The study was carried to determine the labile carbon contents in biochar pyrolyzed under different temperatures and its effect on urea hydrolysis rate. Corn stovers were pyrolyzed at highest temperature of 300°C, 500°C and 700 °C to produce fresh biochar. Soil incubation experiment was conducted with biochar application rate of 0% and 2% with presence of urea. The results showed that biochars accelerated urea hydrolysis, and high temperature pyrolyzed biochar have more significant effect on soil pH enhances and acceleration of urea hydrolysis than biochar pyrolyzed at low temperature. Moreover, biochar produced at 300°C contains relative high concentration of labile carbon, and 43-64% of labile carbons were oxidized within 40 days incubation. The labile carbon in biochars also leaded microbes thrives and resulted in accelerate short-term N turnover. i.e. at early stage of incubation, fresh biochar increased mineralization of soil N by 79-449 mg•kg−1, and matured biochar by 30-61 mg•kg−1, but microbial immobilization effect was observed in fresh biochar-amended soil at the end of incubation. We concluded that aged biochar is suitable for simultaneous soil amendment with urea rather than newly produced biochar as it can promote available N accumulation in short time thus increase the risk of inorganic nitrogen leaching loss.
Maize Stover Biochar Accelerated Urea Hydrolysis and Short-term Nitrogen Turnover in Soil
Zunqi Liu,a,b Tianyi He,a,b Yu Lan,a,b Xu Yang,a,b Jun Meng,a,b,* and Wenfu Chen a
A soil incubation experiment was conducted to investigate the effect of biochar application (2% w/w) on urea hydrolysis and inorganic nitrogen accumulation. Fresh biochars were produced from maize stover that was pyrolyzed at 300 °C, 500 °C, and 700 °C. Then the matured biochars were obtained via a 50 days maturing process. Biochar prepared at 700 °C strongly accelerated the urea hydrolysis and increased soil pH. Fresh biochar, especially when pyrolyzed at low temperature, contained a relatively high concentration of labile carbon and 43% to 64% could be oxidized within 40 days of maturing incubation. The labile carbon in fresh biochars led to microbes thriving and resulted in an accelerated short-term nitrogen (N) turnover, i.e., at an early stage of incubation, fresh biochar increased mineralization of soil organic N by 79 mg·kg−1 to 449 mg·kg−1. However, a reduction of soil available N contents induced by microbial immobilization effect was observed at the end of incubation. The authors concluded that aged biochar was suitable for simultaneous soil amendment with urea rather than newly produced biochar. This is because aged biochar can avoid high soil available N accumulation; thus it can decrease the risk of inorganic N leaching loss.
Keywords: Biochar; Labile carbon; Urea hydrolysis; Available nitrogen
Contact information: a: Agronomy College, Shenyang Agricultural University, 110866, Shenyang, P.R. China; b: Liaoning Biochar Engineering & Technology Research Center, 110866, Shenyang, P.R. China; *Corresponding author: Meng Jun: email@example.com
Burning of crop residues is commonly practiced after harvest in China as well as in other developing countries (Meng et al. 2011; Li and Wang 2013). Although this practice can reduce the amount of crop residuals directly, it results in significant pollution to adjacent regions from smoke and ash, resulting in many cities being covered by smog and haze. Moreover, residue combustion is a source of greenhouse gas emission, associated with global warming. Pyrolyzing stover biochar might be one optimal way for solving this problem without side effects (Meng et al. 2011). Biochar refers to carbon-rich materials produced from the pyrolysis of biomass under limited oxygen and relatively low temperatures. As a soil amendment, biochar has attracted considerable attention due to its ability to improve soil fertility and increase soil carbon sequestration. A biochar carbon sequestration effect mainly benefits from its recalcitrant characteristic formed during pyrolysis procedures (Lehmann 2003) and is beneficial due to its potential to protect soil organic matter from degradation (Whitman et al. 2015). Moreover, it was reported that biochar provides nutrient elements to plants and improves soil quality directly. Specifically, biochar may be enriched with elements including K, Mg, and Ca (Steiner et al. 2007; Cantrell and Martin 2011), and it is capable of improving soil water retention, cation exchange capacity, and nutrient retention capacity (Lehmann et al. 2003; Liang et al. 2006; Laird et al. 2010).
It is generally accepted that the majority of biochar-C is chemically and biologically recalcitrant, but it still contains a minor fraction of bioavailable C (Major et al. 2010; Jones et al. 2011), and the concentration of labile carbon in biochar depends on the conditions during pyrolysis, as well as the composition of the feedstock biomass (Nelissen et al. 2012; Chen et al. 2016). There is also evidence that the labile carbon derived from biochar enhances soil microbial biomass growth and activity (Steiner et al. 2008). From this belief, there can be little doubt that biochar will play an important role on shifting microbial community structure and function involved in nitrogen dynamics.
