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Biotic and abiotic processes of nitrogen immobilization
in the soil-residue interface
Naoki Moritsukaa,*, Junta Yanaic, Keiko Morib, Takashi Kosakic
aEducation and Research Center for Biological Resources, Faculty of Life and Environmental Sciences, Shimane University, Shimane 690-1102, JapanbGraduate School of Agriculture, Kyoto University, Kyoto 606-8502, Japan
cGraduate School of Global Environmental Studies, Kyoto University, Kyoto 606-8501, Japan
Received 23 May 2003; received in revised form 19 February 2004; accepted 23 February 2004
Abstract
The interface between decaying plant residues and soil is a hotspot for microbial immobilization of soil inorganic N. Recent studies on
forest and grassland soils have demonstrated that rapid abiotic immobilization of inorganic N is also induced by the presence of plant
residues. We, therefore, examined (1) how N immobilization varies with distance from the soil-residue interface and (2) whether abiotic
immobilization occurs in agricultural soils. Spatiotemporal changes of N immobilization in the soil-residue interface were evaluated using a
box that enabled soil to be sampled in 2 mm increments from a 4 mm-thick residue compartment (RC). The RC was filled with paddy soil
containing ground plant residue (rice bran, rice straw or beech leaves) uniformly at a rate of 50 g dry matter kg21. Soil in the surrounding
compartments contained no residue. After aerobic incubation for 5, 15 and 30 days at 25 8C, soils in each compartment were analyzed. After
5 days, significant depletion of inorganic N occurred throughout a volume of soil extending at least 10 mm from the RC in all residue
treatments, suggesting extensive diffusion of inorganic N towards the RC. The depletion within 10 mm of the RC amounted to 5.0, 4.3 and
3.4 mg for rice bran, rice straw and beech leaf treatment, respectively. On the other hand, microbial N had increased significantly in the RC of
the rice bran and rice straw treatments (11 mg and 5.5 mg, respectively) and insignificantly in the RC of the beech leaf treatment (0.06 mg).
This increase amounted to 221% (rice bran), 129% (rice straw) and 1.7% (beech leaves) of the decrease in inorganic N within 10 mm of each
RC. Thereafter the rate of N mineralization exceeded that of immobilization, and inorganic N levels had recovered almost to their original
level by 15 days (rice bran) and 30 days (rice straw and beech leaves). These results suggested the predominance of biotic immobilization in
soil near rice bran and rice straw and of abiotic immobilization in soil near beech leaves. No significant increase in both microbial and soluble
organic N in the vicinity of beech leaves after incubation for 5 days further suggested that the abiotic process was responsible for the
transformation of inorganic N into the insoluble organic N.
q 2004 Elsevier Ltd. All rights reserved.
Keywords: Nitrogen immobilization; Plant residue; Microscale heterogeneity; Microbial biomass
1. Introduction
The zone of soil affected by decaying plant residues is a
hotspot of microbial activity in which the supply of
residue-derived C to microorganisms leads to intensive
immobilization of soil inorganic N to meet their metabolic
requirements. Since the supply of residue-C to the
surrounding soil extends for some 3–4 mm (Gaillard
et al., 1999; 2003), the effects of microbial incorporation
of soil inorganic N might be expected to be concentrated in
this area. However, there has been no quantification of the
distance to which residue affects N immobilization in the
surrounding soil.
It is generally assumed that the main process involved
in residue-induced N immobilization in agricultural soils
is a biological one (Mary et al., 1996; Frey et al., 2000).
Therefore it is the availability of organic C to decom-
posing soil microorganisms that usually determines
immobilization capacity (Recous and Machet, 1999).
Although it is known that NH3 can be converted to
non-exchangeable forms through condensation reactions
occurring between NH3 and polyphenols in soil organic
matter (Nommik and Vahtras, 1982), the relative
0038-0717/$ - see front matter q 2004 Elsevier Ltd. All rights reserved.
doi:10.1016/j.soilbio.2004.02.024
Soil Biology & Biochemistry 36 (2004) 1141–1148
www.elsevier.com/locate/soilbio
* Corresponding author. Tel.: þ81-852-34-0311; fax: þ81-852-34-1823.
