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Biology and Fertility of Soils

, Volume 54, Issue 4, pp 481–493 | Cite as

Effects of carbon and phosphorus addition on microbial respiration, N2O emission, and gross nitrogen mineralization in a phosphorus-limited grassland soil

  • Kazi R. Mehnaz
  • Claudia Keitel
  • Feike A. Dijkstra
Original Paper

Abstract

Soil microbes are frequently limited by carbon (C), but also have a high phosphorus (P) requirement. Little is known about the effect of P availability relative to the availability of C on soil microbial activity. In two separate experiments, we assessed the effect of P addition (20 mg P kg−1 soil) with and without glucose addition (500 mg C kg−1 soil) on gross nitrogen (N) mineralization (15N pool dilution method), microbial respiration, and nitrous oxide (N2O) emission in a grassland soil. In the first experiment, soils were incubated for 13 days at 90% water holding capacity (WHC) with addition of NO3 (99 mg N kg−1 soil) to support denitrification. Addition of C and P had no effect on gross N mineralization. Initially, N2O emission significantly increased with glucose, but it decreased at later stages of the incubation, suggesting a shift from C to NO3 limitation of denitrifiers. P addition increased the N2O/CO2 ratio without glucose but decreased it with glucose addition. Furthermore, the 15N recovery was lowest with glucose and without P addition, suggesting a glucose by P interaction on the denitrifying community. In the second experiment, soils were incubated for 2 days at 75% WHC without N addition. Glucose addition increased soil 15N recovery, but had no effect on gross N mineralization. Possibly, glucose addition increased short-term microbial N immobilization, thereby reducing N-substrates for nitrification and denitrification under more aerobic conditions. Our results indicate that both C and P affect N transformations in this grassland soil.

Keywords

15N recovery Denitrification Grassland Gross N mineralization Microbial respiration N2O emission 

Introduction

Nitrogen (N) transformations in soil are mostly mediated by microbes (Robertson and Groffman 2007). While microbial N mineralization supplies plant available N, nitrification and denitrification can result in significant release of nitrous oxide (N2O), which is an important greenhouse gas (Forster et al. 2007) and also responsible for stratospheric ozone depletion (Ravishankara et al. 2009). Indeed, soils are the largest global source of N2O emission representing 56–70% of all N2O sources (Butterbach-Bahl et al. 2013). Therefore, any change in the activity and composition of microbial communities involved in N transformations in soil can affect the ecosystem structure and function, as well as global warming.

In terrestrial ecosystems, the availability and quality of carbon (C) often limit heterotrophic microbial growth and activities, but the use of C substrates can be further limited by availability of nutrients, such as phosphorus (P) (Cleveland et al. 2002; Ilstedt and Singh 2005). Microbial C/P ratios are often smaller than the C/P ratios of the substrates used by microbes, making them susceptible to P limitation (Cleveland and Liptzin 2007). High P adsorption by soil particles and/or precipitation of P as insoluble hydroxy phosphates or various phosphate minerals can exacerbate microbial P limitation in acidic and alkaline soils (Weil and Brady 2016). Microbial P limitation has been observed in highly weathered, old tropical soils depleted in P derived from rocks, as well as in low pH soils containing high iron and aluminum-rich clays that readily fix P into unavailable forms (Cleveland et al. 2002). Calcareous soils with low organic C content also showed P limitation of microbes because of its high P fixing capacity and producing precipitates of insoluble calcium-phosphate minerals (Raiesi and Ghollarata 2006). Available fractions of P were relatively small and tightly cycled thereby limiting productivity in organic C-, N-, and P-rich acidic peatlands (Hill et al. 2014). Often, microbial respiration increases after P addition, suggesting P limitation of microbial activities (Cleveland et al. 2002; Ilstedt and Singh 2005). High P availability could increase microbial respiration by enabling the microbial community to use more recalcitrant forms of soluble C substrates and/or by stimulating a rapid growth and turnover of the microbial community (Cleveland et al. 2002). Wakelin et al. (2017) found increased exogenous C cycling along with an increased size of the microbial community after P addition suggesting a direct effect of P on microbial activity and an indirect effect through supporting plant production and subsequent C inputs to soil for microbial utilization. However, an inhibitory effect of P on microbial respiration has also been observed in low productivity forest soils, attributed to a change in the production or activity of decomposing and/or P-acquiring enzymes and subsequent more efficient utilization of soil C (Kranabetter et al. 2005).

The size and activity of the microbial community also control soil N mineralization (White and Reddy 2000). Microbes release N as a by-product while consuming organic materials as a source of C and energy to support their growth (Robertson and Groffman 2007). However, the quality and quantity of organic matter as well as soil conditions, including temperature and moisture, also determine the rate of N mineralization (Flavel and Murphy 2006; Robertson and Groffman 2007). Organic N mineralization can be increased with acceleration of organic matter decomposition primed by exogenous readily hydrolyzable C sources (Chen et al. 2014), whereas low P availability can restrict soil microbial activity and thus N mineralization in soil (Kranabetter et al. 2005). Enhanced net N mineralization in response to P addition has been observed in tropical forest soils (Chen et al. 2017), pasture soils (Cadisch et al. 1994), volcanic ash soils (Munevar and Wollum 1977), and wetland soils (White and Reddy 2000). However, an increase in inorganic N, accompanied by a decrease in soil respiration and unaltered microbial biomass, after P addition was suggested to be governed by exoenzyme allocation and subsequent depolymerization of organic matter in a P-limited low productivity soil (Kranabetter et al. 2005). Non-significant effects of P amendments on net N mineralization have been observed in other studies (Ross et al. 1995). However, none of these studies have measured gross rates of N mineralization in response to C and P addition, which could help determine the total amount of inorganic N production by soil microbes, and thus provide a better insight into N cycling in response to availability of C and/or P.

Despite the identification of various abiotic and biotic N2O forming processes, it is still considered that N2O is mostly produced by autotrophic ammonia oxidizers and nitrite-oxidizing bacteria responsible for nitrification and heterotrophic denitrifying microorganisms responsible for denitrification (Butterbach-Bahl et al. 2013), with the former process being the main contributor of N2O under aerobic conditions, and the latter being dominant under anaerobic conditions (Liu et al. 2017). Many factors regulate these two microbial processes and hence N2O production in soil, particularly ammonium (NH4+) and nitrate (NO3) concentrations, soil water, temperature, pH, availability of labile organic C, and other nutrients, such as P (Bremner 1997; Dalal et al. 2003; Firestone and Davidson 1989; Signor and Cerri 2013). Soil organic matter as well as manures, plant litters, and root exudates are the major sources of C and energy for heterotrophic denitrifiers (Aulakh et al. 1991; Henry et al. 2008; Murray et al. 2004). Availability of C may support denitrification and hence N2O emission, but high availability of C can also increase the reduction of N2O to dinitrogen (N2), as C availability often limits the final reductive stage of denitrification (Morley and Baggs 2010). However, the N2O/N2 ratio in denitrification may also decrease with reduced soil NO3 concentration and low availability of oxygen (O2), but increase with declining soil pH (Wrage et al. 2001). Availability of P can affect N2O production from nitrification and denitrification by directly influencing the nitrifying and denitrifying microorganisms (Mori et al. 2010, 2013; White and Reddy 1999), and indirectly by affecting organic N mineralization (Falkiner et al. 1993; White and Reddy 2000), plant and microbial N assimilation (Baral et al. 2014; Hall and Matson 1999; Mori et al. 2014), and biological N fixation (Houlton et al. 2008). It has been suggested that alleviation of P limitation of heterotrophic microbes can reduce N resources for N2O production (Hall and Matson 1999; Sundareshwar et al. 2003), while others have suggested that P addition may reduce N2O emission through increasing nitrifying and/or denitrifying respiratory efficiency (Mori et al. 2016). However, several other studies found increased N2O and nitric oxide (NO) emission after P addition without affecting the N2O/NO ratio, due to activation of nitrifying and/or denitrifying bacteria, stimulation of microbial activity in general, and thus other N transformations, such as N mineralization, and/or promotion of anaerobic conditions due to stimulated heterotrophic respiration (Mori et al. 2010, 2013). From the above, it can be expected that high availability of P, in presence of a readily available C source, can stimulate heterotrophic denitrifying activities and hence N2O emission. Nevertheless, a reduction of N2O from nitrification and/or denitrification could also occur due to stimulated N immobilization of heterotrophic microorganisms in general with increasing availability of C and/or P. It is therefore important to understand the interaction of C and P availability on N2O emission in soils, especially in soils limited by nutrients, such as P.

