Nitrification of Different Nitrogen Fertilizers

Illinois Fertilizer Conference Proceedings
January 24-26, 1994

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Nitrification of Different Nitrogen Fertilizers

R.L. Mulvaney¹

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Introduction

In most agricultural soils, ammonium (NH₄⁺) from fertilizer is quickly converted to nitrate (NO₃^-) by the process of nitrification. This process is crucial to the efficiency of N fertilizers and their impact on the environment, because the net effect is a conversion of fertilizer N from a form that is not normally subject to loss from soil (NH₄⁺) into a form that is readily lost by leaching or denitrification (NO₃^-).

Nitrification occurs in two steps: NH₄⁺ is first converted to nitrite (NO₂^-), and the NO₂^- is then converted to NO₃^-. Both reactions are carried out by bacteria present in the soil. Nitrifying bacteria are chemoautotrophic, in that they produce energy by chemical oxidation of NH₄⁺ or NO₂^- and utilize CO₂ as a source of C. Different groups of bacteria are responsible for the two steps involved in nitrification. The NH₄⁺-oxidizing bacteria include species from five genera, the most common being Nitrosomonas; the NO₂^--oxidizing bacteria all belong to the genus, Nitrobacter.

Because there are only a few species of nitrifying bacteria, nitrification is much more sensitive to environmental conditions than are most other N transformations, which are carried out by a more diverse group of microorganisms. All of the nitrifiers are obligate aerobes, which means that they require atmospheric O₂ so nitrification is especially sensitive to soil moisture content and does not occur in waterlogged soils. The rate of nitrification increases with soil temperature up to about 35°C (95°F); below 5°C (40°F) very little NO₃^- is formed. Soil pH is also important. Below a pH of 6.0, nitrification is inhibited by acidity, and the process virtually ceases at a pH of 4.5 to 5.0. Under alkaline conditions, production of NO₃^- is markedly enhanced. The optimum pH is normally between 7.0 and 8.0, but NO₃^- may be formed at a pH of 9.0 or even higher.

Although nitrification in agricultural soils has been studied extensively, most of the studies have been conducted using a single form of N fertilizer, usually ammonium sulfate [(NH₄)₂SO₄]. This fertilizer is an acidic salt, whereas some ammoniacal fertilizers give an alkaline reaction, such as urea or diammonium phosphate [DAP, (NH₄)₂HPO₄]. Early work by Eno and Blue (1957) with three sandy soils from Florida indicated that nitrification of NH₄⁺ derived from urea was much more rapid than nitrification of NH₄⁺ from (NH₄)₂S0₄. The same difference has been observed in subsequent work by Vilsmeier and Amberger (1680), Martikainen (1985), de Boer et al. (1989, and McInnes and Fillery (1989), and has generally been attributed to the rise in pH that results from hydrolysis of urea by soil urease. However, the increased pH of urea-treated soil may also produce a high concentration of ammonia (NH₃), and since Nitrobacter is more sensitive to NH₃ toxicity than is Nitrosomonas, this can lead to an accumulation of NO₂^-, and to subsequent loss of N through chemical reactions involving NO₂^-.

This paper describes studies that were conducted during the second 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 rate of NO₃^- and NO₂^- production from different fertilizers through nitrification; and (ii) to determine whether the differences between fertilizers can be accounted for strictly on the basis of their effects on pH. The following fertilizers were used: urea, DAP, (NH₄)₂SO₄, ammonium nitrate (NH₄NO₃), and monoammonium phosphate (MAP, NH₄H₂PO₄).

Materials and Methods

The soils used (Table 1) were surface (0-15 cm) samples from fields in corn and soybean production. The samples were sieved (-mm screen) in the field-moist condition and were stored in sealed polyethylene bottles in a refrigerator. In the analyses reported in Table 1, 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).

