Labile soil organic matter pools under a mixed grass/lucerne pasture and adjacent native bush in Western Australia.
Much of the wheat produced in Western Australia (WA) is grown on coarse-textured soils (>90% sand). Typically, it is grown in a 3- or 4-course rotation including a break crop (e.g. canola) and annual/perennial grasses with leguminous pasture species. Prior to clearing in the 1950s Proteaceae-dominated woodland and shrubs covered much of the WA wheat-belt. Agricultural practices, such as cultivation, the use of fertilisers, and the establishment of grass/legume pastures have enhanced soil fertility in some parts of WA (Grigg et al. 2000; Mendham et al. 2002), but detrimental effects have also been reported. These include increased compaction, reduced infiltration rates, and decreases in soil organic matter (SOM) in cultivated soils (Abbott et al. 1979; Standish et al. 2006).
Grigg et al. (2000) examined the impact of agricultural management practices on plant production and species diversity in an adjacent area of banksia-dominated native bush at Moora in the WA wheatbelt. This work indicated that the incursion of water and nutrients (especially N) from agricultural land enhance nutrient loads in the adjacent native bush, resulting in enhanced biomass production of banksia trees at the edge of the bush area and decreased diversity and density of native woody plant species. While some of this N enrichment may have been a result of the movement of fertiliser N from agricultural land soon after application, some of it may have been the result of the movement of N derived from the decomposition and subsequent mineralisation of N-rich organic matter.
It is widely accepted that SOM plays a key role in C, N, S, and P cycling and also acts to improve soil structure. Agricultural practices and plant inputs influence both the quantity and quality of SOM, which directly affects soil productivity and its resilience and sustainability. In addition to a substantial proportion of recalcitrant organic matter (Jenkinson 1988), SOM includes soluble and other labile organic matter fractions involved in short-term (1-5 years) changes within the soil. Some of these labile fractions can be isolated (sequentially) using physical fractionation protocols based on density separation. Organic material of low density (< 1.6 g/[cm.sup.3]) isolated in this way is often referred to as the 'free light fraction' (Sollins et al. 1984; Sohi et al. 2001) and is derived from relatively recent plant residue inputs. Golchin et al. (1994) reported that that stability of soil structure is more dependent on these 'active' SOM fractions than the total organic matter. Therefore, the application of advanced analytical techniques such as [sup.13]C CP/MAS NMR to chemically characterise the carbon in whole soils and light fractions may provide valuable information on the effect of changes in land use and management on SOM (Mendham et al. 2002).
Recent studies indicate that soluble organic matter is another relatively labile component of SOM (Murphy et al. 2000; Marschner and Kalbitz 2003) and it has been identified as a key pool in controlling soil nutrient dynamics, especially C and N turnover (Jones et al. 2004; Cookson et al. 2005). Distinctions between the conceptual pools of soluble organic matter (Kalbitz et al. 2000), and the different methodologies used to measure them (Jones and Willett 2006), have resulted in a range of definitions for soluble organic matter in the literature. In this work we measured water-extractable organic C and N (WEOC, WEON) in soil under contrasting land uses. Dissolved organic C (DOC) and N (DON) were measured in soil solutions.
The primary aim of this work was to investigate the extent to which the amount, distribution, and composition of several labile SOM fractions, including plant litter, light fractions, WEOC, WEON, DOC, and DON differed under a grass/lucerne pasture and adjacent native bush at Moora, WA. In addition, we examined the effect of these land uses on the total amount and distribution of both C and N in the soil profile and on N[O.sub.3]-N and N[H.sub.4]-N (SMN) in both soil solution and water extracts.
Materials and methods
The study site comprised an area of flat land containing remnant bushland ('native bush') and an adjacent grass/lucerne pasture ('pasture') established on land cleared in 1962 (Grigg et al. 2000; Ward et al. 2003) near Moora, in the WA wheatbelt (30[degrees]34'38"S, 115[degrees]52'33"E). The region is subject to a Mediterranean-style climate with hot dry summers and cool wet winters. Average annual rainfall at Moora is 463 mm, of which 369 mm fails during the winter cropping period (1 May-31 October). The soil consists of highly uniform siliceous sand to a depth of at least 4.0 m, and is classified (USDA Soil Taxonomy) as a Dystric Xerosamment (McArthur 1991). The surface soil (0-0.15 m) is acidic (pH 5.0) and contains about 4% clay, 0.34).6% C, and 0.02--0.05% N.
The native vegetation was dominated by Banksia prionotes Lindley, which grows to a height of ~4 m, and represents approximately 60% of the total biomass. Grigg et al. (2000) discuss botanic composition in more detail. In August 1998, lucerne (Medicago sativa) was sown on a 200 by 200 m block of cleared land 20 m to the south of the banksia woodland, following application of glyphosate (720 g/ha) on 30 July and a mixture of diquat (150g/ha) and paraquat (250g/ha) on 26 August. Fertiliser, including P (10 kg/ha), K (18 kg/ha), and S (13 kg/ha), was applied within 2 weeks of sowing. The lucerne was heavily grazed in April 1999 and again in August 1999. A mixture of diquat (112 g/ha) and paraquat (187 g/ha) was applied on i 6 June 1999 to control radish (Raphanus raphanistrum) and capeweed (Arctotheca calendula). By August 2000, the lucerne pasture contained a significant amount (about 20-30% of plant ground cover) of grass weed. Prior to 1998, the pasture supported poor annual pastures, with occasional annual crops. Further details of previous cropping and fertiliser inputs are given by Ward et al. (2003) and Anderson et al. (1998a).
In August 2000 soil samples were collected from each of 3 separate locations situated well within the native bush block and grass/lucerne pasture, respectively. The upper soil layers (0--0.15, 0.15--0.35, 0.35--0.65m) were sampled using a 100-mm-diameter sand-auger, and subsoils (0.65--1.00, 1.00--1.40, 1.40--1.50m) were collected using an 80-mm-diameter auger to avoid contamination with surface soil. The midpoint of each sampled layer corresponded closely to the depth at which several quartz-Teflon porous cups had previously been inserted (Ward et al. 2003). Two soil cores were taken at each sampling location. The cores were combined in pairs for each soil layer. To account for the possible effects of the tree canopy and roots on the organic matter content and nutrient distribution in the soil at the native bush site, 1 core was taken under the banksia canopy, close to the foot of a tree, while the second was taken further away, between the tree canopies. Soils were sealed in polythene bags and stored in a cold room (4[degrees]C) before mixing, sieving (<4 mm), and subsampling.