E-mail address: [email protected] (N. Moritsuka).
significance of this abiotic process is presumed to be
much less than that of microbial immobilization (Kelley
and Stevenson, 1987). In contrast, it has been shown that
in other ecosystems, e.g. forests (Schimel and Firestone,
1989; Johnson et al., 2000) and semiarid grasslands
(Barrett et al., 2002), abiotic immobilization of NH4þ can
be an important process. Additionally, there is growing
evidence that rapid abiotic immobilization of NO32 also
occurs in forest soils. Berntson and Aber (2000) examined
the rate of immobilization of 15N-labeled nitrate added to
forest soils where either pine or hardwood stands
prevailed, and observed very rapid immobilization of
NO32 to occur in both soils, apparently by abiotic
processes. Furthermore, Dail et al. (2001) showed that
approximately 40–60% of the 15N-nitrate added to
sterilized soils disappeared from the extractable inor-
ganic-N pool within 15 min, to be subsequently detected
mainly in the soluble organic N (SON) fraction. Compton
and Boone (2002) also reported that only 8–16% of15N-nitrate and 12–18% of 15N-ammonium could be
recovered in an extractable inorganic form 5 min after
solutions of these nutrients were added to forest soils.
From these results, Davidson et al. (2003) proposed a
plausible hypothesis that explains rapid abiotic conversion
of NO32 into dissolved organic N, but the mechanisms
involved are not identified at present.
The objective of this study was to investigate residue-
induced N immobilization with a particular focus on two
points: (1) how N immobilization varies with distance from
the soil-residue interface and (2) whether abiotic immobili-
zation occurs to a significant extent in agricultural soils.
2. Materials and methods
2.1. Preparation of soil and residue materials
Residue materials were added to an air-dried, 2-mm
sieved paddy soil. The soil is classified as Typic Fluvaquent
with the following properties: total C 32.1 g kg21; total N
3.02 g kg21; clay 219 g kg21; silt 318 g kg21; pH (H2O, 1:5
w/v) 5.73; electrical conductivity (1:5 w/v) 0.11 dS m21;
0.5 M K2SO4-extractable inorganic N 57.6 mg kg21 (nitrate
18.8 mg kg21 and ammonium 38.8 mg kg21). Plant residue
materials used were rice straw (Oryza sativa L.), rice bran,
and leaves from beech (Fagus crenata L.). Beech leaves
were included to provide a comparison between agricultural
and deciduous forest residues. Plant material was dried
at 70 8C, ground with a ball mill and sieved to 0.71-mm.
Table 1 shows the concentration of C and N in each type of
residue. Total C concentrations were similar, but hot water-
soluble C and total N varied. The C/N ratio was highest for
beech leaves (45.5), followed by rice straw (25.5) and rice
bran (18.8). The relatively low C/N ratio of rice straw was
due to it being harvested while still green. The percentage of
hot water-soluble C to total C was, on the other hand,
highest for rice bran (30.2%), followed by rice straw
(20.1%) and beech leaves (13.4%), suggesting the differ-
ences in the decomposability of each residue.
2.2. Preparation of residueboxes
A box was prepared to sample soils every few millimeters
from the soil-residue interface. The design shown in Fig. 1 is
similar to that of the rhizobox developed by Youssef and
Chino (1988), and will be referred to as a residuebox in this
paper. The residuebox made from polystyrene was 10 cm in
width, depth and height. It was composed of several
compartments each separated by nylon mesh cloth (mesh
size 20 £ 40 mm2) attached to narrow plastic frames. The
central residue compartment (RC) had a thickness of 4 mm.
To either side of this were five 2-mm-thick compartments,
then a large compartment that extended to the end of the
residuebox (about 40 mm width). The RC was filled with the
paddy soil to which rice bran, rice straw or beech leaves had
been uniformly incorporated at 50 g dry matter kg21 soil.A
control treatment (no residue application) was also prepared.
