Methane and Nitrous Oxide Flux after Biochar Application in Subtropical Acidic Paddy Soils under Tobacco-Rice Rotation

Abstract

Biochar amendment is a good means of mitigating methane (CH₄) and nitrous oxide (N₂O) emissions. However, the effects of biochar amendment on N₂O and CH₄ reduction in soil under rotation with different soil moisture contents is not well understood. To understand CH₄ and N₂O flux from soil with biochar amendment under water-unsaturated and water-saturated conditions, a field experiment was conducted in a tobacco-rice rotation field in subtropical China to investigate N₂O and CH₄ emissions following soil amendment with tobacco straw biochar at rates of 0, 10, 40 and 80 t·ha⁻¹ (B0, B10, B40 and B80, respectively). N₂O and CH₄ emissions were monitored by a closed-chamber method in the water-unsaturated tobacco (UT) and water-saturated rice (SR) seasons during the 2015 planting season. The soil pH increased from 5.4 in the control to 6.1 in the soil amended with biochar at 80 t·ha⁻¹ in the UT season. During both the UT and SR seasons, with biochar amendment at 40 and 80 t·ha⁻¹, the soil bulk density (BD) decreased, while the soil organic matter (SOM) and available potassium (Av. K) contents increased. N₂O flux was significantly greater in UT than in SR in the controls but decreased with the application of biochar during both the UT and SR seasons. The cumulative CH₄ emission decreased with the rate of biochar application and the methanotroph pmoA gene copy number in soils and increased with the methanogenic archaea 16Sr DNA gene copy number in soils during the rice-cropping season. These results indicated that biochar amendment could decrease methanogenic archaea and increase of methanotroph pmoA gene, which are the mechanistic origin for CH₄ reduction.

Introduction

CH₄ and N₂O are important greenhouse gases in the atmosphere^1,2. Due to human activities, the concentrations of N₂O and CH₄ in the atmosphere increased from 270 ppbv and 324 ppb in 1750 to 324 ppbv and 1803 ppb in 2011, respectivelly^2,3. With the growing demand for food, further increases in these greenhouse gasses are projected in the future³. Agricultural soil is the main source of N₂O and CH₄ emissions, accounted for approximately 66% and 50% of total emissions, respectively^1,4. To mitigate global warming, it is necessary to employ strategies that will reduce these gas emissions from agricultural soil⁵.

Reducing N fertilizer input and increasing its use efficiency could decrease N₂O emissions; biochar application might play an important role in this approach. With its feedstock availability and favourable properties, biochar has been considered a good input for improving crop N use efficiency and increasing carbon (C) return to soil^6–8. A meta-analysis using data across 208 peer-reviewed studies showed that symbiotic biological N₂ fixation and plant N uptake were increased by 63% and 11%, respectively, with biochar amendment⁹. It has been found that the crop growth response with biochar is greater in acidic soils than neutral and alkaline soils because the soil nutrient-retention capacity is typically low in acidic soil and applying biochar will increase this capacity¹⁰. Thus, there is an increase in crop growth as the pH of soil amended with biochar is increased¹¹. However, the rate of nitrification and denitrification is improved in acidic soil with increasing pH, and these are the main pathways for N₂O production. Conversely, as the soil pH is increased, the activity of N₂O reductase (N₂O-R) is improved¹². This improvement will lead to more reduction of N₂O to N₂ (which is the last step of the denitrification process), resulting in a decrease in the N₂O emission rate. Additionally, biochar can reduce nirK abundance and increase nosZ abundance, which will inhibit N₂O production and simultaneously increase N₂O consumption in acidic soil^13,14. Therefore, the effects of biochar on N₂O emissions in acidic soil may be different from those in other soil types.