Urea is an industrial product and the main form of nitrogen fertilizer used worldwide. The application of urea has been an important measurement for crop yield improvement and is a significant exogenous nitrogen source for soil. Thus, when biochar is amended into agricultural soil, it inevitably interacts with urea. Therefore, from an agronomic standpoint, the effect of biochar on the behaviours of urea in soil, as well as the combined effect of biochar and urea on soil’s native nitrogen turnover, should be determined. However, only few studies have investigated the interaction between biochar and urea. Anderson et al. (2014) found that the relative proportion of nitrifiers and denitrifiers increases in biochar-amended soils subjected to large influxes of urine-derived N. Results from N-15 isotopic analyses by Reverchon et al. (2014) showed that biochar amendment accelerates the N dynamics and increases N losses, probably through ammonia volatilization or denitrification. Unfortunately, those studies lack the information about how biochar affect urea hydrolysis and the relationship with its labile carbon.
Because the time of urea hydrolysis in soil is usually shorter than 2 weeks, the authors conducted a short-term incubation experiment. The main objectives of this study were to 1) assess the labile carbon contents in biochar pyrolysis under different temperature and its stability; and 2) investigate the urea hydrolysis rate and inorganic N accumulation that was affected by biochar with and without labile carbon. It is hypothesized that 1) biochar can affect urea hydrolysis rate depending on its sorption ability or on improvement of soil quality, and 2) biochar labile carbon plays an important role on shifting soil organic N turnover.
Maize stover biochar was used in this study. After being dried to a constant weight at 65 °C and milled to 1 cm3 to 2 cm3, the maize stover was mixed thoroughly and placed in crucibles covered with lids then pyrolyzed in a programmed muffle furnace (Beijing Zhongxing Laboratory Equipment Inc., Beijing, China) at a heating rate of 10 °C/min from room temperature to 300 °C/500 °C/700 °C and maintained for 30 min. The obtained biochar was marked as FBC300/500/700.
To produce matured biochar that was termed as MBC300/500/700, a 50-day incubation was conducted. The main aim of the maturing process was to remove the labile carbon from biochar. First, the microbial inoculums were mixed with each fresh biochar at a rate of 1:10 (w:w), and then incubated at 35 °C for 50 days. During the incubation process, the same volume of microbial inoculum was added when the mixtures were dried. Subsequently, biochar was air-dried for a future study. The microbial inoculums added were freshly obtained from the extracts of grassland soil, which exhibited high biological activity. Specifically, 100 g of topsoil (0 cm to 10 cm, incubated under 60% water holding capacity (WHC) at 30 °C for 2 d) was mixed with 200 mL of distilled water and shaken for 30 min at 200 rpm. After standing for 12 h the supernatant obtained was microbial inoculums.
Measurement of biochar properties
For elemental analyses, sieved biochar were milled to achieve a homogenous fine powder. C, N, and O contents were measured by using an elemental analyzer (Elementar Macro Cube, Langenselbold, Germany). The H content was calculated by the difference method (Yue et al. 2016). The biochar specific surface area and pore characteristic were determined by using a surface area and pore analyzer (V-sorb 4800, Jinaipu, China). The ash content and volatile matter were determined based on a method described by the IBI Biochar Standards Version 2.1. The biochar pH value was measured at a biochar:water ratio of 1:25.
Biochar’s labile carbon fraction was determined by assessing the CO2 that evolved from biochar over time (40 days) through the method described by Nelissen et al. (2014) and Cheng et al. (2008b). Biochar was thoroughly mixed with sand in a ratio of 1 g of biochar to 20 g of sand and placed into a 450-mL Mason jar. Distilled water, microbial inoculum, and micronutrient solution were added to the jar to maintain biological activity. Approximately 2 mL of microbial inoculum was added to each jar, which was the same as described above to produce matured biochar; then 2 mL micronutrient solution that contained 4 mM NH3NO3, 4 mM CaCl2, 2 mM KH2PO4, 1 mM K2SO4, 1 mM MgSO4, 25 mM MnSO4, 2 mM ZnSO4, 0.5 mM CuSO4, and 0.5 mM Na2MoO4 was also added to each jar. The experiments were initiated immediately after wetting. The evolved CO2 was trapped in 10 mL of 0.1 N NaOH, which was placed in a small vessel inside the sealed incubation jars. The NaOH vessels were removed and replaced with a new vessel on day 1, 3, 5, 7, 11, 15, 23, 31, and 40. The trapped CO2 was precipitated by an excess of 1 M of BaCl2 solution and back-titrated with standard 0.5 M of HCl to quantify the evolved CO2.
Soil incubation experiment
The incubation experiment included the following 8 treatments: (1) no amendment (control); (2) urea amendment (U); (3, 4, and 5) soil amendment with urea and FBC300, FBC500, FBC700, marked as F300, F500, and F700, respectively; and (6, 7, and 8) soil amendment with matured biochar, which was marked as M300, M500, and M700. Each treatment included 3 replications. The soil used was typical brown earth from northeast China, which was collected from a local farmland near Shenyang Agricultural University (41°83′N, 123°58′E). The top 15 cm of soil was collected following a maize harvest. Afterward, the soil was air-dried and sieved by using a 2-mm sieve. The basic properties of the soil were as follows: pH of 7.14 (soil:water = 1:2.5); bulk density of 1.29 g·cm−3; total N content of 1.69 g·kg−1; available N of 41.5 mg·kg−1, and SOC of 10.60 g·kg−1.