Table 1
Concentrations of C and N in the residues used
Total Ca
(g kg21)
Total Na
(g kg21)
C/N
ratio
Hot water-
soluble Cb
(g kg21)
C extractabilityc
(%)
Rice bran 491 26.1 18.8 149 30.2
Rice straw 428 16.8 25.5 86 20.1
Beech leaf 527 11.6 45.5 70 13.4
a Determined by the dry combustion method (Sumigraph NC analyzer
NC-800, Sumika Chem. Anal. Service).b Determined by the method of Quarmby and Allen (1989).c Percentage of hot water-soluble C to total C.
Fig. 1. Schematic diagram (horizontal section) of the residuebox used in the
experiment.
N. Moritsuka et al. / Soil Biology & Biochemistry 36 (2004) 1141–11481142
All other compartments were filled with soil without residue
additions. The weight of soil added to each compartment
was 28 g in the RC, 14 g in each 2-mm-compartment and
700 g in the remaining compartments to keep the soil bulk
density at about 1.0 in the residuebox.
2.3. Incubation experiment
The residueboxes were incubated for 5, 15 or 30 days at
25 8C to evaluate both spatial and temporal changes of
residue-induced N immobilization. The experiment thus
consisted of three residue treatments plus control, each of
which had three incubation periods and three replications.
Prior to the incubation, deionized water was poured onto the
residueboxes filled with air-dry soil to adjust the matric
potential of the soil at 210 kPa. At this potential, the soil
was maintained in relatively aerobic conditions, containing
water at 320 ml kg21 soil and water-filled pore space at
61%. Rapid wetting of soil may cause a flush of microbial
growth, but pre-incubation after soil wetting was not
conducted in order to examine the short-term changes in
soil properties during the incubation. During the incubation,
deionized water was supplied uniformly over the soil
surface once a week in order to maintain the matric potential
of the soil. At the same time, boxes were covered with
aluminum foil to minimize soil water evaporation during the
incubation. After incubation, soils were sampled entirely
from each compartment of the residueboxes. Soil samples
located at the same distance from the RC on both sides were
mixed homogeneously and were stored at 5 8C for 1–2 days
before analysis.
2.4. Soil analyses
Inorganic N (NO32 and NH4
þ) and microbial biomass N
contents of the soil were determined. Inorganic N was
extracted with a 0.5 M K2SO4 solution at a soil:solution
ratio of 1:5 (w/v). The concentration of NH4þ and NO3
2
was determined colorimetrically by the indophenol and the
Griess–Ilosvay methods, respectively (Mulvaney, 1996).
Microbial biomass N was measured by the fumigation–
extraction method using a kEN of 0.57 (Inubushi, 1992).
For both fumigated and non-fumigated soils, total N in
0.5 M K2SO4 extracts was determined colorimetrically at a
wavelength of 220 nm, after extracting the soil with 0.5 M
K2SO4 in the same way as for inorganic N and
then oxidizing the extracts with an alkaline potassium
persulfate solution. The concentration of SON was also
calculated by subtracting the concentration of inorganic N
from that of total N in 0.5 M K2SO4 extracts of
non-fumigated soils.
2.5. Statistical analysis
Data for each incubation period was subjected to an
analysis of variance (ANOVA) to evaluate the effect of
residue application on each soil property using a signifi-
cance level of P , 0:05:
3. Results
3.1. Distribution of inorganic nitrogen
Distribution of NO32, NH4
þ and inorganic N around the
RC is shown in Table 2. After 5 days of incubation,
nitrate became dominated in the control soil. Near the RC
of residue treatments, variable but generally modest
depletion of NH4þ was observed in soil. The NO3
2
concentrations, and hence total inorganic N concentrations
also, declined significantly up to at least 10 mm from the
RC for all residue treatments, suggesting extensive
diffusion of soil inorganic N toward the soil-residue
interface. The high concentrations of NH4þ at 8–50 mm
from the RC of the beech leaf treatment indicate delayed
nitrification, probably due to a slight evaporative drying of
soil. An insignificant depletion of inorganic N was also
found in the 10–50 mm compartment of all residue
treatments.
By 15 days, nitrate dominated in inorganic N pool in all
treatments. In the rice bran treatment, the concentration of
inorganic N in the RC became higher than that in the
control, representing an increase of 66.1 mg kg21 from day
5 to day 15. N mineralization was thus occurring faster than
immobilization at this time. The rice straw and beech leaf
treatments, on the other hand, continued to show significant
depletion of inorganic N up to at least 10 mm from the RC,
but a slight increase in inorganic N in the RC suggests
positive net N mineralization.
