Several laboratory‐based studies have documented the suppression of N 2 O emissions as a result of biochar addition to soils. Cayuela et al. (2010) demonstrated that biochar, produced from green waste and poultry manure, reduced soil N 2 O emissions relative to a control soil when incubated for 60 d at 20°C and 80% water‐filled pore space (WFPS). Rondon et al. (2005) demonstrated reductions in N 2 O emissions of 50 and 80%, following the incorporation of biochar into the soil under soybean and grass systems, respectively. Yanai et al. (2007) concluded that varying the soil moisture status caused biochar to either stimulate or suppress N 2 O emissions due to its effect on soil aeration, since the denitrification mechanism for N 2 O production is aeration dependent. An experiment by Singh et al. (2010) used repacked soil columns, into which were mixed poultry and wood waste biochars. After several wetting–drying cycles, the N 2 O emissions were reduced by up to 72%. Using an unweathered biochar in a laboratory experiment, Clough et al. (2010) found the effect of biochar incorporation (20 t ha −1 ) initially stimulated N 2 O emissions in the presence of ruminant urine, although the cumulative N 2 O flux over time was not significantly different from a urine‐only treatment. Thus, biochar incorporation into soil can affect N 2 O fluxes, but detailed field data are lacking. There have, to our knowledge, been no reports of in situ work performed in pastures under urine treatments.

Intensively managed, grazed pastures receive N inputs as a result of fertilizer application and excreta deposition. A single ruminant urine deposition results in N rates of up to 1000 kg N ha −1 in the urine patch, creating “hot spots,” where soil N concentrations exceed the pasture plants' immediate demands ( Haynes and Williams, 1993 ). Subsequent transformation of this urinary‐N, which is predominantly urea‐N, initially leads to the creation of a significant ammonium (NH 4 + ) pool in the soil. Nitrification of this NH 4 + pool creates nitrate (NO 3 − ). Transformation of these inorganic‐N pools results in N 2 O being produced via nitrification, nitrifier‐denitrification, or denitrification ( Wrage et al., 2001 ).

Anthropogenic emissions of nitrous oxide (N 2 O) are of environmental interest since N 2 O is a greenhouse gas ( Forster et al., 2007 ) and because tropospheric N 2 O emissions are currently the primary source of stratospheric nitrogen oxides. These are involved in catalytic destruction of ozone ( Ravishankara et al., 2009 ). Globally, total anthropogenic emissions of N 2 O are estimated at 6.7 Tg N yr −1 ( Denman et al., 2007 ), with an estimated 1.5 Tg of this total due to the excreta of grazing animals ( Oenema et al., 2005 ).

Statistical analyses were performed using Minitab version 15.1 ( Minitab, 2006 ). Tests for normality showed the N 2 O flux data were skewed and so these were log transformed (In[flux+1]) before statistical analyses, with N 2 O fluxes compared on individual sampling days and for cumulative fluxes over the sampling period. Analysis of variance was used to determine if treatment means were equal. Where significant differences were detected, two‐sample t tests were used to further identify differences between specific treatment means.

Pasture dry matter yields were determined on the gas chamber areas (on Days 21, 43, and 58) by hand harvesting the herbage at a height of 0.05 m. This herbage was dried at 65°C for 48 h, finely ground to <200 μm, and analyzed for its total N content and atom% 15 N enrichment using Dumas combustion and isotope ratio continuous‐flow mass spectrometer (Sercon, UK).

Soil cores (0.025 m diam. × 0.075 m depth) were taken on 17 occasions over the course of the study to monitor θ g and inorganic‐N concentrations. On each occasion, two soil cores were taken from each soil sampling plot, bagged and mixed, and then immediately transported to the laboratory where a subsample of soil was dried at 105°C for 24 h to determine θ g Another 10‐g subsample of moist soil was shaken with 100 mL of 2M KCl for 1 h and then filtered (Whatman No. 42), with the extracts analyzed, using flow injection analysis ( Blakemore et al., 1987 ) for ammonium N (NH 4 + ‐N) and nitrate N (NO 3 − ‐N).

