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a Dep. of Natural Resources and Environ. Sci., Univ. of Illinois, 1102 S. Goodwin Ave., Urbana, IL 61801
b Dep. of Crop Sci., Univ. of Illinois, 1102 S. Goodwin Ave., Urbana, IL 61801
* Corresponding author (mulvaney{at}uiuc.edu)
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ABSTRACT |
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 TOPNOTES ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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Abbreviations: LSD, least significant difference • PPNT, preplant nitrate test • PSNT, presidedress nitrate test • *** significant at the 0.001 probability level
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INTRODUCTION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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Estimation of plant-available N is complicated enormously by the dynamic nature of soil N, owing largely to the effects of temperature and moisture supply on N-cycle processes. Numerous biological and chemical methods have been proposed as an index of soil N availability (Bremner, 1965; Keeney, 1982; Stanford, 1982; Bundy and Meisinger, 1994), but none has been adopted widely for soil testing. Biological methods are necessarily time-consuming because of the need for incubation, and the results represent the net effect of mineralization-immobilization turnover rather than gross mineralization. Chemical methods of estimating potentially mineralizable soil N have been based on an empirical approach, and their use has been very limited because of low correlations with the production of mineral N and crop N uptake.
Soil testing for NO3 is currently considered the best option for identifying sites where N fertilization will be ineffective in producing a yield response by corn (Bundy and Meisinger, 1994). Two soil NO3 tests have been developed that differ in the time and depth of sampling. With the preplant NO3 test (PPNT), profile samples are collected in the early spring to a depth of 60 or 90 cm, to account for carryover of mineral N from previous cropping (e.g., Bundy and Malone, 1988; Roth and Fox, 1990; Schmitt and Randall, 1994). With the presidedress NO3 test (PSNT), soil sampling is done to a depth of 30 cm in late spring, so that soil N mineralization can be taken into account and supplemented, if necessary, by sidedressing (e.g., Magdoff et al., 1984; Fox et al., 1989; Blackmer et al., 1989; Meisinger et al., 1992; Bundy and Andraski, 1993). The PSNT has been recommended more widely than the PPNT in the eastern USA, but usage has been limited by the need to collect soil samples during the growing season, and by the fact that N fertilization must be postponed until after testing and can be ineffective if adverse weather conditions delay sidedressing. Besides logistical problems, an inherent limitation with the PPNT and PSNT arises from the extensive spatial and temporal variability in soil NO3 concentrations, which depend on numerous N-cycle processes, including mineralization, immobilization, nitrification, denitrification, leaching, and plant uptake. Consequently, a one-time test for soil NO3 is apt to be of little value for predicting crop N availability throughout the growing season, particularly in a humid region where these processes occur extensively.
Ideally, a soil test for N would estimate a labile organic fraction that supplies the plant through mineralization. Such an approach would have the major advantage over NO3 testing that soil test levels would depend on fewer N-cycle processes and should, therefore, be less variable. This would make the time of soil sampling much less critical than with NO3 testing, so that soil N availability could potentially be predicted on the basis of a one-time test prior to the growing season.
Numerous attempts have been made to identify a labile pool of soil organic N through chemical fractionation of the N in soil hydrolysates (e.g., Keeney and Bremner, 1964; Porter et al., 1964; Khan, 1971; Smith and Young, 1975; Meints and Peterson, 1977), but with little tangible progress. The stagnation can be attributed, at least in part, to serious defects in methodology that vitiated analyses for amino sugar N and amino acid N. These defects were identified and eliminated through a substantial effort that ultimately led to simple diffusion methods of fractionating the N in soil hydrolysates (Mulvaney and Khan, 2001).
In several recent studies throughout the north-central and northeastern USA, numerous sites have been detected where corn does not respond to N fertilization (e.g., Bundy and Malone, 1988; Blackmer et al., 1989; Fox et al., 1989; Roth and Fox, 1990; Meisinger et al., 1992; Brown et al., 1993; Schmitt and Randall, 1994). Such sites are often associated with recent manuring or the presence of a previous forage legume, but this was not the case with many of the 33 nonresponsive sites detected in an Illinois study that involved 75 site–years (Brown et al., 1993). The newly developed diffusion methods were applied to soil samples collected from responsive and nonresponsive sites in the latter study, so as to ascertain whether a specific fraction of soil organic N might be implicated in nonresponsiveness to N fertilization. The results showed a much higher concentration of amino sugar N for nonresponsive than for responsive soils, whereas no consistent difference was detected in their concentrations of total hydrolyzable N, hydrolyzable NH4-N, or amino acid N (Mulvaney et al., 2001). In subsequent incubation studies reported in the latter publication, nonresponsive soils produced a much larger quantity of mineral N than did responsive soils, and mineralization was accompanied by a net decrease in amino sugar N but not in amino acid N.
