Carbon, nitrogen and sulphur cycling following incorporation of canola residue of different sizes into a nutrient-poor sandy soil

Soil Biology & Biochemistry 38 (2006) 32–42 www.elsevier.com/locate/soilbio

Carbon, nitrogen and sulphur cycling following incorporation of canola residue of different sizes into a nutrient-poor sandy soil Bhupinderpal-Singha,*, Z. Rengela, J.W. Bowdenb a

Soil Science and Plant Nutrition (M087), School of Earth and Geographical Sciences, University of Western Australia, 35 Stirling Highway Crawley, Crawley, WA 6009, Australia b WA Department of Agriculture, P.O. Box 483, Northam, WA 6401, Australia Received 19 October 2004; received in revised form 3 March 2005; accepted 10 March 2005

Abstract No information is available on the role of particle size of canola (Brassica napus) residue in altering C mineralization and nutrient (N, S) cycling in soil. We studied decomposition of canola residue (at 20G1 8C temperature and 10% moisture (w/w) for 6 months to elucidate the effect of its particle size (!1, 5–7, and 20–25 mm) on dynamics of C, N and S turnover following incorporation into a nutrient-poor sandy soil. Averaged over time, particle size of canola residue did not significantly affect C mineralization rate, the size of microbial-C and microbialK N pools, or the extent of CaCl2-extractable S immobilization, but altered the extent of mineral-N ðNHC 4 ; NO3 Þ immobilization and watersoluble organic C (W-SOC) depletion. A rapid decrease in C mineralization rate in the first week matched the rapid depletion of W-SOC, especially for the !1 mm residue treatment. Over 6 months, mineral-N in the amended soils rarely increased beyond the starting level (0.8–1 mg kgK1 soil for all the treatments), whereas nitrate-N increased 19-fold in the non-amended soil. This suggests an occurrence of strong N immobilization in the amended soils; such immobilization was high for the !1 mm residue treatment. On a cumulative basis, 33–35% of C added in canola residues to the soil was respired in 6 months. The microbial-C and microbial-N pools peaked by day 4 for all the residue treatments (compared to time zero, 58–122% increase for microbial-C and 36–57% for microbial-N). Averaged over time, amended soils contained approx. 40% more microbial-C and microbial-N than the non-amended soil. An addition of canola residue (regardless of the size) to soil increased the extractable S significantly (3.4-fold) on day 0; this initially increased S level decreased by one-third over 6 months. In conclusion, particle size of canola residue did not affect temporal pattern of C and S mineralization in a nutrient-poor sandy soil, but altered N cycling. q 2005 Elsevier Ltd. All rights reserved. Keywords: Canola stubble; Decomposition; Microbial biomass; Nutrient cycling

1. Introduction Canola, an oilseed rape (Brassica napus L.), has become an important crop in Western Australia. Canola requires relatively large amount of nitrogen (N) and sulphur (S) per yield unit compared to most grain crops. Canola stems make up a large proportion of the total dry matter returned to soil and contain nutrients, especially S, in larger proportions than in pod walls and seeds (McGrath and Zhao, 1996). However, at present, little is known about decomposition * Corresponding author. Tel.: C61 8 6488 2501; fax: C61 8 6488 1050. E-mail address: [email protected] (BhupinderpalSingh).

0038-0717/$ - see front matter q 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2005.03.025

dynamics of canola residues upon incorporation into nutrient-poor sandy soils of Western Australia. Given that soil microorganisms grow rapidly during the decomposition of plant residues, some nutrients (N, S) present in soil and/or released from residues would be immobilized (Ocio et al., 1991; Wu et al., 1993; Jensen et al., 1997; Magid et al., 1997), especially if low quality plant residues (such as canola stems residue having wide C:N, C:S and lignin:N ratios) were incorporated into a nutrient-poor soil. Plant residue size is a factor that could influence decomposition of added residues to soil, besides a number of other factors, such as soil temperature, soil type, nutrient and water availability in soil, chemical nature and amount of residues and the soil–residue contact (Swift et al., 1979). The literature is unclear on the effect of plant residue

