Published in Soil Sci. Soc. Am. J. 67:1773-1780 (2003).
© 2003 Soil Science Society of America
677 S. Segoe Rd., Madison, WI 53711 USA
DIVISION S-3—SOIL BIOLOGY & BIOCHEMISTRY
Insight into the Active Organic Nitrogen Pool Estimated by Isotopic Equilibrium Approaches
W. J. Wang*,a,
C. J. Smithb and
D. Chenc
a NR&M, 80 Meiers Rd, Indooroopilly, Brisbane, QLD 4068, Australia
b CSIRO Land and Water, GPO Box 1666, Canberra 2601, ACT, Australia
c Dep. of Resource Management and Horticulture, the Univ. of Melbourne, Parkville 3052, VIC, Australia
* Corresponding author (weijin.wang{at}nrm.qld.gov.au).
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ABSTRACT
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A method that reliably estimates the size of the active organic N pool has been desired for predicting N mineralization dynamics. Among the methods proposed to date, the isotopic equilibrium approaches characteristically use N mineralization-immobilization turnover processes to label the biologically labile organic N. In the present study, the implications and usefulness of the active organic N pool estimated with this approach were assessed in a long-term incubation. 15N-labeled NH+4 and glucose were added to soil, and the dynamics of the labeled N in the organic and inorganic phases were monitored by periodic sampling during the incubation. After appropriate revisions, two equations from the literature were used to estimate the active organic N in soil at Time 0 (OT#) and at each sampling during incubation (OT*), respectively. Both OT# and OT* tended to increase with incubation time and were affected by the amounts of labeled N and available C addition. Considerable amounts of mineralized N came from other than OT* fraction of soil organic N, but N mineralization from the OT* pool was much faster than from the bulk soil organic matter. The results supported the concept that soil organic N exists in a continuum of degradability. If the operational procedure is standardized, however, OT# can be used as a relative measure of the active organic N in soil.
Abbreviations: AL, labeled NH+4–N (i.e., fertilizer N in NH+4) • AT, total NH+4–N = AL + AU • AU, unlabeled NH+4–N of soil origin • NL, labeled NO-3–N • nmOT*, net N mineralization from the OT* pool • nmOT, net N mineralization from the OT pool • NT, total NO-3–N = NL + NU • NU, unlabeled NO-3–N of soil origin • OL, labeled organic N (i.e., added fertilizer N in OT) • OT, total organic N • OT*, active organic N in soil at the time of each sampling during incubation • OT#, active organic N of the original soil • Qe, 15N abundance (atom %) of a N pool at the time of sampling during incubation • Qf, 15N abundance (atom%) of labeled fertilizer N • WHC, water-holding capacity
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INTRODUCTION
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IT IS GENERALLY BELIEVED that a part of the soil organic matter is more susceptible to biological decomposition than the rest and therefore plays a prominent role in releasing inorganic N through mineralization (Stanford and Smith, 1972; Duxbury and Nkambule, 1994; Parton, 1996; Molina and Smith, 1998). Although not concisely defined, the concept of an active or labile organic N pool is generally used to describe a portion of soil organic N that participates in short-term biological N cycling (Jansson, 1958; Duxbury et al., 1991), and this concept is often adopted in N transformation models (Dou and Fox, 1995; Bouraoui and Dillaha, 2000). Many chemical, physical, and incubation procedures were proposed for estimating the active organic N in soil, but none has been consistently successful (Dahnke and Vasey, 1973; Keeney, 1982; Bundy and Meisinger, 1994; Wang et al., 2001b).
