Spatial variability and biophysicochemical controls on N2O emissions from differently tilled arable soils
Mohammad Mofizur Rahman Jahangir, Dries Roobroeck, Oswald Van Cleemput & Pascal Boeckx Biology and Fertility of Soils Cooperating Journal of International Society of Soil Science ISSN 0178-2762 Volume 47 Number 7 Biol Fertil Soils (2011) 47:753-766 DOI 10.1007/s00374-011-0580-2
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Author's personal copy Biol Fertil Soils (2011) 47:753–766 DOI 10.1007/s00374-011-0580-2
ORIGINAL PAPER
Spatial variability and biophysicochemical controls on N2O emissions from differently tilled arable soils Mohammad Mofizur Rahman Jahangir & Dries Roobroeck & Oswald Van Cleemput & Pascal Boeckx
Received: 6 February 2011 / Revised: 26 April 2011 / Accepted: 3 May 2011 / Published online: 12 May 2011 # Springer-Verlag 2011
Abstract Nitrous oxide (N2O) emissions, soil microbial community structure, bulk density, total pore volume, total C and N, aggregate mean weight diameter and stability index were determined in arable soils under three different types of tillage: reduced tillage (RT), no tillage (NT) and conventional tillage (CT). Thirty intact soil cores, each in a 25×25-m2 grid, were collected to a depth of 10 cm at the seedling stage of winter wheat in February 2008 from Maulde (50°3′N, 3°43′W), Belgium. Two additional soil samples adjacent to each soil core were taken to measure the spatial variance in biotic and physicochemical conditions. The microbial community structure was evaluated by means of phospholipid fatty acids analysis. Soil cores were amended with 15 kg NO3−-N ha−1, 15 kg NH4+-N ha−1 and 30 kg ha−1 urea-N ha−1 and then brought to 65% water-filled pore space and incubated for 21 days at 15°C, with regular monitoring of N2O emissions. The N2O fluxes showed a log-normal distribution with mean coefficients of variance (CV) of 122%, 78% and 90% in RT, NT and CT, respectively, indicating a high spatial variation. However, this variability of N2O emissions did not show plot scale spatial dependence. The N2O emissions from RT were higher (p< 0.01) than from CT and NT. Multivariate analysis of soil properties showed that PC1 of principal component analysis had highest loadings for aggregate M. M. R. Jahangir (*) Department of Civil, Structural & Environmental Engineering, Museum Building, Trinity College Dublin, Dublin 2, Ireland e-mail:
[email protected] M. M. R. Jahangir : D. Roobroeck : O. Van Cleemput : P. Boeckx Laboratory of Applied Physical Chemistry (ISOFYS), Ghent University, Coupure Links 653, 9000 Ghent, Belgium
mean weight diameter, total C and fungi/bacteria ratio. Stepwise multiple regression based on soil properties explained 72% (pCT>NT. Keywords Spatial heterogeneity . N2O emissions . Aggregate distribution . Denitrification . Microbial community structure
Introduction Nitrification, denitrification and nitrifier-denitrification can generate N2O, a highly potent greenhouse and ozonedepleting gas (Prather et al. 2001). In 2005, the atmospheric concentration of N2O was 319 ppb (10–9 mol mol−1) (IPCC 2007a). Globally, agricultural N2O emissions increased by nearly 17% between 1990 and 2005 (IPCC 2007b). Annual emissions of N2O are estimated at 17.7 Tg N (6.7–36.6 Tg N), about 60% of which is emitted from soils (Ehhalt et al. 2001). However, reliable predictions of N2O emissions from agricultural and natural soils are still not possible (Yanai et al. 2003). Prediction of N2O emissions from agricultural fields must take into account spatial variability because it is generally accepted that emissions within a small area may vary by orders of magnitude (Mathieu et al. 2006). Several studies documented the variability of N2O emissions in different ecosystems, but it has rarely been evaluated within an ecosystem (Van den Heuvel et al. 2009). There is a growing consensus to adopt conservation tillage practices instead of conventional tillage (CT) because of the impact on soil quality and production costs.