In each replication, 100 g of soil was thoroughly mixed with 0 g (control and U treatment) or 2 g of biochar (equal to 55 t·ha-1) and placed into a 450-mL mason jar. Then 10 mL urea solution (6 g·L−1) was applied to the soil (equal to 300 kgN·ha-1, except for the control soil). Distilled water was added to bring the moisture content to 60% of WHC, and the samples were incubated at 27 °C for 3 weeks. The remaining urea in the soil was determined after 12 h, 24 h, 36 h, and 48 h using the method proposed by Douglas and Bremner (1970). The indophenol blue colorimetric method was used to assay the urease activity on day 14. Available nitrogen was measured after 2 days and end of incubation by an alkaline hydrolysis method. Moreover, the microbial biomass carbon (MBC) and microbial biomass nitrogen (MBN) were extracted with 0.5 M K2SO4 and determined by a C/N analyzer (Analytik Jena 3100, Jena, Germany) at the end of incubation.
A one way ANOVA was performed using SPSS version 19 (IBM Corporation, New York, USA) to analyze the data. Significant differences among means were assessed by the least significant difference (LSD) test at 0.05 probability level. Figures shown in this article were made by using Graphpad Prism 5 (GraphPad Software, Inc., La Jolla, USA).
RESULTS AND DISSCUSSION
Characteristics of Biochar
The pyrolysis temperature significantly influenced the basic characteristics of the biochar (Tables 1 and 2). When the pyrolyzing temperatures were increased from 300 °C to 700 °C, the biochar carbon contents increased by 8.4%, while oxygen and hydrogen contents decreased by 9.32% and 4.3%. Accordingly, the ratios of O/C and H/C decreased from 0.39 and 0.12 to 0.2 and 0.4. This observation indicates that the degree of carbonization was accelerated with increasing pyrolysis temperature. During the 50 days of maturing processes, 1.6%, 2.0%, and 0.56% of the total carbon in biochar was lost from FBC300, FBC500, and FBC700, respectively. The biochar total N content exhibited a small change in the maturing processes, only MBC700 increased 0.34% compared with FBC700. The O contents in matured biochar were higher than fresh biochar, one promising reason was some oxygen-containing function groups were formed during maturing incubation (Cheng et al. 2008a).
Table 1. Elemental Composition of the Fresh and Matured Biochar
Table 2. Properties of the Fresh and Matured Biochar
As shown in Table 2, the ash and fixed carbon contents increased from 8.02% and 50.56% (FBC300) to 13.66% and 71.05% (FBC700), respectively. In contrast, biochar’s labile carbon fraction and biochar volatile matter decreased by 18.34% and 4.7 mg·g−1, respectively. After 50 days maturing incubation, the ash contents in matured biochar were 9.18%, 14.8%, and 15.65%, which was 1.16%, 1.7%, and 2% higher than fresh biochar, respectively. Nevertheless, the volatile matter showed no change, thereby the fixed carbon decreased 1.82%, 0.93%%, and 0.87% in MBC300, MBC500, and MBC700, respectively.
Pyrolysis process created small pores and damaged the large pores (Yue et al. 2016), resulting in pore diameter decreased from 49.7 to 27.9 nm when pyrolyzing temperatures were increased from 300 °C to 700 °C. It also can be concluded that more pores were created by increasing pyrolyzing temperature. The total pore volume in FBC700 was 1.69 times as FBC300; consequently, the biochar surface area increase from 24.50 m2·g-1 to 73.68 m2·g-1. A reduction of surface area and pore volumes were detected upon MBC700, possible reasons are that pore destruction or increase in oxygen-containing surface functional groups during the incubation (Ghaffar et al. 2015).
The biochar pH values increased from 7.03 to 10.35 with the increase of pyrolyzing temperature from 300 °C to 700 °C. Furthermore, the maturing processing minimized the range of pH values, which indicated that biochar alkalinity was neutralized by the maturing processes. As shown in Table 2, MBC300 presented a pH of 7.29, and was slightly higher than that of FBC300. However, the pH values for FBC500 and FBC700, which showed strong alkalinity, decreased 0.44 and 0.51 with mature incubation, respectively.
The results showed (Fig. 1) that within fresh biochar, the CO2 evolution occurred during the entire 40 days of incubation and in first 25 days for matured biochar. The biochar pyrolyzed at low temperatures (300 °C and 500 °C) usually contained a considerable amount of carbon that was easily mineralized (Table 2). For instance, the total evolved biochar of CO2-C from FBC300 was 14.6 mg/g of biochar, which was significantly (p < 0.5) higher than FBC500 (11.9 mg·g−1) and FBC700 (9.8 mg·g−1).
Fig. 1. Cumulative carbon mineralized from fresh and matured biochar during 40 days of incubation, (mean ± SE; n = 3)
Smaller amounts of CO2-C evolved from matured biochar (< 5.67 mg·g−1) as its labile carbon was mineralized during maturing processes. No significant differences were observed among MBC300, MBC500, and MBC700.
