the sediment and pore water before the experiment.
Parameters Value
Sediment
pH
Total C (g kg-1)
Total N (g kg-1)
C:N ratio
Potential CH4 production (nmol g-1 d.w hr-1)
Pore water
Electrical conductivity (µS cm-1)
Dissolved organic C (mg L-1)
Dissolved inorganic C (mg L-1)
NH4+ (mg L-1)
PO43- (mg L-1)
N:P ratio
6.92
63.0
5.42
11.6
106.9
853
14.0
3.8
0.43
2.0
0.51
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Table 2. Variables of methanogenic potentials in sediment slurries of different nutrient addition
treatments.
Treatment Variables
Lag phase (hr)* CH4 production rate (umol g-1 hr-1) R2***
Without
C addition
P0 N0 39.5 ± 6.80 0.040 ± 0.003 0.996
N2.5 48.2 ± 6.40 0.006 ± 0.001 0.988
N5.0 52.2 ± 31.8 0.006 ± 0.002 0.978
P1 N0 23.4 ± 10.1 0.045 ± 0.001 0.999
N2.5 74.5 ± 32.0 0.005 ± 0.001 0.978
N5.0 81.6 ± 26.6 0.005 ± 0.002 0.959
P10 N0 12.7 ± 0.90 0.038 ± 0.002 0.998
N2.5 - BD** -
N5.0 - BD -
With C addition P0 N0 23.3 ± 1.20 0.344 ± 0.004 0.994
N2.5 38.0 ± 1.00 0.102 ± 0.003 0.907
N5.0 51.7 ± 0.60 0.087 ± 0.004 0.980
P1 N0 26.0 ± 1.30 0.327 ± 0.007 0.992
N2.5 57.3 ± 0.20 0.108 ± 0.002 0.946
N5.0 74.2 ± 0.70 0.091 ± 0.004 0.969
P10 N0 34.8 ± 2.50 0.126 ± 0.041 0.942
N2.5 109.0 ± 3.900 0.025 ± 0.010 0.951
N5.0 - BD -
* Initiation of linear phase of CH4 production in the data using four time points (extrapolated from linear
regressions).
** BD means below detection limits (CH4 production rate < 0.001).
*** R2 values were estimated from CH4 production in the data.
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Table 3. Three-way ANOVA results. All variables except pH were log-transformed to improve normality.
variable Total CH4
production
CH4 lag-phase Potential
denitrification rate
CH4 production
rate
mcrA nirS pH
(before incubation)
pH
(after incubation)
F P F P F P F P F P F P F P F P
C x N x P 19.91 <0.001 6.392 0.016108.3
3<0.001 0.646 0.427 6.341 0.018 25.164 <0.001 0.119 0.7317 6.69 0.013
N x P 43.25 <0.001 11.638 0.002 85.60 <0.001 8.736 0.006 0.407 0.529 5.936 0.022 6.993 0.011 0.237 0.6289
C x P 15.60 <0.001 42.418 <0.001327.3
1<0.001 20.37 <0.001 0.501 0.485 52.418 <0.001 2.860 0.098 0.024 0.877
C x N 1.08 0.305 0.173 0.680 39.93 <0.001 13.88 <0.001 0.017 0.898 3.769 0.064 0.038 0.564 39.56 <0.001C 258.12 <0.001 10.354 0.003 31.35 <0.001 294.8 <0.001 4.046 0.054 28.208 <0.001 693.7 <0.001 5.392 0.0247
N 250.82 <0.001 63.27 0.001 0.01 0.909 117.7 <0.001 1.540 0.225 10.033 0.00438.49
5<0.001 93.71 <0.001
P 63.22 <0.001 1.715 0.199170.6
1<0.001 29.47 <0.001 0.105 0.748 20.247 <0.001 838.7 <0.001 2404.18 <0.001
Note) F value degrees of freedom = F1; P< 0.05, P< 0.01, and P< 0.001 denotes significance at the 5, 1, and 0.1 % levels, respectively.
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Figure legends
Figure 1. Changes in CH4 concentrations in different nutrient-amended slurries during incubation with
C or without C added. Bars represent standard deviations (n=3). Each panel shows a different P
addition level and the lines per panel show different N addition levels.
Figure 2. Total CH4 production and denitrification potential after incubation. Bars represent standard
deviations (n=3).
