Short-term effects of incubated legume and grass materials on soil acidity and C and N mineralisation in a soil of north-east Australia.
Perennial grass growth forms the basis of beef production systems in northern Australia. Stylosanthes (stylo), a tropical woody legume shrub, has been introduced into these native pastures in order to improve pasture quality and beef production. However, the growth of legumes has been recognised to be a major factor in soil acidification (Haynes 1983; Liu et al. 1989; Bolan et al. 1991; Noble et al. 1997), thus posing a potential risk for land degradation and a reduction in soil fertility.
According to van Breemen et al. (1983), soil acidification is best characterised by the permanent loss of acid neutralisation capacity (ANC). As a consequence of this reduced ability to buffer protons, the pH of soils may decrease. The loss in ANC can be caused by various factors, only some of which are relevant to tropical legume pastures. The most commonly observed process is soil acidification due to nitrate leaching, where ANC is reduced during buffering of acidity produced during nitrate formation and is irreversibly lost when excess nitrate is leached and not taken up by plants. Leaching of nitrate is generally associated with the loss of base cations. They may also be depleted in soils after being taken up by plants, when litter degradation is incomplete and residues accumulate on the soil surface. In pastures, base cation export in forage and animals can also contribute to a decrease in ANC. The relative importance of these processes must be known in order to conceive effective countermeasures against soil acidification.
Due to their base cation content, plant residues added to soils generally increase the pH or ANC initially. Bessho and Bell (1992) interpreted a pH increase as being due to exchange reactions resulting from cation release during plant material decomposition. Pocknee and Sumner (1997) found evidence that organic compounds containing base cations increase soil pH in a similar fashion to mineral lime. There have also been consistent reports that [H.sup.+] is consumed by decarboxylation of organic anions that are added to soil (Barekzai and Mengel 1993; Yan et al. 1996a, 1996b; Tang and Yu 1999; Tang et al. 1999; Marschner and Noble 2000). Additionally, buffering effects occurring immediately following plant material addition were ascribed either to association of protons with functional groups, or to released alkalinity in very soluble forms or in immediately decomposing parts of the residues (Tang et al. 1999; Marschner and Noble 2000). Hoyt and Turner (1975) incorporated organic materials into soil and explained short-term pH increase with N[H.sub.3] production during the incubation period.
The ash alkalinity of plants, also expressed as their organic anion or excess base cation content, plays a crucial role in changing the soil pH, since it has been found to neutralise soil acidity in several incubation studies (Yan et al. 1996a; Noble et al. 1996; Noble and Randall 1999; Tang and Yu 1999; Tang et al. 1999; Marschner and Noble 2000). Since legume residues generally contain high amounts of ash alkalinity (Tang et al. 1999), their incorporation and degradation in soils often also increases ANC, if it is not outweighed by acid production from mineralisation of N-rich plant compounds.
On an experimental site in north-east Australia, soil under stylo showed significantly lower pH values than soil under grass vegetation. In order to examine the processes that have led to soil acidification under stylo, a short-term incubation study was conducted. The objective was to examine the effects of stylo and urochloa residues on C and N mineralisation, acid production, pH, and buffering properties.
Materials and methods
Soil used in study
The soil used in the experiment was collected at Lansdown, Queensland, Australia (19[degrees]37'S, 146[degrees]50'E) from 2 adjacent experimental pasture plots, one dominated by Stylosanthes scabra (stylo) and the other by Urochloa mosambicensis (urochloa), both planted in 1986. The soil examined was classified as Mottled-Subnatric Yellow Sodosol (Australian Soil Classification, Isbell 1996). Five samples were taken from the topsoil (0-10 em) of both plots. The soil was bulked into a composite sample, air-dried, and sieved to pass a 2-mm mesh. Some of the soil properties are shown in Table 1.
Leaves of stylo and tops of urochloa were collected at the study site during the growing season in February 2000. Roots of both species were washed from the soil over a 2-mm mesh. All plant materials were dried at 70[degrees]C and ground to a powder.
Ash alkalinity was determined using the methodology of Jarvis and Robson (1983). Plant samples were heated slowly to 500[degrees]C and held at this temperature for 1 h; 5 mL of 1 M HCl was added to the ash and then titrated against 0.25 M NaOH. The organic C and total N contents of plant material were measured with a C/N analyser (ANA 1500, Carlo Erba). Plant properties are given in Table 2.
