1. Introduction
On a 100-year timescale, the greenhouse gas nitrous oxide (N
2O) has a 265 to 298 times greater global warming potential (GWP) than CO
2. This GHG remains in the atmosphere for 121 years [
1,
2]. The atmospheric N
2O concentration has risen significantly since the pre-industrial period, from 270 ppb in 1750 to 330 ppb in recent years [
3]. Currently, concentrations continue to increase by 0.73 ppb year
–1 [
4], contributing to climate change. By modeling, Silva et al. [
5] predicted such increases in N
2O emissions for the next five decades, regardless of the crop or soil management system, for the conditions of the Brazilian Cerrado.
Nitrous oxide is the most important greenhouse gas emitted from agricultural soils as a byproduct of microbial nitrification and denitrification [
6]. Agriculture and soil management account for 78% of the N
2O anthropogenic emissions [
7]. In Brazil, 87% of the N
2O emissions are attributed to the agriculture and livestock sector [
8]. The main reasons for this high contribution to GHG emissions are organic matter oxidation and microbe-mediated processes, which are sensitive to plant residue management, nitrogen fertilization [
9], and climatic conditions [
6].
Since agriculture contributes significantly to N
2O emissions, it is imperative to find new solutions to increase soil carbon [
10] and crop productivity while reducing the environmental burden of the agricultural system [
11]. To this end, in line with the Paris Agreement, the Brazilian government established strategies to deal with climate change in agriculture, one of which is the expansion of integrated crop–livestock system areas [
12].
In integrated crop–livestock systems that include crops and pasture, N
2O emissions are more effectively mitigated than under traditional management. According to Amadori et al. [
13], integrated farming systems in subtropical regions based on crop/livestock, livestock/forestry, or crop/livestock/forestry reduced soil N
2O emissions by 27–40% compared to livestock only. Under similar climate conditions, Pereira et al. [
14] showed the potential of integrated agricultural systems to reduce N
2O losses when compared to monoculture systems. In an experimental area in the tropics, Sato et al. [
15], observed that even on clayey Oxisols, integrated systems such as integrated crop–livestock are an alternative to mitigate N
2O emissions when compared to continuous cropping due to the accumulation of stabilized C fractions in the soil organic matter (SOM), unavailable to the microorganisms. Based on these recent studies, the potential of integrated farming systems to mitigate soil N
2O emissions is greater than that of livestock/monoculture systems.
Soil use and management influence microbial nitrification and denitrification, which are biogenic sources of N
2O emissions [
16,
17,
18]. In addition, soil microbiological properties play an important role in C and N dynamics, but the influence on N
2O emissions requires further investigation, mainly in diversified systems such as crop/livestock farming. The microbiological properties that represent C and N dynamics are microbial biomass and soil enzymatic activity.
Microbial biomass (MB) is a labile fraction of soil organic matter sensitive to changes in soil management [
9,
19]. Phosphorus and K fertilization can also change the MB. Huang et al. [
20] observed greater microbial abundance after two years of fertilization with high P rates (30 g P m
−2 yr
−1) compared to lower rates. The authors explained this increase in MB by an increase in soil organic carbon. A few studies have addressed the significant correlation between MB and N
2O fluxes [
9,
15,
21].
Most biochemical transformations are mediated by enzymes and influenced by soil management. In other words, the soil enzymatic activity is a sensitive indicator of land use change [
22,
23]. The determination of soil enzymatic activity allows an evaluation of the impact of management practices on soil microbiota and, consequently, on N mineralization and N
2O emissions.
Few studies have assessed the changes in soil C fractions promoted by soil management and their relationship with N
2O fluxes [
15,
24]. The soil particulate fractions of C and N are associated with the formation and stabilization of soil aggregates [
24,
25,
26]. Therefore, these fractions (>53 μm) are sensitive to land use change. They play an important role in nutrient cycling and, consequently, N
2O emission [
9]. Not many studies have related soil carbon fractions with N
2O emissions, but in a recent study, Sato et al. [
15] showed that a high humic acid concentration reduced N
2O emissions. The authors observed that in crop–livestock systems under no-tillage, with
Urochloa grasses, the highest organic carbon contents in soil microaggregates were associated with lower cumulative N
2O emissions.
Studies that correlate N
2O fluxes and soil properties are scarce [
9,
21], and no information about soil enzymatic activity and cumulative N
2O fluxes could be found, in particular for crop–livestock integration systems. Our main hypothesis is that biochemical properties, such as soil enzymes and SOM fractions, are indicators of the N
2O mitigation potential in diversified systems with forages and crops, such as integrated crop–livestock farming systems. These integrated agricultural systems represent a promising option for the Brazilian public policy of low-C agriculture to mitigate GHGs and global climate change. In this context, it is essential to deepen the understanding of the relationships between soil biochemical properties and N
2O emissions in long-term integrated crop–livestock systems.