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

particle size on decomposition and nutrient release dynamics after incorporation into soil (Sims and Frederick, 1970; Ambus and Jensen, 1997; Bending and Turner, 1999), mainly because residue particle size interacts with other factors to influence decomposition. With the reduction in particle size, the ratio of surface area of plant residue to mass of soil would increase (Angers and Recous, 1997), thus finer-sized plant residue could be expected to decompose faster than the coarse ones. However, physical protection of ground plant material by more intimate contact with clay and other soil particles could protect them against rapid decomposition, especially in the early phase after incorporation (Sims and Frederick, 1970; Jensen, 1994). Chemical composition of plant residue is another factor that could interact with particle size to influence decomposition and nutrient cycling in soil (Bending and Turner, 1999). Usually, for crop residues of high quality (low C:N ratio), such as potato shoot (Bending and Turner, 1999) and lentil green manure (Bremer et al., 1991), residue particle size had no effect on the rate of CO2–C respiration by microbes. In contrast, the low quality residues (such as wheat straw with high C:N ratio), smaller-sized residues decomposed faster than larger-sized particles, because of their increased surface area and more uniform distribution in soil volume, resulting in increased accessibility of easily decomposable substrates (and nutrients in them) to microbial attack (Ambus and Jensen, 1997; Angers and Recous, 1997). Summerell and Burgess (1989) compared decomposition of surface-placed and incorporated (by rotary hoeing) wheat straw to a field site in New South Wales (Australia) and attributed enhanced decomposition of rotary-hoed straw to the mechanical breakdown of the lignified tissues that allowed microorganisms an increased access to nutrients and to the parenchymatous tissues of the straw, providing favourable microclimatic conditions for microbial activity. On the other hand, senescent microbial tissues and products formed during decomposition of smaller-sized plant residues may also be stabilised to a greater extent (compared with the larger-sized residues) against further decomposition due to their (i) intimate mixing with mineral soil (Sims and Frederick, 1970; Jensen, 1994) or (ii) reaction with soluble polyphenols (related to humic and fulvic acids) released during decomposition of lignin (Stevenson, 1994). These polyphenols (i) could be released faster from ground residues than larger-sized ones (Bending and Turner, 1999), and (ii) are known to form recalcitrant complexes with a variety of N-containing compounds in decomposing organic matter and dead microbes (Bending and Read, 1996; Bending and Turner, 1999). Further, potentially available S in decomposition products and senescent microbial tissues could become unavailable due to transformation to stable forms in soil (Wu et al., 1993). It is not known how the particle size of canola residue would affect dynamics of C, N and S after incorporation into a nutrient-poor sandy soil, as physical protection of

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decomposing residues would be expected to be small in a sandy soil (Sims and Frederick, 1970) due to lack of sufficient adsorptive surfaces offered by small proportion of silt and clay in the soil. However, differential rates of decomposition of biochemical components (water-soluble C, hemicellulose, cellulose and lignin) of canola residues of different sizes could be expected (Bending and Turner, 1999) and this in turn could alter nutrient dynamics in the residue-amended soil. The main aim of our study was to describe the effect of canola residue and its particle size on the dynamics of CO2–C mineralization, water-soluble organic C (W-SOC) release, microbial-C and microbial-N pools turnover, and plant-available N and S status after incorporation into a nutrient-poor sandy soil.

2. Materials and methods 2.1. Soil and plant material and preparation for incubation A brown sandy soil collected from the top 0 to 10 cm layer of bushland near Lancelin (31801 0 S, 1158 20 0 E), Western Australia, was used in the study. The soil was airdried, and sieved (2 mm sieve), any visible undecomposed plant material was removed, and the soil was analysed for selected properties (pH-H2O 5.21, clay 4.5%, silt 6.5%, sand 89%, total C 7.5 g kgK1, total N 0.21 g kgK1, total S 0.035 g kgK1). The Lancelin soil (750 g, air-dried) was placed in plastic containers (8 cm diameter!12 cm height) (bulk density of approximately 1.5 t mK3 soil) to a depth of 10 cm; moisture content was adjusted to 10% (w/w). The soil in each container was inoculated with 1 g of wet (moisture 40% (w/w)) soil plus green waste based manure (mixed at 60 m3 haK1) applied as a suspension in water. The inoculation was used to raise the initial soil microbial population level in the Lancelin soil. Two such containers (total 1500 g soil) were then placed in a sealed 5 l bucket (representing one treatment within each replicate and time) and pre-conditioned aerobically for a week in the dark at 20 8C. Canola residue used in this study was collected from a recently harvested field at Meckering (31838 0 S, 117800 0 E) in Western Australia, air-dried, and stored at 4 8C in a sealed plastic bag for up to 4 months before use. There were three particle size treatments: the canola residue was either ground (!1 mm) using a grinder, or chopped to 5–7 and 20–25 mm length manually using a pair of scissors. The canola residues mainly consisted of stems (upper and lower), side branches, and pedicles (but no leaves or pod walls). A portion of canola residue of each size was further ground in a coffee grinder and analysed for some basic properties (Table 1). The analytical data were later corrected for moisture content after drying canola residue of different sizes at 60 8C for 96 h.

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Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

Table 1 Some quality parameters (on dry-weight basis) of the canola residues of different sizes (!1, 5–7, 20–25 mm) used in the decomposition experiment Residue quality parameters

Range across residue particle sizes

Total N content (g kgK1 plant material) C:N ratio Total S content (g kgK1 plant material) C:S ratio Ash (%) Water-soluble C (g kgK1 C) Neutral detergent soluble (%) Hemicellulose (%) Cellulose (%) Lignin (%)

3.4–3.9 121–144 1.4–1.6 298–345 3.9–4.1 40–51 16.4a 21.4a 51.1a 11.2a

a The step-wise chemical fractionation was performed on the 5–7 mm cut canola residue according to Goering and van Soest (1970).