By 30 days, the level of inorganic N in the RC of rice
straw and beech leaves had recovered considerably,
although depletion was still significant. In contrast,
inorganic N in the RC of rice bran decreased to control
levels.
3.2. Distribution of microbial biomass nitrogen
Distribution of microbial N around the RC is shown in
Table 3. After 5 days, microbial N increased significantly in
the RC of rice bran and rice straw, but not in the RC of beech
leaves. The values in the RC of rice bran (421 mg kg21) and
rice straw (224 mg kg21) were much higher than those of
inorganic N around the RC (,50 mg kg21), indicating that
microorganisms growing in the close vicinity of rice
residues acted as a strong sink for inorganic N in the
surrounding soil. In the beech leaf treatment, on the other
hand, the level of microbial N in the RC was similar to that
of the control treatment, although a significant depletion of
inorganic N was found around the RC. Microbial N tends to
decrease in zones at a distant from the RC of the rice straw
and beech leaf treatments, but the reason for this was
uncertain.
N. Moritsuka et al. / Soil Biology & Biochemistry 36 (2004) 1141–1148 1143
Table 2
Average concentrations of nitrate, ammonium and inorganic N around the residue compartment of the residueboxes ðn ¼ 3Þ
Soil position After 5-day incubation After 15-day incubation After 30-day incubation
Control Rice bran Rice straw Beech leaf Control Rice bran Rice straw Beech leaf Control Rice bran Rice straw Beech leaf
Nitrate (mg N kg21) RC 38.2 1.22** 2.66** 3.20** 48.2 40.8* 9.68** 12.6** 56.4 49.1 31.1** 37.1*
0–2 mm 40.1 10.4** 13.9** 17.8** 49.8 52.1 20.1** 20.1** 59.7 51.0** 36.6** 26.4**
2–4 mm 40.8 15.2** 18.1** 22.1** 48.1 42.9 21.1** 24.3** 57.3 47.9 31.8** 27.8**
4–6 mm 40.2 19.7** 21.3** 23.1** 46.2 40.4 25.8* 25.4** 59.7 48.7** 35.0** 29.3**
6–8 mm 37.7 23.3** 23.4** 25.3* 48.9 37.4** 27.7** 32.4** 55.8 51.4 32.2** 33.6*
8–10 mm 40.4 30.3* 29.5** 26.7* 50.3 39.1* 35.0** 35.8** 58.2 46.9 41.8 32.8*
10–50 mm 43.5 37.6 32.0 33.8 56.6 45.7* 50.2 51.2 69.3 55.1 44.6** 49.5**
Ammonium (mg N kg21) RC 12.8 7.83 5.41 1.10* 1.96 34.2** 6.04** 1.62 2.06 9.61** 7.85** 2.10
0–2 mm 12.2 2.76* 3.04* 7.80 1.72 1.86 1.39 1.41 1.60 1.45 1.25 1.34
2–4 mm 12.0 3.12* 6.76 12.4 1.48 1.47 1.33 1.33 1.34 1.17 1.15 1.34
4–6 mm 11.4 4.04* 8.96 13.5 1.58 1.11* 1.13 1.73 1.25 1.11 1.11 1.25
6–8 mm 10.8 5.46* 10.8 15.6 1.12 1.22 1.06 1.86 1.15 1.05 1.12 1.23
8–10 mm 10.6 6.53 11.1 16.6* 1.28 1.18 1.20 1.42 1.16 1.08 1.05 1.57
10–50 mm 9.7 8.65 11.7 16.3* 1.27 1.09 1.26 1.40 1.18 1.26 1.04 1.33
Inorganic N (mg N kg21) RC 51.1 9.04** 8.07** 4.30** 50.2 75.1** 15.7** 14.2** 58.5 58.7 39.0** 39.2*
0–2 mm 52.4 13.2** 16.9** 25.6** 51.5 54.0 21.5** 21.5** 61.3 52.5* 37.8** 27.7**
2–4 mm 52.8 18.3** 24.9** 34.5** 49.5 44.3 22.4** 25.6** 58.6 49.1 33.0** 29.1**
4–6 mm 51.7 23.7** 30.3** 36.6** 47.7 41.5 26.9* 27.1** 61.0 49.8** 36.1** 30.5**
6–8 mm 48.5 28.8** 34.2** 40.9* 50.0 38.6** 28.8** 34.3** 56.9 52.4 33.3** 34.8*
8–10 mm 51.0 36.8** 40.6** 43.2* 51.6 40.3* 36.2** 37.2** 59.3 47.9 42.9 34.4*
10–50 mm 53.2 46.2** 43.7** 50.2 57.8 46.8* 51.4 52.6 70.5 56.4 45.7** 50.8**
Marked values with * and ** indicate a significant difference from the corresponding value in the control treatment at the level of P , 0:05 and P , 0:01; respectively (ANOVA). The content of nitrate and
ammonium in soil before incubation was 18.8 and 38.8 (mg N kg21), respectively.