The objective of this study was to evaluate cumulative N2O emissions and their relationships with soil biochemical properties in a two-year phase with soybean in a long-term experiment (26 years) with integrated crop–livestock farming systems fertilized with two P and K rates.
4. Materials and Methods
4.1. Site Description and Experimental Design
The experiment was carried out in a rainfed experimental area of Embrapa Cerrados, Planaltina, Federal District (15°39′ S, 47°44′ W; 1200 m asl). The climate was classified as Aw (tropical savannah), with a dry (winter) and rainy period (summer, October to April), according to Köppen–Geiger’s classification [
61]. Precipitation and temperature data from 2015 to 2017 are shown in
Figure 4.
The soil was classified as a clayey Oxisol based on Embrapa [
62].
The long-term experiment was initiated in 1991 on 40 × 50 m (2000 m2) plots. At that time, 5.8 Mg ha−1 of limestone was applied to achieve 50% base saturation. As a side dressing, 20 kg P2O5 ha−1, 50 kg K2O ha−1, and 60 kg N ha−1 were applied. In the first four years, soybean, maize, rice, and sorghum were cultivated in the area in a conventional system (disk and moldboard plow).
Four years after the implementation of the experiment, Andropogon gayanus was sown in the area. In the continuous pasture treatment, 20 kg ha−1 P2O5, 50 kg ha−1 K2O, and 60 kg ha−1 N were applied as side dressing. In the integrated crop–livestock treatment, no fertilizers were applied in the pasture phase, since enough residual fertilizer from the previous crop was still available for the growth of Andropogon gayanus. In the continuous crop treatment and the crop phase of the integrated crop–livestock system, gradual liming was applied according to the technical recommendations for each crop.
Between 1995 and 2013, the plots were fertilized with the full or half the recommended rate of potassium and phosphorus fertilizers. The experiment was laid out in a randomized block design (two blocks) with a 2 × 2 factorial arrangement (farming systems × P and K rates). The treatments (plots) consisted of continuous crops fertilized with half of the recommended P and K rates (CCF1), continuous crops with the full recommended P and K rates (CCF2), an integrated crop–livestock system with half the recommended P and K rates (ICLF1), and an integrated crop–livestock system at full recommended P and K rates (ICLF2).
The sequence of the crop systems throughout the 26 years is listed in
Table 6.
Soybean cv. BRS 8180 RR was sown in November 2015 of the 2015/2016 growing season and harvested in March 2016, and cv. NS 7200 RR in November 2016 of the 2016/2017 growing season and harvested in February 2017. The soil chemical properties of each treatment were analyzed in January 2016 (
Table 7).
Soybean seeds were inoculated with Bradyrhizobium japonicum and fertilization consisted of 115 kg P2O5 ha−1 and 100 kg K2O ha−1 in both growing seasons (2015/2016 and 2016/2017). The forage was fertilized with 10 kg N ha−1, 50 kg P2O5 ha−1, and 30 kg K2O ha−1.
4.2. Soil Sampling and Analysis
The soil was sampled at soybean flowering in January 2016 and January 2017 (0–10 cm layer) because our objective was to evaluate biochemical properties under optimal conditions of biological activity (rainy season). Five subsamples were taken per plot to blend one composite sample. Part of the samples was stored in an ice box and placed in a refrigerator (±4 °C) for posterior microbiological analysis, and the other part was dried at room temperature for chemical analysis.
The biological and chemical analyses measured soil total C and total N (TC and TN); soil particulate C and N fractions (PC and PN); mineral-associated C and N (MAC and MAN); soil microbial biomass C (MBC) and N (MBN); C and N microbial quotient (CMQ and NMQ); soil available nitrogen (AN); humic fractions of organic C; fulvic-acid (C-FA); humic acid (C-HA); humin (C-HUM); and the soil enzymes β-glucosidase (β-GLU), arylsulfatase (ARYL), and acid phosphatase (PHOS).
4.2.1. Soil Total C and N
The soil samples dried at room temperature were sieved (<2 mm/10-mesh) and ground and sieved through a 0.149 mm/100-mesh sieve. Soil TC and TN were determined in a CHNS/O analyzer (Perkin Elmer 2400 Series II).