The methods employed by Mulvaney et al. (2001) to differentiate between responsive and nonresponsive soils require hydrolysis with 6 M HCl for 12 h, followed by filtration and neutralization of the hydrolysate, and are therefore unsuitable for routine soil testing. The purpose of the work reported here was to develop a much simpler technique, whereby amino sugar N can be readily estimated to detect sites that do not require N fertilization. The soil test developed was evaluated relative to analyses of hydrolyzable amino sugar N, and by comparing test values for soils that differed widely in the yield response of corn to N fertilization.
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MATERIALS AND METHODS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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The 25 sites studied herein are characterized in Table 1, which shows the soil type and physicochemical properties, the previous crop, the tillage system in use, the type and amount of manure applied, PPNT, PSNT, and check-plot corn yield data from Brown (1996), and the percentage yield response by corn to N fertilization at the optimal N rate. Of the chemical analyses reported in Table 1, data for pH, available P, and exchangeable K were obtained from Brown (1996), and organic C and total N were determined as described by Mulvaney and Kurtz (1982). Applications of manure N were estimated on the basis of the quantity of material applied as indicated by farmer records, and the average N concentration according to the Illinois Agronomy Handbook (1998). The percentage yield response by corn to N fertilization was calculated as 100 x (optimum yield - check-plot yield)/check-plot yield, using data reported by Brown (1996) for check-plot yield (i.e., the yield without sidedressing, as reported in Table 1) and optimum yield. The latter value was determined by fitting N rate and corresponding yield data to a quadratic plateau model by nonlinear regression (SAS Institute, 1993).
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Four additional Illinois surface (0–15 cm) soils were used for some of the studies reported, including three Mollisols collected from fields under soybean [Glycine max (L.) Merr.] production and a Histosol obtained from a permanently waterlogged site. Before use, each sample was air dried and crushed to pass through a 2-mm screen. Their physicochemical properties are reported in Table 2.
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Using the diffusion methods described by Mulvaney and Khan (2001), the soil hydrolysates were analyzed for total hydrolyzable N, NH4-N, (NH4 + amino sugar)-N, and amino acid N. Amino sugar N was determined as the difference between (NH4 + amino sugar)-N and NH4-N.
Soil Test Method
Apparati
Diffusion unit
The diffusion unit used consists of a 473-mL (1-pint) wide-mouth Mason jar equipped with a lid that has been modified to support the bottom of a 60-mm (dia.) Pyrex petri dish (Corning Glass Works, Corning, NY). The necessary modifications are described in detail by Mulvaney (1996), Khan et al. (1997), and Mulvaney et al. (1997a). Further information about Mason-jar diffusion methodology can be found therein, including a description of the appropriate cleaning procedures.
Electric hot plate
A commercial griddle (Model 76212; West Bend, West Bend, WI) was used. Before use, the heat control was adjusted so that a temperature of 48 to 50°C was obtained when a thermometer was immersed in 100 mL of deionized water in a Mason jar placed in the center of the griddle.
Microburette or automatic titrator
Titrations were performed using a 5-mL microburette or a Metrohm Model 678 EP/KF Processor equipped with a Model 665 Dosimat (Metrohm, Herisau, Switzerland) and a combination electrode designed for flat-surface measurements (Model 13-620-289; Fisher Scientific, Pittsburgh, PA).
Reagents
Sodium hydroxide solution (2 M)
Reagent-grade NaOH pellets (80 g) were dissolved in 
800 mL of deionized water in a 1-L volumetric flask. After cooling, the solution was diluted to 1 L and mixed thoroughly. The flask was kept tightly stoppered to minimize absorption of atmospheric CO2 during storage of the NaOH solution. Alternatively, 2 M NaOH is available from Fisher Scientific (cat. no. LC24380).