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Moreover, concerns with conventional tillage have been raised for water and air quality (Elmi et al. 2003). Gregorich et al. (2005) proposed conversion from conventional tillage to no tillage (NT) to mitigate N2O emissions, but contrasting results have been reported after changes to minimum or no tillage (Venterea et al. 2005; Beheydt et al. 2008) with lower (Baggs et al. 2003; Drury et al. 2006), equal (Kaharabata et al. 2003; Lemke et al. 1999) and higher N2O emissions (MacKenzie et al. 1997; Ball et al. 1999; Rochette et al. 2008) in NT than in CT. Therefore, the response of N2O emissions under different types of tillage is still in debate and requires a more precise estimation. However, the spatial variability of emissions within individual tillage system poses problem in measuring emissions in specific area of the field either by closed chamber or by incubating soil cores. The spatial variability of soil properties in differently tilled soils can give insights of drivers of spatially different N2O emissions (Choudhary et al. 2002). It is important to determine the spatial variance of biotic and physicochemical conditions in differently tilled field soils, together with the concurrent variability in N2O emissions. Such knowledge will allow better integration of biophysicochemical controls in an accurate prediction of N2O emissions. Contrasting results on some important drivers (e.g. soil pH, bulk density) of N2O emissions have been reported by several authors (Ruser et al. 2006; Simek et al. 2006). However, some drivers such as soil aggregate size fraction (Wang et al. 2008) and microbial community structure (Philippot and Hallin 2005; Calderon et al. 2001) need to be explicitly investigated at the same spatial scale (Van den Heuvel et al. 2009). Our hypothesis is that reduced tillage alters the spatial heterogeneity of soil aggregate mean weight diameter, total C content and microbial community compositions and this influences the spatial variability of N2O emissions. The objectives of the present research were (1) to investigate the spatial variance of N2O emissions from conventionally (CT), reduced (RT) and no tilled (NT) arable soils and (2) to assess the covariance between the spatial variability of (a) the microbial community structure and N2O emissions and (b) soil physicochemical conditions and N2O emissions.
Biol Fertil Soils (2011) 47:753–766
100 years and was converted from CT; mouldboard ploughing to 30 cm depth and harrowing of the top 10 cm to RT; harrowing of the top 10 cm in 1995. In September 2006, one third of the field was re-converted to CT, another third to NT with direct seeding, and the other third was kept in RT. These management practices have been continued in the respective areas of the field. Details of crops grown (winter wheat-green manure-maize) and agricultural management practices were given in Table 1. Sampling was performed on February 19, 2008. Soil core sampling and incubation conditions Intact soil cores (7.1 cm ID and 10 cm height) were collected from a depth of 0 to 10 cm by manually driving PVC cylinders into the soil. Ten soil cores, each in a grid of 25×25 m2, were collected per tillage treatment. This gave a total of 30 cores from three tillage treatments. Soil cores were immediately stored and transported to the laboratory in a cool box. For experiments, all soil cores were maintained at 65% water-filled porosity space (WFPS) throughout the experiment on a weight basis. The initial WFPS of the intact soil cores was determined based on gravimetric water content, bulk density and total pore volume of a replicate core taken adjacent to each experimental soil core. The cores received the following fertilizers: 15 kg NO3−-N ha−1 as KNO3, 15 kg NH4+-N ha−1 as NH4Cl and 30 kg Urea-N ha−1. All nutrients were in liquid form and sprayed uniformly on the surface of core by plastic syringe. Even distribution of N solutions through the entire core from a preliminary analysis was achieved by such application process. Soil cores were incubated in a closed 2-L leak-tight glass jar for day 0 (1.5 h) and days 1, 3, 5, 7, 10, 15 and 20 after fertilizer applications. The soil cores were kept at 15°C (yearly average soil temperature) in a dark incubator throughout the study period, including the 1.5-h incubations. A 12-mL headspace gas sample was collected 0.5, 1 and 1.5 h after headspace closure using an air-tight syringe and stored in an evacuated exetainer until measurement. The N2O concentrations were determined using a gas chromatograph (SHIMADZU 14B, Tokyo, Japan) connected to an electron capture detector. Physical and chemical soil properties
Materials and methods Experimental site and agricultural management The study was conducted in an agricultural field in Maulde, Belgium (50°37′N, 3°34′E). Soils, moderately well-drained silt loam, contained 17.6% sand, 66.3% silt and 16.1% clay and classified as a Luvisol (FAO 2006). The site has a slope of ca 3%. The field site has been cropped for more than