Soil bulk densities (Mg m −3 ) of the main plots were determined 5 mo after pasture renovation on 15 Oct. 2009. These were determined by taking a soil core (0.073 m diam. × 0.075 m deep) and drying the soil at 105°C for 48 h to determine gravimetric moisture content (θ g ) of the sample and calculating the bulk density. Soil surface pH measurements were taken on 33 occasions following urine‐treatment application, from 2 d before urine application until 86 d after urine application, using a flat‐surface pH electrode (Broadley‐James, Irvine, CA), calibrated with appropriate buffer solutions. Gas samples were taken for N 2 O flux determinations on the same days as the soil surface pH measurements. On each gas‐sampling occasion at 0, 15, and 30 min, after positioning the headspace cover, headspace N 2 O samples (10 mL) were taken manually using glass syringes fitted with three‐way taps and compressed into 6‐mL Exetainer tubes (Labco Ltd., High Wycombe, UK). Immediately before analysis, these gas samples were brought to ambient pressure using a double‐ended needle and analyzed on a gas chromatograph (8610, SRI Instruments, Torrance, CA) linked to an autosampler (Gilson 222XL, Middleton, WI), as described by Clough et al. (2010) A further 15‐mL‐headspace gas sample was taken and placed into 12‐mL Exetainer tubes and equilibrated to atmospheric pressure, before analysis for N 2 O‐ 15 N enrichment using automated continuous‐flow isotope‐ratio mass spectrometry, as described by Stevens et al. (1993)

Four biochar‐urine treatments, replicated five times, were set up on the field trial area. Two of these treatments, consisting of nil biochar plus nil urine (control) and nil biochar plus urine (0U), were positioned on the 0 t ha −1 biochar plots ( Fig. 1 ). The biochar at 15 t ha −1 plus urine (15U) and biochar at 30 t ha −1 plus urine (30U) treatments were sited on the 15 and 30 t ha −1 plots, respectively ( Fig. 1 ). Before urine application, pasture was cut to a height of 0.05 m to simulate grazing. Then, urine was collected from dairy cows at the Lincoln University dairy farm (43°39′2″ S, 172°27′13″ E) that had been grazing perennial ryegrass/white clover ( Trifolium repens L.) pasture. The urine contained 5 g N L −1 when collected. This urine was split into two portions. One portion was enriched with 15 N‐labeled urea to 4.963 atom% 15 N, with a final urinary‐N concentration of 10 g N L −1 , before applying it only to pasture within the gas sampling chambers at a rate of 930 kg N ha −1 The second portion had urea, at natural abundance, added so that the concentration of urinary‐N also equaled 10 g N L −1 This was added to the soil sampling plots, adjacent to the gas sampling chambers, at the same N rate. Urine was applied on 26 Nov. 2009.

On 13 Nov. 2009, headspace chamber bases (diam. 0.39 m, stainless steel), which protruded 0.10 m into the soil, were installed. These contained an annular water trough. During gas sampling events, insulated, stainless steel headspace covers with 0.10‐m‐high walls created an 11.6‐L headspace when they were placed on the bases. The headspace cover sat on the annular water‐filled trough, creating a gas‐tight seal. Located immediately adjacent to each gas sampling chamber was a soil sampling plot (0.37 m × 0.43 m) ( Fig. 1 ).