Jansson (1958) proposed an isotopic equilibrium method to measure the active organic N pool by incubating soil with 15N-labeled inorganic N and straw for a period of up to 60 d. It was assumed that equilibrium of 15N between the inorganic and the active organic N pools would be attained through continuous mineralization-immobilization turnover. Therefore, the proportion of labeled N in the remineralized N should approximate the proportion of labeled N in the active organic N pool. This can be expressed as
 | [1] |
or under strongly nitrifying conditions when the AT pool is infinitely small,
 | [2] |
where AL and AT represent the labeled and total NH+4–N, respectively, and OL and OT* represent the labeled and the active organic N, respectively. NL and NT refer to the labeled and total NO-3–N, respectively. Consequently, OT* can be estimated from the measurements of OL, AL, and AT, or NL and NT.
Duxbury et al. (1991) modified the method of Jansson (1958) by adding glucose and a nitrification inhibitor to promote mixing of the active soil organic N and 15N-labeled NH+4 that was applied as fertilizer in the amount of AL0 with an 15N abundance of Qf (atom% 15N). At the end of a 40-d incubation, the 15N abundance (Qe) in the NH+4 pool was measured. Assuming equilibrium of 15N between the inorganic and active organic N pools is achieved, the following equations based on the mass balance were obtained:
 | [3] |
or
 | [4] |
where OT# represents the active organic N pool, to distinguish from that obtained with Eq. [1] or [2].
The isotopic equilibrium approach distinctively relies on microbial activity to mix added 15N with the soil organic N that participates in biologically mediated N turnover (Duxbury et al., 1991). In spite of the long time required for incubation and 15N analysis, these methods have been applied in a number of investigations involving: (i) the effect of cropping and tillage on soil quality (Follett and Schimel, 1989; Duxbury et al., 1991; McCarty et al., 1995); (ii) quantification of the active organic N pool to be used in numerical modeling of N mineralization-immobilization turnover (Smith et al., 1994); and (iii) the relationships of the active organic N to total N and microbial biomass N (McCarty et al., 1995; Liang et al., 1999).
Nevertheless, the operational procedures and incubation conditions in different studies varied significantly. The biological meanings of the data obtained with different equations were not clearly defined. No studies compared the results given by the different equations. The objectives of this study were to: (i) define the meaning of the active organic N pool measured with different equations; (ii) assess the validity of the assumption of an equilibrium of labeled N between the inorganic and the active organic N phases with a prolonged incubation; and (iii) identify the advantages and limitations of this method and propose a more standardized procedure for measuring the biologically labile organic N in soil.
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MATERIALS AND METHODS
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Soils
Soil samples were taken in July 1997 from two sites of the experimental station of the Charles Sturt University, Wagga Wagga, NSW, Australia. Soil 1 was collected from a field planted with lucerne (Medicago sativa L.) since 1994 (Smith et al., 1998), and Soil 2 from a site that had grown annual cereals since 1992 and planted to Canola (Brassica napus L.) in 1997. The surface 20 cm of soil was collected, thoroughly mixed, and passed through a 2-mm sieve to form a composite sample for each paddock. Visible plant residues and gravel in the soils, although rare, were removed. The soils are Palexeralfs according to U.S. classification. Soil 1 had 22% clay, 18% silt, 1.4% organic C, 0.12% total N, pH 5.4 (1:5 soil to water), and a water-holding capacity (WHC) of 35 mL 100 g-1 (oven-dry basis). Soil 2 had 16% clay, 12% silt, 1.6% organic C, 0.13% total N, pH 4.6, and a WHC of 35 mL 100 g-1.
Incubation and Analysis of Soils
The field-moist soils were stored at 4°C for 2 wk before use. The moist samples were mixed and subsamples equivalent to 20 g oven-dry soil were weighed into 250-mL polystyrene jars. The soils was then labeled with 15N-enriched (NH4)2SO4 (82.0 atom % 15N) in three different treatments as follows: Treatment 1, 10 mg N kg-1 soil only; Treatment 2, 10 mg N + 1000 mg glucose-C kg-1 soil; Treatment 3, 100 mg N + 1000 mg glucose-C kg-1 soil.