Soil bulk density was determined by the core sampler method using soil cores collected in Kopecky rings (5 cm ID×5 cm height) at 2.5 to 7.5 cm depth, adjacent to the soil cores collected for the N2O measurement. This gave a total of ten Kopecky rings per tillage treatment. The WFPS was determined as described by Liu et al. (2007). Total pore volume was calculated as 1−(bulk density/particle density; particle density=2.65 gcm−3). Aggregate size distribution
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Table 1 An overview of management practices carried out during 1.5 years time Date
Activities
13 September 2006
Tillage RT: harrowed only up to 10 cm NT: killing cover crop HT: ploughed thoroughly by mouldboard plough and harrowed All fields: sowing winter wheat Fertilization: liquid fertilizer (200 Lha−1) sprayed over the crop as a mixture of 30 kg urea-N ha−1; 15 kg NH4+-N ha−1 and 15 kg NO3−-N ha−1 Fertilization: liquid fertilizer (200 Lha−1) sprayed over the crop as a mixture of 30 kg urea-N ha−1; 15 kg NH4+-N ha−1 and 15 kg NO3−-N ha−1 Harvesting: four different spots of each of 1 m2 were sampled in RT Harvesting: four different spots of each of 1 m2 were sampled in NT Harvesting: four different spots of each of 1 m2 were sampled in HT Organic fertilization: pig manure 32,786 Lha−1; 150 kg ha−1 N, 136 kg ha−1 C; Tillage: RT: harrowed only up to 10 cm NT: killing cover crop and leaving the residues on soil surface HT: ploughed thoroughly by mouldboard and harrowed All fields: sowing green manure 58 kg ha−1 oat; 2 kg ha−1 mustard and 2 kg ha−1 Phacelia Soil sample collection with cores, Kopecky rings and auger for incubation and other parameters. Total N input over the whole management period was 270 kg Nha−1
14 October 2006 04 April 2007 26 April 2007 27 31 01 04
July 2007 July 2007 August 2007 September 2007
12 September 2007 19 February 2008
was determined by sieving 250-g soil over a sequence of sieves ranging from mesh size 4.8, 2.8, 2.0, 1.0, 0.5 to 0.3 mm as described by De Leenheer and De Boodt (1967). Aggregate mean weight diameter (MWD), a measure for soil aggregation, was computed as follows: 0 1 MWD ¼
n B X C B m i C » di n @P A i¼1 mi
ð1Þ
i¼1
where mi and di are respectively the mass and the mean diameter of aggregate size class i. Aggregate stability index (SI) was obtained by determining the mean weight diameter of air-dried soil samples and that of soil samples sieved after being dispersed in water on a planetary shaker for 5 min at 25 rpm. The aggregate stability index was computed as: Aggregate SI ¼
1 MWDðair dryÞ MWDðdispersedÞ
ð2Þ
Soil inorganic N was extracted by 1 M KCl solution of 1:2 (w/w) soil:solution ratio and analysed colorimetrically using a continuous flow auto-analyzer (AA3, Bran & Luebbe, Norderstedt, Germany). Soil pH was measured on the KCl extracts using a glass electrode (pH 3310, Jenway, UK). Total C (percent) and total N (percent) were determined on dry bulk soil samples which were ground with a planetary ball mill (PM400, Germany) and analysed
with an Automated Nitrogen Carbon Analysis-Solids and Liquids (ANCA-SL, PDZ-Europa, Northwick, UK) coupled to an Isotope Ratio Mass Spectrometer (20–20, SerCon, Crewe, UK). Microbial community structure Thirty additional soil samples collected by an auger, adjacent to the soil cores collected for N2O analysis, were used for the microbial analysis. Soil microbial community composition was determined by extraction, transesterification and quantification of PLFAs following the method described by Denef et al. (2009). Phospholipids were methylated by mild alkaline methanolysis (using methanolic KOH) to form fatty acid methyl esters (FAME). The FAME"s were identified and quantified by a capillary gas chromatography-combustion-isotope ratio mass spectrometry (GC-c-IRMS; Delta PLUS XP, Thermo Fisher Scientific Inc., Waltham, USA) via a GC/C III interface. On average, 25 peaks were detected of which 18 were biomarker fatty acids (Frostegard and Baath 1996; Zelles 1997; Stahl and Klug 1996; Boon et al. 1977). For each fatty acid the molar C percentage (Mc%) was calculated as: Mc% ¼
½PLFA Ci 100 n P ½PLFA Ci
i¼1
ð3Þ
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where [PLFA-C]i is the concentration of a fatty acid in PLFA-C L−1. The molar C percentage of a fatty acid biomarker is indicative for the relative abundance of a microbial group.
Biol Fertil Soils (2011) 47:753–766 Table 2 Mean, coefficient of variation (CV) and level of significance of cumulative N2O emissions (milligrams of nitrogen per square meter) during the entire incubation period in intact soil cores from three different tillage treatments Tillage
Statistical and geostatistical analysis The coefficients of variation (CV), computed as the ratio of the standard deviation to the mean, was used as indicator for the spatial variability of a parameter. The normality of the data distribution was checked a priori using the Shapiro–Wilk test (p1.0 mm showed a positive correlation with N2O emissions while aggregate sizes
