Introduction

In terrestrial ecosystems, nitrogen (N) deficiency is the main growth limiting factor, thereby adversely affecting crop productivity1,2,3,4. Therefore, an exogenous supply of N along with phosphorus as fertilizers is key farming practice for improving crop yield3. Conferring to5, a huge quantity of chemically synthetized N compounds will be required to fulfill the food requirements of 9 billion people by 2050. However, the higher land application of N compounds as chemical fertilizers has a negative influence on the environment through the release of N containing gasses such as NH3 and N2O, and its losses to the ground and surface water by nitrate leaching and runoff6,7,8. At current, the N use efficiency of synthetic N fertilizers is too low, only amounting approximately 50%9,10, which adds to the adverse environmental, agronomic and economic impact11,12.

Ammonia volatilization is a major source for N losses and is considered to be the main cause of low N use efficiency9,13. Ammonia is a major alkaline atmospheric pollutant that plays an important role in the formation of aerosols14,15, which badly affect human health16, reduce visibility17,18, alter Earth’s radiative balance, and contribute to a global redistribution of N through atmospheric deposition19,20. Although ammonia is not considered a potential greenhouse gas (GHS), its emission and re-deposition can adversely affect the environment21, and it may act as a secondary cause of N2O emission in soil22,23.

Nitrous oxide is a potential (150 years lifetime) GHS, which is 298 times more efficient than CO2 for its heat-trapping capacity with a 7% contribution to the total GHS emission and 0.26% annual growth rate24. Agricultural soils annually emit 4.1 Tg N to the global atmospheric N2O emission of 14 Tg N24,25. In agricultural soils, N2O is emitted mainly by biological and chemical processes such as nitrification and denitrification26,27,28. Additionally, N2O emissions may also happen via nitrification–denitrification through autotrophic NH3-oxidizing bacteria, where ammonia is oxidized to NO2−, followed by its reduction into NO, N2O and N226,29,30. It can also be produced during a hybrid reaction by co-denitrification where two N atoms are released one each from organic N by mineralization and NO2− by denitrification31. In soil containing high organic matter, it can be released by heterotopic reduction of nitrite by the oxidation of organic N32,33.

Various studies have been aiming to develop sustainable and eco-friendly management practices for reducing N losses, improving NUE and crop yield34,35. Such practices include application of nitrification inhibitors, urease inhibitor (UI), elemental S and polymers36, removal of crop residues from the land in various ratios37,38 and application of biochar39,40. Biochar, a carbon-rich pyrolitic product of organic waste, has received ample attention for its environmental benefits41. Utilization of biochar as a soil amendment/conditioner not only provides a way to recycle environmental waste but also optimizes soil health, crop yield and stimulates soil C sequestration42,43,44,45,46,47,48,49,50. Studies have demonstrated that biochar may be used as a compound fertilizer in conjunction with mineral sources51,52, as a slow-release fertilizer53,54,55, which could enhance fertilizer56 and may increase N use efficiency under different soil conditions51,52. Biochar may have the capacity to sorb NH4+ or NH3 gas released during composting, thereby lowering NH3 emission. Biochar may also adsorb organic N compounds, thus decreasing their mineralisation57 and consequently NH3 emission. This gradual availability of N may be partially explained by lower atmospheric N2O emissions from N fertilizer applied in the presence of biochar58,59.

The application of UI [N-(n-butyl)] and thiophosphoric triamide (nBTPT or NBPT for short) have been reported to reduce urease activity in soil and slow down NH4+ release from urea, thus reducing NH3 emission34,35,60. Urease inhibitor significantly decreases urea hydrolysis, reduced NH4+ concentrations and, thereby limited NO3− supply and nitrification rate thus reducing N2O emission and improves crop yield61,62,63,64.

So far, many experiments have focused the effect of biochar and urease inhibitor sole application on gaseous N emissions, N-transformation processes and crop yield in variable agro-climatic circumstances. However, limited work is documented regarding the influence of urea treated with urease inhibitor and biochar or alone on N2O and NH3 emission from arable agricultural system under hot climate of Pakistan. Therefore, main objectives of current study were to explore the effect of urea together with urease inhibitor and biochar or alone on crop productivity, N2O and NH3 emission and N efficiency.

