Fertilizer Conference Proceedings
R.L. Mulvaney, F. Azam, and F.W. Simmons1
When nitrogen (N) fertilizer is applied to soil, some of the N is taken up by microorganisms in the soil, a process known as immobilization. The immobilized N is incorporated into proteins, nucleic acids, and other organic N constituents of microbial cells and cell walls; as such, it becomes part of the biomass. As the microbes die and decay, some of the biomass N is released as NH4+ through the process of mineralization; the remainder undergoes conversion to more stable organic N compounds, ultimately becoming a part of soil organic matter. The stabilized organic compounds are not readily available to plants; therefore, the net result of immobilization-mineralization is a decrease in the availability of the N added to soil as fertilizer, and also the partial conversion of this N to a form (NH4+) that is not subject to loss from most soils.
The extent to which fertilizer N is immobilized depends largely on the supply of C, which is used by soil microorganisms as an energy source. Of particular importance is the ratio of available C to mineral N (NH4+ and NO3-), or C/N ratio. When this ratio is 20 or lower, mineralization exceeds immobilization, whereas at C/N ratios of 30 or higher, immobilization exceeds mineralization. Also of importance is the type of C source. In the presence of a simple substrate such as glucose, mineral N is quickly consumed and disappears within a few days. With complex substrates, such as cellulose from plant residues, the process is slower, and the supply of mineral N is never completely exhausted.
The microorganisms responsible for immobilization utilize NH4+ in preference to NO3- (Jansson, 1958), which accounts for reports that immobilization-mineralization is more extensive with ammoniacal fertilizers than with NO3- fertilizers (e.g., Wickramasinghe et al., 1985). But like other biological N transformations, immobilization-mineralization is affected by environmental conditions, such as temperature, soil moisture content, and pH, and this may lead to differences between acidic fertilizers such as monoammonium phosphate (MAP) and alkaline or alkalineforming fertilizers such as diammonium phosphate (DAP) or urea.
This paper describes studies that were conducted during the first year of a
3-year project to compare the behavior of different N fertilizers in Illinois
soils. The objectives of these studies were: (i) to estimate the rate and extent
of incorporation of N from different fertilizers into organic forms through
immobilization; and (ii) to compare the immobilization of NH4+-N and NO3--N.
The soils used (Table 1) were surface (0-15 cm) samples from fields in corn and soybean production. Before use, they were air-dried and crushed to pass through a 2-mm screen. In the analyses reported in Table l, pH, organic C, total N, and texture were determined as described in a previous publication (Mulvaney and Kurtz, 1982). Water-holding capacity was determined by the method of Bremner and Shaw (1958). Inorganic N analyses were done by extraction (2 M KCI)-distillation (Keeney and Nelson, 1982).
To estimate the rate and extent of immobilization for different N fertilizers, 15-g samples of soil were placed in 125-mL polyethylene bottles and treated with 3 mL of deionized water containing 7.5 g glucose-C kg-1 soil and 250 mg N kg-1 soil as 15N-labeled urea (3.225 atom % 15N), (NH4)2SO4 (2.082 atom % 15N), (NH4)2HPO4 (2.357 atom % 15N), or NH4NO3 (2.625 atom % 15N). Additional water was applied to achieve a soil moisture content equivalent to 40% of the water-holding capacity, and the bottles were loosely stoppered to allow aeration and incubated at 25°C for 0, 0.5, 1, 2, 4, 8, 16, and 30 d. At regular intervals, the bottles were weighed, and deionized water was added to compensate for moisture loss. After incubation, duplicate samples were treated with 100 mL of 2 M KCl, and mineral N was extracted by shaking for 1 h followed by vacuum filtration. The soil remaining in the bottle was transferred quantitatively to the filter paper by rinsing with 25 mL of 2 M KCl, which was subsequently removed by vacuum filtration. Rinsing was repeated with 25 mL of 2 M KCl to ensure complete removal of mineral N. After drying at room temperature for about 24 h, the soil was passed through a 2-mm screen and mixed thoroughly. Duplicate samples were analyzed by a permanganate-reduced Fe modification of a semimicro-Kjeldahl procedure (Bremner and Mulvaney, 1982). Distillates obtained by steam distillation of the digests were collected in H3BO3-indicator solution for titration with 0.0025 M H2SO4. The distillates were then acidified (0.2 mL of 0.025 M H2SO4) and evaporated to dryness on a hot plate (90°C). The residue was dissolved in 1-2 mL of deionized water, and an aliquot was transferred to a plastic microplate and evaporated to dryness (75°C) for 15N analysis with an automated mass spectrometer (Mulvaney, 1993).