A runout perennial ryegrass ( Lolium perenne L.) pasture situated at Lincoln University (43°38′58″ S, 172°27′53″ E), on a Templeton silt loam soil ( Hewitt, 1998 ), was renovated in May (autumn) 2009 ( Table 1 ). The pasture was cultivated to a depth of 0.30 m, using a rotocultivator. Unweathered biochar, manufactured from Monterey pine ( Pinus radiata ) ( Table 2 ), was then incorporated to a depth of 0.10 m at rates of either 0, 15, or 30 t ha −1 , according to experimental treatment (see below). This was achieved by spreading the biochar material onto the plots and then making a shallow pass with the rotocultivator. The trial area was then rolled with a Cambridge roller to produce a fine seedbed tilth before sowing with a forage perennial ryegrass (cultivar Samson) at a rate of 12.5 kg ha −1 in rows 0.14 m apart. After ryegrass emergence, urea fertilizer was applied twice—83 kg ha −1 on 9 Sept. 2009, and 50 kg ha −1 on 28 Oct. 2009. To suppress broadleaf weed growth, a selective herbicide (Jaguar, Bayer CropScience, Research Triangle Park, NC) was applied (1.5 L ha −1 ) on 21 Oct. 2009. A fungicide to prevent stem rust (Proline, Bayer CropScience) was applied (0.2 L ha −1 ) on 19 Nov. 2009.

As anticipated, the 15 N enrichment of the N 2 O from the urine‐ 15 N treated plots remained higher than in the control treatment throughout the entire period of the study ( Fig. 6 ). When comparing the biochar‐urine treatments, there was a trend for the atom% 15 N enrichment of the N 2 O to be lower with increasing rates of biochar from Day 11 to 33, and this was statistically significant ( P ≤ 0.05) on Days 16 and 28 ( Fig. 6 ). The mean percent recovery of 15 N applied, as N 2 O‐N, equated to 0.86 (0.43), 0.88 (0.84), and 0.23% (0.10) (standard deviation in brackets), with no statistical difference between these values ( P = 0.15).

(a) Geometric mean nitrous oxide emissions from different treatments following urine application (error bars ± one geometric standard error of the mean, n = 5); (b) Nontransformed cumulative loss of N 2 O from urine‐treated soils, showing the amount of N emitted as N 2 O‐N as a percent of the total N applied to the plots. Nil urine is also plotted to show control emissions. (Error bars ± one standard error of the mean, n = 5).

Fluxes of N 2 O were significantly higher ( P ≤ 0.05) than in the control when urine was present, in all or some of the biochar‐urine treatments, from Days 1 to 35, and Days 44, 47, and 54 ( Fig. 5a ). Between Day 4 to approximately Day 35, when comparing only the biochar‐urine treatments, there was a trend for N 2 O fluxes to decrease with increasing rates of biochar, with N 2 O fluxes from the 30U treatment being statistically lower ( P ≤ 0.05) than from the 0U treatment during this period on Days 4, 7, 13, 24, and 26 ( Fig. 5a ). The cumulative N 2 O fluxes were higher under urine deposition when compared with the control. When the biochar‐urine treatments were compared against each other, the N 2 O fluxes from the 30U treatment were lower ( P < 0.05) than in either the 0U or 15U treatments ( Fig. 5b ). When the mean cumulative N 2 O‐N fluxes were expressed as a percent of urine‐N applied, the 0U, 15U, and 30U treatments had respective emissions of 0.15, 0.16, and 0.07%, with statistically lower emissions from the 30U treatment ( P < 0.05). When expressed as an emission factor (N 2 O‐N from the biochar‐urine treatment in question, minus the N 2 O‐N from the control, divided by the urine‐N applied), the 0U, 15U, and 30U treatments had mean emission factors of 0.12, 0.13, and 0.04%, respectively.

The atom% 15 N enrichment of the herbage in the control was at natural abundance, whereas in the biochar‐urine treatments the atom% 15 N enrichment was significantly higher ( P < 0.01), ranging from 3.556 to 3.991 ( Table 3 ). Recovery of applied 15 N in the herbage did not vary due to the addition of biochar at any time with total 15 N recovery in the herbage after 58 d, ranging from 14.3 to 17.5% ( Table 3 ).

Nitrogen uptake by the herbage, a function of N percent and dry matter yield, was higher ( P ≤ 0.05), on all occasions, when urine was present, ranging from 1.3 to 9.0 g m −2 ( Table 3 ). The addition of biochar to the soil had no effect on N uptake in the presence of urine ( Table 3 ).