Boric acid-indicator solution
A reagent containing 40 g of H3BO3 L-1 was prepared as described by Mulvaney (1996), Khan et al. (1997), and Mulvaney et al. (1997a). Alternatively, a suitable reagent may be obtained from Fisher Scientific (cat. no. LC11750).
Dilute sulfuric acid (0.01 M standard)
This reagent was prepared by adding 5.6 mL of concentrated (18 M) H2SO4 to 10 L of deionized water in a 10-L Pyrex solution bottle (Corning Glass Works, Corning, NY). After thorough mixing with a motorized stirrer, the solution was standardized by titrating several 5-mL aliquots of a THAM solution that was prepared by dissolving 0.2430 g of dried, certified THAM (Sigma, St. Louis, MO) in 100 mL of deionized water in a volumetric flask. The endpoint for these titrations was determined as described in the procedure that follows. The molarity of the titrant was calculated as 0.05/V, where V is the mean value for the milliliters of H2SO4 required to reach the endpoint. The calculated molarity was multiplied by 28000 to obtain the titer (µg N mL-1). Alternatively, standard 0.01 M (0.02 N) H2SO4 may be purchased from Fisher Scientific (cat. no. SA226).
Procedure
A 1-g sample of air-dried soil (<2 mm) was weighed into a Mason jar. A petri dish was attached to the jar lid with a cable tie, and 5 mL of H3BO3-indicator solution was dispensed into the dish. The soil sample was then treated with 10 mL of 2 M NaOH, and the jar was swirled to mix the contents, while taking care to minimize soil adherence to the wall of the jar. Within 15 to 30 s, the lid was placed on the jar and sealed with a screw band, and the jar was transferred to the hot plate. After 5 h, the jar was removed from the hot plate and opened, and the petri dish was released from the jar lid. The H3BO3 solution was diluted with 5 mL of deionized water, and subsequently titrated with 0.01 M H2SO4. Prior to titration, 5 mL of H3BO3 solution was dispensed into a petri dish containing 5 mL of deionized water, and the endpoint was established on the basis of the resulting color (for manual titrations) or pH (for automatic titrations). The micrograms of N liberated by diffusion was calculated as S x T, where S is the volume of H2SO4 used in titration of the sample and T is the titer of the titrant (for 0.01 M H2SO4, T = 280 µg N mL-1).
Development of Soil Test Method
The addition of NaOH and the diffusion period specified in the method described were established through comparative studies involving treatment of soil samples (four replicates) with 10 mL of 1, 2, or 5 M NaOH, followed by heating at 48 to 50°C for 1, 2, 3, 4, 5, 6, 7, 8, 12, or 24 h. The soils selected for use in these studies included the responsive and nonresponsive soils that had the highest and lowest concentrations of amino sugar N.
A study was conducted to ascertain whether any recovery of nonexchangeable NH4-N occurs by the method described, by comparing soil test values obtained with KOH versus NaOH. The soils used in this study were selected on the basis of nonexchangeable NH4-N concentration determined as described by Mulvaney (1996), and included two nonresponsive soils, a responsive soil, and three of the soils listed in Table 2. In each case, analyses were performed (four replicates) as specified, or by substituting 2 M KOH for 2 M NaOH.
Recovery tests were performed using 15N to ensure that the method described estimates (NH4 + amino sugar)-N but not NO3-N, NO2-N, or amino acid N. Four soils were used in this study, including one from a manure disposal site. Analyses were performed (four replicates) as specified to serve as a control, or after addition of 1 mL of deionized water containing 300 µg of N as (NH4)2SO4 (1.422 atom % 15N), glucosamine HCl (1.386 atom % 15N), KNO3 (1.646 atom % 15N), NaNO2 (1.534 atom % 15N), or glycine (1.142 atom % 15N). The labeled N was added following NaOH to prevent NH4 fixation, and the jar was then sealed immediately to avoid gaseous loss of NH3. Following quantitative determinations by titration, isotope-ratio analyses were performed on each sample by the Rittenberg process, using an automated system described in detail by Mulvaney et al. (1990), Mulvaney and Liu (1991), and Mulvaney et al. (1997b). Percentage recovery (R) was calculated as
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In a study to evaluate the effect of sampling depth on soil test values, analyses were performed (four replicates) by the method described on profile samples of two responsive and two nonresponsive soils, representing depths of 0 to 15, 15 to 30, and 30 to 60 cm.