2.2. Experimental details One-week after pre-conditioning, canola residue of various sizes was homogenously mixed with the wet soil to a concentration of 3.6 g plant material (air-dry basis) per kg soil, corresponding to 5.4 t air-dry residues per ha to a depth of 10 cm (bulk density 1.5 t mK3); the containers were refilled after mixing. A control treatment with soil, but without additions of the plant material, was also set-up. Following amendments, containers were placed back in the same 5 l sealed plastic buckets containing 100 ml of distilled water to maintain a water-saturated atmosphere and 30 ml of 2 M NaOH to trap microbially respired CO2. The sealed buckets were then wrapped with aluminium foil and placed in a dark room for incubation at 20G1 8C. The weight of the containerCsoilCcanola residues was recorded and adjusted for moisture loss as needed. The experiment comprised three different-sized canola stems and one control, three replicates for each treatment, and seven harvest times (0, 1, 4, 10, 28, 56, and 183 days). Additionally, two extra buckets for each replicate containing no soil (but two sealed empty plastic containers of 600 ml each, 100 ml water and a CO2 trap) were also set-up to account for CO2 in the enclosed space. The three replications for each treatment were staggered in time (10 days). At each harvest, part of the soil from each container was removed for moisture determination after drying at 105 8C for 48 h, another portion (60 g) for W-SOC analysis (see below), and 1100 g soil for evaluating a fractionation procedure for characterizing decomposing residues and organic matter in soil (data not presented). The remaining soil was kept at 4 8C for up to 7 days for various analyses (see below). 2.3. Analyses Soil respiration was monitored by changing NaOH in the sealed buckets on 1, 2, 4, 7, 10, 15, 22, 28, 42, 56, 77, 100,

120, 141, 162, and 183 days and measuring trapped CO2. 2K The HCOK 3 and CO3 ions in the 1 ml of 2 M NaOH were precipitated with 10 ml of 1 M BaCl2 solution, and total respired CO2–C was determined by titrating the residual NaOH with 0.1 M HCl using phenolphthalein as indicator. Water-soluble organic C at each harvest time was determined by shaking 60 g of moist soil with 600 ml distilled water for 1.5 h, filtering supernatant through glass fibre filter (GF,C) and analysing W-SOC with a Shimadzu TOC-5000A analyzer (Shimadzu Corporation, Kyoto, Japan). An aliquot (3.5 ml) of the water extract was preacidified with 20 ml of 4 M HCl before injection of 53 ml of the acidified extract into a combustion furnace heated to 680 8C, with subsequent detection of CO2 using a nondispersive infrared gas detector. Standards for C were made using potassium hydrogen phthalate. Microbial C was determined according to Joergensen and Brookes (1990). Briefly, about 22 g moist soil samples were fumigated for 24 h in the presence of chloroform containing amylene (0.006% v/v) as a stabiliser, followed by extraction with 80 ml of 0.5 M K2SO4. The 0.5 M K2SO4 extract was then diluted 7-fold with MilliQ water prior to analysis by TOC analyser. The diluted extract was pre-acidified with 20 ml of 4 M HCl. A non-fumigated soil sample was also extracted the same way as above, and organic C values were deducted from the fumigated sample to calculate microbial C flush. Microbial ninhydrin-N was analysed in the 0.5 M K2SO4 extract of fumigated and non-fumigated soil (Joergensen and Brookes, 1990). The 0.5 M K2SO4 extract from non-fumigated soil was also analysed for mineral-N K ðNHC 4 ; NO3 Þ by an automated Skalar segmented-flow analyser. The method for the determination of NHC 4 –N is based on the modified Berthelot reaction (Searle, 1984), and for NOK 3 –N on the hydrazinium reduction reaction (Kempers and Luft, 1988). Also, moist soil samples (22 g) were extracted with 40 ml of 0.01 M CaCl2, and total S in the CaCl2 extract was determined by Inductively Coupled Plasma Atomic Emission Spectroscopy (ICP-AES). Microbial C-to-N ratio and metabolic quotient (CO2–C mineralization rate per unit of microbial-C formed) were derived using corresponding measured variables within each replicate. 2.4. Statistics The experiment was set-up in a randomised complete block design. Two-way analysis of variance (ANOVA) was performed using GenStat 7.1 on all the variables, with treatments and time as independent and interactive sources of variance. When significant F-tests were obtained, means separation was achieved using a Least Significant Difference (LSD) test at the 0.05-probability level. All the measured variables, microbial C-to-N ratio and metabolic quotient data were evaluated for normality based on the visual observation of residual and normal probability plots obtained using GenStat 7.1. All the data were found

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

normally distributed, except the CO2 respiration rate, for which only less than 7% data (especially with larger respiration rates) had higher residuals for treatment!time combinations. Furthermore, on comparing ANOVA of nontransformed and log-transformed data (to overcome nonnormal distribution in the data), we found no change in the interpretation of main and interaction effects. Therefore, we present the non-transformed data only.