In this study, we examined the effects of P (20 mg P kg−1 soil) and glucose (500 mg C kg−1 soil), a readily available source of C (to mimic root exudates), and their interactions on microbial respiration, N2O emission, and gross N mineralization in a P-limited grassland soil in New South Wales, Australia. Two soil laboratory incubation experiments were conducted involving P and glucose addition treatments and a 15N tracer (1 mg N kg−1 soil). For the first experiment, we incubated soils for 13 days under high soil moisture (90% water holding capacity, WHC) and NO3 (99 mg N kg−1 soil) conditions to support denitrifying activities. A second experiment was conducted for 2 days under lower soil moisture conditions (75% WHC) and without N addition, to examine treatment effects on gross N mineralization and soil 15N recovery under more aerobic conditions. We hypothesized that addition of P to this P-limited soil will increase N2O emission from denitrification by stimulating the denitrifying microorganisms, which will also be accompanied with an increase in microbial respiration and soil organic N mineralization, possibly due to stimulated organic matter decomposition. We further hypothesized that such effects of P will be even greater in the presence of glucose, providing C to the heterotrophic microbial community, including denitrifiers.

Materials and methods

Study area and soil used

Soil samples were collected in October 2016 from a non-fertilized and moderately grazed grassland located at Westwood farm (latitude 33°59′46″S, longitude 150°39′16″E) close to Camden in New South Wales, Australia. The mean annual precipitation of the area is 790 mm, and the mean air temperature of the month July and January is 10.4 and 23 °C, respectively (Dijkstra et al. 2015). The dominating grass was Paspalum dilatatum Poir. (C4). The soil of this grassland was a P-poor sandy loam soil (sand 63%, silt 19%, and clay 18%; pH (H2O) 6.0; total C 5.6%; total N 0.4% and total P 0.14%; available P 1.91 mg kg−1 soil; exchangeable ammonium (NH4+) 3.60 mg N kg−1 soil; NO31.13 mg N kg−1 soil), classified as Red Kurosol. We collected soil samples at 5–20 cm depth (as P was more depleted at this depth than the upper top soil, Mehnaz and Dijkstra 2016) from five different spots within 10 m2 of the farmland. The five collected soil samples were used as five replicates later in the experiment. Soils were crushed and sieved through a 2-mm sieve and mixed well.

Soil incubations and chemical analyses

Of each replicate, 350-g sieved soil was placed in 1-L plastic buckets. Soils were treated with two different levels of P (0 and 20 mg P kg−1 soil) with or without glucose (Glu; 500 mg C kg−1 soil) and mixed well. We added P as a mixture of monopotassium phosphate (KH2PO4) and dipotassium hydrogen phosphate trihydrate (K2HPO4·3H2O) so that the final solution had a pH similar to the soil pH. All the soils were treated with 100 mg N kg−1 soil as potassium nitrate (KNO3; 99 mg N kg−1 soil) and 15N labeled (99 atom %) ammonium sulfate ((NH4)2SO4; 1 mg N kg−1 soil). Each of four treatments (without P and Glu; with P and without Glu; without P and with Glu; with P and Glu) was replicated five times. The soil water content was adjusted to 90% WHC.

N2O and CO2 emission

Subsamples of treated soils (200 g each) were transferred to 1.89-L glass jars and incubated for 13 days for gas emission analysis. The incubation was performed at 20 °C in the dark. The jars were covered with polyethylene plastic wrap to prevent moisture loss but to allow gas exchange. Gas samples were collected at days 0, 1, 2, 5, and 13 of the incubation. On each day of sampling, the jars were flushed with ambient air and kept open for several minutes before starting gas sample collection. Gas samples were collected at 0, 1, and 2 h intervals after closing the jars with lids equipped with sampling ports. The gas samples were analyzed immediately after collection for N2O and CO2 concentrations using an Agilent greenhouse gas chromatograph (7890 GC, Agilent Technologies Australia Pty Ltd., VIC, Australia) with N2 as the carrier gas. N2O was measured on an electron capture detector (ECD) at 300 °C. CO2 was converted to CH4 with a methanizer and then measured using a flame ionization detector (FID) at 250 °C. Columns were set at 60 °C. The standard gas concentrations used were 0.42 and 4.2 ppm for N2O and 393 and 5060 ppm for CO2. The N2O and CO2 fluxes were calculated as the slope of linear regressions from the measured gas concentrations over time. Cumulative gas emissions (C cum ) were calculated as the total sum of gas produced during the period of the 13-day incubation by multiplying the average gas emission rate between two sampling dates (\( \overline{r_i} \)) by the time interval (∆t i ) between the sampling dates and summing the values for all time intervals i (Mehnaz and Dijkstra 2016):
$$ {C}_{cum}={\sum}_{i=0}^n\left(\overline{r_i}\times \Delta {t}_i\right) $$

After the 13-day incubation, soils (15 g) were extracted with 0.05 M K2SO4 (40 mL) by shaking for 1 h, filtering (Whatman #42), and analyzing exchangeable NH4+, NO3, and dissolved organic C (DOC) on a flow injection analyzer (Lachat Instruments, Loveland, CO, USA) and TOC-VCPN analyzer (Shimadzu Scientific Instruments, Sydney, NSW, Australia), respectively. Net ammonification or nitrification rates were calculated as the difference in NH4+ or NO3 concentrations between days 0 and 13, divided by the time interval (Mori et al. 2010). Available P was determined by extracting the soils with 0.03 N NH4F-0.025 N HCl followed by analyzing the extracts colorimetrically using ammonium paramolybdate and stannous chloride (Olsen and Sommers 1982) on a spectrophotometer at 660 nm (UV-VIS spectrophotometer, Shimadzu, Kyoto, Japan). Soil microbial biomass carbon C, N, and P were determined using the fumigation extraction method (Brookes et al. 1982, 1985; Bruulsema and Duxbury 1996). Soil samples were exposed to chloroform for 24 h for microbial P and 48 h for microbial C and N analysis (shorter fumigation time for microbial P to reduce abiotic fixation of microbial P). Fumigated and non-fumigated samples (for microbial C and N) were extracted with 0.05 M K2SO4 and later analyzed for total C and N using a TOC-VCPN analyzer. Microbial biomass C and N were calculated from the difference of their concentrations in fumigated and non-fumigated samples divided by an extraction efficiency of 0.45 (Beck et al. 1997) and 0.54 (Brookes et al. 1985), respectively. For microbial P, fumigated and non-fumigated samples were treated and analyzed using the same procedure followed for available P determination. Microbial biomass P was calculated using an extraction efficiency of 0.40 (Brookes et al. 1982) and corrected for P adsorption of microbial P during fumigation and extraction using the P-addition-recovery relationship measured for this soil (Dijkstra et al. 2015). Soil pH was measured in a 1:5 soil/deionized water suspension using a pH meter.

