Illinois
Fertilizer Conference Proceedings
January 23-25, 1995
| Home | 1995 Index | Search |
| Home | 1995 Index | Search |
R.L. Mulvaney and S.A. Khan1
Most nitrogen (N) fertilizers supply N in the form of ammonium (NH4+). In some cases, NH4+N is lost through volatilization as ammonia (NH3), but loss of N is more likely to occur by leaching or denitrification of nitrite (NO2-) or nitrate (NO3-) formed by nitrifying microorganisms in the soil. The major products of denitrification are the gases, dinitrogen (N2) and nitrous oxide (N2O), which escape from the soil.
Nitrification proceeds more rapidly with alkaline-forming fertilizers such as urea than with acidic fertilizers such as monoammonium phosphate (MAP, NH4H2PO4)(Eno and Blue, 1957; McInnes and Fillery, 1989; Mulvaney, 1994), and for this reason there should be a difference among fertilizers in the potential for loss of N by denitrification. Moreover, denitrification is favored by alkaline conditions (Firestone, 1982). Of the commonly used N fertilizers, liquid anhydrous NH3 causes the most drastic rise in soil pH, and this is accompanied by some solubilization of soil organic matter (Norman et al., 1987). The solubilized organic matter may be a potential energy source for denitrifying bacteria, as field studies in Iowa (Bremner et al., 1981; Breitenbeck and Bremner, 1986) have indicated emission of N2O to be much greater with anhydrous NH3 than with other N fertilizers. The latter finding was attributed to the production of N2O during nitrification of NH4+ (Bremner and Blackmer, 1978). Unlike N2O, N2 is not produced during oxidation of NH4+ to NO3- by nitrifying microorganisms (Ritchie and Nicholas, 1972), and thus the emission of N2 indicates the occurrence of denitrification. Because N2 is the major constituent of air, 15N-tracer techniques are required for detection of N2 emission into air, along with application of highly labeled fertilizer.
This paper describes studies that were conducted during the third year of a
3-year project concerning factors that affect the efficient use of N fertilizers
in Illinois soils. The objectives of these studies were: (i) to compare the
effects of different N fertilizers on emission of N2 and N2O during denitrification
of NO3- in waterlogged soil; and (ii) to determine whether the differences in
emission are related to fertilizer effects on soil pH or water-soluble organic
C. The following fertilizers were used: ammonium sulfate [(NH4)2SO4], urea,
MAP, diammonium phosphate [DAP, (NH4)2HPO4], ammonium nitrate (NH4NO3), and
anhydrous NH3.
The soil used was a surface (0-15 cm) sample of Drummer silty clay loam soil from a soybean plot at the Agronomy-Plant Pathology South Farm at Urbana. The sample was sieved (2-mm screen) in the field-moist condition, and was stored in a sealed polyethylene bottle in a refrigerator. Analyses as specified in a previous publication (Mulvaney, 1994) gave the following results: pH, 6.0; organic C, 3.15%; total N, 0.285%; sand, 17%; silt, 48%; clay, 35%; water-holding capacity, 480 mL kg-1.
To compare the effects of different fertilizers on emission of N2 and N2O during denitrification in waterlogged soil, 35 samples of field-moist soil, each containing 20 g of oven-dry material, were placed in 237-ml, (8-oz.) wide-mouth, straight-sided glass bottles and treated with: (i) 1 mL of deionized water (control); (ii) 1 ml, of deionized water containing 4 or 20 mg of N as unlabeled (NH4)2SO4, urea, NH4H2PO4 [MAP], (NH4)2HPO4 [DAP], or NH4NO3; or (iii) 1 mL of deionized water, followed by infection of approximately 20 mg of N as unlabeled liquid anhydrous NH3. Injection was accomplished by a syringe technique (Norman and Kurtz, 1986), after sealing the bottle by attaching a Mason jar lid through which had been soldered a brass tube (6.4 mm od, 4.6 mm id, 20 mm long) equipped with a serum stopper to serve as an injection port.
After treatments (i) and (ii), the bottles were sealed with Parafilm®, the Parafilm® was punctured to allow aeration, and the bottles were transferred to an incubator maintained at 25°C and 100% relative humidity. Incubation was carried out under aerobic conditions (soil moisture content = 68.5% of the water-holding capacity) for 0, 2, 4, 7, 14, 21, or 42 d. Samples receiving treatment (iii) were incubated as with treatments (i) and (ii), except the injection lid was left in place for 1 h (for the 0-d aerobic incubation) or 12 h (for all other times) to facilitate retention of gaseous NH3, prior to replacement by Parafilm®.
After each aerobic incubation period, samples were removed (5 replicates per treatment) for anaerobic denitrification assays (3 replicates), and for determination of soil pH and water-soluble organic C (2 replicates).