To elucidate the importance of proper temperature control during the 5-h diffusion period specified, analyses were carried out with heating at 40, 44, 46, 48, 50, 52, 56, or 60°C. In each case, there were four replicate samples of two nonresponsive and two responsive soils. The soils used were selected so that both groups would vary widely in organic C.
The importance of aggregate size in the soil test described was investigated by performing analyses on samples of soil that had been crushed to pass through a 2.0-mm screen, with or without further crushing to <0.15 mm. This investigation was carried out using the four soils listed in Table 2, and involved four replicate determinations.
Data Analysis
Data from replicate determinations were characterized by computing means and standard deviations. In some cases, mean values were compared on the basis of a least significant difference (LSD) at the 0.001 probability level, or by performing pairwise t-tests. Simple correlation or regression analyses were employed to quantify the linear relationship between soil test N and hydrolyzable amino sugar N, using data obtained for the 25 soils listed in Table 1. The ability of the soil test described to differentiate nonresponsive from responsive sites was evaluated by plotting the fertilizer response data for these sites versus their soil test values.
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RESULTS AND DISCUSSION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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45–56 kg ha-1), and exchangeable K (290–335 kg ha-1). Of this group, eight soils had been manured, but the rate of manure N application varied from 85 to >2500 kg ha-1 and far exceeded crop N requirements at three sites that had been used for manure disposal (Soils 1, 2, and 5). Corn followed alfalfa (Medicago sativa L.) at one of the remaining nonresponsive sites (Soil 3), while no-till soybean was the previous crop at another (Soil 12). Soils 8 and 11 were collected from unmanured sites under continuous corn, but a high level of Bray-1 P and/or exchangeable K suggested the possibility of previous manuring or excessive fertilization. On the basis of the proven-yield approach described in the Illinois Agronomy Handbook (1998) to guide N-fertilizer recommendations for corn, six of the 12 nonresponsive sites listed in Table 1 would have been identified correctly for their complete lack of yield response to N fertilization. In each case, the N credit from manure (190 kg manure N ha-1) would have exceeded the N requirement estimated for the yield goal, although this sometimes occurred only because of an additional N credit for corn following soybean (48 kg ha-1). None of the remaining six sites in this group would have been identified by the proven-yield approach as being nonresponsive to N fertilization, and therefore would have been fertilized at an economic loss to the farmer and with the risk of causing environmental pollution. Overfertilization would have been more extensive for Soils 8 and 11, which came from sites where continuous corn was grown without manure, than for Soils 3, 6, 10, and 12, in which case the use of manure or a previous legume led to an N credit.
Soil testing for NO3 was done by Brown et al. (1993) to evaluate the PPNT and PSNT for identifying sites that do not require N fertilization for corn production, as a possible improvement over the proven-yield approach. The data thereby obtained for the 25 soils used in our work are included in Table 1. Assuming a critical value of 16 mg N kg-1 (Schmitt and Randall, 1994), four of the 12 nonresponsive sites were detected by the PPNT. The PSNT was somewhat more effective in identifying six of these sites, relative to a critical value of 21 mg N kg-1 (Fox et al., 1989; Bundy and Andraski, 1993), but provided no improvement over the proven-yield approach, which also identified six of the 12 nonresponsive sites. Most of the soils having a high concentration of NO3 had been manured, although several such sites were not identified by either the PPNT or the PSNT, including one used to dispose of liquid swine (Sus scrofa domesticus) manure (Soil 2). A nonresponsive site where first-year corn followed alfalfa (Soil 3) was identified by the PSNT, but not by the PPNT.