4.2.2. Soil Particulate C and N Fractions
Carbon and N were determined by soil physical fractionation, as proposed by Cambardella and Elliott [
63], with modifications suggested by Bayer et al. [
64] and Bongiovanni and Lobartini [
65]. Soil samples dried at room temperature were sieved (<2 mm/10-mesh) and 20 g of soil was placed in glass tubes with 70 mL of 0.5% sodium hexametaphosphate and shaken (150 rpm) for 15 h in a horizontal shaker. The suspension was sieved through a 270-mesh sieve (<53 µm) and washed thoroughly by spraying with water. The soil retained in the sieve was oven-dried at 45 °C and ground for analysis of total C and N in a CHNS/O analyzer (Perkin Elmer 2400 Series II). The fractions MAC and MAN were calculated by the difference between TC and PC and between TN and PN, respectively.
4.2.3. Soil Microbial Biomass C and N
Microbial biomass carbon and MBN were determined by the fumigation/extraction method [
66,
67,
68]. Soil samples were sieved through an 8 mm sieve to remove roots and plant residues. Prior to fumigation, the sample moisture content was corrected to 80% of the soil retention capacity. The samples were divided into six subsamples of 20 g and incubated for seven days. Half of the samples were fumigated with ethanol-free chloroform for 24 h.
Carbon and nitrogen extraction was performed by adding 5 mol L−1 K2SO4 to fumigated and non-fumigated samples, which were shaken in a continuous shaker at 150 rpm for 40 min and then filtered. Ten mL of extracted solution and 1 mL of substrate were added to glass tubes with 1 mL of distilled water. Microbial biomass carbon and MBN were determined in a CHNS/O analyzer (Perkin Elmer 2400 Series II).
The MBC and MBN contents were determined as the difference between carbon and nitrogen contents in fumigated and non-fumigated samples using a correction factor of Kc = 0.35 and Kn = 0.54 for MBC and MBN, respectively, according to Joergensen [
69]. The MQC and MQN were calculated as the ratio of MBC or MBN by TC or TN, respectively, and expressed as percentages.
4.2.4. Chemical Fractionation of Soil Organic Matter
Soil organic matter was chemically fractionated using 0.1 mol L
−1 NaOH as the extractor (10:1). The fractionation products were the three fractions of soil organic matter: humic acid (C-HA), fulvic acid (C-FA), and humin (C-HUM), determined according to their solubility in basic and acid solution. The C-HUM precipitates in a basic solution (insoluble in basic pH). The fractions C-HUM and C-FA were obtained by extract acidification with 6 mol L
−1 HCl at pH 1.0. The precipitate (C-HUM) and supernatant (C-FA) were separated by centrifugation (4500 rpm) for 30 min [
70]. Total organic C was determined by humic oxidation of organic matter with potassium dichromate in sulfuric acid [
71].
4.2.5. Soil Available N
Soil available N was determined, as suggested by Gianello and Bremner [
72], by measuring the ammonia-N produced from soil organic N after steam sterilization of the soil with Na
3PO
4-borate buffer (pH 11.2). Two grams of soil were transferred to glass tubes with 25 mL of pH 11.2 Na
3PO
4-borate buffer solution (200 g Na
3PO
4·12H
2O + 50 g borax in 2000 mL of distilled water), 0.23 g MgO, 0.1 g Devarda’s alloy, and 10 drops of dimethicone (to reduce foam formation), and the tubes were placed in a micro-distiller.
The distillate was filled into a 50 mL volumetric flask containing 10 mL of 0.05 mol L−1 HCl. For the calculations, we used a calibration curve obtained by distillation of standard N solutions containing 0, 10, 25, 50, 75, and 95 μg N mL−1. The extracted N was determined with a spectrophotometer (Tecnal SP 2000 UV) at 440 nm.
4.2.6. Soil Enzymes
The activities of the soil enzymes acid phosphatase (PHOS), β-glucosidase (β-GLU), and arylsulfatase (ARYL) were determined according to Tabatabai [
73]. The methodology is based on the colorimetric determination of p-nitrophenol released after soil incubation at 37 °C for 1 h in a specific substrate. For each sample, two analytical repetitions plus a control (no enzyme addition) were used.
The soil samples separated to determine the β-glucosidase and arylsulfatase activities, previously stored at 4 °C, were dried and sieved, as proposed by Lopes et al. [
74]. To determine the amount of p-nitrophenol released from the samples, a standard curve was prepared with known p-nitrophenol concentrations (0, 10, 20, 30, 40, and 50 µg p-nitrophenol mL
−1). The enzyme activities were expressed in µg p-nitrophenol h
−1 g
−1 soil.
