Biol Fertil Soils (2001) 34:179–184 DOI 10.1007/s003740100395
O R I G I N A L PA P E R
Torben A. Bonde · Tommy Harder Nielsen Morten Miller · Jan Sørensen
Arginine ammonification assay as a rapid index of gross N mineralization in agricultural soils Received: 9 March 2001 / Published online: 3 July 2001 © Springer-Verlag 2001
Abstract Seasonal dynamics of in situ gross nitrogen (N) mineralization rates were measured using the 15N-NH + isotope dilution method in a Danish soil sub4 jected to four different agricultural practices (set aside, barley, winter wheat and clover). Results were compared to arginine ammonification in the soil samples measured as NH4+ production following addition of excess (1 mM) arginine. In the set aside, barley, winter wheat and clover soils the average annual rates of gross N mineralization (0.29, 0.60, 1.34 and 1.75 µg NH4+-N g–1 day–1, respectively) and arginine ammonification activity (0.21, 0.55, 0.88, and 1.33 µg NH4+-N g–1 h–1, respectively) were well correlated. Furthermore, the seasonal variations of gross N mineralization and arginine ammonification activities were very similar, showing rapid responses to rainfall and generally higher activities in wetted soils. As tested in the laboratory, the arginine ammonification activity correlated well with heterotrophic microbial respiration activity (CO2 production) in soil samples and further displayed a simple, one-component MichaelisMenten kinetics with a high affinity for arginine (Km value of 48 µM ±5 µM) as determined from non-linear parameter estimation. This indicated that arginine ammonification activity was primarily due to microorganisms, and the activity was also shown to be at a minimum in sterile soil samples. All evidence thus supported that our standard assay of arginine ammonification activity provides a good index of gross N mineralization rates by the microorganisms in soil under in situ conditions. T.A. Bonde · T.H. Nielsen · M. Miller · J. Sørensen (✉) Department of Ecology, Section of Genetics and Microbiology, Royal Veterinary and Agricultural University, Thorvaldsensvej 40, 1871 Frederiksberg C, Denmark e-mail:
[email protected] Fax: +45-35-282606 Present addresses: T.A. Bonde, National Environmental Agency, Strandgade 29, 1401 Copenhagen K, Denmark M. Miller, Department of General Microbiology, Institute of Molecular Microbiology, University of Copenhagen, Sølvgade 83H, 1307 Copenhagen K, Denmark
Keywords Arginine ammonification · Gross N mineralization · N mineralization index
Introduction Soil nitrogen (N) mineralization is a key process determining the quantity of N available to a crop and hence the quantity of fertilizer required (Powlson 1997). To allow for rapid assessment in fertilizer planning, several assays determining microbial biomass, activity and specific enzyme pools have been evaluated as possible indices of soil N mineralization (e.g. Keeney 1980; Stanford 1982; Burton and McGill 1992). Microbial biomass was found to predict net N mineralization, representing the actual release of inorganic N from the balance of gross N mineralization and microbial incorporation (Burton and McGill 1992). Recently, Murphy et al. (1998) also found a strong correlation between soil microbial biomass and gross N mineralization in vertical profiles of non-fertilized soils. Both reports thus supported the use of microbial biomass assays as indirect measures of soil N mineralization. In addition to assays of microbial biomass, certain soluble enzyme pools may also relate to N mineralization in soil. Hence, it has been found that protease activity in soil correlates weakly to rates of inorganic N production (Burton and McGill 1992). However, extracellular enzymes such as the proteases are released into the soil water solution and are thus prone to subsequent, partial stabilization in the soil matrix. This stabilized fraction of residual enzyme activity in the soil matrix circumvents both the control by microorganisms per se and by soil factors affecting other reaction steps in N mineralization. The assays based on extracellular enzymes are thus unlikely to reflect the dynamic nature of gross N mineralization (Burns 1982), e.g. short-term events after rainfall or irrigation. Amino acids and peptides are the largest pool of soil organic N identifiable after acid hydrolysis (Stevenson 1982). As ammonia release (ammonification) during ami-