To compare immobilization of NH4+-N and NO3--N,
duplicate samples of soil (20 g) were placed in 125-mL polyethylene bottles
and treated with deionized water containing: (i) 100 mg N kg-1 soil
as 15N-labeled (NH4)2SO4 (2.391
atom % 15N); (ii) 200 mg N kg-1 soil as 15N-labeled
(NH4)2SO4; (iii) 100 mg N kg-1 soil
as 15N-labeled KNO3 (2.696 atom % 15N); (iv)
200 mg N kg-1 soil as 15N-labeled KNO3; (v)
3 g glucose-C kg-1 soil and 100 mg N kg-1 soil as 15N-labeled
(NH4)2SO4; (vi) 6 g glucose-C kg-1
soil and 200 mg N kg-1 soil as 15N-labeled (NH4)2SO4;
(vii) 3 g glucose-C kg-1 soil and 100 mg N kg-1 soil as
15N-labeled KNO3; and (viii) 6 g glucose-C kg-1
soil and 200 mg N kg-1 soil as 15N-labeled KNO3.
In all cases, the volume of water applied was sufficient to achieve a soil moisture
content equivalent to 40% of the water-holding capacity. The applications of
glucose gave a C/N ratio of 30, thereby ensuring complete immobilization of
applied N. The bottles containing the treated soils were loosely stoppered to
allow aeration and incubated at 25°C for 2 weeks. At regular intervals,
the bottles were weighed, and deionized water was added to compensate for moisture
loss. After incubation, 100 mL of 2 M KCl was added to each bottle for
extraction of mineral N. The extracts were steam-distilled with MgO and Devarda's
alloy to determine their content of (NH4+ + NO3-)-N
(Keeney and Nelson, 1982), and the distillates were retained for 15N
analyses, from which recovery of fertilizer N in inorganic forms was calculated.
Analyses for total N and 15N were performed on the residue as described
previously to determine recovery of the applied N in organic forms produced
Table 2 shows the recovery in organic forms from applying N as urea, (NH4)2SO4, DAP, or NH4NO3 to glucose-amended soil, after incubation for various periods up to 30 d. As expected, substantial immobilization occurred during incubation, but some recovery in organic forms was observed even without incubation (i.e., the 0-d sampling). In cases where N was applied as NH4+, the latter finding can be attributed to fixation by clay minerals, because immobilized N was determined by total N analysis of KCl-extracted soils, which would have recovered clayfixed NH4+ as well as organic N. The same explanation holds for urea, since no precautions were taken to prevent hydrolysis by soil urease during extraction of mineral N, which can be appreciable (Keeney and Bremner, 1967).
The increase in recovery observed with continued incubation (Table 2) reflects the accumulation of applied N in the soil biomass during immobilization. Highest recoveries, ranging from 62 to 102%, occurred after incubation for 8 to 16 d. A decline in recovery followed, as some of the immobilized N was remineralized.
Recoveries tended to be lower with DAP than with the other fertilizers tested, which indicates less extensive immobilization. The difference may be related to pH, as unlike the other fertilizers used, application of DAP would have caused an immediate rise in pH (the pH of DAP is about 8.0), which may have retarded microbial activity. Likewise, hydrolysis of urea would have led to an increase in pH, but the increase would have occurred more gradually than with DAP, and this may have facilitated the development of a microbial population better adapted to alkaline conditions. Moreover, the finding that initial recoveries (i.e., after incubation for 0.5-1 d) were similar for urea and (NH4)2SO4 suggests that some urea may have been immobilized without hydrolysis to NH4+.
Table 3 shows the recovery of applied NH4+-N and NO3--N in organic and inorganic forms. In the absence of glucose, recovery in organic forms was higher with NH4+ than with NO3-. In the presence of glucose, immobilization was comparable for NH4+ and NO3- with the Drummer and Flanagan soils, and somewhat more extensive for NO3- than for NH4+ with the Catlin soil. With every treatment studied, a higher percentage of applied N was immobilized in the Drummer soil than in the Catlin or Flanagan soil, but with each soil, the percentage of applied N immobilized was nearly the same at both rates of application (100 and 200 mg N kg-1 soil). Loss of N occurred with both NH4+ and NO3- additions; on average, the largest losses occurred with the Drummer soil.
Existing information indicates that soil microorganisms prefer NH4+ over NO3- as an N source (Jansson, 1958) and that immobilization of NO3- is very limited in fallow soils (Legg and Stanford, 1967). In general, NH4- appears to be preferred over NO3- during microbial assimilation, which is likely attributable to the fact that not all microbes possess NO3- reductase (Payne, 1973). In the present study, the proportion of added N immobilized was smaller with NO3- than with NH4+. However, substantial amounts of NO3--N were transformed into organic forms even when no exogenous C source was provided (Table 3). The difference between immobilization of NH4+ and NO3- was smallest with the Drummer soil, which had the highest content of organic matter (Table 1). The differences could have been due, at least in part, to clay fixation of NH4+, because immobilized N was determined by total N analysis of the KCl extracted soils, which would have recovered fixed NH4+ as well as organic N. Other work in our laboratory has shown that, when incubated as described, the Catlin, Flanagan, and Drummer soils used have the capacity to fix up to 5 % of the N applied as NH4+.