These treatments basically followed that of Jansson (1958), with some modifications including (i) variations in the amounts of N and C addition, and (ii) replacement of straw with glucose. These modifications allowed examination on the effects of the amount of fertilizer N and glucose on the determination of the active organic N pool. Differing from Duxbury et al. (1991), no nitrification inhibitor was used in our study because (i) nitrification does not affect the validity of Eq. [1] and [2], and Eq. [3] and [4] would also be valid for calculating OT# under nitrifying conditions provided corrections are made as shown below; and (ii) a preliminary experiment indicated that NH+4–N accumulation could be insignificant during the early stages of incubation because of strong nitrification, but would become significant later on, which allows recently mineralized NH+4–N to be distinguished from the old NO-3–N.
The (NH4)2SO4 and glucose were applied to the soils in solution drop by drop in such amounts to bring the soil moisture contents to 55% of their WHCs. The jars were covered with Parafilm and incubated at 35°C, which is generally considered close to the optimum temperature for mineralization (Stanford and Smith, 1972). During the course of incubation, the jars were opened every day during the first few weeks and once a week thereafter to maintain aerobic conditions, and replenished with water to maintain constant soil moisture content.
At Days 0, 5, 15, 30, 45, 60, 80, 110, 153, 193, 233, 273, and 333, three jars of each treatment (containing 20 g soil each) were destructively sampled by extracting with 100 mL 2 M KCl for 1 h. NH+4–N in the filtered extracts was determined by steam-distillation with a small amount of carbonate-free MgO; and NO-2 + NO-3–N was determined by a second distillation after adding Devarda's alloy (Keeney and Nelson, 1982). To minimize cross contamination, ethanol distillation was conducted between replicates (Hauck, 1982), and an ethanol-distillation double-distillation procedure (Pruden et al., 1985) was used between samples of different treatments. The titrated distillates were spiked with standard NH+4–N solution if its N content was <150 µg or its 15N abundance was expected to be >10%. Isotope ratios were determined with a magnetic-deflection mass spectrometer (Sira 10, VG Isogas, Middlewich, UK). The procedure for isotope ratio analysis was described by Chen et al. (1990).
The extracted soil was transferred onto a filter paper and washed 6 to 8 times with 0.01 M CaCl2 solution to remove residual mineral N and KCl salt, and dried in an oven at 80°C. The washed soils were then finely ground with a ring grinder (Rocklabs Pty. Ltd., Auckland, New Zealand). Total N content and 15N enrichment were determined by combustion with an automatic CNS analyzer (NA1500, Varian, CA, USA) interfaced with a mass spectrometer (VG Isogas, Sira 10, Middlewich, UK).
Calculation of the Active Organic N Pool
Equation [3] is valid only if (i) initial soil NH+4–N and NO-3–N contents are negligible because Qe could be diluted directly by initial unlabeled NH+4–N and indirectly by initial NO-3–N due to immobilization following glucose addition and subsequent remineralization; and (ii) the labeled N is conserved within the exchangeable NH+4–N and active organic N pools. However, nitrification could remove 15N-labeled N from the NH+4–N pool, and loss of labeled N from the NO-3, NH+4, and organic N pools was often observed shortly after addition of AL because of consumption processes like clay fixation, nitrification/denitrification, and/or ammonia volatilization (Davidson et al., 1991; Liang et al., 1999; Wang et al., 2001a). After 30 d of incubation, total recoveries of the labeled N in NH+4, NO-3, and organic N pools ranged from 80 to 90%, but no further loss of labeled N was observed thereafter (results not presented). Assuming that loss of unlabeled NH+4–N and NO-3–N were small and negligible relative to their total amounts, Eq. [4] should be modified as below with the labeled N pools measured at each sampling during incubation and by subtracting initial mineral N from the active organic N pool.
If no significant amount of NH+4–N was formed under strongly nitrifying conditions, OT# was estimated with NL and Qe of the NO-3–N pool at the time of periodic sampling:
 | [5] |
where NU0 and AU0 refer to unlabeled soil NO-3–N and NH+4–N at time zero, respectively.