To compare the rate of NO₃^- and NO₂^- production for different fertilizers, field-moist samples of soil containing 5 g of oven-dry material were placed in 125-mL polyethylene bottles and treated with 0.5 mL of deionized water containing 5 mg of N as ¹⁵N-labeled urea (2.011 atom % 1sN), (NH⁴)²SO⁴ (2.029 atom % ¹⁵N), MAP (2.176 atom % ¹⁵N), or DAP (1.959 atom % ¹⁵N), or with 10 mg o N as ¹⁵NH₄NO₃ (2.145 atom % ¹⁵N). Additional water was applied to achieve a soil moisture content equivalent to 50% of the water-holding capacity, and the bottles were loosely capped to allow aeration and placed in an incubator maintained at 25°C and 100% relative humidity. After 3, 7, 14, 21, and 42 d triplicate bottles were removed from the incubator, and soil pH was measured with a glass electrode after adding 5 mL of deionized water. Following each measurement, the electrode was rinsed with 50 mL of 2 M KCl containing 250 pg of phenylmercuric acetate (PMA) as a urease inhibitor. The KCl PMA solution from rinsing the electrode was collected in the bottle, and the incubated soil sample was extracted by shaking for 1 h and filtering the resulting suspension under vacuum. The soil on the filter paper was leached twice with 25 mL of 2 M KCl-PMA to ensure complete removal of extractable N, allowed to dry at room temperature for at least 24 h, and then crushed and mixed thoroughly. The soil extract was analyzed for urea and NO₂^- by colorimetric procedures (Mulvaney and Bremner, 1979; Keeney and Nelson, 1982), and for NH₄⁺-N, (NH₄⁺ + NO₃^-)-N, and (NH₄⁺ + NO₃^- + NO₂^-)-N by steam distillation techniques (Keeney and Nelson, 1984 A 0.5- to 1.0-g sample of le extracted soil was digested for total N analysis by a semimicro-Kjeldahl procedure (Bremner and Mulvaney, 1982). Distillates obtained by steam distillation of the digests and extracts were collected in H₃BO₃-indicator solution for quantitative determinations by titration with 0.0025 M H₂SO₄. The titrated distillates were acidified (10-100 µL of 1 M H₂SO₄) and evaporated to dryness on a hot plate (90°C). The residue was dissolved in 0.4-1 mL of deionized water, and an aliquot containing 50-150 µg of NH₄⁺-N was transferred to a plastic microplate and evaporated to dryness in a forced-air oven (70°C) for ¹⁵N analysis with an automated mass spectrometer (Mulvaney, 1993).

In a separate experiment to estimate gaseous loss of fertilizer ¹⁵N as NH₃, triplicate samples of soil containing 10 g of oven-dry material were placed in 250-mL polyethylene wide-mouth bottles and treated with 1 mL of deionized water containing 10 mg of N as ¹⁵N-labeled urea, (NH₄)₂SO₄, MAP, or DAP, or with 20 mg of N as ¹⁵NH₄NO₃ (¹⁵N enrichments as specified previously). Sufficient water was added to raise the soil moisture content to 50% of the waterholding capacity, and the bottles were fitted with an aeration device having an acid trap for absorption of NH₃ evolved on incubation of the soil samples and placed in an incubator maintained at 25°~ and 100% relative humidity. The aeration device, following the design of Bremner and Douglas (1971), consisted of a no. 7 rubber stopper having a central hole fitted with an acrylic tube (110 mm long, 9.5 mm od, 6.4 mm id), to the lower end of which was cemented a 5-mL disposable polystyrene beaker containing 5 mL of 0.25 M H₂SO₄. The bottles were removed from the incubator after 3, 7, 14, 21, and 42 d, and the acid in the beakers was transferred to 100-mL Kjeldahl flasks for steam distillation with 5 mL of 1 M NaOH, followed by acidimetric titration and ₁₅N analysis as described previously. For continued incubation after 3, 7, 14, and 21 d, the bottles were fitted with an aeration device containing fresh H₂SO₄, and then replaced in the incubator.

To ascertain whether the rapid nitrification observed with urea is due entirely to an effect on soil pH, samples of the Catlin soil containing 5 g of oven-dry material were placed in 125-mL polyethylene bottles and treated with deionized water containing: (i) 5 mg of N as ¹⁵N-labeled (NH₄)₂SO₄ (2.051 atom % 15N); (ii) 5 mg of N as ¹⁵N-labeled urea (2.018 atom % ¹⁵N); or (iii) 5 mg o N as ¹⁵N-labeled (NH₄)₂SO₄ , followed by a separate addition of deionized water containing 320 µmol of OH- as NaOH. In all cases, the volume of water applied was sufficient to achieve a soil moisture content equivalent to 50% of the water-holding capacity. The additions of NaOH employed with (NH₄)₂SO₄ were established in preliminary studies as giving approximately the same soil pH values after a 3-d incubation as were obtained with urea. Incubation of the treated soil samples was carried out in triplicate for 3, 7, 14, 21, and 42 d (25°C, 100% relative humidity). Soil pH was then measured as previously described, followed by addition of 50 mL of 2 M KCl-PMA for extraction of inorganic N. The extracts were analyzed colorimetrically for NO₂^-, and steam distillations were performed for N and ¹⁵N analyses of NH₄⁺-N, (NH₄⁺ + N0₃^-)-N, and (NH₄⁺ + NO₃^- + NO₂^-)-N.

The percentage recovery of fertilizer ¹⁵N as exchangeable NH₄⁺, NO₃^-, NO₂^- , NH₃, and organic (residual) N was calculated from the corresponding N and ¹⁵N analyses. Recovery as urea was estimated from colorimetric determinations.