Prior to soil sampling, leaf litter under the native bush was collected from 12 quadrats (each 0.2 by 0.4m) at each of the 3 separate sampling locations. Pasture plant material, including some roots, was taken by hand from 4 quadrats (each 0.2 by 0.4 m) at each of the 3 sampling location. The plant material from each quadrat was combined to give 3 separate samples for each land use. All plant material was dried (35[degrees]C for 72 h) and ground (<2 mm) before analysis.
Soil organic matter fractionation
The light organic matter fraction of density < 1.0 g/[cm.sup.3] (LF 1.0) was isolated from dry (35[degrees]C for 72h) soils (0--0.15m) by extraction and flotation in de-ionised water using a method similar to that described by Osier and Murphy (2005). Briefly, 4 subsamples of soil, each of 37.5 g, were taken from each sample bag. The subsamples were extracted by shaking with 100 mL of water for 1 h. The resulting mixture was centrifuged at 1713g for 10 min. The floating organic material was removed using a vacuum line and collected by filtration on a pre-weighed 20-[micro]m nylon net (Millipore). It was then rinsed with de-ionised water. The filtered water was returned to the centrifuged soil and the exaction procedure was repeated to ensure that all of the LF 1.0 was removed. After the LF 1.0 was removed, the soil was extracted again by shaking with 100 mL of an aqueous solution of Nal adjusted to a density of 1.7 g/[cm.sup.3] (Sollins et al. 1984) to isolate LF 1.7 (1.0-1.7 g/[cm.sup.3]). The LF 1.7 was collected on a 20-[micro]m nylon filter and rinsed with de-ionised water. The soil was extracted twice to ensure all of the LF 1.7 was recovered. The light fractions (LF 1.0 and 1.7) extracted from each of the 4 subsamples were dried (40[degrees]C for 72 h) and weighed separately. Dried subsamples were removed from the nylon mesh and combined so that a single sample of each fraction (LF 1.0 and 1.7) was obtained for each soil sample. The light fractions were stored in sealed glass vials before analyses of total C and N, and characterisation by [sup.13]C CP/MAS NMR.
Measurements of WEOC and WEON were done on freshly sieved soils (<4mm) from the 0-0.15, 0.15-0.35, 0.35-0.65, and 0.65-1.00 m layers only. For each soil layer, 4 subsamples, each of 15 g, were extracted for 1 h on a rotary shaker with 40 mL of de-ionised water. The resulting mixture was centrifuged for 15 min at 2332g. The soluble and paniculate organic matter in the supematant was separated by Millipore filtration under vacuum through a Whatman GMF 2.0-[micro]m glass-fibre filter. Filtrates (<2.0 [micro]m) were stored frozen in acid-washed polyethylene vials for 12 days before further filtrations of a 10-15 mL aliquot of the original filtrate through a 0.22-[micro]m Millipore cellulose-nitrile filter under vacuum. The filtered extracts were stored frozen before analyses of WEOC and WEON. Soil moisture contents (0-1.50 m) were determined gravimetrically by drying at 105[degrees]C for 24 h. Measurements of both WEOC and WEON in each soil layer were adjusted for soil moisture content.
Soil solution sampling
Immediately after soil sampling, soil solutions were collected from quartz-Teflon suction cups inserted at depths of 0.25, 0.50, 0.80, 1.20, and 1.50 m under both the native bush block and the grass/lucerne pasture. Further details of the porous cup installation are given by Ward et al. (2003). The suction cups were evacuated to a partial vacuum of about 50kPa, using an electric vacuum pump, about 4-5 h before sampling. Sample volumes collected ranged from <1 to 30 mL (mean of 13.5 mL). In order to avoid 'hedge' effects (Grigg et al. 2000), soil solutions were collected from about 13 suction cups located about 200 m from the agricultural boundary at each of 3 sampling locations (37 samples in total) in the native bush block. The suction cups were situated close to banksia trees, and between the trees, to account for the effects of tree root distribution on soil solution composition. In addition, soil solutions were collected from 8 or 9 suction cups at each of 3 separate locations (25 samples in total) within the pasture, about 50 m from the edge of the block. Soil solutions were stored frozen (-15[degrees]C) before dilution (range 0-37-fold; mean of 3.8) with de-ionised water and subsequent analyses for DOC, DON, and SMN.
Dried soil (35[degrees]C for 17 days), plant litter, and light organic matter fractions (LF 1.7 and LF 1.0) were finely ground in a disk mill (Tema T100) before determination of total C and N, and [sup.15]N enrichment, by combustion using an automated N and C analyser linked to a mass spectrometer (ANCA-MS, Europa Scientific Ltd, Crewe, UK). The [delta][sup.15]N enrichments of the samples were calculated as shown in Eqn 1:
[delta][sup.15]N = 1000 x ([A.sub.s] - [A.sub.a])/[A.sub.a] (l)
where As is sample atom% [sup.15]N and [A.sub.a] is atom% [sup.15]N of air (0.3663 atom% [sup.15]N).
Total C and N concentrations in soils below 0.65 m were close to the limits of detection. Consequently, only data for the 0-0.15, 0.15-0.35, and 0.35-0.65 m layers are presented. Total C and N in whole soil, light fraction and extractable organic matter were calculated using soil bulk densities of 1.47, 1.64 and 1.78g/[cm.sup.3] for the 0-0.15, 0.15 0.35 and 0.35-1.50m soil layers respectively. Determinations of total C and N in milled plant material and light fractions were corrected for moisture by drying at 80[degrees]C for 18 h before analyses. Calculation of plant litter dry weight (t/ha) and its C and N content (kg/ha) under native bush were done assuming that plant stem areas were negligible. The DOC and WEOC concentrations in soil solutions and filtered soil water extracts were determined using a total oxidisable-C analyser (Shimadzu TOC-5000A, Japan). The corresponding SMN concentrations were determined colourimetrically on an automated flow injection auto-analyser (Skalar Analytical B.V., The Netherlands; Henriksen and Selmer-Olsen 1970; Krom 1980). In addition, total dissolved N (TDN) in soil solutions and water extracts (excluding extracts filtered <0.22 [micro]m) were measured as N[O.sub.3]-N using the Skalar auto-analyser after alkaline persulfate oxidation of soluble organic N (Cabrera and Beare 1993). The DON and WEON in soil solutions and filtered soil extracts were calculated as the difference between TDN and SMN.