If a significant amount of NH+4–N accumulated in soil, N mineralization-immobilization turnover should predominantly occur between the NH+4–N and organic N pools (Jansson, 1958), then OT# was calculated with AL and Qe of the NH+4–N pool:
 | [6] |
where NU is unlabeled NO-3–N content at the time of harvest.
Statistical Analysis
Statistical analyses of data were conducted with the ANOVA procedure of GenStat (Payne, 2000). The difference between treatment means was tested with the LSD at levels of P < 0.05 and P < 0.01.
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RESULTS AND DISCUSSION
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Dynamics of Labeled and Unlabeled Mineral Nitrogen
Net immobilization of mineral N occurred during the initial 5 d in the treatments with glucose addition (Fig. 1)
. Net nitrate immobilization was observed only in Treatment 2, when the NH+4–N pool was inadequate to meet the microbial demand for N following the glucose addition (Fig. 1c and 1d). In contrast, when sufficient NH+4–N was supplied (Treatment 3; added C/N ratio = 10), NO-3–N increased (Soil 1) or remained unchanged (Soil 2) during the net immobilization period. This result corroborated the finding of Jansson (1958) that NO-3–N is not involved in the mineralization-immobilization turnover as long as adequate NH+4–N exists.
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Fig. 1. Dynamics of total NO-3–N (NT) and NH+4–N (AT) accumulation in soils following imposition of different treatments: Treatment 1, 10 mg labeled NH+4–N kg-1 only; Treatment 2, 10 mg labeled NH+4–N + 1000 mg C kg-1; and Treatment 3, 100 mg labeled NH+4–N + 1000 mg C kg-1. The vertical bars represent ±SD.
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With the exception of Treatment 3 for Soil 2, the NH+4–N pool became nearly empty within 5 to 15 d, and remained almost undetectable for 80 to 110 d because of nitrification and immobilization (Fig. 1). After that period, nitrification slowed or ceased probably because of inhibitory effect of the continued high temperature (Robertson et al., 1988), and NH+4 began to accumulate as the result of net mineralization. The accumulation of NH+4 from the mid stage of incubation made it possible to detect minor changes in the isotopic ratios of N mineralized later (see below), which would otherwise have been obscured by the high background levels of NL and NU if nitrification had remained strong. For the same reason, if a nitrification inhibitor (Duxbury et al., 1991) had been used in this prolonged incubation, it would also have been difficult to detect the changes in the isotopic ratios of recently mineralized N in a mixed pool of both old and newly mineralized NH+4–N.
15N-labeled fertilizer N in the mineral N pools was determined from Day 30 (Fig. 2 and 3)
. Labeled NH+4–N was analyzed only when the size of total NH+4–N pool was >5 mg kg-1. Determination of NL was ceased when NO-3 production reached the plateau and/or NH+4–N started to accumulate, based on the assumption that there should be no isotopic exchange between the NO-3 and organic N pools when sufficient NH+4 existed (Jansson, 1958). The amount of NL and/or AL increased continuously for all treatments as remineralization of immobilized labeled N proceeded from Day 30 to the end of the incubation (333 d). When nitrification ceased, NL remained constant, whereas AL continued to accumulate. However, the ratios of labeled to total mineral N decreased consistently during the course of incubation (Fig. 4 and 5)
, suggesting that recently mineralized N had a lower 15N abundance than the bulk mineral N pool. This can be further demonstrated by the significantly lower AL/AT ratios than the NL/NT ratios (P < 0.01) when NH+4 began to accumulate at Day 193 for Treatments 1 and 2 and Day 110 for Treatment 3 of Soil 1, and at Day 110 for Treatments 1 and 2 of Soil 2. The results suggest that the immobilized N had not homogenously mixed with the indigenous mineralizable soil organic N regardless of the treatment applied. Isotopic equilibrium between the inorganic and organic N pools was not achieved during the 333-d incubation.