Soil (50 g) from the stylo and urochloa plots was thoroughly mixed with 0.3 g of finely ground urochloa roots, stylo roots, stylo leaves, or urochloa leaves in experimental jars. The added amount of plant material (6 g/kg) is equivalent to 10 t/ha for 10 cm soil depth at the local bulk density of 1.7 t/[m.sup.3]. A control soil sample with no added plant material was also included in the incubation study. All samples were prepared in triplicate and moistened to 60[degrees],4 of water-holding capacity (WHC) immediately after the plant materials had been mixed into the soils (referred to `before incubation' in the following). WHC was determined by percolating water through a known amount of oven-dried soil several times in a pre-weighed funnel with a moist filter paper until it was saturated. After water completely stopped draining out of the soil, the funnel with filter and soil was weighed again.
Respiration was measured in an automated respirometer (Respicond VI, Nordgren Innovation) as described by Nordgren (1988). During the 25-day incubation at 25[degrees]C, C[O.sub.2] was absorbed in a 0.6 M KOH solution. Respiration was monitored hourly using electrical conductivity measurements of the alkaline solution.
Before and right after incubation, the soil in each of the experimental jars was analysed for ANC and extractable N[O.sub.3.sup.-]-N, N[H.sub.4.sup.+]-N, and dissolved organic carbon (DOC). A modified method of James and Riha (1986) was used to measure the ANC. A 2-g soil sample was weighed into each of 6 polyethylene tubes. Additions of solutions consisting of 0, 0.5, 2.0, 4.0, 10.0, and 20.0 mL of 0.01 M HN[O.sub.3], 1 mL of 1 M KN[O.sub.3], and 0.25 mL of chloroform were made. An appropriate amount of deionised water was added to make the final volume of 25 mL. Samples were equilibrated on an end-over-end shaker for 16 h. pH was measured in the supernatant. ANC was the amount of [H.sup.+] needed to reduce existing pH to a value of 3.
For extraction, 4 g of soil was shaken with 40 mL of 0.01 M Ca[Cl.sub.2] solution for 1 h, followed by membrane filtration (0.45-[micro]m cellulose-nitrate filter, Sartorius). In the soil extracts, DOC was analysed with a total organic carbon analyser (Shimadzu TOC-5050) using high temperature, catalytic oxidation. Analysis of N[O.sub.3.sup.-]-N was undertaken by ion chromatography (Dionex DX 20 autoanalyser). N[H.sub.4.sup.+]-N was determined colorimetrically (Lambda 2, Perkin Elmer) following reaction with sodium dichlorisocyanurate/sodium hydroxide and salicylate citrate solution.
Calculations and statistical analysis
The cumulative amount of C[O.sub.2]-C released ([C.sub.cum]) was derived from the amount of C[O.sub.2] evolved after 25 days of incubation. Mineralised C ([C.sub.min]) was calculated by subtracting the [C.sub.cum] values of the controls from the values of soils treated with plant materials. These results were related to the organic carbon inputs from the various plant materials and expressed as per cent values.
Net nitrogen mineralisation was calculated from the difference of N[H.sub.4.sup.+]-N and N[O.sub.3.sup.-]-N between the extractions before and after incubation.
Production and consumption of [H.sup.+] were derived from ammonification (Eqns 1 and 2) and nitrification (Eqn 3) reactions, as shown in the following equations:
(1) R-N[H.sub.2] + [H.sub.2]O [right arrow] N[H.sub.3] + R-OH
(2) N[H.sub.3] +[H.sup.+] [left and right arrow] N[H.sub.4.sup.+]
(3) N[H.sub.4.sup.+] + 2[O.sub.2] [right arrow] N[O.sub.3.sup.-] + [H.sub.2]O + 2[H.sup.+]
In order to determine the significance of differences between the 2 soils incubated with different plant materials, a t-test for unpaired samples with equal variances was performed. Relationships between 2 variates were evaluated by applying a linear correlation according to Pearson (Brosius 1998).
Compared with the soil under urochloa, the soil under stylo had a lower pH and lower ANC (Table 1). [C.sub.org] was higher in urochloa than in stylo soil. Since the [N.sub.t] content was similar in both soils, the C:N ratio was higher in urochloa soil.
Stylo leaves had the highest ash alkalinity and the lowest C:N ratio of all plant materials (Table 2). In contrast, urochloa roots had the lowest ash alkalinity and the highest C:N ratio. Total N content was lower in urochloa materials than in stylo materials, with maximum values in stylo leaves.