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the cut residues. This largest C mineralization rate in the ground residue-amended soil coincided with the significant decrease (52%) in W-SOC from days 0 to 1 (Fig. 1b). After day 1, W-SOC did not change in the ground residue treatment for up to day 10, then decreased significantly by day 28, and stabilised thereafter. Further, the rate of decrease of C mineralization was steeper for the ground than the two cut residue treatments for up to day 10. The soil treated with the 20–25 mm residue showed a significant increase in the C mineralization from days 1 to 2 before decreasing rapidly by day 10 and then gradually levelling off. The initial rapid decrease in C mineralization rate for the two cut residue-amended treatments was comparable (rZ0.75–0.82*, P%0.05, nZ9) with the rapid depletion of W-SOC in the first 1–10 days (cf. Fig. 1a and b). A plot of CO2–C mineralization rate against cumulative proportion of residue-C respired (Fig. 2), which compares the rates at equivalent degree of decomposition, showed that any difference in C mineralization rates among the residue treatments disappeared after 4–5% of residue-C was mineralised in 10 days. The values of cumulative CO2–C respired from the amended soils were similar for various treatments (data not shown), reaching 33–35% of added residue-C after 183 days of incubation (Fig. 2).

3. Results 3.1. Carbon mineralization and water-soluble organic C The two-way ANOVA showed a significant effect of treatment and time factors as well as a significant treatment!time effect on the CO2–C mineralization rate and W-SOC content during incubation (see insert in Fig. 1a and b). The CO2–C mineralization rate for the soil amended with canola residue was significantly greater (up to 2–3 times) than for the non-amended soil during the period of study (Fig. 1a). On day 1, the CO2–C mineralization rate was greater for the soil amended with the ground (!1 mm) than

mg CO2-C respired kg–1 soil day–1

(a) Control

20 Source of variation 16

C mineralization rate

Treatment Time Treatment × Time Residual

12

df 3 15 45 126

F value 162.8 289.0 11.6 –

<1 mm 5-7 mm

P <0.001 <0.001 <0.001 –

20-25 mm

8

4

0

mg of water-soluble organic C kg–1 soil

(b) 140 120

Source of variation

100

Treatment Time Treatment × Time Residual

80

Water-soluble organic C df 3 6 18 54

F value 22.8 53.5 4.4 –

P <0.001 <0.001 <0.001 –

60 40 20 0 0

5

10

15

20

25

30

40

60

80

100

120 140

160 180

Days after incorporation Fig. 1. Influence of canola residue particle size (!1, 5–7, and 20–25 mm) on temporal pattern of (a) C mineralization rate and (b) water-soluble organic C in amended compared with non-amended soil during 183 days of incubation. Note the change of scale after the break on the x-axis (i.e. days after incorporation). The thick bars are the LSD0.05 for treatment!time ((a)Z1.49 and (b)Z21.1, P%0.05). Inserts in figures are the results of ANOVA comparing C mineralization rate and water-soluble C among residue-amended and non-amended treatments over time.

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

mg CO2-C respired kg–1 soil day–1

36

<1 mm

20

5-7 mm 20-25 mm 15

10

5

0

4

8

12

16

20

24

28

32

36

Cumulative CO2 respired (% of added C) Fig. 2. Rate of C mineralization as a function of cumulative proportion of CO2–C respired from canola residue of different sizes incorporated into sandy soil.

Averaged over time, W-SOC was significantly lower in the non-amended soil (29 mg kgK1 soil) compared to the amended soil. Within the amended treatments, W-SOC in the !1 mm residue-amended soil (46 mg kgK1 soil) was significantly lower than the two cut-residue treatments (56– 58 mg kgK1 soil), which had similar W-SOC content. Comparing across treatments and times, W-SOC was similar in all the residue-amended soil on day 0, which decreased rapidly to 19–21% of the initial value by day 28 and levelled-off thereafter, except for the 5–7 mm treatment. For the !1 mm size treatment, the day 0 W-SOC declined steeply (48–55% decrease) by days 1–4 compared to 27–40% decrease for the 20–25 mm residue and 30–54% for the 5–7 mm residue. 3.2. Microbial biomass C and N The two-way ANOVA showed significant treatment, time, and treatment!time effects for microbial-C, microbial-N (see insert in Fig. 3a and b) and metabolic quotient (Table 2). For microbial C-to-N ratio, only time effect was significant (Table 2). Averaged over time, the residue-amended soil had similar microbial-C and microbial-N, which were significantly greater than in the non-amended soil. In the nonamended soil, microbial-C and microbial-N did not change significantly during incubation. Microbial-C and microbial-N in all the treatments, including non-amended, were similar on day 0 (Fig. 3a and b). Microbial-C peaked in the amended soils by day 4 (between 58 and 122% increase compared to the initial level on day 0, Fig. 3a), and then gradually decreased by day 28 or 56 depending on the treatments, and stabilised thereafter at a consistently higher level than in the non-amended soil. Similarly, microbial-N peaked in the amended soils by day 4 (36–57% increase compared to the initial level on day 0, Fig. 3b), followed by a decrease close to the initial level by day 10 for the 5–7 mm size residue, day 28 for