Results

Soil mineral N dynamics

The NH4+-N concentration observed in urea treated plots on day 1 was significantly (p < 0.05) higher (5–25 mg N kg−1 soil) compared to the control (Fig. 1). In urea treatment, the NH4+-N concentration increased because of the applied urea's fast hydrolysis during the first 3 days after first fertilization. Thereafter, a sharp decrease was observed (Fig. 1). The plots treated with biochar + urea showed maximum soil NH4+-N concentration 28 days after fertilization, which was significantly higher than the plots treated with sole urea (Fig. 1). Furthermore, the application of UI considerably decreased the average soil NO3−-N content. After the first fertilization, soil NO3–-N content slightly increased, and most mineral N was observed as NH4+-N. Soil NO3–-N concentration peaked just after the second urea application in the urea treatment and then displayed a rapidly decreasing trend within 14 days in after fertilizer application (Fig. 1). Additionally, biochar supplementation significantly decreased NO3–-N concentration in the soil.

Figure 1
figure 1

Soil NH4+ and NO3− contents as affected by nitrogen application with and without biochar and/or urease inhibitor. Bars on means represent SE for n = 4. The arrows show time of N application.

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Ammonia volatilization

The results concerning NH3 emission are presented in Fig. 2 and Table 1. A maximum NH3 discharge was detected between days 1 and 3 after every application. The application of BC and UI with urea significantly affected daily and total cumulative NH3 discharge during the initial 10 to 12 days. Total NH3 losses as kg N ha−1 amounted 14.4 with BC and 6.2 with BC + UI compared to sole urea (19.7), representing 27% and 69% reduction by BC and BC + UI, respectively, relative to sole urea application.

Figure 2
figure 2

Soil NH3 Fluxes as influenced by nitrogen application with and without biochar and/or urease inhibitor. Bars on means represent SE for n = 4. The arrows show time of N application. Where, FA (First dose application) and SA (Second dose application).

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Table 1 Nitrogen losses as ammonia volatilization as affected by nitrogen application with and without biochar and/or urease inhibitor.
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Nitrous oxide emissions

The application of urea, BC, and UI significantly affected the daily and cumulative N2O emissions (Fig. 3 and Table 2). The highest and lowest cumulative emissions of N2O were measured for urea and urea + BC + UI treatments, respectively (1.7 and 0.8 kg N2O-N ha−1) (Table 2). Biochar + urea significantly reduced N2O emissions by 24% compared to urea alone. Urea applied in combination with BC and UI reduced N2O emission by 53% over urea alone treatment.

Figure 3
figure 3

Soil N2O flux in response to nitrogen application with and without biochar and/or urease inhibitor. Bars on means represent SE for n = 4. The arrows show time of N application.

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Table 2 Nitrogen loses as nitrous oxide in response to urea supplementation with and without biochar and/or urease inhibitor.
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N-use efficiency, yield and N uptake by wheat

Wheat N uptake, yield and NUE were significantly improved due to biochar and/or UI application (Table 3). Wheat biomass (24%) and grain yields (13%) were improved by BC over sole urea. In contrast, urea's combined application with BC and UI increased wheat biomass and grain yields by 38% and 22% over the sole urea treatment. The highest total N uptake, above ground wheat biomass, contained 101 and 114 kg N ha−1 from urea + BC and urea + BC + UI treated plots, respectively (Table 3). The BC and UI application increased total N uptake by 12% and 27%, respectively, over sole urea (Table 3). Similarly, the highest N uptake and NUE were observed in plots treated with urea + BC + UI, followed by urea + BC (Table 3). The NUE observed for sole urea, urea + BC and urea + BC + UI were 27%, 35%, and 43%, respectively (Table 3).

Table 3 Wheat biomass and grain yields (kg ha−1), N uptake (kg ha−1) and NUE (%) as affected by urea fertilization with and without biochar and/or urease inhibitor.
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Discussion

Ammonia volatilization as affected by biochar and urease inhibitor

A 27% decrease was observed in NH3 volatilization with BC compared to sole urea (Table 1), which can be attributed to its highly porous structure, surface area and high sorption capacities60,61. Furthermore, BC may absorb NH4+-N62 due to the presence of acidic functional groups63, hence decreasing the NH3 volatilization64. Biochar application with N fertilizers could avoid soil alkalization65,66 and decrease organic N mineralization through the adsorption of organic N compounds32,57.