During incubation, more C[O.sub.2]-C ([C.sub.cum]) was released from the unamended stylo soil than from the unamended urochloa soil (Table 3), although the [C.sub.org] content of the stylo soil was lower (Table 1). This resulted in mineralisation of 8% of soil C in the stylo soil and 5% in the urochloa soil. Among the amendments, stylo materials generally were mineralised more effectively than urochloa materials in both soils. The highest values were reached in urochloa soil incubated with stylo leaves. The lowest rates occurred in soils incubated with urochloa roots. With the exception of stylo roots, mineralisation rates of all plant materials were higher in the urochloa soil than the stylo soil. Correlation of [C.sub.cum] with the C:N ratio of added plant residues showed a highly significant negative relationship (r = -0.913).
Respiration peaks in all treatments with urochloa soil were higher and occurred earlier than the corresponding stylo peaks (not shown). Maximum C[O.sub.2] evolution was reached after 2 h for the urochloa soil with no plant addition but only after 7 h in the respective stylo soil. This corresponds with the higher organic matter content of the urochloa soil. Addition of plant materials increased the DOC content in all treatments before incubation (Fig. 1). Changes were most pronounced with stylo leaves. Urochloa roots showed the least effects on DOC. DOC content was significantly higher in urochloa than in stylo soil before incubation, except for the urochloa tops treatment.
[FIGURE 1 OMITTED]
During incubation, a significant decrease in DOC content occurred, down to a level of <50 mg/kg in all treatments. The differences between DOC content in the 2 plant-amended soils were not significant, but the urochloa control value was significantly higher than that of stylo. After the first 2 days of incubation, respiration decreased markedly in both the unamended and amended soils (not shown). The difference in DOC between before and after incubation correlated well with the amount of C[O.sub.2]-C evolved during these first 2 days of incubation (Fig. 2).
[FIGURE 2 OMITTED]
Table 4 summarises the fate of N in the soils during the incubation. Generally, the N[O.sub.3]-N content in the urochloa soil was lower than in stylo soil and N[H.sub.4]-N concentration higher. Comparisons of the concentration of N species before and after incubation showed consistently significant differences. N[O.sub.3]-N increased in all treatments, except with urochloa tops, where the value decreased in both soils. N[H.sub.4]-N content increased in both controls and in soils amended with stylo leaves. In the other treatments N[H.sub.4]-N decreased during incubation.
The highest N mineralisation ([N.sub.min]) was measured in the stylo control. Considering [N.sub.min] without referring them to their corresponding controls, stylo soil with the urochloa roots showed the highest values and urochloa soil with stylo roots the lowest. Added urochloa roots showed higher [N.sub.min] values than stylo roots and incubated stylo leaves brought about similar values for both soils. In general, [N.sub.min] values were higher in the stylo soil than in the urochloa soil.
After incubation, the low [N.sub.min] values in plant-amended soils compared with their corresponding controls showed that added N was immobilised, except when stylo leaves were added to urochloa soil. The highest immobilisation rate was reached in soils incubated with urochloa tops.
Production and consumption of [H.sup.+], as calculated from N transformations, is presented in Fig. 3. The highest [H.sup.+] production occurred in the control soils, due to their higher nitrification rates. Incubated stylo leaves induced a similar mineralisation pattern in both soils; a small amount of acid was produced due to higher nitrification than ammonification rates. Incubated urochloa tops had almost no net effect on N mineralisation or the [H.sup.+] balance. The latter was also true for incubated roots in the urochloa soil.
In the controls and soils amended with stylo leaves, [H.sup.+] was consumed due to net ammonification. In all other treatments, the N[H.sub.4]-N levels before incubation were slightly higher than after incubation, showing acid consumption from ammonification was negligible.
[FIGURE 3 OMITTED]
Soil pH and ANC increased in almost all amended soils immediately after plant material addition (Table 5). The increases in ANC correspond with amounts of added ash alkalinity, indicating that this alkalinity is directly available for acid buffering. When soils were amended with stylo leaves, the increase was more pronounced than with addition of any other plant materials. In general, pH rose proportionally to the added alkalinity of amendments in each soil (r = 0.960 for stylo soil; r = 0.957 for urochloa soil). Relating ANC to the added alkalinity also revealed high correlations (r = 0.957 for stylo soil; r = 0.947 for urochloa soil).