the 20–25 mm size residue, and day 183 for the !1 mm residue treatments. A second peak in microbial-N was found on day 28 for the 5–7 mm residue, or day 56 for the 20–25 mm residue size treatments. Averaged over time, all the residue treatments had similar metabolic quotient values that were significantly higher than in the non-amended soil. The ground residue (!1 mm) treatment had higher metabolic quotient on day 1 followed by the two cut-residue treatments (having similar metabolic quotients) and the non-amended treatment (Table 3). With time, the metabolic quotient decreased by 67% on day 10 for the ground residue and 50% for the other treatments (including non-amended) (P%0.05), and stabilised thereafter (Table 3). Microbial C-to-N ratio was similar for all the residueamended and non-amended treatments (6.4–7.1, averaged over time) (data not shown). Averaged across treatments, microbial C-to-N ratio on day 0 increased (P%0.05) 29– 33% by days 4–10, decreased thereafter and stabilised close to the initial level (Table 3). 3.3. Mineral- and amino-N and CaCl2-extractable S in soil The two-way ANOVA showed significant treatment, time, and treatment!time effects for mineral-N K (NHC 4 ; NO3 , and total) and ninhydrin-reactive N. For the CaCl2-extractable S, only time and treatment effects were significant (Table 2). K Averaged over time, NHC 4 –N; NO3 –N and ninhydrin-N in the non-amended soil were significantly greater than in the residue-amended soils, and within the residue treatments, these N-forms were significantly greater in the 20–25 mm residue than in the 5–7 mm or ground residue K treatments, which had similar NHC 4 –N; and NO3 –N. Ninhydrin-N for the ground residue treatment was significantly greater than for the 5–7 mm treatment. On day 0, NHC 4 –N was similar in all the treatments, except for the soil amended with !1 mm residue, where

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

(a) 180

Microbial-C (mg kg–1 soil)

160

140

Source of variation

Microbial-C

df Treatment 3 Time 6 Treatment × Time 18 Residual 54

F value 12.4 7.7 1.8 –

37

P <0.001 <0.001 0.05 – Control < 1 mm

120

5-7 mm 20-25 mm

100

80

60

Microbial-N (mg kg–1 soil)

(b) 27

Source of variation

Microbial-N

24

df Treatment 3 Time 6 18 Treatment × Time Residual 54

F value P 16.8 <0.001 3.1 0.01 11.6 0.03 – –

21

18

15

12

9 0

10

20

30

40

50

60

90

120

150

180

Days after incorporation Fig. 3. Influence of canola residue particle size (!1, 5–7, and 20–25 mm) on temporal pattern of (a) microbial-C and (b) microbial-N during 183 days of incubation. Note the change of scale after the break on the x-axis (i.e. days after incorporation). The thick bars are the LSD0.05 for treatment!time ((a)Z33.9 and (b)Z4.7, P%0.05). Inserts in figures are the results of ANOVA comparing microbial-C and microbial-N among residue-amended and non-amended treatments over time.

NHC 4 –N was the greatest (Fig. 4a). In the non-amended soil, NHC 4 –N was significantly greater than in the amended soils between days 4 and 28. The size of NHC 4 –N decreased rapidly by day 4 in all the residue treatments; the decrease was significantly greater for the !1 mm residue treatment. After an increase from days 4 to 10 in all the treatments, NHC 4 –N decreased rapidly and levelled-off after day 56, except for the !1 mm residue treatment, where a significant increase occurred on day 183 (Fig. 4a). The ninhydrin-reactive compounds (amino-NCNHC 4 –N) in the amended and non-amended soils followed a similar pattern as for NHC 4 –N until day 28 of incubation (cf. Fig. 4a and b). About a 2-fold increase in the amount of ninhydrinreactive N on days 56 and 183 was found when compared to NHC 4 –N alone, suggesting the contribution of microbial amino-N compounds to the available-N pool in the amended soil (Fig. 4b).

K1 soil) in The size of NOK 3 –N was similar (0.8–1 mg kg all the treatments on day 0, and did not change in any treatment in the first 10 days after incorporation. By day 28 of incubation, NOK 3 –N decreased 9-fold in the !1 mm residue treatment. In the 20–25 mm residue treatment, NOK 3 –N increased from 1 to 1.8 mg kgK1 soil and then levelled off, whereas for the 5–7 mm residue treatment, the change in NOK 3 –N was not significant at any time (Fig. 4c). Interestingly, NOK 3 –N level continued to increase appreciably in the non-amended soil 10 days after day 0 and reached a significantly higher level (16.6 mg kgK1 soil) at the end of incubation in comparison to the levels in amended soils (i.e. 0.3 and 0.8 mg kgK1 soil for the ground and 5–7 mm residue treatments, respectively, which were K1 soil in the significantly lower than the 1.9 mg NOK 3 –N kg 20–25 mm residue treatment). The increase in NOK 3 –N (from 1.2 to 7.8 mg kgK1 soil) in the non-amended soil

!0.001 0.011

46

55.5 2.4 5 15

54

61.5 5.5 6 18

The degrees of freedom (df) in column 2 are common across all the listed variables (except for the metabolic quotient, for which df are presented separately).