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Fig. 2. Dynamics of labeled N in NO-3 (NL) and NH+4 (AL) pools of Soil 1 after different treatments: Treatment 1, 10 mg labeled NH+4–N kg-1; Treatment 2, 10 mg labeled NH+4–N + 1000 mg C kg-1; and Treatment 3, 100 mg labeled NH+4–N + 1000 mg C kg-1. The vertical bars represent ±SD.
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Fig. 3. Dynamics of labeled N in NO-3 (NL) and NH+4 (AL) pools of Soil 2 after different treatments: Treatment 1, 10 mg labeled NH+4–N kg-1; Treatment 2, 10 mg labeled NH+4–N + 1000 mg C kg-1; and Treatment 3, 100 mg labeled NH+4–N + 1000 mg C kg-1. The vertical bars represent ±SD.
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Fig. 4. Changes in the labeled to total N ratios in NO-3 (NL/NT) and NH+4 (AL/AT) pools during incubation of Soil 1, following three different treatments: Treatment 1, 10 mg labeled NH+4–N kg-1; Treatment 2, 10 mg labeled NH+4–N + 1000 mg C kg-1; and Treatment 3, 100 mg labeled NH+4–N + 1000 mg C kg-1. The vertical bars represent ±SD.
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Fig. 5. Changes in the labeled to total N ratios in NO-3 (NL/NT) and NH+4 (AL/AT) pools during incubation of Soil 2, following three different treatments: Treatment 1, 10 mg labeled NH+4–N kg-1 only; Treatment 2, 10 mg labeled NH+4–N + 1000 mg C kg-1; and Treatment 3, 100 mg labeled NH+4–N + 1000 mg C kg-1. The vertical bars represent ±SD.
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Treatment 2 had significantly lower NL/NT ratios than Treatment 1 for both soils because glucose addition promoted immobilization of AL0 (Fig. 4 and 5). However, during the subsequent remineralization of labeled N, the AL/AT ratios were also lower for Treatment 2 than for Treatment 1. This suggested that the initially high C/N ratio (1/100) in the available C and N pools brought about by glucose addition reduced the biological availability of the immobilized labeled N or promoted mixing with the active soil organic N. Of the three treatments, Treatment 3 (C/N = 1/10) had the highest NL/NT ratios because of the high amount of labeled N remaining in the NO-3–N pool, and this treatment also had the highest AL/AT ratios because of the combined effects of a large OL pool and its probably high degradability.
Active Organic Nitrogen Pool
Equations [2] and [5] were used to calculate the sizes of the active organic N pool for Treatments 1 and 2 of Soil 1 sampled from Days 30 to 153, and for Treatment 3 of Soil 1 and Treatments 1 and 2 of Soil 2 sampled from Days 30 to 80 (Fig. 6)
, because the NH+4 pool was then nondetectable or very small for these treatments as a result of rapid nitrification. Thereafter, NH+4 started to accumulate in significant amounts (Fig. 1). Estimation of the active organic N pool was performed with Eq. [1] and [6] because NO-3–N should no longer be involved in N transformations between the organic and inorganic phases (discussed above).
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Fig. 6. The active organic N pool as affected by treatment, incubation time, and the equation used for estimation. Estimates on or after the symbols marked with arrows were based on AL (labeled NH+4–N) and Qe (15N abundance of a N pool at the time of sampling during incubation) of the NH+4 pool. Treatments are as follows: Treatment 1, 10 mg labeled NH+4–N kg-1; Treatment 2, 10 mg labeled NH+4–N + 1000 mg C kg-1; and Treatment 3, 100 mg labeled NH+4–N + 1000 mg C kg-1. The vertical bars represent ±SD.