Incubation had only small effects on pH and ANC (not shown), reflecting the low [H.sup.+] production from nitrification during incubation.
The different C mineralisation rates of the 4 plant materials can be explained by their differences in C:N ratio. The C:N ratio of plant residues has often been taken into consideration as a general index of plant quality, i.e. the C:N ratio having the greatest influence on mineralisation (Heal et al. 1997; Mafongoya et al. 1998; Trinsoutrot et al. 2000; Nicolardot et al. 2001). However, as Vanlauwe et al. (1997) pointed out, it may be misleading to describe decomposition with only a few parameters, because of the complex nature of decaying substrate. In this context, Jansson and Persson (1982) suggested that the C:N ratio is only an approximation for the crucial Energy:N ratio and that this approximation could be deceptive. For example, some constituents of plant material may not be readily available to decomposers (Mtambanengwe et al. 1998). In this short-term study, the C:N ratio of plant material also seemed to be a predictor for the amount of cumulative C[O.sub.2]-C released from plant treated soils, which was reflected in the close inverse relationship between these parameters.
The small differences in C[O.sub.2]-C release from soil with N-rich stylo roots (C:N = 47) and the N-poor urochloa tops (C:N = 90) incubated in the urochloa soil suggest that other quality parameters also influenced mineralisation rates. One is the amount of initially available water-soluble compounds which apparently were preferentially mineralised during the first 2 days of incubation, as indicated by the close relationship between respiration and DOC disappearance. The amounts involved indicate that up to approximately one-third of the C[O.sub.2]-C evolved may have resulted from mineralisation of soluble organic components of the plant materials. Similarly, the studies of Jawson and Elliott (1986) and Marschner and Noble (2000) showed that at much higher rates of plant residue additions, the evolved C[O.sub.2]-C could be explained by the loss of DOC. These soluble compounds probably contain large amounts of carbohydrates and free amino acids, which Marstorp (1996) found to serve as an energy source for the soil microorganisms in the initial phase of decomposition. In agreement with the present study, Mtambanengwe et al. (1998) found the initial dissolved C best described the amount of C mineralised within the first month of decomposition. At the end of their study period, small concentrations of DOC were still present in soil, suggesting that these remaining soluble compounds were less easily degradable.
In contrast to the basal respiration of the control soil, the mineralisation rates of most of the added plant materials (except stylo roots) were higher in the urochloa than in the stylo soil. This may partly be due to the slightly higher pH of the urochloa soil, but does not explain the 1.3-fold elevated mineralisation of stylo leaves, since pH was around 5.7 in both these treatments. Possibly, microorganisms in the urochloa soil are more limited by available C sources than in the stylo soil, as indicated by the different basal respirations. Consequently, fresh plant materials can be readily mineralised while other limitations, such as nutrient deficiency are less relevant. In the stylo soil, some nutrients apart from N may become limiting earlier due to their fixation in the more active microbial population.
Surprisingly, stylo leaves induced a net N-immobilisation in stylo soil, although N-rich fresh legume leaves should promote higher N-mineralisation. Since denitrification is unlikely in this well-aerated incubation system, other processes must have caused N-immobilisation. It could be caused by a higher microbial biomass or by preferential utilisation of N-poor compounds, such as soluble sugars because N-rich compounds may initially be unavailable for decomposition. Tannins present in the plant material could have bound N-rich compounds or suppressed autotrophic bacteria (Paul and Clark 1989). But since N immobilisation and N mineralisation depends on the duration of incubation (Mtambanengwe et al. 1998) it is conceivable that net N-mineralisation would have occurred with stylo leaves if the incubation had been prolonged. In the other treatments, N immobilisation was expected, since the C:N ratios of the residues were well above 20-30, which is generally considered as a threshold value for immobilisation (Haynes 1986; Paul and Clark 1989; Hodge et al. 2000; Seneviratne 2000).
Interestingly, there were distinct differences in mineralisation rates in the 2 unamended soils. In the stylo soil, more C[O.sub.2] was released than in the urochloa soil although its [C.sub.org] content and pH were lower. This could be due to either the higher availability of degradable C sources or the less severe nutrient limitations, or to a combination of both. N availability may be one parameter controlling microbial activity, but the net N-mineralisation in these soils shows that N was not limiting decomposition. On the other hand, the higher microbial respiration in the stylo soil indicates that turnover of soil organic matter is faster in this soil which would account for the lower [C.sub.org]-content of this soil.