The [sup.13]C CP/MAS NMR spectra for plant litter and light organic matter fractions (LF 1.7 and 1.0) were determined as described by Sohi et al. (2001). Briefly, for each material, finely ground subsamples were packed into cylindrical zirconia rotors, each with internal dimensions of 5.6 by 17mm, sealed with Kel-F caps (3M Co., Minneapolis, MN, USA). The samples were analysed on a Bruker MSL 300 spectrometer (Bruker, Coventry, UK). Separate spectra were obtained for each of the 3 replicate samples (i.e. 1 spectrum per replicate) using a frequency of 75.5 MHz, a contact time of 1 ms, a relaxation time of 1 s, a spinning speed of 4.8 kHz with elimination of spinning side-bands using the Total Suppression of Sidebands (TOSS) sequence (Dixon 1982), and line broadening of 50 Hz. The samples were scanned for a period of 3-16h. Chemical shift values were measured with respect to tetramethylsilane. We used the Bruker WinNMR software to measure peak areas for the following chemical shift regions: 0-45ppm (alkyl), 45-65ppm (N-alkyl and methoxy), 65-95ppm (O-alkyl), 95-108ppm (acetal), 108-140ppm (unsubstituted and alkyl-substituted aromatic or aromatic), 140-160ppm (O-substituted aromatic or phenolic), and 160-220ppm (carboxyl, amide, ester, ketone and aldehyde or carboxyl). Actual spectral boundaries were the natural 'valleys' closest to the indicated chemical shift values.
GenStat[R]release 8.2 (VSN International Ltd, Hemel Hempstead, UK) was used to test for significant differences between means of corresponding variates under the different land uses. However, because both the pasture and native bush sites occupied 2 separate blocks of land it was not possible to fully randomise the plant and soil sampling strategy to take account of any systematic variation in soil or plant parameters over the whole study area (i.e. the combined area of both land uses). Therefore, the values obtained for each land use provide pseudo-replication only and it was not possible to take account of the variance due to blocking within subsequent analyses of variance.
Differences between means, including dry weights, total N, [delta][sup.15]N, total C, and C/N ratios of plant materials and light organic matter fractions (LF 1.0 and 1.7) were analysed using a 2-sample t-test. Mean concentrations of WEOC, WEON, DOC, and DON at each of the 3 main sampling locations within each land use were analysed separately using a 2-way analysis of variance (ANOVA) with land use and soil depth as the principal factors. Similarly, ANOVAS were done to examine the effects of land use and depth on soil total C and N, and moisture content. The effect of filtration (<2 [micro]m v. <0.22 [micro]m) on WEOC was examined using a 2-way ANOVA, with filter pore size and soil depth as the principal factors. The effects of land use on the proportion of C present in each of the functional groups measured in each of the labile organic matter fractions, as determined by [sup.13]C CP/MAS NMR of plant material and light fractions, was investigated using a 2-way ANOVA with land use and organic matter fraction as the principal factors.
C and N inputs in plant residues
The total mass of litter under the native bush system was on average more than double that in litter and plant material under pasture (Table 1). The former comprised largely banksia leaf litter, some of which may have dropped a year or two before sampling. In contrast, the latter contained a substantial proportion of fresh plant material in addition to some leaf litter. Leaf litter from the native bush system contained nearly 4 times more C than the pasture plant material, but only about half as much N (Table 1). Consequently, the mean C/N ratio of plant material and leaf litter under the pasture was much narrower than that under the native bush, 12 and 98, respectively (Table 1).
[FIGURE 1 OMITTED]
Whole soil and light fraction organic matter
Total C in soil (0~0.65 m) under pasture and native bush averaged 12.6 and 13.3 t/ha, respectively, but did not differ significantly between land uses (Fig. 1a). In both cases the majority (79-83%) was present in the surface (0-0.15 m) layer. In contrast, only 48-58% of the total soil N (0-0.65 m) under pasture and native bush was present in the surface (0-0.15m) layer (Fig. 1b). However, significantly (P < 0.05) more N was present in the surface soil (0-0.15 m) under pasture compared with that under native bush (0.8 v. 0.4t N/ha). Consequently, the C/N ratio of SOM in the surface soil under pasture was significantly less (P < 0.01) that under native bush (12 [+ or -] 0.49 v. 25 [+ or -] 0.93).
The combined dry weight of both light organic matter fractions (LF 1.0 and 1.7) in the surface soil (0-0.15 m) under the native bush was significantly (P < 0.05) greater than that under the adjacent pasture (12.3 v. 6.5t/ha) (Table 1). In both cases LF 1.7 accounted for the majority (64-68%) of this material (Table 1). The combined weight of both light fractions (LF 1.0 and 1.7) accounted for only 0.34).6% of the total soil mass (0-0.15 m) under both land uses, but contained 27% of the total soil C (0-0.15m) and 22% of the total soil N under native bush, compared with 18% and 17%, respectively, under pasture.
The C/N ratios of both light organic matter fractions (LF 1.0 and 1.7) in the pasture soil (0-0.15m) were significantly (P < 0.05) less than those under native bush (Table 1). This was consistent with the smaller C/N ratio of the plant material and leaf litter collected from the pasture system compared with that under native bush (Table 1), indicating that the light fractions were derived largely from plant residues (Golchin et al. 1995).
The [delta][sup.15]N of the native bush litter averaged -2.0[degrees][per thousand] and was significantly (P < 0.05) less than that measured in the pasture plant material, which averaged 1.8%o (Table 1). The [delta][sup.15]N of LF 1.7 was similar for both land uses and was smaller than that in LF 1.0, but it did not differ significantly from that in plant litter (Table 1) or whole soil (0-0.15 m), which had a [delta][sup.15]N of -2.1[per thousand] averaged over both land uses.
A comparison of the [sup.13]C CP/MAS NMR spectra for the materials obtained from pasture and native bush systems, including plant material, leaf litter inputs, and light organic matter fractions (LF 1.0 and 1.7), indicated that in most cases land use had little impact on the proportion of C in the different functional groups present in these materials (Table 2, Fig. 2a-f), despite their different C/N ratios (Table 1). The carboxyl and acetal groups were the 2 exceptions. On average, the C in the carboxyl groups comprised a significantly (P< 0.01) greater proportion of the total C in all of the organic materials collected from the pasture system, compared with those from the native bush (Table 2). In contrast, the proportion of C in acetal groups in pasture plant material was significantly smaller (P < 0.05) than in leaf litter from the native bush. However, in both systems LF 1.7 contained a significantly (P < 0.05) greater proportion of alkyl-C than that present in LF 1.0 and plant residues, and a significantly (P < 0.01) smaller proportion of O-alkyl-C (Table 2). The increase in the alkyl signal was mainly due to the methylene C in long chain structures, resonating around 30 and 33 ppm (Fig. 2a-f). The proportion of C in aromatic groups was significantly (P < 0.01) greater in both light fractions compared with plant material, and there was an indication that the LF 1.0 contained a greater (P < 0.05) proportion of phenolic groups than either LF 1.7 or plant material (Table 2).