Effect of Biochar on Urea Hydrolysis rate and Available Nitrogen contents
Urea hydrolysis rate
The urea hydrolysis rate was accelerated significantly by biochar addition in 48 h (Fig. 2), particularly for fresh biochar. The residual urea-N in biochar-amended soils was lower than that in the urea addition only soil and decreased rapidly at the start of incubation. After 24 h of incubation, 185.6 mg urea-N·kg−1, 119.8 mg urea-N·kg−1, and 238.8 mg urea-N·kg−1 was hydrolyzed in FBC300-, FBC500-, and FBC700-amended soil, respectively. This was equal to 66.3%, 42.8%, and 85.3% of the total added urea-N, respectively. However, only 28.7% urea-N was hydrolyzed in the control soil (280 mg urea-N·kg−1 soil was added initially). After 48 h of incubation, urea-N in fresh biochar-amended soil was almost completely hydrolyzed, but the U treatment still contained about 50 mg urea-N·kg−1 soil. This result indicated biochar advanced at least one day or longer of the urea completely hydrolysis time than control.
The matured biochars showed a weak effect on accelerating urea hydrolysis, especially for biochar pyrolysis at 700 °C. The remaining urea-N in M700 treatment was 26.3%, 14.6%, and 13.5% higher than that of F700 treatment at 24 h, 36 h, and 48 h, respectively.
Fig. 2. Soil residual urea-N contents of different treatments within 48 h, different letters on the columns of the same incubation time represent significant differences at p < 0.05 level by LSD test; Error bars represent standard errors of the means (n = 3)
Available nitrogen contents
As shown in Fig. 3, at the early incubation stage, the addition of urea could significantly increase the soil available N via a “priming effect”, as only 80.3 mg urea-N·kg−1 was hydrolyzed during one day of incubation (Fig. 2), and the control soil contained available N of approximately 40 mg·kg−1, which indicated there were approximately 221 mg·kg−1[341.7 (total available N) – 40 (from soil) – 80.3 (from urea) = 221.4] soil N mineralized by urea-N addition. Moreover, biochar addition increased the soil N that was mineralized at the early incubation stage. Approximately 671 mg·kg−1, 620 mg·kg−1, and 299 mg·kg−1 N was mineralized from F300, F500, and F700 treatments, respectively. Matured biochar showed a significantly weak mineralization effect on soil N; Approximately 273 mg·kg−1, 251 mg·kg−1, and 282 mg·kg−1 N was mineralized by MBC300, MBC500, and MBC700 addition, respectively.
Fig. 3. Soil available N contents in urea applied soil at early and end of incubation, different letters represent significant differences at p < 0.05 level by LSD test; Error bars represent standard errors of the means (n = 3)
After 21 days of incubation, soil available N contents in biochar treatments (except for F700) were significantly (p < 0.05) lower than that in the U treatment. Among the biochar amended soil, the F500 treatment showed a minimal available N concentration of 59.6 mg·kg−1 among all treatments.
Effect of Biochar on Soil Urease Activity
The urease activity was determined on week 2 of the incubation. As shown in Fig. 4, compared with the control soil, the addition of urea alone inhibited urease activity by 35.5%. This phenomenon may have been caused by the substantial increase in soil pH value during urea hydrolysis, which could lead to further stability of soil organic–inorganic colloid, urease, and urea, and result in urease inactivation (Ran et al. 2000). Biochar potentially increased the urease activity. Among the amendments of biochar, together with urea-N, the urease activity in F500, F700, and M300 was 0.324 mg NH4+–N·kg−1, 0.314 mg NH4+–N·kg−1, and 0.309 mg NH4+–N·kg−1 soil per day, respectively, which was significantly (p < 0.05) higher than that in U treatment (0.191 mg NH4+–N·kg−1 soil per day). No significant differences were detected amongst the fresh and matured biochar.
Fig. 4. Urease activity in control and biochar-amended soil; Different letters represent significant differences at p < 0.05 level by LSD test; Error bars represent standard errors of the means (n = 3)
Effect of Biochar on Soil Microbial Biomass
As shown in Fig. 5a, the addition of FBC300, FBC500, and FBC700 increased soil MBC by 176.7 mg·kg−1, 140.85 mg·kg−1, and 64.11 mg·kg−1, respectively, when compared to U treatment. The MBC contents increased with decreased biochar pyrolysis temperature and a statistical analysis showed a positive correlation (R = 0.96) between fresh biochar’s labile carbon content and soil MBC contents. In matured biochar amended treatments, compared with the U treatment, the MBC contents only increased in MBC300- and MBC500-amended soil by 74% and 57.1%, respectively. The soil MBN contents were also increased by biochar addition (Fig. 5b); nevertheless, the significant difference was only found in FBC500-amended soil, which was 50% higher than the U treatment.