The close link between pH, ANC and added alkalinity from plant residues was consistent with previous studies (e.g. Noble et al. 1996; Noble and Randall 1999; Tang et al. 1999) and shows that this alkalinity is directly available for buffering of acidity.
Unlike several previous incubation studies (e.g. Yan et al. 1996a, 1996b; Marschner and Noble 2000), the present work showed no significant pH increase during incubation, despite the fact that [H.sup.+]-consuming decarboxylation processes are thought to be predominantly responsible for organic matter decomposition (Paul and Clark 1989). However, plant residue additions were much lower than in the above-mentioned studies so that the contribution of decarboxylation processes to acidity may not have been detectable in this well buffered soil.
Contrary to our expectations, N-transformations also had no detectable effect on pH or ANC in the plant-amended soils because [N.sub.min] was not released but was immobilised. This immobilisation was assumed to occur in the growing microbial biomass. However, this may just be a temporary storage pool, since the microbial populations will eventually die when easily degradable substrates are depleted or when environmental conditions become adverse (i.e. drought). The acidity produced from N transformations in the unamended soils may very well reflect this situation, since samples were taken at the end of the dry season. Rewetting of the samples may activate the biomass which then apparently utilises the accumulated N-rich dead microbial cell debris as substrate releasing large amounts of nitrate and [H.sup.+]. If all the nitrate would be leached, then the buffering of acidity produced in the stylo soil would reduce ANC by almost 2%. In the urochloa soil this amounts to only 0.5% due to the lower acid production and higher ANC. Annual acidity production of that magnitude in the field would explain the differences in ANC between the 2 plots, since ANC in the stylo soil is about 17% lower than in the urochloa soil after 13 years field trial.
The results of this incubation study show that the observed soil acidification under stylo plant cover is not directly caused by generation of acidity from N-transformations during degradation of fresh plant materials. Instead, pH and ANC increased proportionally to the amount of ash alkalinity added with the different plant materials. Therefore, other acid generating processes must be effective in the field.
In this short-term study, the acidifying effect of nitrification is negligible in the plant-amended treatments because microorganisms immobilised N due to an abundance of easily degradable C-sources which support microbial growth. This situation is expected to change, when degradation proceeds and these C compounds are depleted. Then microorganisms will die and use larger proportions of organic matter for energy production, both of which will release surplus N available for nitrification and leaching. This was apparently the case in the soil, when it was sampled at the end of the dry season.
Still, for acidification to become effective, acid production from mineralisation processes has to be higher than the alkalinity contained in the plant residues. In the field, the above-ground biomass is not fully returned to the soil, but grazed and exported by cattle. Due to the high ash alkalinity of stylo leaves, more buffering substances will be removed than with the same amount of urochloa biomass. Although N is also exported in this manner, N-inputs from fixation are still higher on the stylo plot as evidenced by the narrow C:N ratio in the soil and the higher amount of [N.sub.min] produced in the control treatments. The enhanced nitrification at the beginning of the growing season makes nitrate leaching very likely, which will thus promote irreversible soil acidification through the associated loss of cations. Which of these processes is more relevant for soil acidification under stylo can only be determined with further field studies where in- and out-puts of all relevant elements are quantified.