On day 1 after application, a rapid increase in NH4+-N content was observed with sole application of urea (Fig. 1), which could be ascribed to the fast hydrolysis of urea. As a result, more NH4+-N and OH− ions are produced and allowing significant NH3 losses (Fig. 1). Urease inhibitor reduce the effect of pH increasing by delaying urea hydrolysis, as demonstrated by lower NH4+-N (Fig. 1) which could significantly reduce NH3 emissions. Slower urea hydrolysis due to UI application could also be associated with increasing pH around urea particles, which prevents hydrolytic action on urea via the enzyme urease10,35,67. It has also been reported that UI slows down the microbial decomposition of ammonia68,69,70,71. Additionally, slower urea hydrolysis due to application of UI could also provide more time for rainfall or irrigation to move the applied urea from the soil surface to the sub-soil layers vertically as well as and laterally consequently protects the applied N from volatilization72,73.

N2O emission as affected by biochar and urease inhibitor

Nitrous oxide emissions are directly related to the amount of mineral N available in the soil. A two-way ANOVA indicated that seasonal N2O emissions during the wheat growing season were significantly affected by biochar application (Fig. 2), in agreement with previous results Zhang et al.33 and Schirrmann et al.74. Our results confirm that the application of biochar reduced N2O emissions, because biochar resulted in lower NO3− contents and higher NH4+ concentrations after N addition (Fig. 2) which affect N availability in the soil, either physically by sorption to surfaces or microbiologically75. These results are in agreement with the findings of Lehmann et al.76 and Kammann et al.77 who reported that after application of N fertilizer, the adsorption of soil NH4+ by biochar on its surface, especially by biochar with a maximum C/N ratio reduce N2O emissions, subsequently resulting a decrease in the processes of ammonification and nitrification.

Nitrous oxide is generated by both nitrification and denitrification processes; therefore, it may be closely related to soil NH4+-N and NO3–-N concentrations78. Under high nitrification activity, when a lot of NO3− is produced in urea-treated soils during the first week of treatment application (Fig. 1) the observed N2O may actually be related to nitrification which produces N2O as a byproduct10. Apart from high water content also a high microbial activity can cause O2 depletion and therefore anoxic conditions that might promote denitrification. Due to the strong nitrification usually taking place during aerobic conditions, NO3− predominates in soils. As a result, denitrification would be prompted by the large amounts of NO3− accumulated in soil, especially under high soil moisture conditions. For a given soil, whether nitrification or denitrification contributed more to N2O emission may be closely related to soil NH4+-N and NO3−-N concentrations when the initial physicochemical factors of the tested soil are almost identical79.

The results showed lower NH4+-N concentrations in plots treated with urea + BC + UI than sole urea treated plots, while NO3-N concentrations were higher in urea treated plots than in urea + BC + UI (Fig. 1 and Fig. 3). It is likely that the combination of biochar and UI significantly decreased NO3− supply and thus reduced the activity of denitrifiers. Here, it is evident that UI application reduced the activity of ammonia oxidase and nitrification process and played a significant role in converting NH4+-N to NO3–-N, which is significantly related to N2O discharge from soil10,36,67. These findings imply that applying either BC or BC + UI plays a significant role in reducing NH3 and N2O emissions from urea fertilized soils. Additional investigations are recommended to elucidate the role of BC and BC + UI in reducing NH3 and N2O emissions at varying soil and climatic conditions.