Some specific elements in pyrolytic materials are disproportionately lost to the atmosphere, fixed into recalcitrant forms, or liberated as soluble oxides during the heating process, which affects the chemical composition of the biochar surface (Chan and Xu 2009). Consequently, the elemental composition of biochar varied with different pyrolyzing temperatures. Consistent with previous research, the current study showed that biochar’s total C contents increase significantly with increasing pyrolyzing temperatures (Bruun et al. 2011; Enders et al. 2012), while O and H contents, O/C and H/C significantly decreased, indicating that the degree of carbonization and aromaticity was accelerated (Kim et al. 2012). In addition, low-temperature-pyrolyzed biochar contained a relatively large proportion of active compounds such as violate matters and labile carbon, but 700 °C biochar contained more fixed carbon. This result can be explained by the fact that during pyrolysis, a series of cleavage and polymerization reactions occur, thereby resulting in thermally stable fixed C structures (Spokas et al. 2012), which are directly related to increased biochar C content. Increasing pyrolyzing temperatures also typically leads to a loss of easily decomposable substances, volatile compounds, and elements (e.g., O, H, N, and S) and thus concentrates other nutrients present in biochar, including C, Ca, Mg, and K (Kim et al. 2012).
Fig. 5. Soil microbial biomass content of a) soil microbial carbon, and b) soil microbial nitrogen; Different letters represent significant differences at p < 0.05 level by LSD test; Error bars represent standard errors of the means (n = 3)
Nitrogen transformation is the most complex process during pyrolysis (Enders et al. 2012); in general, the total N content reaches a maximum between 300 °C and 399 °C and decreases at higher temperatures (Ippolito et al. 2015). This literature result is consistent with the authors’ result that biochar at 300 °C possessed maximum total N contents. However, the minimum total N contents were observed in biochar at 500 °C, which was caused by the increased hydrolysis temperature that led to violated N. Nevertheless, the total N was not concentrated, similar to that in biochar at 700 °C.
Biochar surface area and pore characteristic reflect its adsorption capability, which will affect nutrition cycling and microbial thrive in soil after biochar was applied (Sun et al. 2016). In present study, the author found that biochar pore characteristic was well developed under high pyrolysis temperature. So urea-N adsorbed by biochar may be one potential reason for the low urea-N contents observed in biochar amended soil.
Cheng et al. (2006) and Zimmerman (2010) reported the short-term oxidation of labile carbon fraction from biochar. According to Bruun et al. (2011), 90% of biochar-C losses occurred within the first 20 days of incubation. In the present study, the total CO2-C that evolved from FBC300, FBC500, and FBC700 was 14.5 mg·g−1, 11.9 mg·g−1, and 9.8 mg·g−1 during 40 days of incubation, which was equal to 2.5%, 1.87%, and 1.46% of total biochar carbon losses, respectively. These values matched the range of 0.3% to 3% biochar C losses reported by Zimmerman (2010) and Hamer et al. (2004). The 50 days of maturing processes resulted in 42.9% to 63.9% of biochar’s labile carbon mineralized into CO2, which was equal to 0.6% of total biochar-C in FBC700 and 1.6% of total biochar-C in FBC300.
Only few studies have examined the effect of biochar on available nitrogen concentrations associated with urea-N addition (Gao and Cai 2015; Hangs et al. 2015), and no researchers have detected how biochar affected the urea-N transformation in soil. In the present study, the authors found that biochar significantly accelerated urea hydrolysis and increased the soil available nitrogen content greatly at the beginning of incubation. The accelerated urea hydrolysis can be explained by the following two reasons: i) Increased urease activity. The enhanced urease activity in the presence of biochar is presumably caused by a combination of different mechanisms: a) biochar provides energy-rich organic compounds (Luo et al.2013; Singh and Cowie 2014) and microbial habitat that promote the growth of microorganisms resulting in the production of microbial urease (Zantua and Bremner 1977); and b) urease activity is optimum under slightly alkaline conditions (Dharmakeerthi and Thenabadu 2013), and biochar is a known liming agent; ii) Soil pH corresponds well to urea hydrolysis rate and nitrification processes (Gao and Cai 2015). This is consistent with the authors’ result that the urea hydrolysis rate increased with enhanced biochar pH value (Table 3).
Table 3. pH Values in Soil Amended with or without Urea and Biochar in the End of Incubation (Soil:Water = 1:2.5)
Unlike biochar derived from agricultural wastes, sewage sludge, or manure-based feedstock, crop stover or straw biochar typically exhibit relatively low bioavailable N to plants (< 2%) (Xie et al. 2013). Thus, as mentioned by previous findings that biochar can potentially mineralize N (Anderson et al. 2011; Nelissen et al. 2012), the authors concluded that the large amounts of available N in biochar-amended soil (Fig. 4) originated from the net mineralization of soil N. In detail, the labile carbon in biochar provides carbon sources that promote microbes’ growth and reproduction, and urea-N provided an inorganic N source. As a result, the soil showed a net mineralization effect by biochar and urea addition, especially for biochar that contained a relatively high concentration of labile carbon.