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When OT* was estimated on the basis of NO3–N pool (Eq. [2]), no significant change in the sizes of OT* was observed with time (Fig. 6). The OT* values obtained with Treatments 2 and 3 were consistently higher than those obtained with Treatment 1 (P < 0.01) because of either the significantly lower NL/NT values in Treatment 2 or the higher OL values in Treatment 3 [OT* = OL/(NL/NT)]. When the NH+4 pool became detectable and used to calculate OT* (Eq. [1]), the estimated sizes of the active organic N pool for most treatments increased markedly (Fig. 6; P < 0.01), because of the lower AL/AT ratio than the NL/NT ratio (Fig. 4, 5). As the NH+4 pool was composed of more recently mineralized N, AL/AT should be a better approximation for the ratio of OL/OT* than the NL/NT ratio. The relatively constant and lower values of OT* estimated with NL/NT may be attributable to the insensitivity of the NL/NT ratio to the small amounts of mineralized N relative to the high background levels of NT and NL. With the exception of Treatment 3 of Soil 1, OT* increased steadily with time during the period of NH+4 accumulation (Fig. 6). Although there was no consistent trend in the relative values of OT* estimated by the three treatments, significant differences were often observed, particularly during the early stages of incubation.
Stanford and Smith (1972) proposed that net N production follows first order kinetics and the size of the mineralizable N pool in soil declines with time as net mineralization proceeds. This concept of a discrete potentially mineralizable N pool seems contradictory with the results obtained with the isotopic equilibrium approach in that OT* increased with time. As discussed above, the increasing OT* values resulted from the decreasing proportion of labeled N in the mineralized N. This was explained by Jansson (1958) as an indication of incorporation or equilibration of the active organic N with an increasing amount of indigenous organic N, as well as an indication of OL transferring into a more passive pool or leaving the mineralization-immobilization cycling. It appears, however, more likely that a portion of the indigenous soil organic N never mixed with the labeled N through mineralization-immobilization turnover and released mineral N as a distinct source of decomposable substrate. Continuing release of unlabeled indigenous organic N resulted in the progressive dilution of 15N in the mineral N pool.
The values of OT# increased with incubation time (Fig. 6). Consistent with the values of OT*, significant increases in OT# were obtained for most treatments when its estimation was switched from being based on NO-3–N to NH+4–N pool. Treatment 1 (10 mg N kg-1 only) gave the lowest estimates of OT# (P < 0.01) during the early stages, but the difference in OT# values among different treatments became insignificant toward the end of incubation, except that Treatment 3 gave markedly higher estimate than other treatments for Soil 1.
The estimates of OT# were always higher than the estimates of OT* for the same treatment (Fig. 6; P < 0.01). The OT* pool in Eq. [1] and [2] actually consists of both labeled and unlabeled organic N in the active organic N pool at the time of each sampling during incubation, but does not include the already mineralized soil organic N. To determine the size of the active organic N pool in the original soil before incubation (OT*0), the amount of OL should be subtracted from OT* particularly when a large amount of AL0 was added (e.g., Treatment 3), and the amount of N mineralized from native soil organic N should be taken into account, as they obviously originated from the labile fraction of soil organic matter. Therefore,
 | [7] |
The results given by Eq. [7] were identical to those obtained with Eq. [5] or Eq. [6]; that is, OT*0 = OT#. Therefore, OT# represents the net amount of N mineralized from soil organic N pool plus the indigenous soil organic N in the OT* pool.
Availability of the Active Organic Nitrogen Pool
Assuming that the ratio of OL/OT* could be approximated by the ratio of AL/AT during the late stage of incubation, net N mineralization from OT* (nmOT*) can be calculated from the increase in AL or AL + NL during a time interval, whereas net N mineralization from the total organic N pool (nmOT) can be estimated from changes in mineral N contents.
If nitrification has stopped,
 | [8] |
and
 | [9] |
where subscripts 1 and 2 represent measurements at the beginning and end of a time interval, respectively.