Soils (both initial and incubated) were dried at 60 °C, ground, and analyzed for total N and 15N using an isotope ratio mass spectrometer (IRMS, Delta V Advantage, Thermo Fisher Scientific, Bremen, Germany). The soil 15N recovery (15 Nrec,soil) was calculated using the following equation:
$$ {}^{15}\ {\mathrm{N}}_{\mathrm{rec},\mathrm{soil}}={\mathrm{N}}_{\mathrm{soil}}\times \left({}^{15}\ {\mathrm{N}}_{\mathrm{post},\mathrm{soil}}-{}^{15}\ {\mathrm{N}}_{\mathrm{pre},\mathrm{soil}}\right)/\left({}^{15}\ {\mathrm{N}}_{\mathrm{label}}-{}^{15}\ {\mathrm{N}}_{\mathrm{pre},\mathrm{soil}}\right) $$
where Nsoil is the total N content of soil; 15 Npost,soil and 15 Npre,soil are the 15 N atom% measured in the soil after and before labeling, respectively; and 15 Nlabel is the 15N atom% of the applied label (Dijkstra et al. 2011).

Gross N mineralization

After transferring 200 g soil (treated with four different treatments) to the glass jar, the remaining soil (150 g soil) was incubated in the same 1-L plastic bucket for 48 h to measure gross N mineralization using the 15N pool dilution technique. The incubation was performed at 20 °C in the dark. One sample was collected at the start of the incubation (time 1 = 0) to determine the initial size and 15N atom% in the NH4+-N pool. The second sample was collected after 48 h of incubation for similar analysis. The soil samples collected (10 g) at two different times were extracted with 1 M KCl (40 mL after shaking for 1 h followed by centrifuging and filtering. The NH4+ concentration of the filtrates was analyzed by the flow injection analyzer. The NH4+ in the remaining filtrates was collected using acidified filter paper disks inside polytetrafluoroethylene (PTFE) diffusion traps (Stark and Hart 1996). The diffusion traps were prepared with PTFE tape by sealing inside small filter paper disks (0.5 cm in diameter) soaked with 5 μL of 2.5 M potassium bisulfate (KHSO4). Filtered samples were diffused in small plastic cups for 6 days at room temperature after adding the traps and magnesium oxide (MgO) to convert NH4+ to NH3. The cups were capped tight and swirled regularly to ensure sufficient diffusion. After diffusing, the traps were removed from the suspension, and the filter disks were dried in a desiccator with concentrated sulfuric acid (H2SO4). Dried disks were packed into small tin capsules and analyzed for 15N on the IRMS. The gross N mineralization (GNM) rates were calculated as follows:
$$ \mathrm{GNM}=\left[\left({\mathrm{M}}_1\hbox{--} {\mathrm{M}}_2\right)/\mathrm{t}\right]\times \left[\log\ \left({}^{15}\mathrm{N}\ \mathrm{atom}\%{}_1/{}^{15}\mathrm{N}\ \mathrm{atom}\%{}_2\right)/\log\ \left({\mathrm{M}}_1/{\mathrm{M}}_2\right)\right] $$
where M1 and M2 are the total amount of NH4+ in the soil at time 1 and time 2, respectively; 15N atom%1 and 15N atom%2 are the measured 15N atom% in the NH4+-N pool at times 1 and 2, respectively; and t is the time difference (in hours) between times 1 and 2 (Kirkham and Bartholomew 1954). We also calculated the gross rates of NH4+-N immobilization (GNI) according to Kirkham and Bartholomew (1954):
$$ \mathrm{GNI}=\left[\left({\mathrm{M}}_1\hbox{--} {\mathrm{M}}_2\right)/\mathrm{t}\right]\times \left[\log\ \left\{\left({}^{15}\mathrm{N}\ \mathrm{atom}\%{}_1\times {\mathrm{M}}_1\right)/\left({}^{15}\mathrm{N}\ \mathrm{atom}\%{}_2\times {\mathrm{M}}_2\right)\right\}/\log\ \left({\mathrm{M}}_1/{\mathrm{M}}_2\right)\right] $$

For these calculations, we assumed that the 15N tracer was homogeneously mixed with the background NH4+, that addition of the 15N tracer did not influence gross N mineralization and NH4+-N immobilization, and that the 48-h incubation was short enough to avoid a reflux of the consumed 15N tracer (Braun et al. 2018). We note that gross NH4+-N immobilization does not represent microbial N immobilization, which would include immobilization of NO3-N.

Gross N mineralization (in more aerobic conditions)

We conducted a separate incubation experiment to measure gross N mineralization in more aerobic conditions. Soils were collected from the same site in October 2015, sieved, and mixed well. Subsamples of 200-g sieved soil were placed in 1-L plastic buckets and incubated for 48 h (at 20 °C in dark) after treating the soils with the same P and C treatments used in the experiment described above: 0 and 20 mg P kg−1 soil with and without glucose (500 mg C kg−1 soil). Unlike the first experiment, soil moisture was adjusted to 75% WHC, and no extra source of N was added to the soils except 15N labeled (99 atom %) ammonium sulfate ((NH4)2SO4; 1 mg N kg−1 soil). Each of the four treatments was performed with five replicates. During the incubation, soil samples were collected and analyzed using the same techniques as described above (15N pool dilution technique) to measure gross N mineralization and NH4+-N immobilization rates. After incubation, soils were analyzed for exchangeable NH4+, NO3, microbial biomass C, and N and soil 15N recovery as described above.

Statistical analysis

A two-way ANOVA was used to test for P addition (with and without P) and glucose (Glu) addition (with and without Glu) and their interactions on N2O and CO2 emission rates, cumulative N2O and CO2 emissions, N2O/CO2 ratio, microbial biomass C, N, and P, available P, exchangeable NH4+, NO3, DOC, soil 15N recovery, net ammonification and nitrification rates, gross N mineralization, and NH4+-N immobilization rates and soil pH. For the first experiment, a repeated measures ANOVA was used to test for main effects of P addition, C addition, and time of measurement (0, 1, 2, 5, and 13 days after incubation) and their interactions on N2O and CO2 emissions. Linear regression was used to test for relationship of cumulative N2O emission with soil 15N recovery, gross N mineralization, net ammonification, and net nitrification rates. When treatment effects were significant at P < 0.05, Tukey’s HSD tests were used to compare the means of each treatment combination. P values between 0.05 and 0.1 were considered marginally significant. When necessary, data were log-transformed to improve normal distribution and to reduce heteroscedasticity (examined with the Brown-Forsythe test for equal variance). All statistical analyses were conducted in JMP (v. 10.0.0; SAS Institute, Cary, NC, USA).