However, at the end of incubation, the available nitrogen in biochar amended soils was lower than in the urea-alone treatment. The authors concluded that the reduction of available nitrogen in biochar treatments was a result from a net immobilization effect that caused the thriving of soil microbes (Fig. 5). As former researchers illustrated, the microbial demand for inorganic N increases with biochar addition, thereby immobilizing N biotically (Novak et al. 2010; Nelissen et al. 2012, 2014). Nelissen et al. (2012) demonstrated that low-temperature biochar containing high concentrations of bioavailable C would probably induce net nitrogen immobilization. However, in this study, no differences in inorganic N contents were observed among the fresh and matured biochar treatments. In addition, it was reported that the reduction of inorganic nitrogen may also result from the adsorption by biochar (Ding et al. 2010; Singh et al. 2010; Spokas et al. 2012), but related works to prove this observation were not performed in the present study.
- As pyrolysis temperature increased from 300°C to 700°C, biochar carbonization degree was accelerated and more pores were created, resulting in an increase of biochar surface area from 24.50 m2·g-1 to 73.68 m2·g-1.
- Biochar, especially when newly produced, accelerated the rate of urea hydrolysis, and biochar enhanced soil urease activity and soil pH values were the main reasons for this effect.
- Biochar contained a fraction of labile carbon, which corresponded to 1.5% to 2.5% of biochar total carbon, and this fraction of carbon promoted microbial to thrive, during which lead to soil nitrogen mineralized.
- The highest concentration available of N in fresh biochar amended soil was from urea transformation and soil nitrogen mineralization. Therefore, the authors concluded that aged biochar was suitable for simultaneous soil amendment with urea rather than newly produced biochar, as it can promote available N accumulation in a short time; thus, an increase in the risk of inorganic nitrogen leaching loss.
The authors acknowledge the financial support from National Natural Science Foundation of China (41401325), the Chinese Academy of Engineering Consulting Project (2015-XY-25), and the S&T Program of Liaoning Province (2014215019).
Anderson, C. R., Condron, L. M., Clough, T. J., Fiers, M., Stewart, A., Hill, R. A., and Sherlock, R. R. (2011). “Biochar induced soil microbial community changes: Implications for biogeochemical cycling of carbon, nitrogen and phosphorus,” Pedobiologia 54, 309-320. DOI: 10.1016/j.pedobi.2011.07.005
Anderson, C. R., Hamonts, K., Clough, T. J., and Condron, L. M. (2014). “Biochar does not affect soil N-transformations or microbial community structure under ruminant urine patches but does alter relative proportions of nitrogen cycling bacteria,” Agriculture, Ecosystems & Environment 191, 63-72. DOI: 10.1016/j.agee.2014.02.021
Bruun, E. W., Hauggaard-Nielsen, H., Ibrahim, N., Egsgaard, H., Ambus, P., Jensen, P. A., and Kim, D. J. (2011). “Influence of fast pyrolysis temperature on biochar labile fraction and short-term carbon loss in a loamy soil,” Biomass and Bioenergy 35(3), 1182-1189. DOI: 10.1016/j.biombioe.2010.12.008
Cantrell, K. B., and Martin, J. H. (2011). “Stochastic state-space temperature regulation of biochar production. Part I: Theoretical development,” Journal of the Science of Food and Agriculture 92(3), 481-489. DOI: 10.1002/jsfa.4618
Chan, K. Y., and Xu, Z. H. (2009). “Biochar: Nutrient properties and their enhancement,” in: Biochar for Environmental Management: Science and Technology, J. Lehmann and S. Joseph (eds.), Earthscan, London, UK, pp. 67-84.
Chen, W., Shi, S., Nguyen, T., Chen, M., and Zhou, X. (2016). “Effect of temperature on the evolution of physical structure and chemical properties of bio-char derived from co-pyrolysis of lignin with high-density polyethylene,” BioResources 11(2), 3923-3926. DOI: 10.15376/biores.11.2.3923-3936
Cheng, C. H., Lehmann, J., Thies, J. E., Burton, S. D., and Engelhard, M. H. (2006). “Oxidation of black carbon by biotic and abiotic processes,” Organic Geochemistry 37(11), 1477-1488. DOI: 10.1016/j.orggeochem.2006.06.022
Cheng, C. H., Lehmann, J., and Engelhard, M. H. (2008a). “Natural oxidation of black carbon in soils: Changes in molecular form and surface charge along a climosequence”. Geochimica Et Cosmochimica Acta 72(6), 1598-1610. DOI:10.1016/j.gca.2008.01.010
Cheng, C. H., Lehmann, J., Thies, J. E., and Burton, S. D. (2008b). “Stability of black carbon in soils across a climatic gradient,” Journal of Geophysical Research: Biogeosciences113(G2), 50-55. DOI: 10.1029/2007JG000642
Dharmakeerthi, R. S., and Thenabadu, M. W. (2013). “Urease activity in soils: A review,” Journal of the National Science Foundation of Sri Lanka 24(3), 159-195. DOI: 10.4038/jnsfsr.v24i3.5548
Ding, Y., Liu, Y., Wu, W., Shi, D., Yang, M., and Zhong, Z. (2010). “Evaluation of biochar effects on nitrogen retention and leaching in multi-layered soil columns,” Water Air Soil Pollution 213(1), 47-55. DOI: 10.1007/s11270-010-0366-4
Douglas, L. A., and Bremner, J. M. (1970). “Extraction and colorimetric determination of urea in soil,” Soil Science Society of America Journal 34, 859-862. DOI: 10.2136/sssaj1970.03615995003400060015x
Enders, A., Hanley, K., Whitman, T., Joseph, S., and Lehmann, J. (2012). “Characterization of biochars to evaluate recalcitrance and agronomic performance,” Bioresource Technology 114, 644-653. DOI: 10.1016/j.biortech.2012.03.022
Gao, S., and Cai, Z. 2015. “Nitrogen transformation and nitrous oxide emissions affected by biochar amendment and fertilizer stabilizers,” in: 2015 ASA-CSSA-SSSA International Annual Meeting (CD #369-3), Minneapolis, MN.