Figure 3. Changes in concentrations of nitrate (NO3-), nitrite (NO2
-), and ammonium (NH4+) in pore
water in different nutrient-amended slurries during incubation with C or without C addition. Bars
represent standard deviations (n=3).
Figure 4. Abundances of the mcrA and nirS genes in different nutrient-amended sediments after
incubation. Bars represent standard deviations (n=3).
Figure 5. Effect of N:P ratio on CH4 production and denitrification.
Figure 6. Conceptual scheme of observed and hypothetical effects of the applied treatments in this
study.
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Figure 1.
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Figure 2.
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Figure 3.
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Figure 4.
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Figure 5.
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Figure 6.
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Supporting Information Legends
Figure S1. Experimental scheme for nutrient and substrate additions.
Figure S2. Initial and final pH of incubation in different nutrient-amended sediments. Bars represent
standard deviations (n=3).
Figure S3. Relationships between functional gene abundances and their activities.
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Figure S1.
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• N- (0 mM) • N+ (2.5 mM)• N++ (5.0 mM)
• N- (0 mM) • N+ (2.5 mM)• N++ (5.0 mM)
P++(10 mM)
P+ (1 mM)
• N- (0 mM) • N+ (2.5 mM)• N++ (5.0 mM)
P-(0 mM)
• N- (0 mM) • N+ (2.5 mM)• N++ (5.0 mM)
• N- (0 mM) • N+ (2.5 mM)• N++ (5.0 mM)
Wetlandsediment
C +(10 mM)
C -(0 mM )
P++(10 mM)
P+(1 mM)
• N- (0 mM) • N+ (2.5 mM)• N++ (5.0 mM)
P-(0 mM)
364
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367
Figure S2.
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Figure S3.
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REFERENCES
Bodelier, P.L., 2011. Interactions between nitrogenous fertilizers and methane cycling in wetland and
upland soils. Current Opinion in Environmental Sustainability 3(5), 379-388.
Bouwman, L., Goldewijk, K.K., Van Der Hoek, K.W., Beusen, A.H.W., Van Vuuren, D.P., Willems, J.,
Rufino, M.C., Stehfest, E., 2013. Exploring global changes in nitrogen and phosphorus cycles
in agriculture induced by livestock production over the 1900–2050 period. Proceedings of the
National Academy of Sciences 110(52), 20882-20887.
Cadillo‐Quiroz, H., Bräuer, S., Yashiro, E., Sun, C., Yavitt, J., Zinder, S., 2006. Vertical profiles of
methanogenesis and methanogens in two contrasting acidic peatlands in central New York
State, USA. Environ Microbiol 8(8), 1428-1440.
Cleveland, C.C., Townsend, A.R., Schmidt, S.K., 2002. Phosphorus Limitation of Microbial Processes
in Moist Tropical Forests: Evidence from Short-term Laboratory Incubations and Field Studies.
Ecosystems 5(7), 0680-0691.
Conrad, R., 2002. Control of microbial methane production in wetland rice fields. Nutr Cycl Agroecosys
64(1-2), 59-69.
Conrad, R., 2007. Microbial ecology of methanogens and methanotrophs. Adv Agron 96, 1-63.
Conrad, R., Chan, O., Claus, P., Casper, P., 2007. Characterization of methanogenic Archaea and
stable isotope fractionation during methane production in the profundal sediment of an
oligotrophic lake (Lake Stechlin, Germany). Limnol Oceanogr 52(4), 1393.
D'Angelo, E.M., Reddy, K.R., 1999. Regulators of heterotrophic microbial potentials in wetland soils.
Soil Biology and Biochemistry 31(6), 815-830.
Denman, K., 2007. Denman, K.L., G. Brasseur, A. Chidthaisong, P. Ciais, P.M. Cox, R.E. Dickinson,
D. Hauglustaine, C. Heinze, E. Holland, D. Jacob, U. Lohmann, S Ramachandran, P.L. da
Silva Dias, S.C. Wofsy and X. Zhang, 2007: Couplings Between Changes in the Climate
System and Biogeochemistry. In: Climate Change 2007: The Physical Science Basis.
Contribution of Working Group I to the Fourth Assessment Report of the Intergovernmental
Panel on Climate Change [Solomon, S., D. Qin, M. Manning, Z. Chen, M. Marquis, K.B.