Results

Thirteen-day incubation study

Addition of glucose significantly increased N2O emission at days 0, 1, and 2 of the incubation (P = 0.03, 0.002, 0.0004, respectively) while it marginally significantly (P = 0.07) and significantly (P < 0.0001) decreased N2O emission at days 5 and 13, respectively (Fig. 1a–e). The change in N2O emission rate (both increase and decrease) was faster with glucose addition than without glucose addition (significant time × Glu interactive effect, P < 0.0001, Fig. 2a). Cumulative N2O emission significantly increased with glucose addition (P = 0.002, Fig. 3a). Addition of P did not show any significant effect on N2O emission rate at each sampling day (Fig. 1a–e) or on cumulative N2O emission (Fig. 3a).
Fig. 1

Mean N2O emission rates from soil with and without glucose (Glu) and/or phosphorus (P) addition at days 0, 1, 2, 5, and 13 of the 13-day incubation (a–e). Error bars represent standard error (n = 5). P values for ANOVA are reported when significant (P < 0.05) and marginally significant (P < 0.1)

Fig. 2

Mean N2O (a) and CO2 (b) emission rates from soil with and without glucose (Glu) addition during the 13-day incubation. Gas emission rates are in log scale. Error bars represent standard error (n = 5). P values for ANOVA are reported when significant (P < 0.05)

Fig. 3

Mean cumulative N2O (a) and CO2 (b) emissions and the ratio of N2O and CO2 emission rates (c) from soil with and without glucose (Glu) and/or phosphorus (P) addition of the 13-day incubation. Error bars represent standard error (n = 5). P values for ANOVA are reported when significant (P < 0.05) and marginally significant (P < 0.1)

Unlike N2O emission, CO2 emission significantly increased with glucose addition at each sampling day, days 0, 1, 2, 5, and 13 (P = 0.004, 0.001, < 0.0001, < 0.0001, 0.0002, respectively, Fig. 4a–e). There was a pronounced temporal dynamic with glucose addition which was absent without glucose, causing a significant time × Glu interactive effect (P < 0.0001, Fig. 2b). As expected, cumulative CO2 emission significantly increased with glucose addition (P < 0.0001, Fig. 3b). Addition of P had no effect on CO2 emission rate at days 0, 1, and 2 but showed a significant (P = 0.01) and marginally significant (P = 0.07) reduction on CO2 emission rate at days 5 and 13, respectively (Fig. 4a–e). P addition had no significant effect on cumulative CO2 emission (Fig. 3b). We also calculated the ratio of N2O to CO2 emission rates to normalize N2O emission by separating treatment effects on denitrifiers from treatment effects on the whole microbial community. We found a marginally significant P × Glu interactive effect on N2O/CO2 ratio (P = 0.08, Fig. 3c). P addition increased the N2O/CO2 ratio when glucose was not added but decreased the ratio when glucose was present.
Fig. 4

Mean CO2 emission rates from soil with and without glucose (Glu) and/or phosphorus (P) addition at days 0, 1, 2, 5, and 13 of the 13-day incubation (a–e). Error bars represent standard error (n = 5). P values for ANOVA are reported when significant (P < 0.05) and marginally significant (P < 0.1)

Glucose addition showed a significant decrease in soil available P (P = 0.008) suggesting a possible increase of microbial immobilization of this nutrient. Surprisingly, microbial biomass P was not affected by P and glucose (Table 1). A non-responsive behavior of microbial biomass P could be because of the high P adsorption capacity of this soil (reported in Dijkstra et al. 2015). However, soil available P significantly increased (P < 0.0001) with P addition, and the large amount of remaining P in the P amended soil compared to the non-amended soil (Table 1) could suggest that the availability of P was high enough to relieve P limitation of microbes if there was any. P addition also did not affect microbial biomass N and C, whereas glucose addition significantly increased microbial biomass N (P = 0.002) and marginally significantly increased microbial biomass C (P = 0.09) (Table 1). Exchangeable NH4+ and NO3 were lower in the soils added with glucose (P = 0.08 and < 0.0001, respectively, Table 1). Although P addition did not affect exchangeable NH4+ in the glucose-added soil, it increased the NH4+ concentration in the soil without glucose (significant P × Glu interactive effect, P = 0.005). Soil NO3 showed significant main P treatment (P < 0.0001) and P × Glu interactive effects (P < 0.0001) where addition of P did not affect NO3 concentration in the soil with glucose but decreased NO3 concentration in soil without glucose. Net ammonification rates were not affected with glucose or P addition, whereas net nitrification rates were significantly decreased with glucose (P < 0.0001) and P (P = 0.03) addition (Table 1). Although P addition did not change net nitrification rates in presence of glucose, it significantly reduced these rates when glucose was not present, causing a significant P × Glu interactive effect, P = 0.02. Although net ammonification rates had no relationship with cumulative N2O emission, net nitrification rates showed a negative relationship with cumulative N2O emission (r2 = 0.46, P = 0.002). As expected, addition of glucose significantly increased DOC (P < 0.0001, Table 1).
Table 1

Mean ± standard error (SE, n = 5) of soil available P, exchangeable NH4+, NO3, dissolved organic C (DOC), net ammonification and nitrification rates, microbial biomass C, N and P, and soil pH after the 13-day soil incubation, in response to glucose (Glu) and/or phosphorus (P) addition

Treatments

Available P

Exchangeable NH4+

Net ammonification rate

NO3

Net nitrification rate

DOC

Microbial C

Microbial N

Microbial P

pH

Glu

P

(mg P kg−1)

(mg N kg−1)

(mg N kg−1 day−1)

(mg N kg−1)

(mg N kg−1 day−1)

(mg C kg−1)

(mg C kg−1)

(mg N kg−1)

(mg P kg−1)

Without

Without

1.95 ± 0.10 c

4.94 ± 0.95 a

0.09 ± 0.08

92.46 ± 5.59 a

0.22 ± 1.22

52.11 ± 1.69 c

379.39 ± 22.71

26.31 ± 2.80 ab

16.51 ± 2.03

5.72 ± 0.10 b

Without

With

7.50 ± 0.44 a

1.90 ± 0.44 b

− 0.10 ± 0.07

35.53 ± 4.19 b

− 4.27 ± 0.64

51.03 ± 1.82 c

339.99 ± 24.03

21.53 ± 2.60 b

18.93 ± 2.93

5.58 ± 0.07 b

With

Without

1.68 ± 0.22 c

1.63 ± 0.31 b

− 0.10 ± 0.05

0.20 ± 0.03 c

− 6.93 ± 0.47

64.35 ± 2.25 a

416.08 ± 34.19

37.33 ± 5.01 ab

14.20 ± 2.61

6.32 ± 0.06 a

With

With

5.28 ± 0.44 b

2.67 ± 0.42 b

− 0.03 ± 0.05

0.41 ± 0.21 c

− 6.58 ± 0.87

58.61 ± 1.56 ab

402.31 ± 26.63

39.10 ± 4.72 a

17.60 ± 3.76

6.15 ± 0.02 a

ANOVA P values

 Glu

 

0.008

0.03

ns

< 0.0001

< 0.0001

< 0.0001

0.09

0.002

ns

< 0.0001

 P

 

< 0.0001

0.08

ns

< 0.0001

0.03

ns

ns

ns

ns

0.04

 Glu × P

 

ns

0.002

ns

< 0.0001

0.02

ns

ns

ns

ns

ns

Not significant, ns. Different letters in each column indicate significant differences among treatment combinations (P < 0.05, Tukey’s HSD test)

Both P and glucose addition had no effect on gross N mineralization, but gross NH4+-N immobilization significantly increased (P = 0.03) with P addition (Table 2). However, gross N mineralization had no relationship with cumulative N2O emission. We observed significant main and interactive effects of P and glucose addition on soil 15N recovery (Table 2). Soil 15N recovery was increased by P addition but reduced by glucose addition (P = 0.0001 and < 0.0001, respectively). Although P addition did not change soil 15N recovery in soils without glucose, soil 15N recovery was significantly higher after P addition in soils treated with glucose (P × Glu interactive effect, P = 0.0002). Soil 15N recovery showed a negative relationship with cumulative N2O emission (r2 = 0.47, P = 0.001; Fig. 5). Both P and glucose addition treatment caused a significant effect on soil pH measured at the end of incubation (Table 1). Although soil pH increased in glucose-added soils (P < 0.0001), it was reduced when P was added (P = 0.04).
Table 2