As expected, immobilization was increased considerably by addition of glucose.
Of particular interest is the fact that with glucose, very little difference
occurred in immobilization of NH4+ and NO3-
(Table 3). Several studies have shown that
NO3- is actively immobilized in the presence of an ample
supply of easily oxidizable C (e.g., Azam et al., 1988) and that remineralization
occurs more rapidly when soils are treated with NO3- than
with NH4+, either in the presence or absence of exogenously
applied C (Nommik, 1981). The latter finding can probably be attributed to more
rapid utilization of mineralizable C in NO3--treated soils,
and suggests that the supply of C has a greater effect on immobilization of
NO3- than NH4+, presumably because
of the additional energy requirement associated with microbial reduction of
NO3-. This effect should be greatest when the period of
incubation is limited, as was the case in the present study (2 weeks). During
extended incubation, a wider variety of microorganisms may be involved in the
turnover of N, since immobilized NO3--N will be quickly
remineralized upon exhaustion of available C sources, and the resulting accumulation
of NH4+ will promote the growth and proliferation of microorganisms
that lack NO3- reductase.
A laboratory incubation experiment to compare the immobilization of different N fertilizers in glucose-amended soil showed that DAP was immobilized to a lesser extent than urea, (NH4)2SO4, or NH4NO3. The difference was attributed to the fact that application of DAP causes an immediate increase in pH, which suppresses microbial acitivity.
A study to compare the immobilization of NH4+-N and NO3--N
showed that, in the absence of glucose, immobilization was greater with NH4+
than with NO3-, whereas in the presence of glucose, there
was very little difference between the two forms of N. This finding was attributed
to a higher energy requirement for immobilization of NO3-
than NH4+, owing to the need for microbial reduction of
NO3- to NH4+.
Table 1: Analyses of soils
Table 2: Recovery in organic forms of N applied as various fertilizers
Table 3: Recovery of applied NH4+-N and NO3--N in organic and inorganic forms
Azam, F., T. Mahmood, and K. A. Malik. 1988. Immobilization-remineralization of N03-N and total N balance during decomposition of glucose, sucrose and cellulose in soil incubated at different moisture regimes. Plant and Soil, 107:159-163.
Bremner, J. M., and C. S. Mulvaney. 1982. Nitrogen - Total. In: Methods of Soil Analysis (A. L. Page et al., ed.) Agronomy Monograph 9, Part 2, 2nd ed. American Society of Agronomy, Madison, WI pp. 595-624.
Bremner, J. M., and K. Shaw. 1958. Denitrification in soil. I. Methods of investigation. Journal of Agricultural Science, 51:22-39.
Jansson, S. L. 1958. Tracer studies on nitrogen transformations in soil with special attention to mineralisation-immobilisation relationships. Annals of the Royal Agricultural College of Sweden, 24:101-361.
Keeney, D. R., and J. M. Bremner. 1967. Determination and isotope-ratio analysis of different forms of nitrogen in soils: 7. Urea. Soil Science Society of America Proceedings, 31:317-321.
Keeney, D. R., and D. W. Nelson. 1982. Nitrogen - Inorganic forms. In: Methods of Soil Analysis (A. L. Page et al., ed.) Agronomy Monograph 9, Part 2, 2nd ed. American Society of Agronomy, Madison, Wisconsin. pp. 643-698.
Legg, J. 0., and G. Stanford. 1967. Utilization of soil and fertilizer N by oats in relation to the available N status of soils. Soil Science Society of America Proceedings, 31:215-219.
Mulvaney, R. L. 1993. Mass spectrometry. In: Nitrogen Isotope Techniques (R. Knowles and T. H. Blackburn, ed.). Academic Press, San Diego, CA pp. 11-57.
Mulvaney, R. L., and L. T. Kurtz. 1982. A new method for determination of "N-labeled nitrous oxide. Soil Science Society of America Journal, 46:1178-1184.
Nommik, H. 1981. Fixation and biological availability of ammonium in soil clay minerals. In: Terrestrial Nitrogen Cycles (F. E. Clark and T. Rosswall, ed.) Ecological Bulletin (Stockholm), 33:273-279.
Payne, W. J. 1973. Reduction of nitrogenous oxides by microorganisms. Bacteriological Reviews, 37:409-452.
Wickramasinghe, K. N., G. A. Rodgers, and D. S. Jenkinson. 1985. Transformations of nitrogen fertilizers in soil. Soil Biology & Biochemistry, 17:625-630.
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