Wheat yield and N uptake as affected by biochar and urease inhibitor

Both BC and UI significantly improved total N uptake and yield of wheat (Table 3). Application of BC and/or UI with N fertilizers can reduce N losses as GHS and may prevent NO3− leaching, thus enhancing the bioavailability of N36,48,60. Dawar et al.13 reported that the retention of mineral N in the form of NH4+, rather than NO3−, for several days after urea application increasing N uptake and thus improved crop yield. The results also show that BC improved soil WHC, allowing the wheat to retain a proper moisture level between irrigation periods, with a subsequent positive impact on final grain yields80. Furthermore, BC + UI could significantly improve soil conditions such as soil organic content, pH, and total N content81, improving N availability by reducing NH3 and N2O from the soil. Urease inhibitor and BC retain NH4+ in the soil for a longer time and improve its subsequent uptake by wheat, thus may improve crop yield. This not only offers environmental benefits by preventing and NO3− leaching and sinking NH3 and N2O release13,36, but also includes economic benefits using an increase in NUE. The plant uses relatively less energy to absorb NH4+ than NO3− as the transformation of NO3− into NH4+ via amides, amines, and amino acids is energy-consuming. Thus, higher availability of NH4+ facilitates improved crop yield and nutrition10.

Conclusions

It was observed that BC and UI have the highest potential to reduced NH3 and N2O emissions in urea fertilized soils. Furthermore, applying BC and/or UI with urea significantly improved wheat biomass and grain yield compared to sole urea. Therefore, the application of BC and/or UI with urea plays a significant role in mitigating NH3 and N2O emissions from cultivable land and improving crop yield under the hot and semi-arid agro-climatic conditions of Pakistan.

Materials and methods

Location

This experiment was executed at the National Institute of Food and Agriculture (NIFA), Peshawar (34.01°N, 71.71°E), Pakistan. The study area is semi-arid to arid and humid in the north to dry in the southern parts with a mean yearly precipitation of 384 mm and temperature of 22.7 °C (Fig. 4). The field had been conventionally cultivated with maize-wheat cropping system for approximately 10 years. The main soil (0–15 cm) properties of the composite sample are provided in Table 4.

Figure 4
figure 4

Soil (0–10 cm) temperature, moisture and precipitation throughout the growing season.

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Table 4 Characterization of experimental field.
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Experiments

The field experiment including five treatments [control; urea (150 kg N ha−1); BC (10 t ha−1); urea + BC and urea + BC + UI] was designed following a randomized complete block design (RCBD) with 4 replications. The ‘Pirsabak-2013’ variety of wheat was sown in 5 m × 3 m plots containing 10 rows with row-to-row distance was kept 30 cm apart at a seed ratio of 120 kg ha−1 on 15th Nov, 2017. The conventional tillage was implemented using a mouldboard plow (40–50 cm deep) followed by disking (20–25 cm deep) and land leveller operations for seed bed preparation. Before seeding, 90 kg P ha−1 and 60 kg K2O ha−1 as single super phosphate and sulphate of potassium, respectively, were applied as a basal dose. The biochar used was produced from eucalyptus wastes (branches) pyrolyzed at 350 °C and sieved through a 5 mm sieve. Nitrogen was supplemented in the form of urea as a top dressing in two splits, half each a sowing and tillering. Urease inhibitor was supplemented as 0.1% solution. Granular urea with UI containing 25% NBPT was added at 150 kg N ha–1 as a surface application followed by surface incorporation and irrigation (10 mm) to ensure its proper distribution in the root zone.

N2O measurement

The N2O discharge from soil was measured from November, 2017 to May, 2018 using opaque manual spherical static chambers, following the procedure described in Sanz-Cobena et al.34. For sample collection, the cylindrical (radius 12.5 and height 20 cm) chambers were inserted in the soil at 15 cm depth 24 h before sampling. The chambers were properly sealed, and samples were taken after 0, 30 and 60 min once a week in the morning between 7:00 and 10:00 by a butyl septum installed on the upper part of the chamber. The samples were injected in 20 ml glass vials (Agilent Technologies, USA) via syringes (50 ml) equipped with inner three-way stopcocks (0.7-mm). The air and soil temperatures inside the chambers were recorded during gas sampling via a portable digital thermometer. Gas samples were analyzed on gas chromatographs (Varian Aerograph Series 2800 in NZ; Perkin Elmer Auto system XL B5902) equipped with 63Ni electron capture detectors (Pye Unicam) and two manual switching valves (Valco Instruments Co., Inc.)