At Day 21, only the herbage in the 0U and 15U treatments had a N level higher than in the control ( P < 0.05); but by Days 43 and 58, all biochar‐urine‐treated herbage had higher ( P < 0.01) N levels than in the control ( Table 3 ). Comparing only the biochar‐urine treatments, there were no significant differences in dry matter N percent as biochar rate was increased, at any time, although the trend was for lower N contents with increasing biochar rate on Days 21 and 43 ( Table 3 ).

No statistical differences in dry matter yield, due to treatment, occurred on Day 21, although the trend was for higher dry matter yield when urine was present ( Table 3 ). By Day 43, dry matter yields were higher ( P < 0.01) under biochar‐urine treatments than in the control; but by Day 58, only the 15U treatment had a higher ( P < 0.05) dry matter yield than in the control ( Table 3 ). When comparing just the biochar‐urine treated plots, increasing the biochar rate had no significant effect on dry matter yields ( Table 3 ). There was insufficient growth for harvesting of dry matter at Day 86 when the final N 2 O flux measurements were made.

The inorganic‐N concentrations increased significantly ( P < 0.01) with urine addition and NH 4 + ‐N reached maximum mean concentrations of 180 to 234 μg g −1 dry soil between Days 4 to 7 ( Fig. 4 ). There was a trend for soil NH 4 + ‐N concentrations to be higher with increasing rates of biochar‐urine after Day 21, but this was not statistically significant ( P ≥ 0.09; Fig. 4 ). By Day 65, soil NH 4 + ‐N concentrations were still elevated in the 30U treatment when compared with the 0U treatment ( P < 0.05; Fig. 4 ). Mean soil NO 3 − ‐N concentrations were significantly higher ( P < 0.05) under biochar‐urine treatments than in the control, from Day 9 onward, peaking at 72 μg g −1 dry soil in the 0U treatment on Day 26. Between Days 11 to 44, there was a trend for soil NO 3 − ‐N concentrations to be lower with increasing biochar rate and this was statistically significant on Days 11, 18, and 26 ( Fig. 4 ).

Surface soil pH became elevated following urine application and remained higher ( P < 0.01) in plus‐urine treatments, when compared with the control, until Day 86 ( Fig. 3 ). When comparing only the biochar‐urine treatments, there was a trend for the soil surface pH to be higher with increasing biochar rate when sampled between Days 7 to 47, but statistically significant differences ( P ≤ 0.05) only occurred on Day 16 and between Days 38 to 47, when the 30U treatment had a surface soil pH higher than in the 0U treatment ( Fig. 3 ). Soil bulk densities did not change significantly, despite the addition of biochar, with 0U, 15U, and 30U treatments having bulk densities (± standard deviation) of 1.29 (0.12), 1.29 (0.12), and 1.29 (0.10) Mg m −3 , respectively.

Soil moisture (θ g ) did not differ significantly due to biochar or urine treatments for any given sampling day, averaging 18.8, 19.3, 17.9, and 16.7% for the control, 0U, 15U, and 30U treatments, respectively, with maximum and minimum mean values of 26.7 and 6.5%, respectively, corresponding to WFPS values of 67 and 16% ( Fig. 2 ). As the summer season progressed, θ g declined over time, due to evapotranspiration exceeding sporadic and infrequent rainfall, but θ g increased following irrigation or substantial rainfall events. The average daily soil temperature (0.10 m depth) ranged from 13.2 to 25.6°C, following trends in the average daily air temperature, which ranged from 8.7 to 23.6°C ( Fig. 2 ).

Discussion

Increases in soil surface pH, and the duration of the pH increase, were typical of what is expected following ruminant urine deposition onto pasture (Jarvis and Pain, 1990). This increase occurs due to urea hydrolysis, whereas the subsequent decline in pH is a result of H+ ions being released during ammonia (NH 3 ) volatilization (Sherlock and Goh, 1984) and nitrification (Wrage et al., 2001).