Water extractable organic matter (WEOM)
Overall, WEOC concentrations measured in soil water extracts filtered <2.0 [micro]m did not differ significantly from those in extracts filtered <0.22 [micro]m. Therefore, in most cases particulate (0.22-2.0 [micro]m) organic C and N were considered negligible and only data for extracts filtered <2.0 [micro]m is presented. However, WEOC concentrations in the surface soil (0-0.15m) under native bush indicated that a significant (P < 0.05) proportion (c. 47%) of this material was in particulate (0.22-2.0 [micro]m) forms. The source of this material is unknown, but it may have been partly derived from the significantly greater (P < 0.05) pool of light fraction organic matter (LF 1.0 and 1.7) present in soil under the native bush (Table 1).
Mean WEOC (<2.0 [micro]m) concentrations in soils ranged from 2 to 38 mg C/kg (Fig. 3a). Concentrations declined significantly (P < 0.01) with increasing depth under both native bush and pasture, but did not differ significantly between land uses. Mean WEON concentrations ranged from 1 to 5 mg N/kg (Fig. 3b), but did not differ significantly between land uses or soil layers. Total amounts of WEON and WEOC in the soil profile (0-1.00m) under both systems averaged 56 and 140kg/ha, respectively. Mean C/N ratios of WEOM in the soil profiles ranged from 1 to 19, but did not differ significantly between land uses or soil depths.
Dissolved organic matter (DOM)
Overall, mean DOC concentrations in soil solutions collected from porous cups inserted at 0.25, 0.50, 0.80, 1.20, and 1.50m under pasture were significantly (P<0.001) greater than those under native bush (Fig. 4a). The differences were especially marked in solution collected at 0.25 and 0.50m, but decreased with increasing depth. In addition, there was a reasonable correlation ([r.sup.2] = 0.53) between DOC in soil solution collected at 0.25, 0.50, and 0.80 m under pasture and WEOC in filtered (<2.0 [micro]m) soil extracts from corresponding (0.15-0.35, 0.35-0.65, and 0.65-1.00 m) soil layers (Fig. 3a). In contrast, the corresponding correlation under the native bush system was poor ([r.sup.2] = 0.11).
[FIGURE 2 OMITTED]
Mean DON concentrations in soil solutions under the pasture ranged from 8 to 55 mg N/L and were in all cases significantly (P <0.05) greater than the 0.4-1.0mgN/L measured under native bush (Fig. 4b). The DON concentrations in soil solutions collected at 0.25 m under pasture were significantly (P < 0.05) greater than those in deeper soil layers, but there was no clear effect of depth on DON concentrations under native bush. The mean C/N ratios of DOM in soil solutions under the native bush (0-1.50 m) ranged from 11 to 27 and were significantly (P < 0.001) greater than those under pasture, which ranged from 1 to 4. However, measurements of organic C in porous cup solutions and soil extracts were sometimes close to the limits of detection. Similarly, in some cases measurements of total soluble N in soil solutions were only slightly greater than blank values. Consequently, the low C/N ratios (<5) measured in both DOM and WEOM, especially in subsoils, are subject to cumulative errors and should be regarded with caution.
Soil mineral N
Mineral N (N[H.sub.4]-N and N[O.sub.3]-N) concentrations in soil solutions, although very variable, were significantly (P < 0.001) greater under pasture than native bush (Fig. 5). In both cases, the majority of this mineral N was present as nitrate in most of the samples. In contrast, SMN concentrations measured in soil extracts were negligible (data not shown).
Soil mineral N, DOC, and DON concentrations in soil solutions under both land uses may have been influenced by soil moisture. In particular, the greater DON, DOC, and SMN concentrations measured in soil solutions collected at 0.25 m under the pasture compared with native bush may, in part, have been because the surface soil was significantly (P < 0.05) drier. However, DON and DOC concentrations in subsoil solution (below 0.25 m) were also greater under pasture than native bush, despite the fact that these soils were significantly (P < 0.05) wetter. Therefore, soil moisture was not the primary factor controlling C and N concentrations in soil solution. Overall, the pasture soil (0-1.50m) contained on average 89mm of moisture, compared with 77 mm under native bush, indicating that before sampling the bush vegetation may have utilised water in deeper soil horizons more effectively than the pasture (Ward et al. 2003).
[FIGURE 3 OMITTED]
[FIGURE 4 OMITTED]
[FIGURE 5 OMITTED]
Total soil C and N
The significantly greater N content of the pasture soil (0-0.15 m) compared with that under native bush was almost certainly a result of the accumulation of N in previous crop residue inputs and past N fertiliser applications. However, total C (0-0.65 m) did not differ significantly between land uses (Fig. la). In contrast, Dalal and Mayer (1986a) reported substantial decreases in both total C and N in cultivated soils compared with undisturbed soils from adjacent areas under native vegetation in southern Queensland. The apparent discrepancy between the findings reported here and those of Dalal and Mayer (1986a) may in part be due to the low C and N contents of the Moora soil together with differences in soil texture and climate compared with those in Queensland. The indigenous N contents of the deep sands and duplex soils found in south-west Australia are inherently low (0.01-0.03% of dry weight). Consequently, it is only through regular inputs of nitrogen fertilisers and/or biologically fixed N that crop production in this region can be sustained. Danso et al. (1988) reported that annual N fixation by lucerne grown in mixed lucerne-ryegrass swards in Austria was 104-108 kg/ha, 82-90% of the total plant N. In a study at 6 sites in south-west Australia, Unkovich et al. (1994) reported that at peak biomass production lupins contained 199-372 kg N/ha, and of this, 86% was derived from the atmosphere. Of this fixed N it was estimated that 32-96 kg/ha was returned to the soil in crop residues. However, it was considered that following losses through stubble grazing, erosion, and nitrate leaching, the N supplied by these residues would only be sufficient to support a modest increase in grain yield of the subsequent cereal crop. Anderson et al. (1998a) reported that N fixed by subterranean clover grown in a mixed pasture at Moora (WA) amounted to 29-162 kg/ha, similar to the 90-151 kg/ha fixed by lupins alone. They concluded that, although the inclusion of legumes in rotations with cereals was able to add sufficient N in residues to support above-average yields, the asynchrony between the soil N supply and crop demand resulted in the inefficient use of this N by the subsequent cereal crop. Thus, it seems that a significant proportion of the N input by legumes in pasture-wheat rotations may be at risk to loss over the summer and autumn when crops and pastures are not grown. Consequently, in the longer term, N enrichment of agricultural land may lead to enhance N enrichment of adjacent native systems, resulting in significant effects on their productivity and biodiversity (Anderson et al. 1998b; Grigg et al. 2000).