CH₄ is primarily produced in an exclusively anaerobic process by methanogenic archaea^4,15. However, under unsaturated conditions, soils are considered sinks for CH₄⁷. Powlson et al.¹⁶ reported that non-flooded upland soils are an important CH₄ sink and that approximately 15% of global CH₄ oxidation is attributed to this sink. Microbial oxidation of CH₄ is the main pathway of soil CH₄ uptake, which is driven by methanotrophs. Some methanotrophs feed solely on CH₄, while others are facultative methanotrophs, which include Methylocella and Methylocapsa^17,18. It has been reported that CH₄ uptake in soil may be increased by biochar amendment due to the adsorption of CH₄ onto the surfaces of biochar¹⁹. Additionally, soil aeration may be affected when biochar is added, and this may increase diffusive CH₄ uptake²⁰. Biochar has also been observed to stimulate methanotrophic CH₄ consumption in anoxic environments, particularly at oxic/anoxic interfaces²¹. For example, most CH₄ is produced in anoxic sediment in saturated soils, and some CH₄ is consumed at the aerated root interface⁸. It was reported by Feng et al.²² that under saturated conditions, biochar significantly increased the strength of CH₄ sink properties compared to the controls via decreasing the ratio of methanogenic archaea to methanotrophic bacteria⁸. Furthermore, biochar can provide a C substrate for microbial CH₄ oxidation in soils²⁰, and the labile C of biochar may be used as a methanogenic substrate in anoxic environments, promoting CH₄ production²³. However, the effects of biochar on CH₄ emissions from soils under saturated and unsaturated conditions are not fully understood.

Additionally, uncertainty exists as to whether biochar’s GHG mitigation effects persist for one cropping season or longer. Lentz et al.²⁴ suggested that the GHG mitigation effects of biochar application may be long-lived, whereas Spokas²⁵ indicated that they were mainly short-term (a few days to several weeks). Therefore, further study is required to determine the period of GHG mitigation effects resulting from biochar application.

Tobacco followed by rice cropping is a common agricultural practice in South China. However, tobacco straw is discarded beside the field after harvest, and this may lead to disease outbreaks and infections. Instead of simply discarding tobacco straw, incorporating this material into the soil as biochar has been widely recommended. This will help improve soil organic matter and reduce environmental pollution caused by straw littering.

To investigate the effect of biochar made from tobacco straw on N₂O and CH₄ emissions in acidic soil under rotation with different water regimes, a field experiment with tobacco and rice rotations was conducted in subtropical China. Specifically, we hypothesize that soil amendment with biochar (i) may not reduce the N₂O emission rate, given that stimulating N₂O reduction may be counteracted by improved nitrification and denitrification in acidic soil due to increased soil pH after biochar application, and (ii) affects the emissions of N₂O and CH₄ early in the first planting season but not in the second planting season because the active surfaces of the biochar become saturated over time.

Results

Soil properties

The soil pH determined after tobacco harvest increased with an increased rate of biochar application; the highest pH was found in B80 (Table 1). However, no significant differences were recorded in pH after the rice harvest (Table 1). Soil BD was decreased from 1250.0 kg·m⁻³ in the control to 1170.0 kg·m⁻³ and 1160.0 kg·m⁻³ in B40 and B80 in the UT season, and from 1360.0 kg·m⁻³ in the control to 1210.0 kg·m⁻³ and 1190.0 kg·m⁻³ in B40 and B80 in the SR season. However, the SOM and Av. K contents increased with increasing rates of biochar in both the UT and SR seasons.

Table 1.

Soil properties after biochar amendment.

Treat- ment	UT season						SR season
	pH	SOM g·kg−1	BD kg·m−3	Av. N mg·kg−1	Av. P mg·kg−1	Av. K mg·kg−1	pH	SOM g·kg−1	BD kg·m−3	Av. N mg· kg−1	Av. P mg·kg−1	Av. K mg·kg−1
B0	5.4c	22.7b	1250.0 a	122.2a	34.9a	117.0b	5.5a	22.8b	1360.0ab	111.5b	24.6b	103.4c
B10	5.6bc	25.5b	1210.0ab	127.3a	37.3a	148.8b	5.4a	24.5b	1310.0 a	120.9b	46.6a	137.8c
B40	5.6bc	32.2a	1170.0bc	122.2a	36.3a	303.3b	5.2a	30.2a	1210.0 b	121.7b	51.0a	385.6b
B80	6.1a	35.3a	1160.0 c	120.3a	39.5a	481.3a	5.6a	34.9a	1190.0 b	153.0a	51.5a	467.6a

The soil samples were collected after tobacco and rice were harvested. Data in the table are the means, and different letters indicate significant differences at 5%; B0, B10, B40 and B80 are no biochar applied and biochar applied at the rate of 10, 40 and 80 t·ha⁻¹, respectively; UT is unsaturated tobacco cropping; SR is saturated rice cropping; ccSOM is soil organic matter; BD is bulk density; Av. N is alkali-hydrolysable nitrogen; Av. P is available phosphorus; and Av. K is available potassium.