Elevation of soil NH 4 +‐N concentrations resulted from the hydrolysis of urine‐derived urea. The small increase in soil NH 4 +‐N concentrations in the control treatment (water only) was possibly due to mineralization of organic matter. Soil NH 4 + is in chemical equilibrium with aqueous NH 3 in the soil and significant volatilization of NH 3 may occur when the soil pH is elevated (>7.0), as occurs under urine patches. Loss of NH 3 was not measured in the current experiment and we can only speculate the NH 3 loss amount. For urine patches in grazed pastures, NH 3 loss is commonly thought to be 10 to 20% of urine‐N deposited (Sherlock et al., 2008). The remaining soil NH 4 + pool can be taken up by pasture plants, become immobilized by soil microbes, or be oxidized further to NO 3 −‐N. The latter process explains the observed increase in soil NO 3 −‐N concentrations under the biochar‐urine treatments.

In this current study, biochar addition clearly influenced soil inorganic‐N dynamics, with lower NO 3 −‐N concentrations at the highest rate of biochar (30U), when compared with the 0U treatment, and trends for higher NH 4 +‐N under the 30U treatment. It is well recognized that biochar materials are able to promote adsorption of NH 3 (Clough and Condron, 2010; and references therein). Thus, biochar in the soil under a urine patch potentially creates a sink for the NH 3 We propose that one possible mechanism for the reduced NO 3 − concentrations and lower N 2 O emissions observed under the 30U treatment was the uptake and adsorption of NH 3 by the biochar. Adsorption of urinary‐derived NH 3 by the biochar would have increased with increasing biochar rate. This would serve to reduce the soil NH 4 +‐N pool available to nitrifiers and the NO 3 −‐N pool subsequently formed. If such adsorbed NH 3 is extractable with 2M KCl, it would explain the observed trend for higher NH 4 +‐N concentrations at the highest biochar rate (30U).

A further factor demonstrating that biochar altered the soil inorganic‐N pool was the reduced 15N enrichment of the N 2 O flux in the 30U treatment, indicating that the source of the inorganic‐N that the N 2 O was derived from came from an inorganic‐N pool with a lower proportion of urine‐N than in the 0U treatment.

Thus, we propose that under the highest rate of biochar, NH 3 formation and its subsequent adsorption onto and/or into the biochar reduced the inorganic‐N pool available for nitrifiers and thus NO 3 −‐N concentrations were reduced. Then, since the NO 3 −‐N pool had a lower concentration, the 14N dilution arising from soil mineralization was relatively greater, thus lowering the 15N enrichment of the N 2 O source pool(s). Consistent with this is the lower N 2 O flux from the 30U treatment, as a percent of urine‐N applied, and as a cumulative N 2 O flux. Soil N 2 fluxes were not measured in this study and the relatively alkaline nature of biochar, when compared with soil, may possibly have favored further reduction of N 2 O to N 2 This could explain a lower N 2 O flux but not the differences in 15Nenrichment observed.

An alternative theory to explain lower soil NO 3 −‐N concentrations in the presence of biochar was drawn by Singh et al. (2010), following the incorporation of either poultry‐ or woodchip‐derived biochar to soil columns and a 5‐mo incubation with three wetting–drying cycle differences in NH 4 +‐N and NO 3 −‐N leaching being observed. Singh et al. (2010) concluded that these differences could have been due to increases in the sorptive properties of the biochars. Other studies have demonstrated increases in soil cation exchange capacity (CEC) over longer periods (600–8700 yr) due to the oxidation of biochar surfaces or the adsorption of organic matter to the biochar particles (Liang et al., 2006). It seems unlikely that after only 14 mo such mechanism would have occurred, but these cannot be ruled out and future studies need to examine short‐term changes in CEC.