Table 1. Selected soil properties of soils collected from the stylo and urochloa plots (n = 5) Soil pH ANCA (0.01 M Ca ([mmol. [C.sub.org] [N.sub.t] C:N [Cl.sub.2]) sub.c]kg) (%) (%) Stylo 4.5 34 0.61 0.09 7 Urochloa 4.9 41 0.87 0.09 10 (A) Acid neutralisation capacity. Table 2. Selected plant properties of leaves and roots collected from the two sites Ash Plant material alkalinity (A) [C.sub.org] (B) [N.sub.t] (B) C:N ([mmol.sub.c]/kg) (%) (%) Stylo leaves 1140 43.7 2.48 17 Stylo roots 465 41.9 0.94 47 Urochloa tops 433 42.0 0.43 90 Urochloa roots 215 41.4 0.40 106 (A) n=5. (B) n=2. Table 3. Organic carbon ([C.sub.org]) inputs, cumulative C[O.sub.2]-C (C[O.sub.2]-[C.sub.cum]) and relative C-mineralisation of the added plant materials ([C.sub.min) in stylo and urocbloa soil incubated with different plant materials Stylo soil [C.sub.org] Plant material input C[0.sub.2]- (mg/kg) [C.sub.cum] [C.sub.min] (mg/kg) (%) Control 0 513 -- Stylo leaves 2518 1281 30 Stylo roots 2623 1386 33 Urochloa tops 2482 846 13 Urochloa roots 2511 738 9 Urochloa soil Plant material [C.sub.cum] [C.sub.min] (mg/kg) (%) Control 391 * -- Stylo leaves 1418 * 41 ** Stylo roots 1111 * 27 Urochloa tops 988 *** 24 *** Urochloa roots 747 14 * Significance of differences between stylo and urochloa soil, using t-test: * P [less than or equal to] 0.05; ** P [less than or equal to] 0.01; *** P [less than or equal to] 0.001. Table 4. Nitrogen inputs, N[H.sub.4]-N and N[O.sub.3]-N before and after incubation, and N mineralised ([N.sub.min]) in stylo and urochloa soil incubated with different plant materials Values in mg/kg Plant N Stylo soil material input Before After N[H.sub.4]-N N[O.sub.3]-N N[H.sub.4]-N Control 0 0.4 0.8 0.9 Stylo leaves 149 0.4 0.8 2.2 Stylo roots 56 0.4 0.8 0.2 Urochloa tops 27 0.4 0.7 0.1 Urochloa roots 24 0.3 0.5 0.1 Values in mg/kg Stylo soil Plant material After [N.sub.min] N[O.sub.3]-N Control 10.3 10.0 Stylo leaves 4.7 5.7 Stylo roots 4.6 3.6 Urochloa tops 0.1 -0.9 Urochloa roots 8.0 7.3 Values in mg/kg Urochloa soil Plant material Before After N[H.sub.4]-N N[O.sub.3]-N N[H.sub.4]-N Control 0.5 <0.1 ** 1.9 ** Stylo leaves 0.6 0.2 *** 2.2 Stylo roots 0.5 ** 0.3 *** 0.2 Urochloa tops 0.5 0.1 *** 0.1 Urochloa roots 0.5 ** 0.2 ** <0.1 Values in mg/kg Urochloa soil Plant material After [N.sub.min] N[O.sub.3]-N Control 4.1 ** 5.5 ** Stylo leaves 4.3 5.7 Stylo roots 0.7 *** 0.1 ** Urochloa tops 0.1 -0.4 ** Urochloa roots 1.4 *** 0.7 *** Significance of differences between stylo and urochloa soil, using t-test: ** P [less than or equal to] 0.01; *** P [less than or equal to] 0.001. Table 5. Added alkalinity, pH, and acid neutralising capacity (ANC) in stylo and urochloa soil mixed with different plant materials before incubation Stylo soil Plant material Added alkalinity ([mmol.sub.c]/kg) pH (A) ANC ([mmol.sub.c]/kg) Control 0 4.8 34 Stylo leaves 6.8 5.7 * 42 * Stylo roots 2.8 4.9 * 35 Urochloa tops 2.6 5.0 ** 36 * Urochloa roots 1.3 4.9 34 Plant material Urochloa soil pH (A) ANC ([mmol.sub.c]/kg) Control 5.3 41 Stylo leaves 5.6 *** 47 Stylo roots 5.3 44 Urochloa tops 5.4 ** 45 Urochloa roots 5.3 42 Significance of differences between controls and plant amended soils, using t-test: * P [less than or equal to] 0.05; ** P [less than or equal to] 0.01; *** P [less than or equal to] 0.001. (A) In 0.04 M KN[O.sub.3], 1:12.5 soil:solution ratio.
This study was generously supported by the Meat and Livestock Australia through project NAP3.218. The authors would like to thank the laboratory staff at CSIRO Land and Water in Townsville and at the Ruhr-University of Bochum for technical assistance.
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Manuscript received 13 November 2001, accepted 6 May 2001
M. Marx (A), B. Marschner (BD), and P. Nelson (C)
(A) Institute for Soil Ecology, GSF, 85764 Neuherberg, Germany.
(B) Ruhr Universitat Bochum, Institut fur Geographic, 44780 Bochum, Germany.
(C) PNGOPRA, PO Box 28, Popondetta, Oro Province, Papua New Guinea.
(D) Corresponding author, email: firstname.lastname@example.org
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|Author:||Marx, M.; Marschner, B.; Nelson, P.|
|Publication:||Australian Journal of Soil Research|
|Date:||Dec 1, 2002|
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