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no acid degradation (e.g. deamination reaction involving the primary amino-N group of amino acids) is anticipated to take place intracellularly in the soil microorganisms, it may be suggested that ammonification activity is more closely correlated with N mineralization in soil than the activity of extracellular enzymes such as proteases. In the model proposed by Burton and McGill (1992), histidine ammonification (NH4+ production when histidine was added at saturating concentrations) was suggested to serve as an index of soil gross N mineralization. A comparable enzyme assay based on arginine ammonification activity (NH4+ production after addition of saturating arginine concentrations) was suggested by Alef and Kleiner (1986) to serve as a general index of overall soil microbial activity as verified by measurements of O2 respiration. The short-term assay of arginine ammonification convincingly demonstrated that only the actual enzyme content was determined, without influence from de novo synthesis during the assay. It was further demonstrated that only the soil microorganisms (and not plant fragments or fauna) were responsible for the arginine ammonification. Subsequent studies have found that the arginine ammonification assay is useful as a rapid assessment of soil microbial biomass (Alef and Kleiner 1987; Lin and Brookes 1999). Even if arginine ammonification correlates with microbial biomass and microbial activity (O2 respiration) in soil, there has never been direct evidence that arginine ammonification provides a suitable index of gross N mineralization. However, as already anticipated from the model of Burton and McGill (1992) based on histidine ammonification, arginine ammonification could be such an index of gross N mineralization in soil and be even more sensitive than the assay based on histidine. Hence, Alef and Kleiner (1986) showed that arginine ammonification gave the highest rate of NH4+ production when a number of amino acids were compared in agricultural soil. This could either be due to the high N content (a total of four amino-N groups) of arginine or the occurrence of several pathways of arginine ammonification in soil. The reactions responsible for arginine ammonification can be grouped in two categories: (1) arginine deamination (or deimination) activity resulting in immediate release of one NH4+ group and (2) arginase activity with production of urea, which can be rapidly hydrolysed by urease activity with release of NH4+. Urease activity in agricultural soils was recently demonstrated to be high and quantitatively important for in situ N mineralization (Nielsen et al. 1998). In the present work we compared the arginine ammonification assay directly with a 15N-NH4+ isotope dilution technique providing a measure of gross N mineralization in soils with different agricultural crops (set aside, barley, wheat and clover). We report that the rapid and sensitive assay of arginine ammonification provides a good index of gross N mineralization in these soils, reflecting both soil-type differences associated with cropping systems, seasonal patterns in the fields, and shortterm fluctuations of activity after rainfall in one of the selected fields (barley).
Materials and methods Soil sampling and characteristics Soils were collected during 1994–1995 at the “Højbakkegård” Experimental Station belonging to the Royal Veterinary and Agricultural University, Copenhagen, Denmark. The soil was collected from the upper 30 cm of a course sandy soil consisting of 3.0% clay, 10.0% silt, 39.5% fine sand and 47% coarse sand. The total C and N contents were 1.0% and 0.1%, respectively. Three of the soils were designated according to their crop during the 1994 growing season, which was spring barley (Hordeum vulgare “Alexis”), white clover (Trifolium repens “Milka”) and a field with no crops designated “set-aside”. During the growing season of 1995 the barley, clover and set-aside soils were left uncropped while soil samples in the fourth soil were collected between rows of winter wheat (Triticum aestivum “Hussar”). The four sampling sites were situated within 700 m2. In the laboratory, soil samples were left at the in situ temperature for a few hours (max. 8 h) and then homogenized gently to a particle size below 4 mm. Larger pieces of plant fragments were removed before the experiments. Arginine ammonification assay Soil samples (0.1 g), weighed into 1.5-ml Eppendorf vials in two sets of four replicates were treated with 20 µl 10 mM L-arginine (Merck)and 180 µl Milli-Q purified water (Millipore). The arginine solution was not added to control soils. Samples tested for arginine ammonification activity received 20 µl 10 mM L-arginine solution (final concentration of 1 mM arginine) to initiate the assay. All samples were then Whirly-mixed (3 s) and incubated for 1–2 h at 20°C without further agitation. The incubation time was set to allow for less than 20% of the added arginine to mineralize during the incubation. After incubation, control samples also received 20 µl 10 mM L-arginine solution. Addition of 