N₂O and CH₄ emissions

The N₂O flux from all treatments was greater during the UT season than in the SR season (Fig. 1). The highest rate of N₂O emission in most of the flux peaks was found in the B0 treatment during the experiment (Fig. 1). The cumulative N₂O emissions during the UT season was 9.8 to 11.3 kg N·ha⁻¹, which was significantly higher than that during the SR season by ≈10 times (Fig. 2). There was no significant difference among the treatments during the UT season (Fig. 2a), whereas the cumulative N₂O flux during the SR season from B80 was significantly lower than that from B0 (Fig. 2b). There was a negative relationship between cumulative N₂O flux and the rates of biochar applied during the UT and SR seasons (Fig. 3a,b, respectively).

Figure 1.

Temporal pattern of N₂O fluxes for the different treatments during the tobacco and rice growth periods. The period from 3/20 (MM/DD) to 7/20 was the unsaturated tobacco growth season (UT), and the period from 7/20 to 11/20 was the saturated rice growth season (SR); B0, B10, B40 and B80 are no biochar applied and biochar applied at a rate of 10, 40 and 80 t·ha⁻¹, respectively.

Figure 2.

Cumulative N₂O missions during the tobacco (a) and rice (b) growth periods. Values with same letter are not significantly different (p < 0.05); B0, B10, B40 and B80 are no biochar applied and biochar applied at the rate of 10, 40 and 80 t·ha⁻¹, respectively.

Figure 3.

Relationship between the rate of biochar application and cumulative N₂O emission during the tobacco (a) and rice (b) growth periods and between the rate of biochar application and cumulative CH₄ uptake during the tobacco growth season (c) and emission during the rice growth period (d). Positive values of CH₄ flux are CH₄ emission, and negative values are CH₄ uptake.

Unlike N₂O, CH₄ was taken up during the UT season, and the flux was also recorded during the SR season (Fig. 4). In the first two months of the SR season, the greatest CH₄ emission flux was observed for B0, followed by B10, B40 and B80 (Fig. 4). This indicated that CH₄ was affected by the rate of biochar amendment. The cumulative CH₄ uptake during the UT season in the B80 treatment was −8.5 kg·ha⁻¹, which was significantly greater than the values obtained for the B0 and B10 treatments (Fig. 5a). The cumulative CH₄ emission during the SR season in B0 was 159.3 kg·ha⁻¹, which was significantly higher than the values obtained from other treatments (Fig. 5b). Compared to N₂O, the cumulative CH₄ uptake during the UT season increased with increasing rates of biochar amendment (Fig. 3c). The cumulative CH₄ emission during the SR season decreased with an increasing rate of biochar application (Fig. 3d).

Figure 4.

Temporal pattern of CH₄ fluxes for the different treatments during the tobacco and rice growth periods. The period from 3/20 (MM/DD) to 7/20 was the unsaturated tobacco growth season (UT), and the period from 7/20 to 11/20 was the saturated rice growth season (SR); B0, B10, B40 and B80 are no biochar applied and biochar applied at the rate of 10, 40 and 80 t·ha⁻¹, respectively.

Figure 5.

Cumulative CH₄ uptake during the tobacco growth season (a) and CH₄ emissions during the rice growth period (b). Values with the same letter are not significantly different (p < 0.05); B0, B10, B40 and B80 are no biochar applied and biochar applied at the rate of 10, 40 and 80 t·ha⁻¹, respectively. Positive values of CH₄ flux are CH₄ emission, and negative values are CH₄ uptake.

Soil 16Sr DNA and pmoA abundance

Methanogenic archaea 16Sr DNA and methanotroph pmoA gene copy numbers were determined after the rice harvest. Figure 6 shows that the methanogenic archaea 16Sr DNA gene copy number in the B0 treatment was 5.3 × 10⁶ copies·g⁻¹ soil; this decreased with biochar application, but there were no significant differences at p ≤ 0.05 due to large variations in the same treatments. The highest methanotroph pmoA gene copy numbers were found in the B80 treatment, and the lowest was found in the B0 treatment, with no significant difference recorded among the different treatments (Fig. 6). The cumulative CH₄ emission during the SR season significantly increased with the methanogenic archaea 16Sr DNA gene copy number and decreased significantly with the methanotroph pmoA gene copy number (Fig. 7).