Ghaffar, A., Ghosh, S., Li, F., Dong, X., Di, Z., Min, W., and Li H. (2015). “Effect of biochar aging on surface characteristics and adsorption behavior of dialkyl phthalates,” Environmental Pollution 206, 502-509. DOI.org/10.1016/j.envpol.2015.08.001
Hamer, U., Marschner, B., Brodowski, S., and Amelung, W. (2004). “Interactive priming of black carbon and glucose mineralisation,” Organic Geochemistry 35(7), 823-830. DOI: 10.1016/j.orggeochem. 2004.03.003
Hangs, R. D., Ahmed, H. P., and Schoenau, J. J. (2015). “Influence of willow biochar amendment on soil nitrogen availability and greenhouse gas production in two fertilized temperate prairie soils,” Bioenergy Research 9(1), 157-171. DOI: 10.1007/s12155-015-9671-5
IBI Biochar Standards Version 2.1 (2015). “Standardized product definition and product testing guidelines for biochar that is used in soil,” International Biochar Initiative, (http://www.biochar-international.org/sites/default/files/IBI_Biochar_Standards_ V2.1_Final.pdf), 23 November 2015.
Ippolito, J. A., Spokas, K. A., Novak, J. M., Lentz, R. D., and Cantrell, K. B. (2015). “Biochar elemental composition and factors influencing nutrient retention,” in: Biochar for Environmental Management: Science and Technology, J. Lehmann and S. Joseph (eds.), Earthscan, London, UK, pp. 67-84.
Jones, D. L., Murphy, D. V., Khalid, M., Ahmad, W., Edwards-Jones, G., and DeLuca, T. H. (2011). “Short-term biochar-induced increase in soil CO2 release is both biotically and abiotically mediated,” Soil Biology and Biochemistry 43(8), 1723-1731. DOI: 10.1016/j.soilbio.2011.04.018
Kim, K. H., Kim, J.-Y., Cho, T.-S., and Choi, J. W. (2012). “Influence of pyrolysis temperature on physicochemical properties of biochar obtained from the fast pyrolysis of pitch pine (Pinus rigida),” Bioresource Technology 118, 158-162. DOI: 10.1016/j.biortech.2012.04.094
Laird, D., Fleming, P., Wang, B., Horton, R., and Karlen, D. (2010). “Biochar impact on nutrient leaching from a midwestern agricultural soil,” Geoderma 158(3-4), 436-442. DOI: 10.1016/j.geoderma.2010.05.012
Lehmann, J., Silva, J. P. D., Steiner, C., Nehls, T., Zech, W., and Glaser, B. (2003). “Nutrient availability and leaching in an archaeological anthrosol and a ferralsol of the central amazon basin: Fertilizer, manure and charcoal amendments,” Plant Soil 249(2), 343-357. DOI: 10.1023/A:1022833116184
Li, F., and Wang, J. (2013). “Estimation of carbon emission from burning and carbon sequestration from biochar producing using crop straw in China (in Chinese),” Transactions of the Chinese Society of Agricultural Engineering 29(14), 1-7. DOI: 10.3969/j.issn.1002-6819.2013.14.001
Liang, B., Lehmann, J., Solomon, D., Kinyangi, J., Grossman, J., O’Neill, B., Skjemstad, J. O., Thies. J., Luizäo, F. J., Petersen, J., et al. (2006). “Black carbon increases cation exchange capacity in soil,” Soil Science Society America Journal 70(5), 1719-1730. DOI: 10.2136/sssaj2005.0383
Luo, Y., Durenkamp, M., Nobili, M. D., Lin, Q., Devonshire, B. J., and Brookes, P. C. (2013). “Microbial biomass growth, following incorporation of biochars produced at 350 °C or 700 °C, in a silty-clay loam soil of high and low pH,” Soil Biology and Biochemistry 57, 513-523. DOI: 10.1016/j.soilbio.2012.10.033
Major, J., Lehmann, J., Rondon, M., and Goodale, C. (2010). “Fate of soil-applied black carbon: Downward migration, leaching and soil respiration,” Global Change Biology 16, 1366-1379. DOI: 10.1111/j.1365-2486.2009.02044.x
Meng, J., Zhang, W., Wang, S., Xu, Z., and Chen, W. (2011). “Development and prospect of carbonization and returning technology of agro-forestry residue (in Chinese),” Journal Shenyang Agricultural University 42(4), 387-392. DOI: 10.3969/j.issn.1000-1700.2011.04.001
Nelissen, V., Rütting, T., Huygens, D., Ruysschaert, G., and Boeckx, P. (2014). “Temporal evolution of biochar’s impact on soil nitrogen processes – A 15N tracing study,” GCB Bioenergy 7(4), 635-645. DOI: 10.1111/gcbb.12156
Nelissen, V., Rütting, T., Huygens, D., Staelens, J., Ruysschaert, G., and Boeckx, P. (2012). “Maize biochars accelerate short-term soil nitrogen dynamics in a loamy sand soil,” Soil Biology and Biochemistry 55, 20-27. DOI: 10.1016/j.soilbio.2012.05.019
Novak, J. M., Busscher, W. J., Watts, D. W., Laird, D. A., Ahmedna, M. A., and Niandou, M. A. S. (2010). “Short-term CO2 mineralization after additions of biochar and switch grass to a typic kandiudult,” Geoderma 154(3-4), 281-288. DOI: 10.1016/j.geoderma.2009.10.014
Ran, W., Shen, Q., and Zheng, J. (2000). “Effect of urea concentration, incubation time and temperature on urea hydrolysis in 3 different soils” Journal of Nanjing Agricultural University 23(2), 43-46.