!0.001 0.484 9.3 1.0 0.024 0.972 2.7 0.4 !0.001 !0.001 33.7 5.2 !0.001 !0.001 40.3 54.9 !0.001 !0.001 132 122

19.6

!0.001 !0.001

P F

14.3 3

df P

!0.001 200

F P

0.586

F

0.7

P

0.002 F

5.6 !0.001

P F

262 !0.001

P F

331

F

3

Treatment Time Treatment! time Residual

P

NOK 3 –N NHC 4 –N df Source of variation

!0.001

Metabolic quotient CaCl2-extractable S Microbial C-to-N ratio Ninhydrin-N Mineral-N ðNHC 4C NOK 3Þ

!0.001

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42 Table 2 Results of two-way analysis of variance (ANOVA) comparing mineral-N, ninhydrin-N, microbial C-to-N ration, extractable S, and metabolic quotient among residue-amended (!1, 5–7, and 20–25 mm) and non-amended treatments over time

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between days 10 and 56 coincided with the decrease in K1 NHC soil) during this period, 4 –N (from 3.7 to 0.2 mg kg K explaining 53% increase in NO3 –N in the soil (cf. Fig. 4a and c). Averaged over time, the size of the total mineral-N pool was larger in the non-amended soil (6.8 mg kgK1 soil) followed by 20–25 mm residue (3.0 mg kgK1 soil), 5–7 mm residue (2.3 mg kgK1 soil) and then !1 mm residue-treated soil (1.9 mg kgK1 soil) (Fig. 4d). For various residue treatment and time combinations, mineral-N in the ! 1 mm and 5–7 mm residue treated soil was similar at all the times and was significantly (P!0.001) smaller than in the 20–25 mm residue treated soil 4 days after incorporation (except on day 28 for the 5–7 mm residue treatment) (Fig. 4d). Depending on the tissue-S content (Table 1), canola residue incorporation into the soil was expected to supply 4.7–5.3 mg S kgK1 soil. However, on average, 77% of added S in the canola residues was immediately (on day 0) extracted in 0.01 M CaCl2, thus increasing the extractable-S level in all the amended soils by approximately 3.4-fold (P!0.001) compared with the non-amended soil (Fig. 5). Averaged over time, the residue treatments had similar amounts of extractable-S that were significantly greater than in the non-amended soil. Averaged across treatments (including non-amended), the amount of extractable-S gradually decreased (P!0.001) to about two-thirds of the initial level over the 6 month period.

4. Discussion The results of the present study suggest that the particle size of canola residues did not significantly alter temporal patterns of C and S mineralization, but did so for the N mineralization. The lack of significant differences (except in the first week) among the residue treatments for C mineralization was not expected, because by reducing the particle size, a greater surface area of contact with soil (cf. Angers and Recous, 1997) could increase accessibility of substrates in the residues to microbial attack (Jensen, 1994; Ambus and Jensen, 1997). It was also thought that physical protection of decomposing organic matter (especially likely for the ground residues) would be relatively low in the sandy soil in the present study (e.g. Sims and Frederick, 1970). Both the soil and the canola residue used were low in N content. Thus, one possibility for the lack of treatment differences could be the limitation to microorganisms of available N that would have decreased the overall decomposition rate for all the treatments. Increasing N availability in soil through external N applications would possibly promote decomposition of relatively labile substrates of plant materials during early stages, but may suppress decomposition of recalcitrant substrates, such as lignin at later stages (Wang et al., 2004). It is possible that any N-induced differences in the particle-size treatments

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

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Table 3 Influence of particle size of canola residue on metabolic quotient (mg CO2–C respired mgK1 microbial-C formed 24dK1) and microbial C-to-N ratio Metabolic quotienta

Days after incorporation

Microbial C-to-N ratiob, averaged across all treatments

Canola residue particle sizes 0 1 4 10 28 56 183

Non-amended

!1 mm

5–7 mm

20–25 mm

NA 2.2c 1.5c 1.1b 0.8b 0.9a 0.6b

NA 4.2a 1.8bc 1.4ab 1.6a 1.1a 1.1ab

NA 3.5b 2.2ab 1.7a 1.3ab 1.3a 1.2a

NA 2.9b 2.5a 1.5ab 1.8a 1.2a 1.2ab

5.8 7.0 7.6 7.8 6.6 5.6 6.3

Microbial C-to-N ratio was averaged across treatments because of no significant differences among treatments. Comparing across residue-amended and nonamended treatments, the metabolic data followed by the same letter (a, b, c) are not significantly different at P%0.05. NA, not applicable. a LSD0.05 for treatment!timeZ0.6. b LSD0.05 for time effectZ1.4.