Figure 6.

Methanogenic archaea 16Sr DNA gene (a) and methanotroph pmoA gene (b) copy numbers in the rice growth period. Values with the same letter are not significantly different (p < 0.05); B0, B10, B40 and B80 are no biochar applied and biochar applied at the rate of 10, 40 and 80 t·ha⁻¹, respectively.

Figure 7.

Relationship between the amount of CH₄ emission during the rice growth period and methanogenic archaea 16Sr DNA gene (a) and methanotroph pmoA gene (b) copy numbers.

Discussion

N₂O emissions decreased with the rate of biochar application in both the UT and SR seasons

In contrast to our hypothesis that amendment with biochar may not reduce the N₂O emission rate in acidic soil. We observed that N₂O emissions decreased with the biochar application rate in both the UT and SR seasons (Fig. 3a,b). In line with the finding of Cayuela et al.²⁶, there was a direct negative correlation between the rate of biochar application and N₂O emission reductions. Both nitrification and denitrification have been identified as the predominant pathways for N₂O production. It has been reported that biochar mitigation of N₂O emissions via nitrification may be due to improvements in soil biological properties, physical properties, and chemical properties^11,20. Furthermore, reducing nitrification substrate by NH₄⁺ adsorption and the inhibition of potential microbial metabolic pathways (e.g. various polyphenolic and monoterpene compounds) play important roles in inhibiting nitrification and subsequent N₂O emission^27,28. Zhang et al.²⁹ and Fidel et al.³⁰ reported that amendment with biochar produced at 400 and 600 °C increased the NH₄⁺ adsorption capacity by 62–81% and was maximized with low pyrolysis temperature (400 °C), leading to a significant decrease in soil inorganic N. The biochar used in our experiment was produced at 450 °C; thus, the adsorptive capacity for NH₄⁺ related to nitrification could be a key factor in mitigating N₂O emissions.

There are several mechanisms that have been suggested to explain the reasons for biochar mitigation of N₂O emissions via denitrification. These mechanisms include NO₃⁻ immobilization, aeration regulation and biochar toxicity³¹. It has been reported that adsorption and retention of NH₄⁺ are improved in soils amended with biochar, indirectly leading to reductions in the amount of available N for denitrification, which is considered one of the important reasons for reducing N₂O emissions via denitrification by biochar-amended soils³². Additionally, biochar has been proposed as a reducing agent for soils containing redox-reactive Fe(III) and Mn(IV) to compete with NO₃⁻, reducing denitrification and promoting the reduction of N₂O to N₂³³.

It has been reported that biochar can reduce the abundance of nitrite-reducing bacteria (carrying the nirK and nirS genes) but also increase the abundance of N₂O-reducing bacteria (carrying the nosZ gene); thus, the mitigation of N₂O emissions by biochar may be attributed to the fact that biochar can inhibit N₂O production and simultaneously promote N₂O consumption^14,34,35. The reduction of N₂O to N₂ in the last step of denitrification may be improved, which would lead to a decrease in the N₂O emission rate¹³. However, work performed by Ameloot et al.³⁶ showed that a reduction in the N₂O/(N₂O + N₂) ratio was not observed via the acetylene inhibition method, which suggested that biochar did not stimulate nosZ and the reduction of N₂O to N₂. These contradictory phenomena may be attributed to soil and biochar properties. It has been reported that biochar predominantly promotes the last step of denitrification in fine-textured soil³². In acidic soil, the abundance of nitrite-reducing bacteria and N₂O-reducing bacteria increases as the soil pH is increased³⁷. This may not only enhance the reduction of NO₃⁻ to N₂O but will also lead to stronger and more complete N₂O reduction to N₂, culminating in a balance in soil N₂O emissions. Additionally, Cayula et al.^33,38 suggested that the effect of biochar on the denitrification of N₂O was mostly depend on its pH and the ratios of C/N and H/Corg.