There is much evidence to indicate that, in general, N-rich plant residues with small C/N ratios (e.g. legumes) decompose rapidly and enhance net N mineralisation in soil (Waksman and Tenney 1927; Thorup-Kristensen 1994). In contrast, the incorporation of residues with wider C/N ratios and significant lignification is often associated with decreased net N mineralisation in soil and/or immobilisation (Powlson et al. 1985; Muller et al. 1988; Rahn and Lillywhite 2002). Therefore, in all probability, the smaller C/N of the plant residue inputs (Table 1) to the soil under the pasture systems enhanced net N mineralisation rates to a greater extent than those under native bush, which had a much greater C/N ratio and almost certainly contained a longer proportion of lignified material (Mendham et al. 2002). This is consistent with the work reported by Ward et al. (2003), which showed that between August and November 2000, net N mineralisation in the 0-0.20 m layer under lucerne pasture at the Moora site was 3 times greater than that under adjacent native bush. Some of this N may have been derived from the mineralisation of N fixed in the lucerne root nodules (Anderson et al. 1998a).
Light fraction organic matter
The significantly greater quantity of light fraction material (LF 1.0 + LF 1.7) present in soil (0-0.15 m) under native bush compared with that under pasture (Table 1) may be the result of the more rapid decomposition of these organic matter fractions following the clearance of native vegetation and cultivation of land. Dalai and Mayer (1986a) reported that the light fraction (<2 g/[cm.sup.3]) C content in soils cultivated for 20-70 years decreased by 25-67% compared with that present in soil under adjacent native vegetation in southern Queensland. The loss of C from the light fraction was 2-11 times more rapid than the heavier organic matter fractions (>2 g/[cm.sup.3]), and the difference in the rate of C loss between these fractions was greater in soils with larger clay contents (Dalai and Mayer 1986b). In addition, in some cases, plant residue inputs under native system may be greater than under agricultural systems, where crops are harvested or grazed. Low and Lamont (1990) reported a root/shoot ratio of 2.35, to a depth of 2.5 m, in a banksia dominated scrub-heath in WA. Rootstocks, laterals, and proteoid roots made major contributions to below-ground phytomass in the 0-0.15 m soil layer and litter contributed 19% of above-ground dead plus live phytomass, and was poorly decomposed.
The C/N ratio of both the light organic matter fractions present in soil under native bush and the corresponding litter inputs were greater that those under pasture, providing evidence to support the widely held view that the light fraction material is derived largely from partially decomposed plant residues (Sohi et al. 2001). Thus it seems the relatively recalcitrant nature of the plant residue returns under the native bush may also be partly responsible for the larger quantity of light fraction present in the soil. The greater density of LF 1.7 compared with LF 1.0 was probably a result of its closer association with the soil mineral component, because of its more advanced stage of decomposition (Hassink 1995a; Sohi et al. 2001). Although, in both systems the light fraction (LF 1.0 + LF 1.7) accounted for < 1% of the soil mass, it contained 18-27% of the total soil C and 17-21% of the total soil N content. This was broadly consistent with work reported by Dalai and Mayer (1986b), in which the light organic matter fraction (<2.0 g/[cm.sup.3]) in soils (0-0.10 m) cultivated for 20-70 years accounted for 15-32% of the total soil C, but only 1.8-3.2% of the soil mass. Thus, changes in land use and/or management which influence the light fraction content of the coarse-textured soils in WA have considerable potential to enhance or decrease their total C and N stocks.
Hassink (1995b) reported correlation coefficients of 0.75-0.94 for the relationship between the N content of macroorganic matter fractions (< 1.13, 1.13-1.37, and > 1.37 g/[cm.sup.3]) and net N mineralisation rates in arable soils, but correlations were weaker in grassland soils. Similarly, Barrios et al. (1996) found that the N content of the light organic matter (<1.13 g/[cm.sup.3]), but not dry weight or C content, was correlated with both anaerobic and aerobic N mineralisation in whole soil incubation studies. They also reported that the dry weight, and N and C content of the heavier fraction (<1.7 g/[cm.sup.3]) did not correlate well with whole soil N mineralisation. Cookson et al. (2005) reported that light fraction organic material (<1.0 g/[cm.sup.3]) accounted for only 9% of the potentially mineralisable N in coarse-textured grassland soils amended with NPK fertiliser and/or grass hay, and acted as a sink for mineral N. Thus, in this study, it seems unlikely that the mineralisation of the light fraction material (LF 1.0 and LF 1.7) alone could fully account for the apparently greater net N mineralisation rates in the pasture soil compared with that under native bush, as reported by Ward et al. (2003). Therefore, it seems more likely that it was a consequence of the larger N returns in plant residues with small C/N ratios (Table 1) and the long-term effects of N fertiliser applications on the accumulation of soil organic N (Glendining and Powlson 1995; Macdonald et al. 2002) rather than N mineralised from the 'light' organic matter fraction alone.
The similarities in the [sup.13]C CP/MAS NMR spectra for the organic materials obtained from the 2 different land uses indicated that this approach may be of limited value for differentiating between materials with different C/N ratios, and hence different decomposability. Decreases in the intensity of the O-alkyl-C signal coupled with an increase in the alkyl-C signal reflect the rapid decomposition of the O-alkyl-C present in plant polysaccharides during the decomposition of plant residues in soil, coupled with the relatively slow decay of organic matter rich in alkyl, aromatic, and carbonyl C (Skjemstad et al. 1997). Consequently, the O-alkyl:alkyl-C ratio is widely regarded as an indicator of the relative degree of decomposition of soil organic matter fractions (Mathers et al. 2000; Sohi et al. 2001; Chen et al. 2004). The similarity between the O-alkyl:alkyl-C ratios measured in LF 1.0 and recent organic matter inputs provided further evidence to indicate that LF 1.0 comprised largely plant material in the early stages of decomposition. The smaller O-alkyl:alkyl-C ratio of LF 1.7 (Table 2) indicates that it was at a more advanced stage of decomposition (Golchin et al. 1995).