Averyt, M.Tignor and H.L. Miller (eds.)]. Cambridge University Press, Cambridge, United
Kingdom and New York, NY, USA.
27 | P a g e
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
Francis, C.A., Beman, J.M., Kuypers, M.M.M., 2007. New processes and players in the nitrogen cycle:
the microbial ecology of anaerobic and archaeal ammonia oxidation. ISME J 1(1), 19-27.
Galand, P.E., Fritze, H., Yrjälä, K., 2003. Microsite‐dependent changes in methanogenic populations
in a boreal oligotrophic fen. Environ Microbiol 5(11), 1133-1143.
Glissman, K., Chin, K.-J., Casper, P., Conrad, R., 2004. Methanogenic pathway and archaeal
community structure in the sediment of eutrophic Lake Dagow: effect of temperature. Microb
Ecol 48(3), 389-399.
Howarth, R.W., Marino, R., 2006. Nitrogen as the limiting nutrient for eutrophication in coastal marine
ecosystems: evolving views over three decades. Limnol Oceanogr 51(1), 364-376.
Inglett, P.W., Inglett, K.S., 2013. Biogeochemical changes during early development of restored
calcareous wetland soils. Geoderma 192(0), 132-141.
IPCC, 2013. Climate Change 2013: The Physical Science Basis. Contribution of Working Group I to
the Fifth Assessment Report of the Intergovernmental Panel on Climate Change [Stocker,
T.F., D. Qin, G.-K. Plattner, M. Tignor, S.K. Allen, J. Boschung, A. Nauels, Y. Xia, V. Bex and
P.M. Midgley (eds.)]. Cambridge University Press, Cambridge, United Kingdom and New
York, NY, USA, 1535 pp.
Kluber, H.D., Conrad, R., 1998. Inhibitory effects of nitrate, nitrite, NO and N2O on methanogenesis by
Methanosarcina barkeri and Methanobacterium bryantii. Fems Microbiol Ecol 25(4), 331-339.
Kraft, B., Tegetmeyer, H.E., Sharma, R., Klotz, M.G., Ferdelman, T.G., Hettich, R.L., Geelhoed, J.S.,
Strous, M., 2014. The environmental controls that govern the end product of bacterial nitrate
respiration. Science 345(6197), 676-679.
Krause, S., Le Roux, X., Niklaus, P.A., Van Bodegom, P.M., Lennon, J.T., Bertilsson, S., Grossart, H.-
P., Philippot, L., Bodelier, P.L., 2014. Trait-based approaches for understanding microbial
biodiversity and ecosystem functioning. Frontiers in Microbiology 5, 251
210.3389/fmicb.2014.00251
Liu, Y., Whitman, W.B., 2008. Metabolic, Phylogenetic, and Ecological Diversity of the Methanogenic
Archaea. Annals of the New York Academy of Sciences 1125(1), 171-189.
Makino, W., Cotner, J.B., Sterner, R.W., Elser, J.J., 2003. Are bacteria more like plants or animals?
Growth rate and resource dependence of bacterial C : N : P stoichiometry. Funct Ecol 17(1),
121-130.
28 | P a g e
405
406
407
408
409
410
411
412
413
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
433
434
Medvedeff, C.A., Inglett, K.S., Inglett, P.W., 2014. Evaluation of direct and indirect phosphorus
limitation of methanogenic pathways in a calcareous subtropical wetland soil. Soil Biology and
Biochemistry 69(0), 343-345.
O'Connor, F.M., Boucher, O., Gedney, N., Jones, C.D., Folberth, G.A., Coppell, R., Friedlingstein, P.,
Collins, W.J., Chappellaz, J., Ridley, J., Johnson, C.E., 2010. Possible role of wetlands,
permafrost, and methane hydrates in the methane cycle under future climate change: A
review. Reviews of Geophysics 48(4), RG4005.
Payne, W.J., 1981. Denitrification. John Wiley & Sons Inc.
Philippot, L., Spor, A., Henault, C., Bru, D., Bizouard, F., Jones, C.M., Sarr, A., Maron, P.A., 2013.
Loss in microbial diversity affects nitrogen cycling in soil. Isme Journal 7(8), 1609-1619.
Piña-Ochoa, E., Álvarez-Cobelas, M., 2006. Denitrification in Aquatic Environments: A Cross-system
Analysis. Biogeochemistry 81(1), 111-130.