Mean ± standard error (SE, n = 5) of gross N mineralization (GNM) and gross NH4+-N immobilization (GNI) rates, and soil 15N recovery measured during and at the end of soil incubations (13 days and 48 h), respectively, in response to glucose (Glu) and/or phosphorus (P) addition

Treatment

After 13-day incubation

After 48-h incubation

GNM rate

GNI rate

Soil 15N recovery

GNM rate

GNI rate

Soil 15N recovery

Glu addition

P addition

(mg N kg−1 day−1)

(mg N kg−1 day−1)

(%)

(mg N kg−1 day−1)

(mg N kg−1 day−1)

(%)

Without

Without

1.47 ± 0.20

1.17 ± 0.38

91.39 ± 2.24 ab

3.36 ± 0.28

5.48 ± 0.92

85.92 ± 2.81 ab

Without

With

2.15 ± 0.92

2.58 ± 0.92

91.54 ± 2.93 a

3.82 ± 0.41

5.96 ± 0.99

78.56 ± 1.76 b

With

Without

1.40 ± 0.12

1.10 ± 0.24

59.55 ± 1.23 c

3.27 ± 0.31

4.64 ± 0.55

92.56 ± 5.15 ab

With

With

1.59 ± 0.11

2.96 ± 0.65

82.35 ± 1.72 b

2.99 ± 0.39

5.50 ± 0.94

98.14 ± 5.54 a

ANOVA P values

 Glu

 

ns

ns

< 0.0001

ns

ns

0.008

 P

 

ns

0.03

0.0001

ns

ns

ns

 Glu × P

 

ns

ns

0.0002

ns

ns

ns

Not significant, ns. Different letters in each column indicate significant differences among treatment combinations (P < 0.05, Tukey’s HSD test)

Fig. 5

Relationship of soil 15N recovery with cumulative N2O emission in response to glucose (Glu) and/or P addition, after the 13-day incubation

Forty-eight-hour incubation study (at 75% WHC):

Gross N mineralization and NH4+-N immobilization rates measured under more aerobic conditions (at 75% WHC) were not affected by P or glucose addition (Table 2). However, soil 15N recovery was significantly higher with glucose addition (P = 0.008; Table 2). Both exchangeable NH4+ and NO3 were significantly decreased with glucose addition (P = 0.003 and < 0.0001, respectively) but were not affected by P addition (Table 3). Addition of glucose significantly increased microbial biomass C (P = 0.003) and marginally significantly increased microbial biomass N (P = 0.09, Table 3). As in the previous experiment, P addition had no effect on microbial biomass C or N.
Table 3

Mean ± standard error (SE, n = 5) of soil exchangeable NH4+, NO3, microbial biomass C and N after the 48-h incubation, in response to glucose (Glu) and/or P addition

Treatments

Exchangeable NH4+

NO3

Microbial C

Microbial N

Glu addition

P addition

(mg N kg−1)

(mg N kg−1)

(mg C kg−1)

(mg N kg−1)

Without

Without

1.38 ± 0.30 ab

8.65 ± 2.23 a

538.96 ± 58.58 a

81.44 ± 2.34

Without

With

1.48 ± 0.27 a

9.22 ± 2.36 a

574.83 ± 31.98 a

80.05 ± 3.64

With

Without

0.84 ± 0.13 ab

0.51 ± 0.11 b

736.81 ± 53.28 a

90.58 ± 8.54

With

With

0.66 ± 0.13 b

0.52 ± 0.16 b

732.90 ± 57.30 a

91.88 ± 5.07

ANOVA P values

 Glu

 

0.003

< 0.0001

0.003

0.09

 P

 

Ns

Ns

ns

ns

 Glu × P

 

Ns

Ns

ns

ns

Not significant, ns. Different letters in each column indicate significant differences among treatment combinations (P < 0.05, Tukey’s HSD test)

Discussion

Effect of C and P addition on N2O emission, respiration, and gross N mineralization (at 90% WHC)

Addition of glucose showed no effect on gross N mineralization (Table 2) but caused a significant increase in cumulative N2O emission (Fig. 3a) suggesting that the denitrifying microbes were limited by labile organic C in this soil, but not the heterotrophic microbial community in general. In the presence of high NO3 concentration, availability of labile organic C can stimulate N2O production through denitrification (Firestone 1982; Luo et al. 1999). Although greater availability of C often decreases the N2O/N2 ratio, the significant increase of N2O emission rate (at days 0, 1, and 2) as well as cumulative N2O emission suggests an overall increase of microbial denitrification. Addition of glucose may have influenced denitrification directly by supplying donor electrons to the denitrifiers and/or indirectly by lowering O2 concentrations with the stimulation of heterotrophic microbial respiration (Morley and Baggs 2010; Signor and Cerri 2013; Robertson and Groffman 2007). The CO2 emission rates at all sampling dates (Fig. 4a–e) as well as the cumulative CO2 emission (Fig. 3b) significantly increased with glucose addition, indicating increased respiration activities with glucose addition.

Addition of glucose also showed a temporal effect on N2O emission (Fig. 2a). Although initially (at days 0, 1, and 2) N2O emission significantly increased with glucose addition, it decreased at the later stage of incubation (at days 5 and 13) (Fig. 1), which could suggest that denitrifiers were no longer limited by C, but rather by NO3 in the soil. In presence of readily available C substrates, a rapid initial NO3 respiration of denitrifiers may have subsequently reduced the NO3 concentration of soil in the glucose added pots (Mori et al. 2013) and hence reduced denitrification and N2O emission at the later stages of incubation. Significantly reduced soil NO3 concentration was observed with glucose addition (Table 1) at the end of the 13-day incubation. However, the higher concentration of NO3 (~ 100 mg N kg−1) compared to the NH4+ concentration (~ 5 mg N kg−1) at the start of the incubation may have stimulated NO3 immobilization more than NH4+ immobilization after glucose addition, further reducing the soil NO3 concentration for N2O production. No significant effect of glucose on gross NH4+-N immobilization (Table 2) but significantly higher soil microbial biomass N with glucose addition (Table 1) at the end of the 13-day incubation can support this hypothesis. However, the reduced N2O emission at the later stage of incubation could also be due to the increased reduction of N2O to N2. The increasing percentage of NO3 lost via denitrification and/or a possible lowering of partial pressure of O2 resulting from increased microbial respiration after glucose addition could stimulate the final reductive step of denitrification, hence decrease N2O production and make N2 the principle gas evolved from denitrification (Firestone and Davidson 1989; Weier et al. 1993). Unfortunately, we did not measure N2 emission.