4.2.7. Nitrous Oxide Sampling and Analysis
The N
2O fluxes were evaluated during soybean growth in the growing seasons of 2015/2016 and 2016/2017. In the total evaluation period, 78 samplings were carried out in 603 days [
27]. After sowing and N fertilization, N
2O fluxes were measured for up to five consecutive days. After rainfall, tilling, or harvesting, N
2O fluxes were measured for 2–3 consecutive days. During the dry season, N
2O fluxes were evaluated every 15 days. Several calibration measurements were carried out in the same experimental area [
15,
27].
The N
2O fluxes were measured in closed static chambers using the methodology described by Sato et al. [
75]. On each plot, two static chambers were maintained from November 2015 to July 2017, one installed in the planting row and the other in between the rows. Each chamber consisted of a rectangular hollow metal frame (38 × 58 × 6 cm) inserted to a depth of 5 cm into the soil. The top of each chamber was covered with a polyethylene tray coupled to the base during gas sampling. To avoid airtightness during N
2O sampling, soft rubber was added and both ends were fixed to the metal base. A three-way Luer valve was attached to the top part of the tray to fasten the syringes for gas sampling. The N
2O samples were collected in 60 mL polypropylene syringes and transferred to 20 mL pre-evacuated glass vials.
The N
2O concentration was determined by gas chromatography (Thermo Scientific Model Trace 1310, Milan, Italy), with “Porapak Q” columns comprising a backflush system connected to an electron capture detector. The N
2O fluxes (FN
2O) were measured based on the linear variation in gas concentration in relation to incubation time in the chambers, calculated with the following equation:
where δC/δt is the change in N
2O concentration in the chamber during the incubation period; V and A are the chamber volume and the covered soil area, respectively; M is the molecular weight of N
2O; and Vm is the molecular volume at sampling temperature.
Cumulative emissions were estimated by plotting the mean N
2O fluxes and time scale on a graph and calculating the resulting area under the integration curve using Sigmaplot
® Version 10 (Systat Software Inc., Chicago, IL, USA, 2007). The cumulative N
2O fluxes from each plot were estimated by the integrated trapezoidal area of the daily N
2O flux over time, considering that fluxes change linearly between measurements [
76].
4.3. Statistical Analysis
The data were tested for normality of residuals and homogeneity of variances and then subjected to one-way ANOVA using the R program.
The effects of farming systems on cumulative N2O fluxes were compared using Tukey’s test (p < 0.05). In the model, a nested design was assumed for each land use. The data on chemical and microbiological soil properties were statistically analyzed by the Tukey test (p < 0.05). The interaction between years of evaluation was not analyzed due to the drought stress that affected the 2016/2017 growing season.
Prior to principal component analysis (PCA), measures of sampling adequacy were calculated by the Kaiser–Meyer–Olkin (KMO) and Bartlett’s sphericity tests (
p < 0.05). When the KMO was higher than 0.5–1.0, the PCA application was considered acceptable [
77]. Bartlett’s sphericity test checks the independence of the tested variables. Principal component analysis was performed to group the dataset into new variables that resume the information in principal components (PC). The analysis also helps to avoid multicollinearity between the original variables. Principal component analysis (PCA) was performed using the R program and was applied to a data matrix with 16 rows corresponding to the cultivation systems (CCF1, CCF2, ICLF1, and ICLF2 in four repetitions) and 17 columns representing soil properties (TC, TN, PC, PN, MAC, MAN, MBC, MBN, MQC, MQN, AN, C-FA, C-HA, C-HUM, β-GLU, ARYL, and PHOS) to identify which properties most affected cumulative N
2O fluxes, represented by cumulative emissions.
5. Conclusions
The TC values were highest in ICLF2 in both growing seasons, and the particulate and mineral-associated fractions were highest in 2016 and 2017, respectively. Also, the microbial biomass fraction was highest in ICLF2 in both years. In terms of enzyme activity, ICLF1 and ICLF2 stood out with higher acid phosphatase and aryl sulfatase contents.
The soil properties correlated with cumulative N2O emission were TC, TN, PN, AN, MBC, and MBN. The integrated systems induced C storage in more stable fractions and reduced available N, decreasing N2O emissions and promoting lower cumulative N2O emissions, which could mitigate N2O.
The production systems ICLF1 and ICLF2 were more closely correlated with soil enzymatic activity (β-glucosidase, arylsulfatase, and acid phosphatase), particulate fraction of soil organic matter, and mineral-associated organic carbon in both growing seasons, whereas enzyme activity was not correlated with N2O emissions. In addition, ICLF2 promoted an accumulation of C in the more stable soil organic matter (SOM) fractions, which are unavailable to the microorganisms and resulted in lower N2O emissions.