Pate et al. (1994) reported that the [delta][sup.15]N of most non-[N.sub.2] fixing pasture species was greater than that found in [N.sub.2]-fixing species. However, in an earlier survey (Pate et al. 1993), which included non-fixing woody species, shrubs, and herbs, it was not possible to distinguish between [N.sub.2] fixing and non-fixing species using measurements of [delta][sup.15]N in plant shoots. In this study, the non-fixing grasses in the pasture plant material may have slightly increased its overall [delta][sup.15]N. In both cases the [delta][sup.15]N of LF 1.0 (Table 1) was significantly greater (P < 0.05) than that for plant litter and whole soil ([delta][sup.15]N -2.2, 0-15cm). This may be a result of differences in the isotopic composition of the readily decomposable and recalcitrant organic material in the plant residues, and/or isotopic discrimination against the heavier isotope by the microbial processes controlling residue decomposition. The [delta][sup.15]N of LF 1.7 was almost certainly influenced by its association with the more recalcitrant organo-mineral soil fractions. This may in part explain why it was between that of LF 1.0 (Table 1) and the whole soil (0-0.15 m).
DOM, WEOM, and SMN
Both DON and DOC concentrations in soil solution were greater under pasture than native bush. In contrast, land use had little impact on the soil WEOM content. Overall, WEON concentrations were similar to those measured by McNeill et al. (1998) in a loamy sand (0-0.10 m) under continuous wheat and pasture in the WA wheatbelt. In contrast, WEOC concentrations reported here are rather less than those measured by McNeill et al. (1998). This may have been in part because the latter were measured in the 0-0.10 m layer in February, during the summer fallow period. Our measurements were made in the 0-0.15 m layer, during the cooler and wetter winter period. The greater apparent effects of land use on DOM compared with WEOM may indicate that the former is linked more closely with dynamic C and N pools influenced by shorter term effects associated with changes in land use and management, while WEOM may be more closely linked with changes in the recalcitrant soil organic matter fractions. Thus, in soils of inherently low fertility, C and N dynamics in soil solution may be more sensitive indicators of the impact of changes in land use and management on labile soil organic matter pools than soil extracts. However, the reasonable correlation ([r.sup.2] = 0.53) between DOC in soil solution collected under pasture and WEOC in filtered (<2.0 [micro]m) soil extracts indicates a link between the soluble C present in the mobile soil solution and that in the whole soil. In contrast, the corresponding correlation under the native bush system was poor ([r.sup.2] = 0.11). This may have been because the pasture soil samples were taken closer to the porous cup locations than was the case under the native bush. It may also have been a consequence of non-equilibrium between WEOM and DOM under native bush.
The greater N enrichment of the whole soil and the light fraction organic matter under pasture compared with native bush was associated with a greater SMN concentration in soil solution, and may be indicative of enhanced N mineralisation under the pasture system. However, the difference was short lived, because although Ward et al. (2003) measured similar concentrations at the Moora site a few weeks later, there were no significant differences between land uses at this time. In contrast, SMN concentrations in whole soils under both land uses were negligible. This was consistent with the small concentrations (0.1-1.1 mg N/kg) measured (0-3.5 m) by Ward et al. (2003).
It is apparent from this work that some labile fractions of SOM (LF 1.0, LF 1.7, and surface litter) were smaller under pasture than native bush, but that others (soluble C and N in soil solution) were greater. The total soil C (0-0.65 m) did not differ between land uses, but the significantly greater total N content of the surface soil (0-0.15 m) under pasture provided evidence of N enrichment. In all probability this occurred as a consequence of the change in land use from native bush to agriculture, after the land was cleared in the 1960s. The increase in both total soil N and N enrichment of labile soil organic matter fractions under pasture were almost certainly derived in part from N inputs in leguminous plant residues and residual organic N from past fertiliser N inputs. This may well have enhanced the N mineralisation potential of the soil (Ward et al. 2003) and in time may contribute to increased N[O.sub.3]-N concentrations in the soil and surrounding environment. The [sup.13]C CP/MAS NMR spectra for plant material and light fractions obtained from the 2 different land uses were similar, indicating that this approach may be of limited value for differentiating between materials with different C/N ratios, and hence different decomposability.
The work of A. Macdonald was supported in part by a fellowship from the Organisation for Economic Co-operation and Development under the Co-operative research programme on Biological Resources Management for Sustainable Agricultural Systems. D. V. Murphy was supported by an Australian Grains Research and Development Corporation Research Fellowship. Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council of the United Kingdom. Additional support was provided by the UK Natural Environmental Research Council. We thank Nui Milton for assistance with the analytical work.
Manuscript received 27 August 2006, accepted 14 June 2007
Abbott I, Parker CA, Sills ID (1979) Changes in the abundance of large soil animals and physical properties of soils following cultivation. Australian Journal of Soil Research 17, 343-353. doi: 10.1071/SR9790343
Anderson GC, Fillery IR, Dolling PJ, Asseng S (1998a) Nitrogen and water flows under pasture-wheat and lupin-wheat rotations in deep sands in Western Australia. 1. Nitrogen fixation in legumes, net N mineralisation, and utilisation of soil-derived nitrogen. Australian Journal of Agricultural Research 49, 329 344. doi: 10.1071/A97142
Anderson GC, Fillery IR, Dunin FX, Dolling PJ, Asseng S (1998b) Nitrogen and water flows under pasture-wheat and lupin-wheat rotations in deep sands in Western Australia. 2. Drainage and nitrate leaching. Australian Journal of Agricultural Research 49, 345-361. doi: 10.1071/A97142
Barrios E, Buresh RJ, Sprent JI (1996) Nitrogen mineralization in density fractions of soil organic matter from maize and legume systems. Soil Biology & Biochemistry 28, 1459-1465. doi: 10.1016/S0038-0717(96)00155-1
Cabrera ML, Beare MH (1993) Alkaline persulphate oxidation for determining total nitrogen in microbial biomass extracts. Soil Science Society of America Journal 57, 1007-1012.
Chen CR, Xu ZH, Mathers NJ (2004) Soil carbon pools in adjacent natural and plantation forest of subtropical Australia. Soil Science Society of America Journal 68, 282-291.
Cookson WR, Abaye D, Marschner P, Murphy DV, Stockdale EA, Goulding KWT (2005) The contribution of soil organic matter fractions to carbon and nitrogen mineralization and microbial community size and structure. Soil Biology & Biochemistry 37, 1726-1737. doi: 10.1016/j.soilbio.2005.02.007
Dalal RC, Mayer RJ (1986a) Long-term trends in fertility of soils under continuous cultivation and cereal cropping in southern Queensland. I. Overall changes in soil properties and trends in winter cereal yields. Australian Journal of Soil Research 24, 265-279. doi: 10.1071/SR9860265
Dalal R, Mayer R (1986b) Long-term trends in fertility of soils under continuous cultivation and cereal cropping in southern Queensland. IV. Loss of organic carbon from different density fractions. Australian Journal of Soil Research 24, 301-309. doi: 10.1071/SR9860301
Danso SKA, Hardarson G, Zapata F (1988) Dinitrogen fixation estimates in alfalfa-ryegrass swards using different nitrogen-15 labelling methods. Crop Science 28, 106-110.