could be nullified or reversed over the long term. Thus, especially for ground residues, N supply could either increase decomposition in the short term, due to increased residue-soil contact or suppress decomposition in the longterm, due to formation of recalcitrant lignin (polyphenols)N complexes (Bending and Turner, 1999), because lignin (polyphenols) would be more exposed by grinding of lignified tissues. This interactive effect of particle size with increasing N availability may be difficult to interpret and merits detailed examination in further studies. In our study, the rapid decrease (52%) in W-SOC by days 1 and a subsequent slow decrease until day 10 for the !1 mm (b)

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residue treatment compared to the two cut residue treatments could explain the initial significant differences among treatments for the CO2–C mineralization rates (i.e. slower rate for the !1 mm residue) (cf. Fig. 1a and b). It is likely that grinding of intact lignified tissues of canola residues made soluble organic matter in them more accessible to microbes immediately following incorporation, as consistent with the results of Ambus and Jensen (1997). The soil amended with the 5–7 mm residue showed a significant increase in W-SOC at the end of incubation, which may have occurred due to breakdown of some complex C components (such as cellulose) to water-soluble compounds by

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C Fig. 4. Influence of the canola residue particle size (!1, 5–7, and 20–25 mm) on temporal pattern of (a) NHC 4 –N, (b) ninhydrin-reactive compounds (NH4 –N K C K and amino-N), (c) NO3 –N, and (d) total mineral-N ðNH4 –C NO3 –NÞ during 183 days of incubation. Note the change of scale after the break on the x-axis (i.e. days after incorporation). The thick bars are the LSD0.05 for treatment!time ((a)Z0.71, (b)Z0.74, (c)Z0.76 and (d)Z1.04, P%0.05).

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Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

Control <1 mm 5-7 mm 20-25 mm

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Days after incorporation Fig. 5. Influence of the canola residue particle size (!1, 5–7, and 20– 25 mm) on temporal pattern of 0.01 M CaCl2-extractable S during 183 days of incubation. Note the change of scale after the break on the x-axis (i.e. days after incorporation). Bars on the data points represent CSE (nZ3). The treatment!time interaction was not significant at P%0.05. Treatment and time effects were significant (LSD0.05 for treatmentZ0.34 and for timeZ0.44).

the microorganisms (Summerell and Burgess, 1989). Why this happened only for the 5–7 mm residue than for the other two residue sizes was not known. We found no effect of canola residue particle size on the size and pattern of microbial C and N during incubation, in agreement with the results of Ambus and Jensen (1997). Microbial biomass remained larger in the amended versus non-amended treatment throughout (Fig. 3a and b), suggesting their slow but sustained growth on the other components of canola residue after the depletion of W-SOC. The increase in microbial C by 58–122% and microbial N by 34–57% (range covers residue particle sizes) following incorporation in the first 4 days (Fig. 3a and b) compares well with a 2-fold increase in microbial C and N 7 days after field incorporation of winter wheat straw (at 10 t haK1, C:N ratio 48:1), as observed by Ocio et al. (1991). This initial rapid increase in microbial C or N is likely occurring on labile substrates (including water-soluble organic matter). The subsequent significant increases were observed for microbial-N on 28 or 56 days after residue incorporation, but not for microbial-C, suggesting a greater demand for available N in soil by microbial populations because of their greater need for the synthesis of extracellular enzymes during decomposition of relatively recalcitrant substrates at later stages (Stewart et al., 1966). Azam et al. (1988) and Williamson and Johnson (1994) found that, though the initial rate of N immobilization by microbes was greater with glucose, addition of cellulose caused a slow but sustained and long-lasting immobilization of N in microbial biomass. Chapman (1997) observed the same effect of cellulose versus glucose degradation on S immobilization by microbes. Metabolic quotient was high (i.e. low microbial substrate use efficiency) initially, but then decreased as