Chemicals that inhibit nitrification lead to a prolonged occurrence of NH 4 +‐N in the soil and lower NO 3 −‐N concentrations—a trend observed in the 30U treatment in the current study. It has been previously noted that biochar contains volatile organic compounds (VOCs) that are known nitrification inhibitors. For example, Clough et al. (2010) found α‐pinene in an unweathered biochar. In the current study, ethanol was the only VOC detected in the biochar before its incorporation into the soil, which was several months before urine deposition. Thus, it is assumed that the effect of ethanol, if any, would have been negligible by the time urine was applied. Other nonvolatile microbially inhibiting compounds may have existed in the biochar. Spokas et al. (2010) hypothesized that ethylene, a known microbial inhibitor, was microbially produced from biochars and that ethylene may be the cause of the observed changes in microbial dynamics and N 2 O suppression. This interesting theory needs testing with respect to longevity of ethylene production in the soil. The biochar in the current study had been in the soil for approximately 7 mo (May–November) before urine treatment.

The addition of fire‐derived charcoal to forest soils has been shown to enhance native soil organic matter mineralization (Wardle et al., 2008a), albeit in forest soils, highlighting the current lack in our understanding of how biochar might affect native soil C pools (Wardle et al., 2008b). At the high rate of biochar used here, the dilution of the 15N pool, supplying the N 2 O flux, could potentially have occurred as a result of enhanced mineralization of soil organic matter. But had this been the case, we might have expected to observe an increase in the size of the inorganic‐N pool under the 30U treatment and this was not observed.

The soil microbial‐15N pool was not measured in the current study and this should be included in future studies to further elucidate the mechanisms of biochar perturbance of the N cycle.

Soil moisture conditions were consistent with summer soil conditions, and given that denitrification is expected to dominate at WFPS values >60%, the soil moisture conditions (WFPS range of 16–67%) predominantly favored nitrification mechanisms as N 2 O‐forming pathways. However, denitrification and nitrifier‐denitrification may still operate at anaerobic microsites under aerobic soil conditions (Müller et al., 2004; Russow et al., 2009). During Days 15 to 35, when the N 2 O‐15N enrichment was generally less in the 30U treatment, there was also considerable rainfall or irrigation, and the highest N 2 O fluxes occurred. Denitrification of NO 3 −‐N as an N 2 O production mechanism is definitely plausible under these conditions. As a percent of the urine‐N applied, the cumulative N 2 O fluxes were low when compared with the New Zealand specific N 2 O emission factor for urine‐N, which is currently set at 1.0% of urine‐N excreted (Kelliher et al., 2005) and with other studies as noted below. This was most likely a function of the drier summer soil conditions, despite the irrigation, and this study needs to be repeated under winter conditions to see if similar reductions in cumulative N 2 O emissions occur under the 30U treatment.

The fact that dry matter yields were not detrimentally affected by biochar addition indicates that there are no apparent negative effects of biochar incorporation under the conditions of this trial. The lower percent N of the dry matter harvested from the 30U treatment on Day 21 is consistent with what was seen in the inorganic‐N pool at this time, with less inorganic‐N equaling less N uptake at this time. Had this been under a grazing regime, this would have produced a positive feedback since lower dry matter N would have resulted in lower dietary N intake and subsequent urine‐N excretion. Lower rates of urine‐N excretion would reduce subsequent derived N 2 O emissions.

Surprisingly, the addition of biochar did not translate into statistically different soil bulk densities. At the highest rate of biochar (30 t ha−1), the area of the soil‐sampling core, used to determine bulk density, would have received 12.6 g of biochar, which at its measured bulk density of 0.4 Mg m−3 equates to 3.13 × 10−5 m3 of biochar. Assuming all this was equally distributed within the target depth of 0.10 m, the soil bulk density core, with a depth of 0.075 m, could have contained 2.35 × 10−5 m3 of biochar. A theoretical bulk density under such ideal mixing, and using the nil biochar soil as a reference, would equal 1.23 Mg m−3, which is within one standard deviation of the bulk density determined at 30 t ha−1 Thus, further replication of the bulk density sampling, or changes in the method, are required if changes in bulk density at biochar rates up to 30 t ha−1 are to be determined. It should be noted, however, that even a change in soil bulk density from 1.30 to 1.23 Mg m−3 is sufficient to potentially affect soil processes.