When NO-3–N and NH+4–N accumulate simultaneously,
 | [10] |
and
 | [11] |
Given the above assumption, nmOT* was compared with nmOT during the period of Days 233 to 333 (Table 1).
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Table 1. Comparison of degradability of the active organic N (OT*) and total organic N (OT) during the late stage of incubation (Days 233 to 333).
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The mineralization rate of the active organic N (nmOT*/OT*) was approximately twice as fast as that of total soil organic N (nmOT/OT), as shown in Table 1. This was consistent with the findings of Ladd et al. (1977), but differed from those of Blackmer and Green (1995) that little of the labeled N immobilized following plant residue application was remineralized during the subsequent net N mineralization period. Tusneem & Patrick (1971) also found that recently immobilized N was not always more decomposable than that of native soil organic N. In the present study, N mineralized from OT* accounted for 47.1 to 85.0% of the total amount of net N mineralization for Soil 1, and 40.3 to 54.3% for Soil 2. Therefore, the hypothetical active organic N pool OT* could not represent the entire source of mineralizable N. The proportion of net N mineralization from OT*(nmOT*/nmOT) increased in the order Treatment 1 < Treatment 2 < Treatment 3.
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SUMMARY
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The size of the active organic N pool estimated with the isotopic equilibrium approach varied with the experimental conditions, such as glucose addition and its amount, the amount of labeled N, glucose-C/fertilizer-N ratio, and the incubation duration. During the long-term incubation, complete isotopic mixing between the inorganic and the mineralizable organic N pools was not achieved. It is legitimate to speculate that as long as net N mineralization does not stop (i.e., mineralization >immobilization), a real isotopic equilibrium would not be reached, and estimates of the size of the active organic N pool (OT* or OT#) would continue to increase with time. The results support the concept that soil organic matter is composed of a continuum of heterogeneous compounds that have various molecular recalcitrance and physical accessibility to microbial decomposition (Bosatta and Agren, 1991; Duxbury and Nkambule, 1994). Therefore, the existence of a discrete active organic N pool of fixed size in soil appears unlikely. Like all other physical, chemical, and incubation techniques, the active organic N estimated with the isotopic equilibrium approach should be interpreted as a relative index of N mineralization capacity. However, OT* or OT# must be measured under standardized conditions for a certain period so that the results from different studies would be comparable. The procedure described by Duxbury et al. (1991) has at least two significant advantages over the original method of Jansson (1958): (i) the soil was incubated for a fixed short period, and (ii) glucose was used to replace plant materials that may contain a substantial amount of N. However, some significant modifications of the operationally defined procedure of Duxbury et al. (1991) are warranted: (i) soil moisture is controlled at a near optimum level for N mineralization (e.g., 55% WHC or field capacity); (ii) the soil samples are incubated at 35°C, which was considered the near-optimum temperature for net N mineralization and is generally used in incubation of soils to determine N mineralization potential (Stanford and Smith, 1972); and (iii) addition of nitrification inhibitor is unnecessary and Eq. [5] and [6] instead of Eq. [3] should be used to calculate OT#, especially for soils that initially contain significant amounts of mineral N or have high 15NH+4 consumption capacities by processes other than biological immobilization such as clay fixation or ammonia volatilization.
The most prominent feature of the isotopic equilibrium approach in comparison with conventional short- or long-term incubation techniques lies in the fact that this approach included a portion of labile organic N (OT* - OL) not mineralized by the time of sampling during incubation. Further experimentation is required to evaluate whether this feature of OT# provides real advantage over conventional incubation techniques for predicting long-term net N mineralization or for parameterizing N turnover models.
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ACKNOWLEDGMENTS
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The authors gratefully acknowledge the support of the University of Melbourne and CSIRO in providing funds and facilities. We are indebted to Dr. P.M. Chalk (IAEA, Vienna), who offered great support for the initiation of this research.
Received for publication August 1, 2002.
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