The mean variation in gas concentration was calculated through linear regression while the ideal gas law was applied for quantification of gas-fluxes as follows:

$${\text{F}} = \uprho \times \left( {{\text{P}} / {76}0} \right) \times \left( {{\text{V}} / {\text{A}}} \right) \times \left( {\Delta {\text{C}} / \Delta {\text{t}}} \right) \times \left[ {{273} / \left( {{273} + {\text{T}}} \right)} \right]$$
(1)

where F, P, V, A, Δc/Δt and T represent N2O flux (µg m−2 h−1), density (mg m−3), chamber size (m3), base size (m2) of the chamber, mean rate of variation in gas concentration per unit time (mg kg−1 h−1) and chamber inner temperature (°C), respectively. The acceptable range of R2 for N2O fluxes was taken as 0.80 for the static chamber, except where the maximum change in concentration was lower than the gas-specific GC detection limit (< 10 ppb for N2O), where no filtering criterion was adopted (Järveoja et al.82). Using these criteria, 10% of fluxes (N2O) were subtracted from succeeding data analysis.

The following formula was used for the determination of cumulative N2O emissions (Ec, kg N2O-N ha−1):

$$\sum_{i=1}^{n}\left({F}_{i}+{F}_{i+1}\right) /2\times ({t}_{i+1}-{t}_{i})\times 24$$

where Fi and Fi + 1 is the ith and (i + 1)th measured value, respectively, of N2O flux (µg N2O-N m−2 h−1 ); ti and ti + 1 is the day when the ith and (i + 1)th measurement of N2O flux is taken, respectively (d); and n is the total number of the measurements.

Ammonia measurement and analysis

Ammonia volatilization was quantified by a 5 L (plastic bottle) semi-static open chamber, according to Araujo et al.83. In each plot, a single chamber was installed 5 mm above the ground surface and was randomized daily as advised by Jantalia et al.84. A foam strip pre-soaked in 1 molar H2SO4 and 4% (V/V) glycerol was kept moist throughout sampling duration by immersing one end into a polypropylene jar containing 15 mL 1 molar H2SO4 solution suspended inside the chamber. In each plastic bottle, the solution and foam strip were initially replaced by fresh solution after 12 h and then every 24 h and 14 days and processed for NH4+-N concentration via the steam distillation method85.

The following equation was applied for the quantification of NH3 fluxes (kg N ha−1 d−1):

$$\mathrm{F}=\frac{2 \times \mathrm{C}\times \mathrm{V}\times 14\times {10}^{-2}}{\uppi \times {\mathrm{r}}^{2}}\times \frac{24 }{\mathrm{t}}$$

where; C, V, t and r represent the molar concentration of H2SO4, amount of H2SO4 consumed during titration (ml), sampling time (hours) and chamber radius (m). The cumulative NH3 flux was determined by adding the NH3 fluxes for all sampling days for their respective treatment plots.

This method showed 57% NH3 recovery by calibration with 15N isotope equilibrium technique (Araújo et al.83). Therefore, for the accurate estimation of cumulative NH3 emissions and flows, a correction factor of 1.74 was applied (Araújo et al.86). According to Jantalia et al.84, this method for quantification of NH3 emissions is more suitable than the wind tunnel procedure35 for comparison of different treatments.

Soil and plant analysis

From each plot, five soil samples (0–10 cm) were randomly collected, well mixed, sieved (2 mm) and analyzed for key soil properties (Table 4). To determine mineral N (NO3− and NH4+), extraction was done with 1:5, 2 M KCl for 1 h on a rotary shaker10, filtered and analyzed via ultraviolet spectrophotometry (Jenway, 6305 UV/Vise, UK). Soil moisture was measured by oven-drying and was transformed to water-filled pore space (WFPS) using the formula of Li et al.85.