Results and Discussion

Tables 2, 3, 4, 5 and 6 show the recovery in nongaseous forms from applying labeled N to soils as urea, (¹⁵NH₄)₂HPO₄, (¹⁵NH₄)₂SO₄, ¹⁵NH₄NO₃, or ¹⁵NH₄H₂PO₄, after incubation for various periods up to 6 weeks. Table 7 summarizes the soil pH values after each incubation interval.

Examination of the data in Tables 2, 3, 4, 5 and 6 reveals marked differences in nitrification among the five fertilizers studied. Conversion of NH₄⁺-N to (NO₃^- + NO₂^-)-N generally decreased in the order: urea > DAP > (NH₄)₂SO₄ > NH₄NO₃ > MAP. The differences can be attributed, at least in part, to an effect on soil pH, because urea and DAP give an alkaline reaction when applied to soil, whereas (NH₄)₂SO₄, NH₄NO₃, and MAP are acidic salts. Some support for this view is provided by Table 7, which shows that the highest values for soil pH were obtained by incubation with urea for 3 d and that substantially lower pH values were obtained after the same period with DAP, (NH₄)₂SO₄, NH₄NO₃, and MAP. With the Cisne, Blount, and Drummer soils, the pH after 3 d was higher with DAP than with (NH₄)₂SO₄, NH₄NO₃ or MAP, and a higher rate of NO₃^- production was observed in the DAP-treated samples. With each soil studied, MAP gave the lowest pH after incubation for 3 d and the lowest rate of nitrification throughout the 6-week study period.

Not all of the differences observed between fertilizers can be attributed to an effect of soil pH on nitrification. For example, the pH with NH₄NO₃ was usually higher than with (NH₄)₂SO₄, and sometimes higher than with urea or DAP (Table 7). Yet nitrification was considerably slower for NH₄NO₃, as can be seen by comparing Tables 2, 3, or 4, with Table 5. This finding suggests that NO₃^- may have an inhibitory effect on nitrification. If so, inhibition must occur during the oxidation of NH₄⁺ to NO₂^-, as no accumulation of NO₂^- was observed after incubations of the Cisne, Blount, Catlin, or rummer soil with NH₄NO₃, and only limited accumulation occurred with the Harpster soil (Table 5).

Total recoveries of NH₄⁺-N applied as (NH₄)₂SO₄, MAP, DAP, or NH₄NO₃ ranged from 87 to 102% and usually exceeded 94% (Tables 3, 4, 5 and 6) Recoveries with urea were considerably lower, particularly with the Cisne and Blount soils, in which case total recovery after incubation for 42 d was less than 50% (Table 2). The results of a study to estimate gaseous losses of fertilizer ¹⁵N as NH₃ (Table 8) show extensive losses of urea N from the Cisne, Blount, and Catlin soils during the first week of incubation, and leave no doubt that the deficits reported for these soils in Table 2 can be attributed largely, if not entirely, to NH₃ volatilization. The data in Table 8 indicate little, if any, potential for volatilization of NH₄⁺-N applied as (NH₄)₂SO₄, MAP, DAP, or NH₄NO₃.

Table 2 provides ample evidence of the tendency for soils to accumulate substantial amounts of NO₂^- when treated with urea. Maximal recovery of urea N as NO₂^- occurred after incubation for 7-21 d, and ranged from 4 to 64%. The highest percentages were obtained with the calcareous Harpster soil, which can be attributed to a high pH during incubation (Table 7) and to limited loss of urea N as NH₃ (Table 8). The lowest percentages were obtained with the Drummer soil; however, total recovery decreased from 98 to 82% during incubation from 14 to 42 d (Table 2). This decrease cannot be attributed to NH₃ volatilization, which was negligible with the Drummer soil (Table 8), or to biological demtrification, which would have been minimal under the aerobic conditions of incubation. The most likely explanation is that gaseous loss of N occurred through chemical decomposition of NO₂^- at a pH < 5, as existed in ureatreated samples of the Drummer soil during the latter four weeks of the study period (Table 7).

No NO₂^- was detected upon incubation of the Cisne, Blount, Catlin, or Drummer soils with (NH₄)₂SO₄ , MAP, DAP, or NH₄NO₃ (Tables 3, 4, 5 and 6). Substantial levels of NO₂^- were found when samples of the Harpster soil were incubated with (NH₄)₂SO₄ (Table 4), and considerably lower accumulations in samples treated with NH₄NO₃ (Table 5), but NO₂^- was not detected during incubations of the Harpster soil with DAP or MAP (Tables 3 and 6). The latter finding may be related to the fact that MAP and DAP gave a lower soil pH than the other three fertilizers (Table 7).