0.8 ml icecold 2 M KCl solution to both sets terminated the incubations. After ten end-over-end rotations by hand, all samples were immediately centrifuged (20,000 g for 10 min at 4°C) and the NH4+ concentration of the supernatants were analysis by a standard colorimetric assay presented by Werdouw et al. (1977) and modified by Højberg et al. (1996). Briefly, three stock solutions were used: (A) 100 g sodium salicylate (Merck), 10 g potassium sodium tartrate (Fluka), and 100 g tri-sodium citrate (Merck) in 1 l Milli-Q water; (B) 100 ml 5% sodium hypoclorite (Aldrich) and 100 ml 5 M sodium hydroxide (Merck) in 1 l Milli-Q water; (C) 0.5 g sodium nitroprusside (Sigma) in 1 l Milli-Q water. From the supernatant, 0.5 ml was pipetted into 1-ml micro-cuvettes containing 0.4 ml solution A, after which 0.08 ml of both solutions B and C were added. The samples were immediately mixed and allowed to stand for 0.5 h before the absorbance was read at 625 nm. Arginine ammonification activity was expressed in µg NH4+-N g–1 dw soil h–1, using a correction for soil water content determined by oven drying at 105°C overnight. To determine the kinetics constants of arginine ammonification in the different soils, different concentrations (0.01, 0.05, 0.1, 0.25, 0.5, 0.75, 1, 3, and 5 mM) of L-arginine were incubated as described for the standard assay above. Response curves of enzyme activity versus substrate concentration were fitted using simple Michaelis-Menten kinetics. The non-linear regression analysis was performed using the Marquardt procedure of PROC NLIN in the SAS/STAT Guide (SAS Institute, N.C., USA). Respiration assay A respiration assay of rewetted soil samples from the barley field was based on the rate of CO2 accumulation in closed vials (Miller et al. 2001). Soils samples were first desiccated to approximately 2% water content (w/w) and stored for 2 days before rewetting to approximately 12% water content (w/w). The rewetted soil (15 g)
181 was then immediately transferred to five incubation flasks (100 ml) which were sealed with rubber stoppers. At the sampling time 5-ml gas samples were withdrawn from the headspace of the flasks and CO2 accumulations were determined on a gas chromatograph (Hewlett Packard 5890 Series II) equipped with a TCD detector. Gross N mineralization assay The 15N-isotope dilution method to measure gross N mineralization in field soil samples has been described in detail by Nielsen et al. (1998). The soil (150 g) was weighed into a polyethylene bag and then sprayed with a 24 mM (NH4)2SO4 solution (98% 15N; Cambridge Isotope Laboratories, Mass.), under gentle mechanical mixing. The water content increased by less than 2% by the 15NH4+ addition. Labelled soil samples (8 g) were weighed into 50-ml centrifuge tubes with loosely attached screw caps and incubated at the in situ temperature for 24–72 h, depending on activity level. Triplicate samples were subsequently sacrificed on four or five occasions during the incubation period, in order to verify that gross N mineralization activities were constant (Nielsen et al. 1998). After each sampling, 24 ml 2 M KCl solution (ice cold) were added and samples were shaken on an orbital shaker (100 rpm) for 1 h, before the supernatant was recovered by centrifugation (4,500 g). The supernatant was filtered through GF/C filters pre-washed in 2 M KCl solution and then stored at –20°C before being analysed for NH4+ concentration using the standard colorimetric assay described above for the arginine ammonification assay. The 15N content (atom%) of the NH4+ pool was determined on a Tracermass mass spectrometer (Europa Scientific, Crewe, UK) after conversion of the NH4+ pool to N2 gas by the micro-diffusion technique of Risgaard-Petersen et al. (1995). The 15N-atom percentage of the NH4+ pool was calculated by the method of Nielsen (1992). Gross N mineralization rates were calculated using the isotope dilution technique described by Blackburn (1979). The method involves a determination of gross N mineralization, i.e. total NH4+ release (d) and total consumption of NH4+ (i) according to two equations: P(t ) = P0 + (d − i)t
(1)
[
]
(d − i)t + P0 ln( R − 15n) = ln( R0 − 15n) − d × ln d −i P0
(2)
where P0 and P(t) are the 14+15N pools at time 0 and time t, respectively, 15n is the natural abundance of 15N (0.37 atom %), and R0 and R are the relative abundances of 15N expressed as 15N/15N + 14N at time 0 and time t, respectively. When i and d are constant, P(t) in Eq. 1 is linear with time. Equation 2 describes that ln (R–15n) is linear with ln {[(d–i)t + P0]/P0}, represented by the slope –d/(d–i) and the intercept ln (R0–15n). The gross N mineralization (d), expressed in µg NH4+-N g–1 dw soil day–1, can be calculated by combination of Eqs. 1 and 2.