Denitrification assays were carried out by treating samples with 5 ml, of deionized water containing 5 mg of N as 15N-labeled KNO3 (73.7 atom % 15N). Water was added to increase the soil moisture content to 300% of the water-holding capacity, and the bottles were then sealed with Mason jar lids fitted with brass shut-off valves and placed in an incubator maintained 25°C. After 5 d, the bottles were removed from the incubator, and atmospheric samples were collected in 60-mL evacuated tubes fitted with high-vacuum stopcocks. Analyses were performed by isotope-ratio mass spectrometry to determine 15N-labeled N2 and N2O (Mulvaney and Kurtz, 1982; Mulvaney and Boast, 1986).
To measure soil pH and water-soluble organic C, samples were treated with deionized
water to obtain a soil:water ratio of 1:1, the bottle was swirled to mix the
resulting suspension, and pH was measured with a glass electrode. After each
measurement, the electrode was rinsed with 20 mL of deionized water, which was
collected in the bottle, giving a soil:water ratio of 1:2. The bottle was sealed
with a screw-lid, and then shaken gently with an orbital shaker for 15 min.
Most of the liquid was transferred to a 100-mL polyethylene centrifuge tube
using a 50-mL pipette with a wide tip, treated with 1 g of K2SO4
as a clarifying agent, and centrifuged at approximately 15,000 g for 5 min.
The clear supernatant was filtered under vacuum through 0.2mm Metricel®
membrane filter (Gelman Instrument Co., Ann Arbor, Michigan). The extract was
analyzed for soluble organic C using a dichromate oxidation method (Burford
and Bremner, 1975).
Figures 1, 2, 3, 4, 5, 6, and 7 show the rates of labeled N2 and N2O emission during a 5-d anaerobic incubation of 15NO3-amended soil, following aerobic incubation for up to six weeks with or without unlabeled anhydrous NH3, urea, DAP, (NH4)2SO4, NH4NO3, or MAP. Figures 8 and 9 show soil pH values and concentrations of water-soluble organic C, respectively, determined after each interval of aerobic incubation. The unlabeled fertilizers were applied at a rate of 1000 mg N kg-1 soil, so as to simulate a band application. Except for anhydrous NH3, incubations were also carried out with 200 mg N kg-1 soil, to approximate a broadcast application. In Fig. 8 and 9, all data reported for the various fertilizer treatments were obtained with 1000 mg N kg-1 soil.
Comparison of Fig. 2 to 7
with Fig. 1 shows that all of the fertilizers
studied increased the emission of N2 and N2O from waterlogged
soil during incubation with 15NO3-, and that
the increases were usually larger for N2O than for N2.
These emissions can be attributed to bacterial denitrification, because although
some production of N2O (but not N2) may have occurred
during nitrification of NH4+ in fertilized samples of
soil, any N2O thereby produced would have been unlabeled and therefore
undetectable by the mass spectrometric procedures employed for atmospheric analysis.
Examination of Fig. 2 to 7
reveals marked differences among the fertilizers in their effect on denitrification
of NO3-. Emission of N2 and N2O
decreased in the order: anhydrous NH3 > urea >> DAP >
(NH4)2SO4 > NH4NO3
> MAP. The differences are probably due, at least in part, to an effect on
soil pH, as denitrification is favored by alkaline conditions (Firestone, 1982).
Of the fertilizers studied, anhydrous NH3 and urea gave the highest
values for soil pH, whereas application of (NH4)2SO4,
NH4NO3, or MAP, which are acidic salts, led to an immediate
decrease in soil pH (Fig. 8).
Denitrification requires oxidizable C, and this is often a limiting factor under field conditions, particularly with agricultural soils that have been treated with N fertilizer. Soil organic matter is a major source of this oxidizable C, but only becomes available when dissolved in the soil solution (Burford and Bremner, 1975). The solubility of organic matter increases markedly under alkaline conditions (Stevenson, 1994), and this undoubtedly accounts, at least in part, for the finding that injection of anhydrous NH3 leads to an increase in the concentration of soluble organic C (e.g., Norman et al., 1987). Further evidence for the ability of anhydrous NH3 to solubilize soil organic C is provided by Fig. 9, which shows that elevated levels of soluble organic C were also observed with urea and MAP. The effect of urea can likely be attributed to an increase in pH resulting from hydrolysis by soil urease (Fig. 8). The effect of MAP may be related to the high content of PO43-, as PO43- reagents (particularly Na4P2O7) have been used successfully as extractants of soil organic matter (Stevenson, 1994). However, acidity inhibits denitrification, and this probably accounts for the fact that emissions of N2 and N2O were much lower with MAP than with anhydrous NH3 or urea.
Table 1 lists the coefficients obtained by correlation of N2 and/or N2O emissions with soil pH or water-soluble organic C. Most of the correlations were statistically significant at the 0.1% level.
With all fertilizers tested, highest emissions of N2 and N2O were observed when the soil was waterlogged within one week after fertilizer application. Longer periods of aerobic incubation caused a marked decrease in denitrification, presumably because of the drop in soil pH caused by nitrification of NH4+ (Fig. 8). Likewise, aerobic incubation of the unfertilized control soil led to a decrease in emission of N2, and N2O during anaerobic incubation with 15NO3- (Fig. 1), probably owing to a decline in the availability of soluble organic C. After aerobic incubation for six weeks, very little denitrification occurred in the control soil, whereas substantial emission of N2O was observed in all cases involving application of fertilizer at a rate of 1000 mg N kg-1 soil.