N.
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During the 5–15 day period, the levels of microbial N in
the RC decreased from 421 to 218 mg kg21 in the rice bran
treatment, remained relatively constant in the rice straw
treatment, and increased from 30 to 96 mg kg21 in the beech
leaf treatment. This small and slow increase in microbial N
in the RC of beech leaves may be related to the poor
decomposability of beech leaves as suggested by its low C
extractability (Table 1). Decreases of microbial N in the RC
of rice bran during this period corresponded well with the
increase in inorganic N near the RC, indicating that
microbially immobilized N in the vicinity of rice bran was
remineralized, nitrified and diffused away from the RC in
the form of NO32. In contrast to this, the increase of
microbial N in the RC of the beech leaf treatment coincided
with the decrease of inorganic N in 2–10 mm from the RC.
During the 15–30 day period, microbial N decreased in
the RC of all residue treatments, suggesting that readily
decomposable C had been severely depleted by 30 days. The
decrease of microbial N in this period coincided with the
increase of inorganic N near the RC of the rice straw and
beech leaf treatments (Table 2). Decreases of inorganic N in
the RC of rice bran, on the other hand, might be due to
denitrification.
3.3. Relationship between the decrease of inorganic N
and the increase of microbial N
To compare the decrease of inorganic N with the increase
of microbial N quantitatively, each of them was calculated
for the 5-day incubation results in which severe depletion of
inorganic N was observed (Table 4). The amount of
decrease in inorganic N was calculated by summing up
the product of soil weight in each compartment and
the difference of inorganic N concentrations between the
residue and control treatments. On the other hand, the
amount of increase in microbial N was calculated by
multiplying soil weight in the RC (28 g) by the difference of
microbial N concentrations between the residue and control
treatments. Changes in microbial N occurred mainly in the
RC (Table 3), and thus those observed in other compart-
ments were neglected in the calculation.
Table 4 indicates that the increase in microbial N in the
RC was largest in the rice bran treatment (11.0 mg),
followed by the rice straw (5.49 mg) and beech leaf
(0.06 mg) treatments. On the other hand, the amount of
depletion in inorganic N within 10 mm of the RC was
4.97 mg (rice bran), 4.27 mg (rice straw) and 3.42 mg
(beech leaves). The increase in microbial N in the RC thus
explained 221% (rice bran), 129% (rice straw) and 1.73%
(beech leaves) of the depletion of inorganic N occurring
within 10 mm of the soil-residue interface. If the statisti-
cally insignificant depletion of inorganic N in the
10–50 mm compartments is also taken into account, this
percentage drops to 111% (rice bran), 50.2% (rice straw)
and 1.06% (beech leaves). These relationships indicate that
the amount of increase in microbial N was comparable to
that of the depletion in inorganic N in the rice residue
treatments, but not in the beech leaf treatment.