Addition of P did not show any significant effect on N2O emission rates at each sampling date (Fig. 1) as well as on cumulative N2O emission (Fig. 3a), and this contradicts other studies where N2O emission was decreased (Sundareshwar et al. 2003) or increased (Mori et al. 2010, 2013; Mehnaz and Dijkstra 2016) with added P. It was suggested that alleviation of P limitation can stimulate microbial N immobilization and hence reduce N substrates for nitrification and denitrification, resulting in reduced N2O emission (Hall and Matson 1999). In contrast, others suggested that high availability of P can increase N2O emission by (i) stimulating N mineralization leading to increased inorganic N to be utilized in nitrification and denitrification, (ii) developing anaerobic conditions suitable for denitrification by increasing heterotrophic respiration, and/or (iii) alleviating P limitation of nitrifying and/or denitrifying bacteria (Mori et al. 2010, 2013, 2017). Although gross NH4+-N immobilization (Table 2), measured within the first 2 days of incubation, was increased with P addition, and soil NO3 was reduced in combination with glucose addition after 13 days (Table 1), it did not affect N2O emission. P addition had no effect on the gross rate of N mineralization in this soil, suggesting no increase in inorganic N to support nitrifiers and denitrifiers for N2O production. Unlike other studies (Cleveland et al. 2002; Ilstedt and Singh 2005), P addition did not stimulate microbial respiration, suggesting that P addition did not increase O2 consumption to provide reducing conditions to facilitate denitrification-N2O loss. The lack of a P addition effect on cumulative CO2 emission (Fig. 3b), gross N mineralization (Table 2), and CO2 emission rates during the first 2 days of incubation (Fig. 4a–c) also suggests that the microbial activity, possibly including heterotrophic denitrifying activity, was not limited by P in this soil. No significant effect of P on microbial biomass P and C (Table 1) supports this suggestion (although our microbial biomass measurements using fumigation-extraction may not be very sensitive to short-term changes in P availability). Similarly, the lack of a P addition effect on N2O emission suggests that the nitrifying and/or denitrifying microorganisms were not limited by P in this soil incubated at 90% WHC. However, this result is opposite to our previous study (Mehnaz and Dijkstra 2016) conducted with soil taken from the same grassland but incubated at 45% WHC with smaller amount of added NO3 (15 mg N kg−1 soil), where P addition stimulated CO2 emission as well as N2O emission from denitrification. Possibly, the denitrifying community is more sensitive to P addition when N availability and/or soil moisture are low.

The significant reduction in soil pH with P addition (Table 1) could also have affected biomass, activity (Pietri and Brookes 2008), as well as composition of the microbial community (Bååth and Anderson 2003). It is unclear why P addition reduced soil pH, since P was added in solution with the same pH as in the soil. A possible explanation could be an increase in nitrification rate after P addition. Hue and Adams (1984) found slow rates of nitrification with low P concentration in soil solution, with greater effect on the NH4+—than the nitrite (NO2)—oxidizing bacteria. In this study, high availability of P might have stimulated NH4+ oxidation and thus reduced soil pH by releasing protons. The NH4+ concentration of soil decreased with P addition at the end of incubation (Table 1); however, the decrease was only significant without C addition. The reduced soil pH with P addition may have reduced the activity of the whole heterotrophic microbial community, as well as inhibited the growth of denitrifiers (Wang et al. 2017), which could possibly offset the stimulated N2O emission from denitrification. The reduction in microbial activity with P addition can be supported by the decrease in CO2 emission observed after P addition during the last 9 days of incubation (Fig. 4d, e). Although the ratio of N2O to N2 frequently increases with declining pH (Wrage et al. 2001), an overall decrease in denitrification rate (Šimek and Hopkins 1999) may have caused a net reduction in N2O production in response to reduced soil pH after P addition.

Although P addition had no effect on N2O loss, 15N recovery in glucose-added pots was significantly reduced without P addition (Table 2), while addition of glucose overall reduced 15N recovery in the soil (Table 2). The significant negative relationship between soil 15N recovery and cumulative N2O emission (Fig. 5) suggests that the loss of 15N in the soil was caused by denitrification. However, there was no relationship of cumulative N2O emission with gross N mineralization and net ammonification rates, but a significant negative relationship with net nitrification rates, suggesting that the loss of N as N2O was mostly from heterotrophic denitrification, as we expected due to the high soil moisture condition of the soil. While the increase in denitrification with glucose addition likely caused increased loss of 15N from the soil, we are unclear why the largest 15N loss (lowest 15N recovery) occurred without P addition. Since N2O emission was not affected by P addition, we speculate that gaseous loss of N after C addition in this soil with high moisture content occurred mostly as N2, but that N2 emission was significantly reduced after P addition and increased the N2O/N2 ratio. Unfortunately, we did not measure N2 emission. Further study should be conducted to understand the effect of P on different product ratios of denitrification.

As N2O emission in this soil with high moisture percentage (90% WHC) seemed to occur mostly from denitrification, we calculated the ratio of N2O and CO2 emission rates (Fig. 3c), to separate the treatment effect on denitrifiers from the effect on the overall heterotrophic community (although this normalization is not absolute as N2O could also be produced from autotrophic processes, such as nitrification). The increase in this ratio with P addition without glucose addition could suggest that P availability may have favored denitrifying activities (supported by reduced net nitrification rates after P addition without glucose addition) compared to the overall heterotrophic activities, at least in the absence of sufficient C, supporting the findings of our previous study (Mehnaz and Dijkstra 2016). A decrease in this ratio after P addition with glucose addition could suggest a possible decrease in denitrifying activities, compared to the overall heterotrophic activities, may be because of increased microbial N immobilization (supported by the greater soil 15N recovery with P addition in presence of glucose). A change in the microbial community composition may have occurred after P addition in the presence and/or absence of glucose due to a change in the elemental stoichiometry of soil (Zechmeister-Boltenstern et al. 2015). Further studies should be conducted on microbial community composition to understand the P addition effect on N2O emission.

Effect of C and P addition on gross N mineralization and soil 15N recovery (at 75% WHC)

Results from the 48-h incubation conducted under 75% WHC suggest that the gross N mineralization and thus soil organic matter decomposition was not limited by P and glucose in this grassland soil under more aerobic conditions either (Table 2). However, the increased soil 15N recovery with glucose addition in this short-term incubation study (Table 2) suggests a short-term reduction in gaseous N loss from the soils treated with glucose (as no leaching occurred in the pots), which is opposite to what we found in our 13-day incubation study, possibly due to the different C/N stoichiometry prevailing in soils of the two separate experiments. The high availability of soluble C, in relatively low soil moisture (75% WHC) and low inorganic N environment (no extra N was added except 1 mg 15NH4+-N kg−1 soil) compared to the 13-day incubation study, may have stimulated a short-term heterotrophic microbial growth and immobilization of N which ultimately reduced the nitrifying and denitrifying activity provided with reduced N substrates (Luo et al. 1999). Increased microbial biomass C after glucose addition (Table 3) confirms the stimulating effect of soluble C on microbial growth. The lower exchangeable NH4+ and NO3 in the glucose-added soils observed at the end of incubation (Table 3) could suggest increased immobilization of inorganic N by heterotrophic microbes. In fact, our data also showed that glucose addition increased microbial biomass N in the soil (Table 3). However, glucose addition had no effect on gross NH4+-N immobilization (Table 2), suggesting a rapid production of 15NO3 from 15NH4+ through nitrification (favored by the relatively low soil moisture condition in this study soil), which subsequently was taken up by heterotrophic microbes in the presence of high C availability.

Conclusion

Our results showed that denitrification was mostly responsible for the N2O loss from this soil with high moisture content (90% WHC), especially in the presence of glucose suggesting C limitation of denitrifiers. Although high availability of labile C, in the presence of sufficient NO3, can initially increase N2O emission, a rapid denitrification and NO3 immobilization can subsequently reduce soil NO3 concentrations and N2O production. However, a higher C/N stoichiometry and a relatively low moisture content (75% WHC) in soil could change the N loss and turnover cycle, at least in the short term, by increasing immobilization and hence soil recovery. As N2O emission was not affected by P, a reduced N loss after P addition in presence of glucose could suggest a potential increase in N2O/N2 ratio with high P availability. A change in the C/P stoichiometry of soil after glucose or P addition could have possibly changed the relative contribution of denitrifiers compared to the overall heterotrophic community by changing the microbial community composition.

Notes

Acknowledgements

We thank Hero Tahaei and Janani Vimalathithen for their assistance in soil chemical analyses. This research was financially supported by the Australian Research Council (FT100100779).