In our observations, the soil pH was increased by biochar in the UT season from 5.4 to 6.1, whereas in the SR season, the pH returned to the initial range of values between 5.2 and 5.6 (Table 1). This was an indication that the addition of biochar to the soil had a rapid effect on soil pH, and this observation is in line with the findings of Castaldi et al.³⁹. They reported that soil pH was significantly higher in soils treated with char incorporation than the control; however, the effect of biochar on pH was transient within the first three months, and the pH later returned to the initial value after 14 months³⁹. As the soil pH was increased by biochar application, the cumulative N₂O emissions from the treatments with different rates of biochar application were not significantly different because the inhibition of N₂O emissions by biochar was likely offset by the stimulated pH increase.

Biochar decreased CH₄ emissions and increased CH₄ uptake in the UT season and SR season

In contrast to our hypothesis that biochar would decrease CH₄ emission in the first season and not in the second season, biochar decreased CH₄ emissions in the second cropping (SR season). Similarly, Feng et al.²² and Chen et al.⁴⁰ also observed a decline in CH₄ flux following biochar application in paddy soils and suggested that the effects of biochar on CH₄ emission were long-lived. However, contrary to this finding, Zhang et al.⁴¹ reported that CH₄ emission was increased after biochar was added to paddies, while Xie et al.⁴² reported that there was no significant difference in seasonal cumulative CH₄ emission between treatments. In the present study, we observed that the abundances of methanogenic archaea in the paddy were decreased by biochar application, while the methanotroph abundances were increased after biochar was applied, although the differences were not significant due to the large variations in a given treatment (Fig. 7). This showed that biochar addition decreased CH₄ emissions, which may be due to decreased methanogenic archaea abundance; hence, CH₄ could be utilized by methanotrophs. Feng et al.²² reported that methanogenic archaea were not inhibited by biochar amendments but there was a decreased ratio of methanogenic to methanotrophic microorganisms in paddy soils. An increase in methanotrophic abundance implies that methanogenic activity is been stimulated under biochar amendment (Fig. 7).

Biochar improved the sink capacity for CH₄ (Figs. 3c and 5a), which may be directly attributed to the decrease in the bulk density and soil aeration that occurred during the UT season (Table 1). Environments with a CH₄ sink capacity are suitable for methanotroph growth; however, Fungo et al.¹¹ reported that biochar reduced the sink capacity of CH₄. They attributed this to the easily mineralizable C provided by biochar, which was a substitute for methanotrophic bacteria⁴³. Biochar acts as a slow C release source, and the relationship between biochar and C mineralization is dependent on the production procedure of the biochar⁴⁴. The chemical properties and the type of biomass used for pyrolysis may also have influenced soil C and N cycling⁴⁵. Additionally, biochar has high porosity, which may be able to increase soil aerobic micro-sites, affecting aeration and improving the supply and distribution of CH₄ and O₂^46,47 When soil aeration increases as bulk density decreases (Table 1), the CH₄ oxidation activity of methanotrophs is greatly enhanced, which results in CH₄ emission mitigation^22,48.

Conclusion

Biochar application affected soil pH in the short term during the tobacco cropping season; however, in the saturated rice growth season, the pH reverted back to the initial value. The available potassium and SOM contents were improved, and BD was decreased by biochar application during the tobacco and rice growth seasons. N₂O flux during the UT season was significantly greater than that in the saturated rice growth season and decreased with the rate of biochar application. The soils were sinks for CH₄, and the cumulative CH₄ uptake was increased with the rate of biochar application in the tobacco cropping season. However, there was considerable CH₄ flux during the rice growth season, and the cumulative CH₄ emission decreased with an increased rate of biochar application. Cumulative CH₄ emissions had a negative relationship with methanotroph pmoA gene copy numbers and a positive relationship with methanogenic archaea 16Sr DNA gene copy numbers in the soils, indicating that stimulating methanotrophs and depressing methanogenic archaea are the mechanisms for CH₄ reduction upon biochar amendment. Therefore, to prevent environmental pollution and maintain the soil organic matter content, tobacco straw could be used as a biochar feedstock to reduce N₂O and CH₄ flux from soil. The rate of biochar application played important roles in N₂O and CH₄ flux, and further research should be conducted to study the relationship between biochar and the sink capacity for CH₄.