Results Arginine ammonification assay in laboratory tests Dry (desiccated, approximately 4% water content) and wet (field moist, approximately 15% water content) samples of soil from the barley field were prepared for comparison of arginine ammonification activities at different arginine concentrations. Figure 1 shows that both soil samples displayed simple Michaelis-Menten kinetics with a high affinity for arginine (Km 48 µM ±5 µM) as determined from non-linear parameter estimation (Robinson 1985). The results indicated that the standard
Fig. 1 Arginine ammonification in moist and dry soils amended with different concentrations of arginine. The activity was fitted by a non-linear regression analysis including simple MichaelisMenten kinetics (n=3 ± SD)
Fig. 2 Arginine ammonification and microbial respiration after rewetting of air-dried soil to a final water content of 12% (n=3 ± SD)
assay, using 1 mM arginine, gave a constant maximum (Vmax) rate of NH4+ release during the incubation time of 1–2 h at 20°C. Further, according to the Vmax rates measured, a maximum of 20% of the added arginine was mineralized during the incubation. The results strongly supported that arginine ammonification was associated with microbial activity (CO2 respiration) in the tested soil. Rewetting of dry soil from the barley field was used to test if arginine ammonification activity displayed a response with transient increase of activity, as has been reported for C and N mineralization (e.g. Powlson and Jenkinson 1976). The results from the experiments shown in Fig. 2 demonstrated that arginine ammonification indeed responded quickly to the increase in soil moisture. The barley soil, which had been dried and stored for 2 days at 2% water content and then re-wetted to 12% water content, showed a peak level of arginine ammonification activity within the first 2 days. A similar, transient increase of respiration activity was observed and the results thus confirmed that arginine ammonification was due to heterotrophic microbial activity.
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Fig. 3 Seasonal variation of arginine ammonification and soil water content in a barley field during July to September of 1994 (n=3 ± SD)
Arginine ammonification assay in the field study In the field study, soil from the barley field was analysed on 16 occasions during an intensive sampling period in July–August 1994. Since temperature was relatively high and constant during the summer period, results from the standard assay performed at 20°C were recorded. As shown in Fig. 3, this period was generally very dry and interrupted by only three rainfall events in mid-July, late July and mid-August, respectively. As expected, arginine ammonification activity was at a minimum of approximately 1 µg N g–1 h–1, during the dry period when the soil water content was only 2–3 % (w/w). The three rainfalls resulted in short, transient increases of soil water content, but most interestingly, there were similar, transient increases of arginine ammonification activity associated with the rainfall events. Hence, while short bursts of arginine ammonification were associated with the two short (1 day) rainfalls in July, there was a continued increase of activity during the several days of rainfall occurring in mid-August. At this time, the activity reached a maximum of approximately 2.5 g N g–1 h–1. It seemed that each of the three rainfall events succeeding dry periods of 1–2 weeks were able to induce a marked increase of arginine ammonification, presumed to reflect an increase of gross N mineralization in soil. The results from the field bear similarity to those from the laboratory (Fig. 2), which showed that arginine ammonification (gross N mineralization) increases shortly after rewetting of dry soil, presumably due to a flush of substrate made available for microbial activity, and hence N mineralization, in the soil. The direct comparison of arginine ammonification activity and gross N mineralization determined by the isotope dilution method was performed in the subsequent season, March–September 1995. The study was made in the same field as in 1994, but this time without barley crop vegetation. Average soil temperatures increased from approximately 4°C to 20°C during this period and results from the standard arginine ammonification assay