the decomposition progressed (Table 3). This suggests that although microbes may grow rapidly on simpler W-SOC substrates than on cellulose and lignin (Williamson and Johnson, 1994; Tate, 1995; Mueller et al., 1998), being faced with an immediate supply of readily available substrates, microbial communities tend to use up (waste) energy in rapid uncoupled growth and turnover (Tempest and Neijssel, 1984; Tate, 1995). Thus, a low N requirement of microbes during the degradation of easily decomposable substrates may result from a rapid turnover of microbial-N. Microbial C-to-N ratio varied between a narrow range (5.6–7.8, Table 3) during incubation despite the incorporation of a wide C:N ratio canola residue. Jensen et al. (1997) also did not find marked variation in the microbial C-to-N ratio during decomposition of oilseed rape straw during 1 year. A significant increase in microbial C-to-N ratio in the first 4–10 days followed by a decrease by day 56 (Table 3) may imply a rapid, but for a short term, colonisation of canola residue by fungi, as observed by Strong et al. (2004) for wheat husks. The changes in microbial-N during decomposition were not distinctly reflected in the pools of total mineral-N C or NOK 3 –N (except for NH4 –N) in the amended soils (Fig. 4a,c,&d). Further, amino-N of microbial origin (measured as an indicator of microbial-N turnover) became distinct 56 days after the residue incorporation in soil (Fig. 4b). Prior to day 56, microbial-N that turned over could have been rapidly re-metabolised by the growing microbial biomass or transformed to stable organic-N fractions upon reaction with soluble polyphenols (Bending and Read, 1996). Furthermore, increases in microbial-N in the first 56 days were larger in magnitude compared to corresponding decreases in total mineral-N in all the amended soils (after taking into account the corresponding levels in the nonamended soil), suggesting that residue-N was also immobilized (cf. Figs. 3b and 4d). Thus, changes in microbial biomass may not serve a direct measure of a source or sink for available-N in soil, as also reported by Jensen et al. (1997). The initial high NHC 4 –N in the non-amended soil on day 4 could have possibly resulted from mineralization of N in dead microbes that became available immediately after initial mixing of the soil (e.g. microbial N decreased, albeit non-significant, by 4 mg kgK1 soil between days 1 and 4, see Fig. 3b). This available NHC 4 –N in the nonamended soil decreased rapidly because of its continuous K oxidation to NOK 3 –N and NO2 –N (autotrophic nitrification) (Fig. 4c). The brown sandy soil used in the present study, having a low native organic matter content (Table 1), would not support fast growth of heterotrophs that could cause NHC 4 –N immobilization (Jones et al., 2004; Cookson and Murphy, 2004). In this situation, autotrophic nitrifies could compete successfully for NHC 4 –N resulting in NOK 3 –N accumulation (Fig. 4c). On the other hand, residue-amended soil with a supply of readily available C substrates supported greater heterotrophic growth and use of N (Fig. 3b).

Bhupinderpal-Singh et al. / Soil Biology & Biochemistry 38 (2006) 32–42

The present study shows a pattern of increased N immobilization with the decrease in the residue particle size (Fig. 4a–d). However, microbial-N was similar among the residue treatments, suggesting that greater N immobilization other than in microbial biomass also occurred in the !1 mm residue treated soil compared to the two cut residues. This could be due to greater stabilisation and protection of N-containing monomers (organic matter) from further microbial attack upon reaction with soluble polyphenols (Bending and Turner, 1999) or even some adsorption on to a small amount of clay in the soil; such immobilization of N seems to be smaller than in microbial biomass. Jensen (1994), using pea residues (C:N 19.5), observed a greater immobilization of N in a sandy loam soil (clay 11.4%) amended with residues of smaller (1 mm) than larger (10 mm) particle sizes. We observed this effect even for canola residue with a wide C:N ratio (132:1) added into a sandy soil, suggesting that although residue particle size may not affect C mineralization, it could cause greater N immobilization in soil. Preservation of N in soil by the application of a wide C:N ratio residue, such as canola, could be especially important in conditions of leaching and denitrification, usually experienced in coarse-textured soils and during a period of high rainfall. Williamson and Johnson (1994) found 40% decrease in nitrate leaching in a wheat-straw amended compared to non-amended soil receiving an equivalent of 20 mm of precipitation each week. In our study, although K NHC 4 –N showed a decrease, NO3 –N rarely increased above the initial day 0 level in the canola-amended soil during incubation (Fig. 4d). Thus, nutrient-poor soils would require application of fertiliser N, where canola residues are to be incorporated, so that next crop should not experience N limitation from soil. Nevertheless, especially !1 mm residue treatment did show a trend towards N mineralization at the end of incubation (Fig. 4d). A longer incubation period (O6 months) would be required to elucidate the timing of release of immobilised-N in the canola residue amended sandy soil. The immediate (i.e. on day 0) 3.4-fold increase in the CaCl2-extractable S upon incorporation of canola residue to soil indicates that much of the tissue S existed in soluble inorganic sulphate and readily degradable organic-S forms (S in proteins, amino acids). Most of the extractable S was in sulphate-S form (85–102% for the amended and 69–87% for the non-amended soil) during incubation (data not shown). Wu et al. (1993) also found a rapid and significant release of S from rape leaves (50%) and barley straw (42%) as sulphate-S upon first extraction) of amended soils with 0.1 M KH2PO4 on day 5. In the present study, the initial increased extractable S level on day 0 in the amended soils was gradually decreased by ca. 33% over 6 months, suggesting that available S, like N, was either immobilised by microbial biomass, transformed to stable soil organic S (Wu et al., 1993), and/or re-incorporated into decomposing residue by microflora (Salas et al., 2003).

41

In conclusions, particle size of canola residue did not affect C and S mineralization patterns in a nutrient-poor sandy soil, and thus would not have practical relevance for a greater storage or loss of C in sandy soils in the long-term. Canola residue incorporation into a nutrient-poor soil could result in a net immobilization of available N and S over time in microflora, and through transformation to stable organic forms in soil. Canola residue addition can rapidly increase the size of available S pools in soil. Further, the smaller the particle size of canola residue, the greater the potential for immobilization of N in soil.

Acknowledgements The authors thank Australian Research Council and Agriculture Western Australia for financial support.

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