Materials and Methods

Biochar production

Biochar was produced from dried tobacco straw; the straw was cut into small segments (<50 mm length) before being placed into the reactor. The reactor was heated by a step-wise procedure. The heating temperature was increased to 450 °C under anaerobic conditions and maintained for approximately 8 h. Before applying biochar to the field, the biochar was further reduced to smaller sizes of <5 mm. The concentrations of N, P, K, and organic C and pH (H₂O) in the biochar were 15.0 g·kg⁻¹, 1.4 g·kg⁻¹, 20.1 g·kg⁻¹, 475.92 g·kg⁻¹ and 9.7, respectively.

Field site description

The field experiment was conducted in 2015 in Jinan County, Fujian Province, China (119°36′86″E, 26°17′33″N). The mean annual temperature and precipitation in this region are 18.3 °C and 1500 mm (over 30 years), respectively, and the region is characterized by a subtropical monsoon climate. The soil is defined as an Anthrosol (WRB Soil Taxonomy), and the average concentrations of SOM, total N (TN), total phosphorus (TP), total potassium (TK), alkali-hydrolysable nitrogen (Av. N), available phosphorus (Av. P), and Av. K and pH (H₂O) were 25.6 g·kg⁻¹, 1.4 g·kg⁻¹, 0.7 g·kg⁻¹, 20.0 g·kg⁻¹, 181.4 mg·kg⁻¹, 58.0 mg·kg⁻¹, 443.0 mg·kg⁻¹ and 5.3, respectively. The cropping sequence at the site was as follows: tobacco was planted in mid-March, then rice was planted in mid-July, for more than 15 years. The root and straw of tobacco were removed before tilling by machine. The treatments included three rates of biochar application and a control that did not receive any biochar amendment: no biochar applied (B0); biochar applied at the rate of 10 t·ha⁻¹ (B10); biochar applied at the rate of 40 t·ha⁻¹ (B40); and biochar applied at the rate of 80 t·ha⁻¹ (B80). Three replicate plots (24 × 6 m) of each treatment were established in a randomized block design. The biochar was applied on 1^st March 2015, before tobacco seedlings were transplanted. Except for the biochar, all treatments received N, P and K fertilizers at a recommended rate divided into three separate applications, which are given in Table 2. Compound fertilizers, urea and potassium nitrate were applied as N sources for tobacco; ammonium bicarbonate and urea were applied as N sources for rice.

Table 2.

Annual fertilizer application rates in the field experiment (kg·ha⁻¹).

Fertil- izers	UT season					SR season					Total amount
	Base fertilizers	Seedling stage	Rosette stage	Vigorous growth stage	Total	Base fertilizers	Green stage	Tillering stage	Heading stage	Total
N	104.4	3.9	7.6	11.5	127.5	66.7	66.2	16.6	16.6	166.0	293.5
P2O5	97.0	—	1.0	1.0	99.0	29.9	—	—	—	29.9	128.9
K2O	266.4	13.3	23.3	74.2	377.2	36.0	36.0	—	—	72.0	449.2

UT is unsaturated tobacco cropping, and SR is saturated rice cropping.

CH₄ and N₂O emission monitoring

Greenhouse gas emissions were monitored using static closed chambers²¹. Gas samples were collected between 9 and 11 am in a 7-day interval during the UT and SR seasons. Two chambers (0.5 m × 0.5 m × (0.5 + x) m) were placed on a fixed steel frame (stainless) in each plot after transplanting in the tobacco and rice growing seasons. One tobacco or six rice plants were closed in the chamber, and the height of the chamber was increased according to the height of the plant (x = 0.5 m or 1.0 m). To seal the rim of the chamber with a level surface, a groove (50 mm depth) along the top edge of each steel frame was filled with water. To minimize air temperature variation inside the chamber during the sampling period, the chambers were wrapped with a layer of porous insulation and aluminium foil. A circulating fan, humidity meter and temperature meter were equipped inside each chamber. After chamber closure, a syringe was used to collect gas samples at 0, 20, 40, and 60 min throughout the UT season and at 0, 10, 20, and 30 min during the SR season. The concentrations of N₂O and CH₄ were simultaneously analysed by a gas chromatograph (Agilent 7890B, USA), which was equipped with an electron capture detector (ECD) for N₂O and a flame ionization detector (FID) for CH₄ analysis²¹.