Fig. 4 Seasonal variation of arginine ammonification and gross N mineralization activities in a barley soil during April to September of 1995. Arginine ammonification activity was corrected to field soil temperatures using a Q10 value of 2.0. Soil temperature and soil water content are indicated (n=3 ± SD)
performed at 20°C were corrected to field soil temperatures using a Q10 value of 2.0. This value is reasonable based on microbial activity variations in the temperature regime of 4°C to 20°C in soil. The 15N assay of gross N mineralization was performed at field soil temperatures. Figure 4 shows both the in situ soil temperature and water contents and the recorded activities of arginine ammonification and gross N mineralization. While the temperature increased over the sampling season in spring and summer, there was a concurrent decrease of soil water content from wet conditions of approximately 18% (early April) to very dry conditions of approximately 2% (late August). The seasonal study clearly demonstrated that both arginine ammonification and gross N mineralization increased steadily from early April when minimum activities of approximately 0.2 g N g–1 h–1 and 0.4 g N g–1 day–1 were recorded for the two activities, respectively. The increase took place until mid-July when the soil temperature reached a maximum at approximately 20°C and the recorded activities were at their maxima of approximately 1.0 g N g–1 h–1 for arginine ammonification and approximately 1.0 g N g–1 day–1 for gross N mineralization. However, as desiccation continued and soil water content became lower than approximately 5%, there were dramatic decreases of first gross N mineralization and then arginine ammonification. Gross N mineralization even reached an undetectable level at the lowest soil water content of approximately 2% in late August. In the dry soil with less than 5% water content, the gross N mineralization thus became markedly lower than the arginine ammonification. The apparent uncoupling of arginine ammonification and gross N mineralization seems to be an exceptional characteristic of very
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Fig. 5 Comparison of annual means of arginine ammonification and gross N mineralization activities in four different agricultural fields in 1995. Arginine ammonification was corrected to field soil temperatures using a Q10 value of 2.0. Sampling sites were uncropped during the study, but were designated by their cropping plant in 1994 (n=7 or 9 ± SD)
dry conditions in the barley soil, however, and there was generally a good match between the two activities. In the seasonal study conducted on seven to nine samplings during 1995, we compared the seasonal variations and annual averages of both arginine ammonification and gross N mineralization in the four different soils (set-aside, barley, clover and wheat). Seasonal variations in the soils representing set-aside, clover and wheat (data not shown) were quite similar to that reported for barley (Fig. 4). Furthermore, when comparing the seasonal averages of arginine ammonification and gross N mineralization in all four soils, there was a very good correlation between the two activities as shown in Fig. 5. The arginine ammonification assay proposed to reflect gross N mineralization thus resulted in the same ranking of the four fields as did the direct measures of gross N mineralization by the 15N isotope dilution technique.