4. Discussion
4.1. Spatiotemporal changes of nitrogen immobilization
in the vicinity of residues
It has been recognized that the soil is an entity with
high spatial and temporal variability at every scale of
observation. Like the rhizosphere, the soil-residue interface
Table 3
Average concentrations of microbial biomass N (mg kg21) around the residue compartment of the residueboxes ðn ¼ 3Þ
After 5-day incubation After 15-day incubation After 30-day incubation
Soil position Control Rice bran Rice straw Beech leaf Control Rice bran Rice straw Beech leaf Control Rice bran Rice straw Beech leaf
RC 27.9 421** 224** 30.0 26.7 218** 189** 95.7** 8.93 86.7** 82.2** 47.4**
0–2 mm 41.1 49.5 21.4 18.4 27.1 61.2** 45.7* 38.1* 11.5 18.1* 31.1** 17.8
2–4 mm 36.2 36.1 16.6* 18.0* 24.5 44.2 44.9 41.0 15.3 16.5 21.4 18.0
4–6 mm 32.0 33.0 13.3** 22.0 32.6 41.6 37.6 38.1 9.64 14.3 22.2 18.7
6–8 mm 44.3 26.4 18.6* 16.9* 37.7 38.7 38.3 36.9 15.3 19.0 12.0 15.8
8–10 mm 27.0 23.7 19.0 19.3 29.4 37.1 36.0 36.5 14.7 16.5 25.2 18.5
10–50 mm 35.6 22.6 21.0 12.4 31.8 32.2 33.0 36.0 16.1 15.9 19.3 22.1
Marked values with * and ** indicate a significant difference from the corresponding value in the control treatment at the level of P , 0:05 and P , 0:01;
respectively (ANOVA).
Table 4
Amounts (mg N) of decrease of inorganic N and increase of microbial N
after 5 day-incubation
Decrease of
inorganic N
(total)a
Decrease of
inorganic N
(,10 mm)b
Increase
of microbial
Nc
Rice bran 9.88 4.97 11.0
Rice straw 10.9 4.27 5.49
Beech leaf 5.57 3.42 0.06
a Total decrease of inorganic N from a residuebox.b Decrease of inorganic N within 10 mm of the RC.c Increase of microbial N in the RC.
N. Moritsuka et al. / Soil Biology & Biochemistry 36 (2004) 1141–1148 1145
is a biologically active region within the soil, but its
ecological and agronomical importance has been paid much
less attention than has the rhizosphere. Some research has
been carried out on carbon mineralization (Gaillard et al.,
2003), residue-induced carbon, nitrogen, and microbial
gradients (Gaillard et al., 1999), and enzyme characteristics
(Kandeler et al., 1999) in the vicinity of plant residues.
Spatial changes in N immobilization near residues have
rarely been investigated, although temporal changes
have been researched extensively (Mary et al., 1996;
Recous and Machet, 1999; Recous et al., 1999; Trinsoutrot
et al., 2000).
In our experiment, N immobilization extended to at least
10 mm from the soil-residue interface after 5 days of
incubation, regardless of type of residue (Table 2). The
region of N depletion was larger than that of N release from
mature wheat straw, i.e. 4–5 mm from the soil-residue
interface (Gaillard et al., 1999). This implies that the zone
affected by decomposing residues after 5 days of incubation
could be separated into an inner region extending a few
millimeters from the soil-residue interface which is
affected by both immobilization of soil inorganic N and
mineralization of residue-derived N, and an outer region
extending a few centimeters from the interface which is
affected mainly by immobilization. By longer incubation,
immobilization of soil inorganic N was followed by
its mineralization. The net N mineralization occurred
faster and more intensively in the rice bran treatment,
probably due to the low C/N ratio and high microbial
decomposability of rice bran (Table 1).
From agronomic viewpoint, our results suggest that the
effect of N immobilization and mineralization on crop
growth following the application of residues to soil depends
not only on timing of application and type of residue but
also on the position of residues in soil in relation to growing
roots.
4.2. Relative significance of biotic and abiotic
immobilization
It should be noted beforehand that the relative import-
ance of biotic and abiotic immobilization could be estimated
only roughly. Since our experiment was conducted without
either a sterile control or the use of 15N, we could not
examine the release of inorganic N through mineralization
of plant-derived N and microbial N as well as microbial
incorporation of plant-derived N, each of which would
affect the decrease of soil inorganic N and the increase of
microbial N. But small and slow increase in microbial N
near beech leaves (Table 3) suggests that such microbial
processes proceeded very slowly at least in the beech leaf
treatment during the 0–5 day period of incubation. Besides
this, denitrification following NO32 reduction might have
caused the depletion of inorganic N in the soil-residue
interface. It is generally recognized, that the emission
of N2O induced by residue application decreases with
the increase in the C/N ratio of residues (Aulakh et al., 1991;
Kaiser et al., 1998). The highest C/N ratio of beech leaf
(Table 1) implies the least possibility of denitrification.