Soil samples

Soil samples were collected before the experiment and after the tobacco and rice had been harvested for property analyses. Soil organic matter was analysed by wet digestion with H₂SO₄-K₂Cr₂O₇, and total N was analysed using semi-micro Kjeldahl digestion with Se, CuSO₄ and K₂SO₄ as catalysts⁴⁹. A pH detector (Quark Ltd, Nanjing, China) was used to measure soil pH with a ratio of soil to water of 1:2.5 (w/v). Soil BD was analysed via the cutting ring method. Soil TP and TK were determined by the colorimetric and flame photometer methods after wet digestion with a mixture of H₂SO₄ and HClO₄ and a mixture of HF and HClO₄, respectively⁵⁰. Soil Av. N was diffused with 1.0 mol·L⁻¹ NaOH and trapped with 0.32 mol·L⁻¹ H₃BO₃. Soil Av. P was extracted with a mixture of 0.025 mol·L⁻¹ HCl and 0.03 mol·L⁻¹ NH₄F, while soil Av. K was determined by the 1.0 mol L⁻¹ CH₃COONH₄ extraction method^51,52.

A PowerSoil® DNA Isolation kit (MO BIO Laboratories, Inc., Carlsbad, USA) was used to extract DNA from 0.25 g fresh soil following the manufacturer’s instructions. The quality and quantity of DNA were checked by a NanoDrop spectrophotometer (NanoDrop Technologies Inc., Wilmington, USA). The copy number of methanogenic archaea 16Sr DNA genes and methanotroph pmoA genes were enumerated by quantitative PCR using primer sets 1106 F/1378R⁵³ and A189/m661²² with a CFX96 Optical Real-Time Detection System (Bio-Rad Laboratories, Inc. Hercules, CA). The qPCR standard was produced by plasmid DNA from representative clones including the methanogenic archaea 16Sr DNA gene and methanotroph pmoA gene. The 25.0 µL reaction mixture contained 12.5 µL of SYBR Premix Ex Taq (TaKaRa Biotech, Dalian), 1.0 µL of each primer, 0.5 μL Rox Reference Dye II (50×), and 1.0 µL template. The thermal conditions of quantitative PCR for the methanogenic archaea 16Sr DNA genes and methanotroph pmoA genes were those given by Feng et al.²² and Watanabe et al.⁵³. Specific amplification of the 16Sr DNA and pmoA genes was checked by confirming a single peak in melting-curve analysis. Copy numbers of genes are reported per dry weight of soil.

Calculation

The rates of GHG emission from soil were calculated using Eq. (1), as follows:

$$F=\rho \times h\times dc/dt\times 273/(273+T)\times t$$ 1

where F is the CH₄ or N₂O emission rate from soil (mg·m⁻²·h⁻¹), ρ is the density of CH₄ or N₂O under standard atmospheric pressure (0.714 and 1.96 kg·m⁻³, respectively), h is the height of the static closed chambers (m), dc/dt is the rate of change in CH₄ or N₂O concentration, T is the temperature inside the chamber (°C), and t is the time of chamber closure (h).

The amounts of CH₄ and N₂O emissions were calculated using Eq. (2), as follows:

$$C=\sum _{i=1}^n(\frac{{\mathit{F}}_{\mathit{i}}+{\mathit{F}}_{\mathit{i}+\mathit{1}}}{\mathit{2}}\times \mathit{24}\times D)$$ 2

where C is the amount of CH₄ or N₂O emission (kg·ha⁻¹), F_i and F_i + 1 are the CH₄ or N₂O emission rates at times i and i + 1, respectively (mg·m⁻²·h⁻¹), and D is the number of days between times i and i + 1.

Statistical analysis

The differences in the rates and amounts of CH₄ and N₂O emissions and soil properties were examined by one-way ANOVA. The significant differences among treatments were identified by Duncan’s test (where p < 0.05).

Author contributions

Y.H. wrote the main manuscript. C.W., C.L., Y.Z., X.C., L.T. did the field experiment. C.L. and Q.C. analyzed samples. M.O. and T.S. analyzed data. All reviewed and commented on the paper.

Competing interests

The authors declare no competing interests.