References

  1. Aulakh MS, Walters DT, Doran JW, Francis DD, Mosier AR (1991) Crop residue type and placement effects on denitrification and mineralization. Soil Sci Soc Am J 55:1020–1025CrossRefGoogle Scholar
  2. Bååth E, Anderson TH (2003) Comparison of soil fungal/bacterial ratios in a pH gradient using physiological and PLFA-based techniques. Soil Biol Biochem 35:955–963CrossRefGoogle Scholar
  3. Baral BR, Kuyper TW, Van Groenigen JW (2014) Liebig’s law of the minimum applied to a greenhouse gas: alleviation of P-limitation reduces soil N2O emission. Plant Soil 374:539–548CrossRefGoogle Scholar
  4. Beck T, Joergensen R, Kandeler E, Makeschin F, Nuss E, Oberholzer H, Scheu S (1997) An inter-laboratory comparison of ten different ways of measuring soil microbial biomass C. Soil Biol Biochem 29:1023–1032CrossRefGoogle Scholar
  5. Braun J, Mooshammer M, Wanek W, Prommer J, Walker TWN, Rütting T, Richter A (2018) Full 15N tracer accounting to revisit major assumptions of 15N isotope pool dilution approaches for gross nitrogen mineralization. Soil Biol Biochem 117:16–26CrossRefGoogle Scholar
  6. Bremner JM (1997) Sources of nitrous oxide in soils. Nutr Cycl Agroecosyst 49:7–16CrossRefGoogle Scholar
  7. Brookes PC, Powlson DS, Jenkinson DS (1982) Measurement of microbial biomass phosphorus in soil. Soil Biol Biochem 14:319–329CrossRefGoogle Scholar
  8. Brookes PC, Landman A, Pruden G, Jenkinson D (1985) Chloroform fumigation and the release of soil nitrogen: a rapid direct extraction method to measure microbial biomass nitrogen in soil. Soil Biol Biochem 17:837–842CrossRefGoogle Scholar
  9. Bruulsema T, Duxbury J (1996) Simultaneous measurement of soil microbial nitrogen, carbon, and carbon isotope ratio. Soil Sci Soc Am J 60:1787–1791CrossRefGoogle Scholar
  10. Butterbach-Bahl K, Baggs EM, Dannenmann M, Kiese R, Zechmeister-Boltenstern S (2013) Nitrous oxide emissions from soils: how well do we understand the processes and their controls? Phil Trans R Soc B 368:20130122CrossRefPubMedPubMedCentralGoogle Scholar
  11. Cadisch G, Giller KE, Urquiaga S, Miranda CHB, Boddey RM, Schunke RM (1994) Does phosphorus supply enhance soil-N mineralization in Brazilian pastures? Eur J Agron 3:339–345CrossRefGoogle Scholar
  12. Chen R, Senbayram M, Blagodatsky S, Myachina O, Dittert K, Lin X, Blagodatskaya E, Kuzyakov Y (2014) Soil C and N availability determine the priming effect: microbial N mining and stoichiometric decomposition theories. Glob Chang Biol 20:2356–2367CrossRefPubMedGoogle Scholar
  13. Chen H, Zhang W, Gurmesa GA, Zhu X, Li D, Mo J (2017) Phosphorus addition affects soil nitrogen dynamics in a nitrogen-saturated and two nitrogen-limited forests. Eur J Soil Sci 68:472–479CrossRefGoogle Scholar
  14. Cleveland CC, Liptzin D (2007) C:N: P stoichiometry in soil: is there a “Redfield ratio” for the microbial biomass? Biogeochemistry 85:235–252CrossRefGoogle Scholar
  15. Cleveland CC, Townsend AR, Schmidt SK (2002) Phosphorus limitation of microbial processes in moist tropical forests: evidence from short-term laboratory incubations and field studies. Ecosystems 5:680–691CrossRefGoogle Scholar
  16. Dalal RC, Wang W, Robertson GP, Parton WJ (2003) Nitrous oxide emission from Australian agricultural lands and mitigation options: a review. Soil Res 41:165–195CrossRefGoogle Scholar
  17. Dijkstra FA, Hutchinson GL, Reeder JD, LeCain DR, Morgan JA (2011) Elevated CO2, but not defoliation, enhances N cycling and increases short-term soil N immobilization regardless of N addition in a semiarid grassland. Soil Biol Biochem 43:2247–2256CrossRefGoogle Scholar
  18. Dijkstra FA, He M, Johansen MP, Harrison JJ, Keitel C (2015) Plant and microbial uptake of nitrogen and phosphorus affected by drought using 15N and 32P tracers. Soil Biol Biochem 82:135–142CrossRefGoogle Scholar
  19. Falkiner RA, Khanna PK, Raison RJ (1993) Effect of superphosphate addition on N mineralization in some Australian forest soils. Soil Res 31:285–296CrossRefGoogle Scholar
  20. Firestone MK (1982) Biological denitrification. In: Stevenson FJ (Ed) Nitrogen in agricultural soils. Am Soc Agron, Madison, WI, USA, pp. 289–326Google Scholar
  21. Firestone MK, Davidson EA (1989) Microbiological basis of NO and N2O production and consumption in soil. In: Andreae MO, Schimel DS (eds) Exchange of trace gases between terrestrial ecosystems and the atmosphere. Wiley, New York, pp 7–21Google Scholar
  22. Flavel TC, Murphy DV (2006) Carbon and nitrogen mineralization rates after application of organic amendments to soil. J Environ Qual 35:183–193CrossRefPubMedGoogle Scholar
  23. Forster P, Ramaswamy V, Artaxo P et al (2007) Changes in atmospheric constituents and in radiative forcing. In: Solomon S, Qin D, Manning M et al (eds) Climate change 2007: the physical science basis. Contribution of working group I to the fourth assessment report of the intergovernmental panel on climate change. Cambridge University Press, Cambridge, pp 129–234Google Scholar
  24. Hall SJ, Matson PA (1999) Nitrogen oxide emissions after nitrogen additions in tropical forests. Nature 400:152–155CrossRefGoogle Scholar
  25. Henry S, Texier S, Hallet S, Bru D, Dambreville C, Chèneby D, Bizouard F, Germon JC, Philippot L (2008) Disentangling the rhizosphere effect on nitrate reducers and denitrifiers: insight into the role of root exudates. Environ Microbiol 10:3082–3092CrossRefPubMedGoogle Scholar
  26. Hill BH, Elonen CM, Jicha TM, Kolka RK, Lehto LL, Sebestyen SD, Seifert-Monson LR (2014) Ecoenzymatic stoichiometry and microbial processing of organic matter in northern bogs and fens reveals a common P-limitation between peatland types. Biogeochemistry 120:203–224CrossRefGoogle Scholar
  27. Houlton BZ, Wang YP, Vitousek PM (2008) A unifying framework for dinitrogen fixation in the terrestrial biosphere. Nature 454:327–330CrossRefPubMedGoogle Scholar
  28. Hue NV, Adams F (1984) Effect of phosphorus level on nitrification rates in three low-phosphorus ultisols. Soil Sci 137:324–331CrossRefGoogle Scholar
  29. Ilstedt U, Singh S (2005) Nitrogen and phosphorus limitations of microbial respiration in a tropical phosphorus-fixing acrisol (ultisol) compared with organic compost. Soil Biol Biochem 37:1407–1410CrossRefGoogle Scholar
  30. Kirkham DON, Bartholomew WV (1954) Equations for following nutrient transformations in soil, utilizing tracer data. Soil Sci Soc Am J 18:33–34CrossRefGoogle Scholar
  31. Kranabetter JM, Banner A, Groot AD (2005) An assessment of phosphorus limitations to soil nitrogen availability across forest ecosystems of north coastal British Columbia. Can J For Res 35:530–540CrossRefGoogle Scholar