Total N in soil and plant samples was determined by the Kjekdhal method of Keeney and Nelson86. In this method, 0.2 g of finely ground samples of dry materials were digested with 3 ml of concentrated H2SO4 in the presence of 1.1 g digestion mixture containing CuSO4, K2SO4 and Se on a heating mantle for about 1 h. The digest was transferred quantitatively to the distillation flask and distilled in the presence of 10 ml of 10 MNaOH solutions. The distillate was collected in 5 ml boric acid mixed indicator solution and then titrated against 0.01 MHCl solution by adding 5 ml boric acid mix Indicator. Using the follow formula total N was calculated.

$$\text{Total} \; \text{Nitrogen} \; \%=\frac{({\text{Sample}}-\text{Blank}) \times 0.005 \times 0.014 \times 100 } {{\text{Weight}} \; {\text{of}} \; {\text{soil}} \times {\text{volume}} \; {\text{made}}}$$

For obtaining the data on grain and biological yield, the central four rows were harvested from each plot at physiological maturity and data were recorded on various agronomical traits (biomass, grain yield and straw yield) and total N uptake in crop. Biomass yield was separated into grain and above ground plant tissue (i.e. shoot and leaves) and record their fresh bulk weight immediately. Five randomly chosen plant tissue sub-samples (ca. 1000 g fresh weight) from each sub-plot; were transferred to sealable plastic bags, and transferred to lab in container with ice to ensure no water losses occur from collected plant tissue. After transporting the plant tissue samples to the lab, fresh weight was immediately recorded. After recording the fresh weight, harvested material was placed in pre-weighed paper bags and dried at 65 °C for 7 days. Dry weights of the plant tissue after 7 days were recorded in order to calculate its moisture content or fraction. The grain yield was adjusted for moisture fraction, prior to obtaining its dry weight, using a moisture tester. For N uptake by above ground plant tissues (i.e. shoot and leaves) and by the grain, the two tissues separately samples were ground to a fine powder (for determination of the total N.

Grains yield was recorded after threshing of wheat plants taken from central four rows of each treatment and then converted into kg ha−1 by using the following formula.

$${\text{Grain yield}}\,{\text{(kg}}\,{\text{ha}}^{{ - 1}} {)} = \frac{{\text{Grain yield in four central rows}}}{{{\text{Row}}\, - \,{\text{row }}\,{\text{distance}}\,\times\,{\text{Row}}\,{\text{length}}\,\times\,{\text{No}}{.}\,{\text{of}}\,{\text{rows}}}} \times {{10000}}$$

Biological yield was recorded by harvesting 4 central rows in each plot, dried and weighed and then weight was converted into kg ha−1 using the following formula;

$${\text{Biological}}\,{\text{yield}}\,{\text{(kg}}\,{\text{ha}}^{{ - 1}} {)} = \frac{{\text{Biological yield in four central rows}}}{{{\text{Row}}\, - \,{\text{row }}\,{\text{distance}}\, \times\,{\text{Row}}\,{\text{length}}\,\times\,{\text{No}}{.}\,{\text{of}}\,{\text{rows}}}} \times {{10000}}$$

Nitrogen uptake and NUE were then calculated as follows:

$$\text{Total} \; \text{ N} \; \text{uptake}=\frac{{\% \text{N} \; \text{in} \; \text{grains} }\times \text{grain} \; \text{yield } \; (\text{kg } \; {\text{ha}}{^{-1}})}{100}$$
$$\text{NUE} \; (\%) = \frac{\text{Total} \; \text{N}.\text{ uptake }\left(\text{kg} \; \text{ha} {^{{-1}}}\right) \; \text{ in} \; \text{fertilized} \; \text{plot }-\text{ Total} \; \text{N}.\text{ uptake }\left(\text{kg} \; \text{ha} {^{-1}}\right)\; \text{ in} \; \text{control} \; \text{plot }}{\mathrm{N}.\mathrm{ applied } \; \left(\text{kg} \; \text{ha} {^{-1}}\right)}\times 100$$

Statistical analysis

Standard error (SE) of mean (n = 4) was quantified via descriptive statistics. The replicated data were processed using analysis of variance (ANOVA) followed by least significant difference (LSD) test (P < 0.05) using general linear model (GLM)87.

Plant material collection and use permission

No permission is required for plant material as it was purchased from certified dealer of local area.

Ethics approval and consent to participate

We all declare that manuscripts reporting studies do not involve any human participants, human data, or human tissue. So, it is not applicable.

Complies with international, national and/or institutional guidelines

Experimental research and field studies on plants (either cultivated or wild), comply with relevant institutional, national, and international guidelines and legislation.