Examination of Fig. 3 to 7 shows that emission of N2 and N2O during denitrification of 15NO3 was usually lower with fertilizer applications of 200 mg N kg-1 soil than with those supplying 1000 mg N kg-1 soil. The difference can be attributed to the lower application rate having a lesser effect on soil pH and soluble organic C (data not reported). In a few cases, mainly involving MAP or NH4NO3, emission of N2 was higher with 200 mg N kg-1 soil than with 1000 mg N kg-1 soil. This can probably be explained by the fact that the lower application rate gave a higher,soil pH, with the result that denitrification proceeded more rapidly and produced a larger proportion of N2, as compared to N2O (Firestone, 1982).
A laboratory incubation experiment using 15NO3 to estimate gaseous loss of N as N2 and N2O showed .that N fertilizers promote denitri.fication in waterlogged soil. Emission of N2 and N2O decreased in the order: anhydrous NH3 > urea >> DAP > (NH4)2SO4 > NH4NO3 > MAP. Highest emissions were observed when the soil was waterlogged within one week after fertilizer application. A lower rate of fertilizer application tended to decrease the emission of N2 and N2O. Emissions of N2 and N2O were significantly correlated with fertilizer effects on soil pH and water-soluble organic C.
Figure 1. Emission of N2 and N2O from control soil (no treatment with unlabeled fertilizer)
Figure 2. Emission of N2 and N2O from anhydrous NH3-treated soil (1000 mg N kg-1)
Figure 3. Emission of N2 and N2O from urea-treated soil
Figure 4. Emission of N2 and N2O from DAP-treated soil
Figure 5. Emission of N2 and N2O from (NH4)2SO4-treated soil
Figure 6. Emission of N2 and N2O from NH4NO3-treated soil
Figure 7. Emission of N2 and N2O from MAP-treated soil
Figure 8. Fertilizer effects on soil pH (0 or 1000 mg N kg-1)
Figure 9. Fertilizer effects on water-soluble organic C (0 or 1000 mg N kg-1)
1Professor and Research Associate, Department of Agronomy, University of Illinois, Urbana, IL
Breitenbeck, G. A., and J. M. Bremner. 1986. Effects of various nitrogen fertilizers on emission of nitrous oxide from soils. Biology and Fertility of Soils, 2:195-199.
Bremner, J. M., and A. M. Blackmer. 1978. Nitrous oxide: Emission from soils -during nitrification of fertilizer nitrogen. Science, 199:295-296.
Bremner, J. M., G. A. Breitenbeck, and A. M. Blackmer. 1981. Effect of anhydrous ammonia fertilization on emission of nitrous oxide from soils. Journal of Environmental Quality, 10:77-80.
Burford, J. R., and J. M. Bremner. 1975. Relationships between the denitrification capacities of soils and total, water-soluble and readily decomposable soil organic matter. Soil Biology & Biochemistry, 7:389-394.
Eno, C. F., and W. G. Blue. 1957. The comparative rate of nitrification of anhydrous ammonia, urea, and ammonium sulfate in sandy soils. Soil Science Society of America Proceedings, 21:392-396.
Firestone, M. K. 1982. Biological denitrification. In: Nitrogen in agricultural soils (F. J. Stevenson et al., ed.). Agronomy Monograph 22. American Society of Agronomy, Madison, Wisconsin. pp. 289-326.
McInnes, K. J., and I. R. P. Fillery. 1989. Modeling and field measurements of the effect of nitrogen source on nitrification. Soil Science Society of America Journal, 53:1264-1269.
Mulvaney, R. L. 1994. Nitrification of different nitrogen fertilizers. In: 1994 Illinois Fertilizer Conference Proceedings (R. G. Hoeft, ed.). pp. 83-96.
Mulvaney, R. L., and C. W. Boast. 1986. Equations for determination of nitrogen-15 labeled dinitrogen and nitrous oxide by mass spectrometry. Soil Science Society of America Journal, 50:360-363.
Mulvaney, R. L., and L. T. Kurtz. 1982. A new method for determination of 15N-labeled nitrous oxide. Soil Science Society of America Journal, 46:1178-1184.
Norman, R. J., and L. T. Kurtz. 1986. Laboratory apparatus for preparing and dispensing nitrogen-15-labeled anhydrous ammonia. Soil Science Society of America Journal, 50:238242.
Norman, R. J., L. T. Kurtz, and F. J. Stevenson. 1987. Solubilization of soil organic matter by liquid anhydrous ammonia. Soil Science Society ofAmerica Journal, 51:809-812.
Ritchie, G. A. F., and D. J. D. Nicholas. 1972. Identification of the sources of nitrous oxide produced by oxidative and reductive processes in Nitrosomonas europaea. Biochemistry Journal, 126:1181-1191.
Stevenson, F. J. 1994. Humus chemistry. Genesis, composition, reactions. 2nd ed. John Wiley and Sons, New York.