Dixon WT (1982) Spinning-sideband-free and spinning-sideband-only NMR spectra in spinning samples. The Journal of Chemical Physics 77, 1800-1809. doi: 10.1063/1.444076
Glendining MJ, Powlson DS (1995) The effects of long continued applications of inorganic nitrogen fertilizer on soil organic nitrogen-A review. In 'Advances in soil science'. (Eds R Lai, BA Stewart) pp. 385-446. (CRC Press, Inc: Boca Raton, FL)
Golchin A, Oades JM, Skjemstad JO, Clarke P (1994) Soil structure and carbon cycling. Australian Journal of Soil Research 32, 1043-1068. doi: 10.1071/SR9941043
Golchin A, Oades JM, Skjemstad J O, Clarke P (1995) Structural and dynamic properties of soil organic matter as reflected by [sup.13]C natural abundance, pyrolysis mass spectromerty and solid state [sup.13]C NMR spectroscopy in density fractions of an oxisol under forest and pasture. Australian Journal of Soil Research 33, 59-76. doi: 10.1071/SR9950059
Grigg AM, Pate JS, Unkovich MJ (2000) Responses of native woody taxa in Banksia woodland to incursion of groundwater and nutrients from bordering agricultural land. Australian Journal of Botany 48, 777-792. doi: 10.1071/BT99078
Hassink J (1995a) Decomposition rate constants of size and density fractions of soil organic matter. Soil Science Society of America Journal 59, 1631-1635.
Hassink J (1995b) Density fractions of soil macroorganic matter and microbial biomass as predictors of C and N mineralization. Soil Biology & Biochemistry 27, 1099-1108. doi: 10.1016/0038-0717(95)00027-C
Henriksen A, Selmer-Olsen AR (1970) Automatic methods for determining nitrate and nitrite in water and soil extracts. The Analyst 95, 514-518. doi: 10.1039/an9709500514
Jenkinson DS (1988) Soil organic matter and its dynamics. In 'Russell's soil conditions and plant growth'. (Ed. A Wild) pp. 564-607. (Longman Scientific and Technical: London)
Jones DL, Shannon D, Murphy D, Farrar J (2004) Role of dissolved organic N (DON) in soil N cycling in grassland soils. Soil Biology & Biochemistry 36, 749-756. doi: 10.1016/j.soilbio.2004.01.003
Jones DL, Willett VB (2006) Experimental evaluation of methods to quantify dissolved organic nitrogen (DON) and dissolved carbon (DOC) in soil. Soil Biology & Biochemistry 38, 991-999. doi: 10.1016/j.soilbio.2005.08.012
Kalbitz K, Solinger S, Park JH, Michalzik B, Matzner E (2000) Controls on the dynamics of dissolved organic matter in soils: a review. Soil Science 165, 277-304. doi: 10.1097/00010694-200004000-00001
Krom MD (1980) Spectrophotometric determination of ammonia: a study of a modified Berthelot reaction using salicylate and dichloroisocyanurate. The Analyst 105, 305-316. doi: 10.1039/an9800500305
Low AB, Lamont BB (1990) Aerial and below-ground Phytomass of Banksia Scrub-heath at Enebha, South-western Australia. Australian Journal of Botany 38, 351-359. doi: 10.1071/BT9900351
Macdonald AJ, Poulton PR, Stockdale EA, Powlson DS, Jenkinson DS (2002) The fate of residual [sup.15]N-labelled fertilizer in arable soils: its availability to subsequent crops and retention in soil. Plant and Soil 246, 123-137. doi: 10.1023/A:1021580701267
Marschner B, Kalbitz K (2003) Controls of bioavailability and biodegradability of dissolved organic matter in soils. Geoderma 113, 211-235. doi: 10.1016/S0016-7061(02)00362-2
Mathers NJ, Mao XA, Xu ZH, Saffigna PG, Berners-Price SJ, Perera MCS (2000) Recent advances in the application of [sup.13]C and [sup.15]N NMR spectroscopy to soil organic matter studies. Australian Journal of Soil Research 38, 769-787. doi: 10.1071/SR99074
McArthur WM (1991) Reference soils of south western Australia. Department of Agriculture and Australian Society of Soil Science, WA.
McNeill AM, Sparling GP, Murphy DV, Braunberger P, Fillery IRP (1998) Changes in extractable and microbial C, N and P in a Western Australian wheatbelt soil following simulated summer rainfall. Australian Journal of Soil Research 36, 841-854. doi: 10.1071/S97044
Mendham DS, Mathers NJ, O'Connell, Grove TS, Saffigna PG (2002) Impact of land-use on soil organic matter quality in south-western Australia--characterisation with [sup.13]C CP/MAS NMR spectroscopy. Soil Biology & Biochemistry 34, 1669-1673. doi: 10.1016/S0038-0717(02)00151-7
Muller MM, Sundman V, Soininvaara O, Meriliainen A (1988) Effect of chemical composition on the release of nitrogen from agricultural plant materials decomposing in soil under field conditions. Biology and Fertility of Soils 6, 78-83. doi: 10.1007/BF00257926
Murphy DV, Macdonald A J, Stockdale EA, Goulding KWT, Fortune S, Gaunt JL, Poulton PR, Wakefield JA, Webster CP, Wilmer WS (2000) Soluble organic nitrogen in agricultural soils. Biology and Fertility of Soils 30, 374-387. doi: 10.1007/s003740050018
Osler GHR, Murphy DV (2005) Oribatid mite species richness and soil organic matter fractions in agricultural and native vegetation soils in Western Australia. Applied Soil Ecology 29, 93-98. doi: 10.1016/ j.apsoil.2004.09.002
Pate JS, Stewart GR, Unkovich M (1993) [sup.15]N natural abundance of plant and soil components of a Banksia woodland ecosystem in relation to nitrate utilization, life form, mycorrhizal status and [N.sub.2]-fixing abilities of component species. Plant, Cell & Environment 16, 365-373. doi: 10.1111/j.1365-3040.1993.tb00882.x
Pate JS, Unkovich MJ, Armstrong EL, Sanford P (1994) Selection of reference plants for [sup.15]N natural abundance assessment of [N.sub.2] fixation by crops and pasture legumes in South-West Australia. Australian Journal of Agricultural Research 45, 133-147. doi: 10.1071/AR9940133
Powlson DS, Jenkinson DS, Pruden G, Johnston AE (1985) The effect of straw incorporation on the uptake of nitrogen by winter wheat. Journal of the Science of Food and Agriculture 36, 26-30. doi: 10.1002/jsfa.2740360105
Rahn CR, Lillywhite R (2002) A study of quality factors affecting the short term decomposition of field vegetable residues. Journal of the Science of Food and Agriculture 82, 19-26. doi: 10.1002/jsfa.1003
Skjemstad JO, Clarke P, Golchin A, Oades JM (1997) Characterisation of soil organic matter by solid-state [sup.13]C NMR Spectroscopy. In 'Driven by nature: plant litter quality and decomposition'. (Eds G Cadisch, K Giller) pp. 253-271. (Cab International: Wallingford, UK)
Sohi SP, Mahieu N, Arah JRM, Powlson DS, Madari B, Gaunt JL (2001) A procedure for isolating soil organic matter fractions suitable for modelling. Soil Science Society of America Journal 65, 1121-1128.