Discussion It may be argued that assays based on the terminal reactions steps for intracellular amino acid (arginine, histidine, etc.) degradation in microorganisms are better suited than assays based on proteolysis because the process represents the actual production and release of mineral N in the soils. Hence, it may be expected that the amino acid ammonification assays may reflect the in situ N mineralization in soil, including both the overall variation between different soil types and cropping systems, seasonal variation within a specific field or crop production, and short-term fluctuations after irrigation or rainfall at a field site. Representing one of the amino acid ammonification assays, the arginine ammonification assay was demonstrated to be rapid, sensitive and suitable for assessing the variation of heterotrophic microbial activity (respiration) in different agricultural soils (Alef and Kleiner
1986). Our initial laboratory tests thus confirmed that the assay is simple and easy, and sensitively reflects a heterotrophic microbial activity in the soil samples. The kinetic plots of soil arginine ammonification activity in both dry and moist barley soils shown in Fig. 1 thus displayed a simple, one-component Michaelis-Menten relationship. This is in contrast to another assay of amino acid degradation in soil, histidine ammonification, which was also proposed to reflect N mineralization; Burton and McGill (1989) thus reported that this activity had two kinetically distinct components. The high-affinity kinetics (Km value approximately 50 µM) further supported that arginine ammonification reflected intracellular, microbial enzyme activity rather than activity associated with the soil matrix. Finally, in the laboratory tests with rewetted soil samples, we demonstrated that arginine ammonification activity correlated strongly with heterotrophic microbial activity (CO2 production). The rapid, but transient increases of both activities occurring approximately 1–2 days after rewetting also gave the first evidence that even short-term fluctuations of heterotrophic microbial activity could be detected by the assay. The biochemical pathways of arginine ammonification include both the arginine deamination (or deimination) reaction to citrulline and ornithine known in several soil bacteria, e.g. Pseudomonas spp. (Krieg and Holt 1984), and the arginine hydrolysis (arginase) reaction to ornithine and urea, which in turn is completely mineralized by another hydrolysis (urease) reaction. It has recently been demonstrated that urea turnover may in fact be quantitatively important in gross N mineralization in both marine sediments (Pedersen et al. 1993) and agricultural soils (Nielsen et al. 1998). In this case, the degradation (ammonification) activity of the precursor amino acids, e.g. arginine and ornithine, for urea production should indeed be suitable indices of the overall, gross N mineralization in soil. Direct measurements of gross N mineralization are rarely performed because they involve a complicated 15N isotope dilution technique. In the present work, we compared such direct determinations of gross N mineralization by isotope technique (Nielsen et al. 1998) with results from our standard arginine ammonification assay. The strong correlation between annual (1995) averages of the two parameters tested in four different soils (setaside, barley, clover and wheat) gave good evidence that the arginine ammonification assay was indeed a suitable index of gross N mineralization. Further evidence came from the detailed seasonal progress of the two parameters in the barley field. The measurements performed throughout the 1995 season thus showed a steady increase of both arginine ammonification and gross N mineralization during spring and summer; eventually, however, both activities declined in high summer (July– August) when dry weather conditions resulted in soil water contents dropping below approximately 5% (w/w). We concluded from these field studies that arginine ammonification was indeed reflecting gross N mineralization and was quite able to pick up differences in soil type
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(soil under different cropping systems) and both annual and seasonal variations in a selected barley field. As demonstrated already in laboratory tests, the arginine ammonification assay was quite sensitive and responded quickly enough to detect the flush of heterotrophic microbial activity occurring 1–2 days after rewetting of a dry soil. A comparison to similar conditions in the field soil was available from the very dry season in summer 1994, when long periods (1–2 weeks) of drought were interrupted only by three short events of rainfall. The short and transient increases of arginine ammonification activity occurring immediately or with a very short lag (few days) after the rainfalls clearly confirmed the evidence from laboratory tests: hence, the assay was adequately able to also pick up these short-term fluctuations of heterotrophic activity. The short periods of increased soil water may have resulted in rapid stimulation of metabolism in bacteria made inactive by desiccation and/or a rapid increase of substrate availability for their metabolism. In either case, the recorded events may both be quantitatively important in overall N cycling and for maintenance of active bacteria in the dry soil. None of the previously reported assays of microbial biomass or specific enzyme pools would have been sensitive enough to pick up these short-term variations in soil. We concluded from these detailed recordings that the arginine ammonification assay may be useful to assess not only the short-term variations in gross N mineralization expected to occur at fluctuating soil temperatures and water contents. The assay may also be useful for detection of variations occurring locally at hot-spots in the soil, e.g. the rhizosphere of living plant roots and the residuesphere of degrading plant fragments. Acknowledgements We acknowledge Henrik Saaby Johansen for valuable assistance in measuring 15N. The work was supported by the Danish Strategic Environmental Research Programme (1992–1996).