Reverchon, F., Flicker, R. C., Yang, H., Yan, G., Xu, Z., Chen, C., Bai, S. H., and Zhang, D. K. (2014). “Changes in δ15N in a soil–plant system under different biochar feedstocks and application rates,” Biology and Fertility of Soils 50(2), 275-283. DOI: 10.1007/s00374-013-0850-2
Singh, B. P., and Cowie, A. L. (2014). “Long-term influence of biochar on native organic carbon mineralisation in a low-carbon clayey soil,” Scientific Reports 4, #3687. DOI: 10.1038/srep03687
Singh, B. P., Hatton, B. J., Singh, B., Cowie, A. L., and Kathuria, A. (2010). “Influence of biochars on nitrous oxide emission and nitrogen leaching from two contrasting soils,” Journal of Environmental Quality 39(4), 1224-1235. DOI: 10.2134/jeq2009.0138
Spokas, K. A., Novak, J. M., and Venterea, R. T. (2012). “Biochar’s role as an alternative N-fertilizer: ammonia capture,” Plant and Soil 350(1), 35-42. DOI: 10.1007/s11104-011-0930-8
Steiner, C., Teixeira, W. G, Lehmann, J., Nehls, T., Macêdo, J. L. V., Blum, W. E. H., and Zech, W. (2007). “Long term effects of manure, charcoal and mineral fertilization on crop production and fertility on a highly weathered central Amazonian upland soil,” Plant and Soil 291(1), 275-290. DOI: 10.1007/s11104-007-9193-9
Steiner, C., Das, K. C., Garcia, M., Förster, B., and Zech, W. (2008). “Charcoal and smoke extract stimulate the soil microbial community in a highly weathered xanthic ferralsol,” Pedobiologia 51(5–6), 359-366. DOI: 10.1016/j.pedobi.2007.08.002
Sun, D., Hale, L., and Crowley, D. (2016). “Nutrient supplementation of pinewood biochar for use as a bacterial inoculum carrier”. Biology and Fertility of Soils, 52(4), 1-8. DOI: 10.1007/s00374-016-1093-9
Whitman, T., Singh, B. P., and Zimmerman, A. R. (2015). “Priming effects in biochar-amended soils: Implications of biochar-soil organic matter interactions for carbon storage,” in:Biochar for Environmental Management: Science and Technology, J. Lehmann and S. Joseph (eds.), Earthscan, London, UK, pp. 67-84.
Xie, Z., Xu, Y., Liu, G., Liu, Q., Zhu, J., Tu, C., Amonette, J. A., Cadishch, G., Yong, J. W. H., and Hu, S. (2013). “Impact of biochar application on nitrogen nutrition of rice, greenhouse-gas emissions and soil organic carbon dynamics in two paddy soils of China,” Plant and Soil 370(1), 527-540. DOI: 10.1007/s11104-013-1636-x
Yue, Y., Lin, Q., Irfan, M., Chen, Q., Zhao, X. (2016). “Characteristics and potential values of bio-oil, syngas and biochar derived from Salsola collina pall. in a fixed bed slow pyrolysis system,” Bioresource Technology 220, 378. DOI: 10.1016/j.biortech.2016.08.028
Zantua, M. I., and Bremner, J. M. (1977). “Stability of urease in soils,” Soil Biology and Biochemistry 9(2), 135-140. DOI: 10.1016/0038-0717(77)90050-5
Zimmerman, A. R. (2010). “Abiotic and microbial oxidation of laboratory-produced black carbon (biochar),” Environment Science and Technology 44(4), 1295-1301. DOI: 10.1021/es903140c
Article submitted: March 17, 2017; Peer review completed: June 10, 2017; Revised version received: June 21, 2017; Accepted: June 22, 2017; Published: July 6, 2017.