Sollins P, Spycher G, Glassman CA (1984) Net nitrogen mineralization from light- and heavy-fraction forest soil organic matter. Soil Biology & Biochemistry 16, 31-37. doi: 10.1016/0038-0717(84)90122-6
Standish RJ, Cramer VA, Hobbs RJ, Kobryn HT (2006) Legacy of land-use evident in soils of Western Australia's wheat belt. Plant and Soil 280, 189-207. doi: 10.1007/s11104-005-2855-6
Thorup-Kristensen K (1994) An easy pot incubation method for measuring nitrogen mineralization from easily decomposable organic material under well defined conditions. Fertilizer Research 38, 239-247. doi: 10.1007/BF00749697
Unkovich MJ, Pate JS, Hamblin J (1994) The nitrogen economy of broadacre lupin in southwest Australia. Australian Journal of Agricultural Research 45, 149-164. doi: 10.1071/AR9940149
Waksman SA, Tenney FG (1927) The composition of natural organic materials and their decomposition in the soil II. Influence of age of plant upon the rapidity and nature of its decomposition--rye plants. Soil Science 24, 317-333.
Ward PR, Fillery IRE Maharaj EA, Dunin FX (2003) Water budgets and nutrients in a native Banksia woodland and an adjacent Medicago sativa pasture. Plant and Soil 257, 305-319. doi: 10.1023/A:1027331712165
A. J. Macdonald (A,E), D. V. Murphy (B), N. Mahieu (c), and I. R. P. Fillery (D)
(A) Soil Science Department, Rothamsted Research, Harpenden, Herts AL5 2JQ, UK.
(B) School of Earth and Geographical Sciences, Faculty of Natural and Agricultural Sciences, The University of Western Australia, Crawley, WA 6009, Australia.
(C) Queen Mary, University of London, London E1 4NS, UK.
(D) Commonwealth Scientific Industrial Research Organisation, PO Box Wembley, WA 6193, Australia.
(E) Corresponding author. Email: email@example.com
Table 1. Total C and N contents, and [[delta].sup.15] of plant material, leaf litter, and light fractions (0-0.15m) collected from native bush and grass/lucerne pasture Data are means of 3 replicates, with standard errors in parentheses Sample type Dry wt Total C Total N Native bush (t/ha) (kg/ha) (kg/ha) Plant litter 5.5 (0.52) 2575 (254.2) 26.4 (2.36) LF 1.0 4.4 (0.29) 1048 (94.5) 46.5 (5.74) LF 1.7 7.9 (0.94) 1890 (315.6) 47.9 (15.33) Pasture Plants and litter 2.2 (0.39) 670 (99.0) 54.2 (8.64) LF 1.0 2.1 (0.40) 483 (57.1) 47.6 (9.12) LF 1.7 4.4 (0.13) 1083 (102.4) 82.3 (8.58) [delta] Sample type [sup.15]N [per C/N Native bush thousand] ratio Plant litter -2.0 (1.23) 97.5 (3.81) LF 1.0 33.2 (6.46) 22.6 (2.07) LF 1.7 4.8 (3.34) 39.4 (6.90) Pasture Plants and litter 1.8 (0.52) 12.4 (0.18) LF 1.0 13.9 (3.29) 10.1 (0.80) LF 1.7 7.7 (2.27) 13.2 (2.50) Table 2. Carbon present in functional groups as a percentage of the total C in vegetation and light fractions under native bush and adjacent grass/lucerne pasture, as determined by [sup.13]C CP/MAS NMR spectroscopy Data are means of 3 replicates with standard errors in parentheses Sample type Alkyl N-alkyl & O-alkyl Acetal methoxyl Native bush Plant litter 20 (1.5) 14 (0.3) 34 (0.8) 13 (0.5) LF 1.0 19 (2.0) 15 (0.6) 28 (0.5) 11 (0.4) LF 1.7 33 (1.2) 13 (0.4) 20 (1.9) 8 (0.3) Pasture Plants and litter 17 (0.7) 15 (0.1) 33 (0.5) 10 (0.3) LF 1.0 16 (0.7) 16 (0.8) 30 (1.5) 11 (0.9) IT 1.7 33 (2.0) 14 (0.5) 19 (1.1) 7 (0.6) Sample type Aromatic Phenolic Carboxyl O-alkyl/ alkyl Native bush Plant litter 9 (0.1) 6 (0.1) 5 (0.2) 1.8 (0.18) LF 1.0 13 (0.6) 7 (0.7) 6 (0.8) 1.5 (0.19) LF 1.7 12 (0.6) 6 (0.4) 7 (0.5) 0.6 (0.08) Pasture Plants and litter 9 (0.2) 5 (0.2) 12 (0.3) 1.9 (0.10) LF 1.0 13 (1.5) 7 (0.9) 8 (l.2) 1.9 (0.08) IT 1.7 12 (0.3) 6 (0.2) 10 (0.8) 0.6 (0.07)
|Printer friendly Cite/link Email Feedback|
|Author:||Macdonald, A.J.; Murphy, D.V.; Mahieu, N.; Fillery, I.R.P.|
|Publication:||Australian Journal of Soil Research|
|Date:||Aug 1, 2007|
|Previous Article:||Gravity segregation during miscible displacement--re-investigation and re-interpretation.|
|Next Article:||Effects of 15 years of conservation tillage on soil structure and productivity of wheat cultivation in northern China.|