The rather arbitrary choice made for the coefficient used in
the calculation of microbial N (kEN(0.57) could also cause
an error in estimating the microbial biomass, since kEN may
vary with soil type between 0.3 and 0.8 (Joergensen and
Mueller, 1996).
Even though such limitations and artifacts in our
experimental approach were taken into account, there was
an apparent difference between the rice residue and beech
leaf treatments. The increase of microbial N following the
application of beech leaves occurred more slowly than the
depletion of soil inorganic N (Tables 2–3) and explained
less than 2% of the N depletion on day 5 (Table 4), whereas
the increase of microbial N following the application of rice
residues was large enough to be comparable to the decrease
of inorganic N (Table 4). Owing to this contrasting result, it
was suggested strongly that the dominant N immobilization
process was abiotic in soil near beech leaves and biotic in
soil near rice residues.
The abiotic immobilization in the beech leaf treatment
supports the findings of field studies that report an absolute
increase in the N content of decomposing beech leaves
(Osono and Takeda, 2001; Pardo et al., 1997). In one of
these studies, the amount of N in the fungal biomass of the
decomposing beech leaves was estimated to be less than 2%
of total N in the leaves; a level insufficient for the N increase
to be explained as N incorporation by fungi (Osono and
Takeda, 2001). Similar results have been reported for
decomposing leaves from a marsh grass (Spartina alterni-
flora L.) (Lee et al., 1980) and a mangrove (Rhizophora
mangle L.) (Hernes et al., 2001), suggesting the widespread
existence of this phenomenon. Abiotic immobilization of
inorganic N near rice residues seemed to be insignificant in
this study, but such process might be dominated in other
type of agricultural residues.
In summary, residue-induced immobilization of soil
inorganic N is driven by both biotic and abiotic processes,
with the relative importance of each process being
determined by residue type.
4.3. Process of abiotic immobilization in soil near beech
leaves
The dominance of abiotic immobilization suggested in
soil near beech leaves may be due to the limited biotic
immobilization caused by the low availability of C to
microbes (suggested by low water-soluble C content in
Table 1). In addition to this, a specific reaction for abiotic
N immobilization might be present. The possible pro-
cesses are transformation of inorganic N to K2SO4-SON,
K2SO4-insoluble organic N or K2SO4-insoluble inorganic
N. In this case, however, the last transformation, i.e.
fixation of NH4þ into the interlayer of clay minerals,
N. Moritsuka et al. / Soil Biology & Biochemistry 36 (2004) 1141–11481146
is practically impossible, since the immobilization
occurred as a result of residue application.
The distribution of SON in soil after 5 days of incubation
shown in Table 5 indicated that SON decreased significantly
in the close vicinity of beech leaves, in contrast to the rice
residue treatments. The SON concentration in soil at a few
millimeters away from the soil-residue interface was not
much affected by residue application. These results
suggested that the process of abiotic immobilization in
soil near beech leaves was not transformation of inorganic N
into SON, but probably its transformation into 0.5 M
K2SO4-insoluble organic N.
Our result differs from previous ones. Dail et al. (2002)
reported that, after abiotic incorporation of 15N-nitrate into a
C-rich (25% organic C) acid forest soil, the immobilized
nitrate was found mainly in SON and less than 5% was
recovered as insoluble organic N. Compton and Boone
(2002) also suggested that rapid incorporation of15N-ammonium and 15N-nitrate into forest soils was due
to their rapid conversion into the SON fraction. It is
uncertain whether our results arose from a special trait in
beech leaf composition or from differences in experimental
design. More research is required to determine what kind of
reactions are involved in the abiotic immobilization, since
the involved reaction site and mechanisms were almost
completely unknown.
Acknowledgements
The authors would like to thank Mr Matthew Turner for
reviewing the manuscript and Dr Osono Takashi for helpful
discussion.
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