References Alef K, Kleiner D (1986) Arginine ammonification, a simple method to estimate microbial activity potentials in soils. Soil Biol Biochem 18:233–235 Alef K, Kleiner D (1987) Applicability of arginine ammonification as indicator of microbial activity in different soils. Biol Fertil Soils 5:148–151 Blackburn TH (1979) Method for measuring rates of NH4+ turnover in anoxic marine sediments, using a 15N-NH4+ dilution technique. Appl Environ Microbiol 37:760–765
Burns RG (1982) Enzyme activity in soil: location and possible role in microbial ecology. Soil Biol Biochem 14:423–427 Burton DL, McGill WB (1989) Role of enzyme stability in controlling histidine deaminating activity in soil. Soil Biol Biochem 21:903–910 Burton DL, McGill WB (1992) Spatial and temporal fluctuation in biomass, nitrogen mineralizing reactions and mineral nitrogen in a soil cropped to barley. Can J Soil Sci 72:31–42 Højberg O, Binnerup SJ, Sørensen J (1996) Potential rates of ammonium oxidation, nitrite oxidation, nitrate reduction and denitrification in young barley rhizosphere. Soil Biol Biochem 28:47–54 Keeney DR (1980) Prediction of soil nitrogen availability in forest ecosystems: a literature review. For Sci 26:159–171 Krieg NR, Holt JG (eds) (1984) Bergey’s manual of systematic bacteriology, 9th edn, vol 1. Williams & Wilkins, Baltimore London Lin Q, Brookes PC (1999) Arginine ammonification as a method to estimate soil microbial biomass and microbial community structure. Soil Biol Biochem 31:1985–1997 Miller M, Nielsen TH, Bonde T, Sørensen J (2001) Fluctuations of formamide hydrolase activity in a barley soil after drying and rewetting. Appl Soil Ecol 16:159–167 Murphy DV, Sparling GP, Fillery IRP (1998) Stratification of microbial biomass C and N and gross N mineralization with soil depth in two contrasting Western Australian agricultural soils. Aust J Soil Res 36:45–55 Nielsen LP (1992) Denitrification in sediment determined from isotope pairing. FEMS Microbiol Ecol 86:357–362 Nielsen TH, Bonde TB, Sørensen J (1998) Significance of microbial urea turnover in N cycling of three Danish agricultural soils. FEMS Microbiol Ecol 25:147–157 Pedersen H, Lomstein BA, Blackburn TH (1993) Evidence for bacterial urea production in marine sediments. FEMS Microbiol Ecol 12:51–59 Powlson DS (1997) Integrating agricultural nutrient management with environmental objectives – current state and future prospects. The Fertiliser Society Proceedings No. 402. The Fertiliser Society, York, UK Powlson DS, Jenkinson DS (1976) The effect of biocidal treatments on metabolism in soil. II. Gamma irradiation, autoclaving, air drying and fumigation. Soil Biol Biochem 8:179–188 Risgaard-Petersen N, Rysgaard S, Revsbech NP (1995) Combined microdiffusion-hybrobromite oxidation method for determining nitrogen-15 isotope in ammonium. Soil Sci Soc Am J 59: 1077–1080 Robinson JA (1985) Determining microbial kinetic parameters using nonlinear regression analysis. Advantages and limitations in microbial ecology. Adv Microb Ecol 8:61–144 Stanford G (1982) Assessment of soil nitrogen availability. In: Stevenson FJ (Ed) Nitrogen in Agricultural Soils. (Agronomy monographs, no. 22) American Society of Agromomy, Madison, Wis., pp 651–688 Stevenson FJ (1982) Organic forms of nitrogen. In: Stevenson FJ (Ed) Nitrogen in Agricultural Soils. (Agronomy monographs, no. 22) American Society of Agronomy, Madison, Wis. pp 67– 122 Werdouw H, Van Echteld CJA, Dekkers EMJ (1977) Ammonia determination based on indophenol formation with sodium salicylate. Water Res 12:399–402