  32. Liu R, Hayden HL, Suter H, Hu H, Lam SK, He J, Mele PM, Chen D (2017) The effect of temperature and moisture on the source of N2O and contributions from ammonia oxidizers in an agricultural soil. Biol Fertil Soils 53:141–152CrossRefGoogle Scholar
  33. Luo J, Tillman RW, Ball PR (1999) Factors regulating denitrification in a soil under pasture. Soil Biol Biochem 31:913–927CrossRefGoogle Scholar
  34. Mehnaz KR, Dijkstra FA (2016) Denitrification and associated N2O emissions are limited by phosphorus availability in a grassland soil. Geoderma 284:34–41CrossRefGoogle Scholar
  35. Mori T, Ohta S, Ishizuka S, Konda R, Wicaksono A, Heriyanto J, Hardjono A (2010) Effects of phosphorus addition on N2O and NO emissions from soils of an Acacia mangium plantation. Soil Sci Plant Nutr 56:782–788CrossRefGoogle Scholar
  36. Mori T, Ohta S, Ishizuka S, Konda R, Wicaksono A, Heriyanto J, Hardjono A (2013) Effects of phosphorus addition with and without ammonium, nitrate, or glucose on N2O and NO emissions from soil sampled under Acacia mangium plantation and incubated at 100% of the water-filled pore space. Biol Fertil Soils 49:13–21CrossRefGoogle Scholar
  37. Mori T, Ohta S, Ishizuka S, Konda R, Wicaksono A, Heriyanto J (2014) Phosphorus application reduces N2O emissions from tropical leguminous plantation soil when phosphorus uptake is occurring. Biol Fertil Soils 50:45–51CrossRefGoogle Scholar
  38. Mori T, Yokoyama D, Kitayama K (2016) Contrasting effects of exogenous phosphorus application on N2O emissions from two tropical forest soils with contrasting phosphorus availability. SpringerPlus 5:1237CrossRefPubMedPubMedCentralGoogle Scholar
  39. Mori T, Wachrinrat C, Staporn D, Meunpong P, Suebsai W, Matsubara K, Boonsri K, Lumban W, Kuawong M, Phukdee T, Srifai J, Boonman K (2017) Effects of phosphorus addition on nitrogen cycle and fluxes of N2O and CH4 in tropical tree plantation soils in Thailand. Agric Nat Resour 51:91–95Google Scholar
  40. Morley N, Baggs EM (2010) Carbon and oxygen controls on N2O and N2 production during nitrate reduction. Soil Biol Biochem 42:1864–1871CrossRefGoogle Scholar
  41. Munevar F, Wollum AG (1977) Effects of the addition of phosphorus and inorganic nitrogen on carbon and nitrogen mineralization in Andepts from Colombia. Soil Sci Soc Am J 41:540–545CrossRefGoogle Scholar
  42. Murray PJ, Hatch DJ, Dixon ER, Stevens RJ, Laughlin RJ, Jarvis SC (2004) Denitrification potential in a grassland subsoil: effect of carbon substrates. Soil Biol Biochem 36:545–547CrossRefGoogle Scholar
  43. Olsen SR, Sommers LE (1982) Phosphorus. In: Pace AL, Miller RH, Keeney DR (eds) Methods of soil analysis. Part 2. Chemical and microbiological properties, 2nd edn. Am Soc Agron, Inc., Madison, WI, pp 403–430Google Scholar
  44. Pietri JA, Brookes PC (2008) Relationships between soil pH and microbial properties in a UK arable soil. Soil Biol Biochem 40:1856–1861CrossRefGoogle Scholar
  45. Raiesi F, Ghollarata M (2006) Interactions between phosphorus availability and an AM fungus (Glomus intraradices) and their effects on soil microbial respiration, biomass and enzyme activities in a calcareous soil. Pedobiologia 50:413–425CrossRefGoogle Scholar
  46. Ravishankara A, Daniel JS, Portmann RW (2009) Nitrous oxide (N2O): the dominant ozone-depleting substance emitted in the 21st century. Science 326:123–125CrossRefPubMedGoogle Scholar
  47. Robertson GP, Groffman PM (2007) Nitrogen transformations. Soil Microbiol Ecol. Biochemist 3:341–364Google Scholar
  48. Ross DJ, Speir TW, Kettles HA, Mackay AD (1995) Soil microbial biomass, C and N mineralization and enzyme activities in a hill pasture: influence of season and slow-release P and S fertilizer. Soil Biol Biochem 27:1431–1443CrossRefGoogle Scholar
  49. Signor D, Cerri CEP (2013) Nitrous oxide emissions in agricultural soils: a review. Pesq Agrop Trop 43:322–338CrossRefGoogle Scholar
  50. Šimek M, Hopkins DW (1999) Regulation of potential denitrification by soil pH in long-term fertilized arable soils. Biol Fertil Soils 30:41–47CrossRefGoogle Scholar
  51. Stark JM, Hart SC (1996) Diffusion technique for preparing salt solutions, Kjeldahl digests, and persulfate digests for nitrogen-15 analysis. Soil Sci Soc Am J 60:1846–1855CrossRefGoogle Scholar
  52. Sundareshwar P, Morris J, Koepfler E, Fornwalt B (2003) Phosphorus limitation of coastal ecosystem processes. Science 299:563–565CrossRefPubMedGoogle Scholar
  53. Wakelin SA, Condron LM, Gerard E, Dignam BE, Black A, O’Callaghan M (2017) Long-term P fertilisation of pasture soil did not increase soil organic matter stocks but increased microbial biomass and activity. Biol Fertil Soils 53:511–521CrossRefGoogle Scholar
  54. Wang Q, Liu YR, Zhang CJ, Zhang LM, Han LL, Shen JP, He JZ (2017) Responses of soil nitrous oxide production and abundances and composition of associated microbial communities to nitrogen and water amendment. Biol Fertil Soils 53:601–611CrossRefGoogle Scholar
  55. Weier KL, Doran JW, Power JF, Walters DT (1993) Denitrification and dinitrogen/nitrous oxide ratio as affected by soil water, available carbon dioxide and nitrous oxide production in tilled and nontilled soils. Soil Sci Soc Am J 48:1267–1272Google Scholar
  56. Weil RR, Brady NC (2016) The nature and properties of soils, 15th edn. Pearson, Boston, p 912Google Scholar
  57. White J, Reddy K (1999) Influence of nitrate and phosphorus loading on denitrifying enzyme activity in Everglades wetland soils. Soil Sci Soc Am J 63:1945–1954CrossRefGoogle Scholar
  58. White J, Reddy K (2000) Influence of phosphorus loading on organic nitrogen mineralization of Everglades soils. Soil Sci Soc Am J 64:1525–1534CrossRefGoogle Scholar
  59. Wrage N, Velthof G, Van Beusichem M, Oenema O (2001) Role of nitrifier denitrification in the production of nitrous oxide. Soil Biol Biochem 33:1723–1732CrossRefGoogle Scholar
  60. Zechmeister-Boltenstern S, Keiblinger KM, Mooshammer M, Peñuelas J, Richter A, Sardans J, Wanek W (2015) The application of ecological stoichiometry to plant–microbial–soil organic matter transformations. Ecol Monogr 85:133–155CrossRefGoogle Scholar

Copyright information

© Springer-Verlag GmbH Germany, part of Springer Nature 2018

Authors and Affiliations

  • Kazi R. Mehnaz
    • 1
  • Claudia Keitel
    • 1
  • Feike A. Dijkstra
    • 1
  1. 1.Sydney Institute of Agriculture, School of Life and Environmental SciencesThe University of SydneyCamdenAustralia

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