Recent work in our laboratory suggests that a soil test could be developed to estimate the supply of plant-available N, based on chemical analyses for a specific fraction of soil organic N that is highly mineralizable. This fraction was identified by comparing N-distribution analyses for soils that differed in whether N fertilization had effected a yield response by corn, using diffusion methods developed by Mulvaney and Khan (2001). The results showed 11 nonresponsive soils to be significantly higher than seven responsive soils in their concentrations of amino sugar N, whereas no consistent difference was observed for total hydrolyzable N, hydrolyzable NH4-N, or amino acid N (Mulvaney et al., 2001). The same finding applies in the present study, which involved a larger number of soils representing additional nonresponsive and responsive sites. This is apparent from Table 3, in that the two groups were completely resolved (P < 0.001) on the basis of amino sugar N. The lowest value for any nonresponsive soil was 34% higher than the highest value for any responsive soil, and on average, the difference in amino sugar N was more than 200%.
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To optimize reaction conditions for liberating amino sugar N from soil, studies were conducted to compare different concentrations of NaOH and different diffusion periods, in which diffusions were performed at 48 to 50°C on a hot plate to promote the alkaline decomposition of amino sugars (Mulvaney and Khan, 2001). The results (Fig. 1) were obtained using the two responsive and the two nonresponsive soils that had the highest and lowest concentrations of amino sugar N. As expected, a larger amount of N was liberated when diffusion was performed with a higher concentration of NaOH or for a longer period. A 2 M reagent and a 5-h diffusion period were adopted in the soil test described as the best compromise in terms of speed, convenience, sensitivity, and resolution. A longer diffusion period was required with 1 M NaOH to clearly differentiate responsive from nonresponsive soils, whereas the diffusion period was much more critical with 5 M NaOH because of less temporal stability in soil test values. With 2 M NaOH, a 5-h diffusion period was adequate to easily resolve the two nonresponsive soils from the two responsive soils, and even provided sufficient resolution to clearly distinguish among all four of these soils. Moreover, the data in Fig. 1 show that soil test values obtained with 2 M NaOH do not increase appreciably if diffusion is continued beyond the 5-h period specified, although this practice should be avoided, since prolonged heating may lead to drying of the H3BO3 solution used to absorb gaseous NH3 and thereby vitiate the analysis. The latter problem does not occur if heating is discontinued after 5 h, as very little, if any, change has been observed in soil test values by leaving the jar unopened overnight at room temperature (25°C). This could be a valuable option in processing a large number of soil samples, as is often necessary in soil testing laboratories.
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The soil test described could have been easily modified to recover (NO3 + NO2)-N through addition of Devarda's alloy as a reducing agent, but this modification was omitted deliberately, so as to avoid two problems that result from the dynamic nature and mobility of soil NO3. One of these problems is the extensive spatial and temporal variability that occurs in soil NO3 concentrations (Lockman and Storer, 1990; Cahn et al., 1994; Hergert et al., 1995; Everett and Pierce, 1996), which would reduce soil test reliability in detecting sites that do not need N fertilization. The other is the need for profile sampling to account for downward movement of NO3 through leaching. Such sampling is incompatible with routine soil testing for pH, P, and K, which normally involves sampling to a depth of 15 to 18 cm. The soil samples used in our work were collected from the 30-cm surface; however, a study to compare N-test values for different profile depths (Table 6) showed that the highest values were obtained for the 15-cm surface, and that a decrease occurred with greater depth. This is exactly what would be expected for an organic fraction of soil N that is subject to little, if any, transport by leaching. Table 6 suggests that standard sampling techniques would be appropriate for the soil test described, and that testing could be done in conjunction with normal soil tests for pH, P, and K. Table 6 also indicates that testing could be done with any tillage system, since N-test values for the four soils used followed the same order, regardless of sampling depth.
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The soil samples used in the present project were ground much more finely than samples for routine soil testing, which are usually screened to <2 mm to remove organic detritus or inert rock fragments. A study to compare soil test values for samples differing in aggregate size (Table 7) showed no appreciable difference with and without fine grinding (to <0.15 mm), as would be expected from the fact that soil aggregates quickly form a slurry when treated with 10 mL of 2 M NaOH. This suggests that the soil test described will require no special processing of soil samples.
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SUMMARY AND CONCLUSIONS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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The soil test described has obvious value for improving N-fertilizer efficiency, increasing the profitability of corn production, and reducing the adverse environmental effects of excessive N fertilization.
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ACKNOWLEDGMENTS |
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NOTES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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Received for publication February 20, 2001.
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REFERENCES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONSUMMARY AND CONCLUSIONSREFERENCES |
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