Qin, S.P., Hu, C.S., Oenema, O., 2012. Quantifying the underestimation of soil denitrification potential
as determined by the acetylene inhibition method. Soil Biol Biochem 47, 14-17.
Roy, R., Conrad, R., 1999. Effect of methanogenic precursors (acetate, hydrogen, propionate) on the
suppression of methane production by nitrate in anoxic rice field soil. Fems Microbiol Ecol
28(1), 49-61.
Schrier-Uijl, A., Veraart, A., Leffelaar, P., Berendse, F., Veenendaal, E., 2011. Release of CO2 and
CH4 from lakes and drainage ditches in temperate wetlands. Biogeochemistry 102(1-3), 265-
279.
Scott, J.T., McCarthy, M., Gardner, W., Doyle, R., 2008. Denitrification, dissimilatory nitrate reduction
to ammonium, and nitrogen fixation along a nitrate concentration gradient in a created
freshwater wetland. Biogeochemistry 87(1), 99-111.
Smith, M.S., 1982. Dissimilatory Reduction of NO2− to NH4+ and N2O by a Soil Citrobacter sp.
Applied and environmental Microbiology 43(4), 854-860.
Steinberg, L.M., Regan, J.M., 2008. Phylogenetic comparison of the methanogenic communities from
an acidic, oligotrophic fen and an anaerobic digester treating municipal wastewater sludge.
Applied and environmental microbiology 74(21), 6663-6671.
Sterner, R.W., Elser, J.J., 2002. Ecological stoichiometry: the biology of elements from molecules to
the biosphere. Princeton University Press.
29 | P a g e
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
Throbäck, I.N., Enwall, K., Jarvis, Å., Hallin, S., 2004. Reassessing PCR primers targeting nirS, nirK
and nosZ genes for community surveys of denitrifying bacteria with DGGE. Fems Microbiol
Ecol 49(3), 401-417.
Tiedje, J.M., 1988. Ecology of denitrification and dissimilatory nitrate reduction to ammonium. Biology
of anaerobic microorganisms 717, 179-244.
van de Leemput, I.A., Veraart, A.J., Dakos, V., de Klein, J.J., Strous, M., Scheffer, M., 2011. Predicting
microbial nitrogen pathways from basic principles. Environ Microbiol 13(6), 1477-1487.
Veraart, A.J., 2012. Denitrification in Ditches, Streams and Shallow Lakes. Wageningen University.
Veraart, A.J., Steenbergh, A., Ho, A., Kim, S.Y., Bodelier, L.E.P., 2015. Beyond nitrogen: The
importance of phosphorus for CH4 oxidation in soils and sediments, Geoderma (in revision).
Wang, J., Krause, S.M.B., Muyzer, G., Meima-Franke, M., Laanbroek, H.J., Bodelier, P.l.E., 2012.
Spatial patterns of iron- and methane-oxidizing bacterial communities in an irregularly flooded,
riparian wetland. Frontiers in Microbiology 3, 64. doi: 10.3389/fmicb.2012.00064.
Wang, W., Sardans, J., Zeng, C., Zhong, C., Li, Y., Peñuelas, J., 2014. Responses of soil nutrient
concentrations and stoichiometry to different human land uses in a subtropical tidal wetland.
Geoderma 232–234(0), 459-470.
Wang, Z.P., DeLaune, R.D., Patrick, W.H., Masscheleyn, P.H., 1993. Soil Redox and pH Effects on
Methane Production in a Flooded Rice Soil. Soil Sci. Soc. Am. J. 57(2), 382-385.
WMO, 2014. Greenhouse gas bulletin, Geneva.
Ye, R., Jin, Q., Bohannan, B., Keller, J.K., McAllister, S.A., Bridgham, S.D., 2012. pH controls over
anaerobic carbon mineralization, the efficiency of methane production, and methanogenic
pathways in peatlands across an ombrotrophic–minerotrophic gradient. Soil Biology and
Biochemistry 54(0), 36-47.
Yin, S., Chen, D., Chen, L., Edis, R., 2002. Dissimilatory nitrate reduction to ammonium and
responsible microorganisms in two Chinese and Australian paddy soils. Soil Biology and
Biochemistry 34(8), 1131-1137.
30 | P a g e
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490