To further clarify whether the rapid nitrification observed with urea may be due entirely to an effect on soil pH, a study was conducted using (NH₄)₂SO₄ with and without addition of NaOH to simulate the pH obtained with urea. The results gable able 9) indicate that, although the use of NaOH led to some production of NO₂^- from (NH₄)₂SO₄, the levels did not approach those observed for the urea-treated soils. No explanation can be given for this finding, but attention should be drawn to the fact that C02 is liberated during hydrolysis of urea, and this may have promoted nitrification and thereby contributed to the accumulation of NO₂^-. In recent work by Kinsbursky and Saltzman (1990), elevated concentrations of atmospheric C02 were found to increase the population of nitrifying microorganisms and the rate of NH₄⁺ oxidation during incubation of urea-treated soil.

Summary

A laboratory incubation experiment to compare the rate of nitrification for different N fertilizers showed that production of (NO₃ + NO₂^-)-N decreased in the order: urea > DAP > (NH₄)₂SO₄ > NH₄NO₃ > MAP. This decrease was attributed in part to an increase in acidity, but the difference between urea and (NH₄)₂SO₄ was not eliminated by pH adjustment with NaOH. The low rate of nitrification observed with NH₄NO₃ was attributed largely to inhibition of NH₄⁺ oxidation by NO₃^-.

Extensive accumulation of NO₂^- occurred in soils treated with urea, and in some cases this led to gaseous loss through chemical decomposition of the NO₂^-. Substantial loss of urea N also occurred through volatilization of NH₃. Of the fertilizers tested, MAP has special value because: (i) nitrification is retarded; (ii) NO₂^- does not accumulate; and (iii) there is no risk of N loss by NH₃ volatilization.

Acknowledgements

Appreciation is expressed to Barbara Brozak, Diejun Chen, Ben Leak, and Tony White for performing various aspects of the work described.

Tables and Figures Referenced

Table 1: Analyses of soils

Table 2: Recovery after incubation for various periods of N added to soils as ¹⁵N-labeled urea (1 g N kg^-1 soil)

Table 3: Recovery after incubation for various periods of N added to soils as (¹⁵NH₄)₂HPO₄ (1 g N kg^-1 soil)

Table 4: Recovery after incubation for various periods of N added to soils as (¹⁵NH₄)₂SO₄ (1 g N kg^-1 soil)

Table 5: Recovery after incubation for various periods of N added to soils as ¹⁵NH₄NO₃ (1 g NH₄⁺-N kg^-1 soil)

Table 6: Recovery after incubation for various periods of N added to soils as ¹⁵NH₄H₂PO₄ (1 g N kg^-1 soil)

Table 7: Fertilizer effects on soil pH

Table 8: Gaseous losses of fertilizer ¹⁵N as NH₃

Table 9: Effect on ¹⁵N recovery in Catlin soil of adding base with (¹⁵NH₄)₂SO₄ to stimulate the pH of samples treated with ₁₅N-labeled urea

Footnotes and References

¹ Associate Professor, Department of Agronomy, University of Illinois, Urbana, IL.

Bremner, J. M., and L. A. Douglas. 1971. Decomposition of urea phosphate in soils. Soil Science Society ofAmerica Proceedings, 35:575-578.

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 ofAgricultural Science, 51:22-39.
de Boer, W., H. Duyts, and H. J. Laanbroek. 1989. Urea stimulated autotrophic nitrification in suspensions of fertilized, acid heath soil. Soil Biology & Biochemistry, 21:349-354.

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 ofAmerica Proceedings, 21:392-396.

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.

Kinsbursky, R. S., and S. Saltzman. 1990- C02-nitrification relationships in closed soil incubation vessels. Soil Biology & Biochemistry, 22:571-572.

Martikainen, P. J. 1985. Nitrification in forest soil of different pH as affected by urea, ammonium sulphate and potassium sulphate. Soil Biology & Biochemistry, 17:363-367.

McInnes, K. J., and I. R. P. Fillery. 1989. Modeling and field measurements of the effect of nitrogen source on nitrification. Soil Science Society ofAmerica Journal, 53:1264-1269.

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 J. M. Bremner. 1979. A modified diacetyl monoxime method for colorimetric determination of urea in soil extracts. Communications in Soil Science and PlantAnalysis, 10:1163-1170.

Mulvaney, R. L., and L. T. Kurtz. 1982. A new method for determination of 15N-labeled nitrous oxide. Soil Science Society ofAmerica Journal, 46:1178-1184.

Vilsmeier, K., and A. Amberger. 1980. Umsetzung von Cynamamid, Harnstoff and Ammonsulfat in Abhangigkeit von Temperatur and Bodenfeuchtigkeit. Zeitschrift far Pflanzenern4hr and Bodenkunde, 143:47-54.