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Appl Environ Microbiol. 2004 November; 70(11): 6363–6369.
PMCID: PMC525158

Dynamics of a Pasture Soil Microbial Community after Deposition of Cattle Urine Amended with [13C]Urea

Abstract

Within grazed pastures, urine patches are hot spots of nitrogen turnover, since dietary N surpluses are excreted mainly as urea in the urine. This short-term experiment investigated 13C uptake in microbial lipids after simulated deposition of cattle urine at 10.0 and 17.1 g of urea C m−2. Confined field plots without or with cattle urine amendment were sampled after 4 and 14 days, and soil from 0- to 5-cm and 10- to 20-cm depths was analyzed for content and composition of phospholipid fatty acids (PLFAs) and for the distribution of urea-derived 13C among individual PLFAs. Carbon dioxide emissions were quantified, and the contributions derived from urea were assessed. Initial changes in PLFA composition were greater at the lower level of urea, as revealed by a principal-component analysis. At the higher urea level, osmotic stress was indicated by the dynamics of cyclopropane fatty acids and branched-chain fatty acids. Incorporation of 13C from [13C]urea was low but significant, and the largest amounts of urea-derived C were found in common fatty acids (i.e., 16:0, 16:1ω7c, and 18:1ω7) that would be consistent with growth of typical NH4+-oxidizing (Nitrosomonas) and NO2-oxidizing (Nitrobacter) bacteria. Surprisingly, a 20‰ depletion of 13C in the cyclopropane fatty acid cy17:0 was observed after 4 days, which was replaced by a 10 to 20‰ depletion of that in cy19:0 after 14 days. Possible reasons for this pattern are discussed. Autotrophic nitrifiers could not be implicated in urea hydrolysis to any large extent, but PLFA dynamics and the incorporation of urea-derived 13C in PLFAs indicated a response of nitrifiers which differed between the two urea concentrations.

Urine deposition is an important source of N in grazed pastures. Nitrogen turnover in urine patches is intense, and the potential for environmental losses is significant, depending on urine composition (31, 49). Excess N in the diet of cattle is mainly excreted as urea in the urine (23), and understanding the regulation of urea turnover in urine patches is therefore of particular interest. Urea typically disappears from the soil solution within 24 to 48 h and is replaced by high concentrations of NH4+ (18, 36). The accompanying increase in osmotic pressure and free NH3 concentrations can lead to N2O emissions and other microbial stress reactions (12, 40).

Urea hydrolysis to NH4+ and CO2 may occur both on plant surfaces and in the soil (20). Hydrolysis takes place through the action of free or colloid-bound extracellular ureases or intracellularly following microbial uptake (20, 39). Klose and Tabatabai (26) determined extracellular and intracellular urease activities to be on average 54 and 46% of the total potential activity in various cropping systems. Nielsen et al. (33) found that urea turnover was comparable in magnitude to gross N mineralization and hypothesized that direct uptake and intracellular metabolism of urea are quantitatively important aspects of N cycling in agricultural soil. Different bacteria have shown a capacity for urea uptake, including Klebsiella and Alcaligenes spp. (21) and autotrophic NH4+-oxidizing bacteria (2).

Stable-isotope analysis of microbial lipids has recently been introduced in community-level studies. Even though isotope fractionation during lipid biosynthesis is a matter of concern (5), this approach can potentially link specific substrates with individual populations in situ (6, 16, 22). Substrate-derived 13C has been traced in membrane lipids of heterotrophic (1, 7) and autotrophic (27) organisms. Further, 13C enrichment of phospholipid fatty acid (PLFA) profiles has been used to investigate microbial community dynamics (9).

Here we report on the turnover of [13C]urea in pasture soil and the associated incorporation of 13C in PLFAs after simulated deposition of cattle urine at two different urea concentrations. It was hypothesized that the dynamics and labeling of PLFAs could provide information about the fate of urea and the response of nitrifiers to the sudden increase in N availability.

MATERIALS AND METHODS

Site information.

The field experiment was carried out between 20 September and 4 October 2001. The site was an 8-year-old grass-clover pasture near the Danish Institute of Agricultural Sciences, Tjele, Denmark (55°52′N, 9°34′E). The soil was a sandy loam and was classified as a Typic Hapludult. It contained 2.7% C and 0.18% N, the equation M1was 5.5, and total cation exchange capacity was 8.7 cmol kg−1. Air and soil temperatures (10-cm depth) averaged 12.8 and 13°C, respectively, during the experiment, and total precipitation was 56 mm (27 mm within the first 48 h).

Experimental design.

The experimental design contained three randomized blocks, each with three treatments, i.e., a control with no amendments (CTL), standard cattle urine (LU), and urine with approximately double the urea concentration (HU). Urine was collected from dairy cows during milking 3 days before start of the experiment and was stored at 2°C until used; the urea content was determined prior to storage. On day 0 of the experiment, the cattle urine was amended with [13C]urea (99 atom%; CK Gas Products, Hook, United Kingdom) and unlabeled urea (HU treatment only), to give concentrations of 12.0 and 20.4 g of urea liter−1 with [13C]urea concentrations of 11.0 atom% (LU) and 18.3 atom% (HU). The labeled urine was applied to 25- by 35-cm frames, installed to a 15-cm depth, at rates corresponding to 10.0 and 17.1 g of urea C m−2 in for the LU and HU treatments, respectively.

Sampling.

Emissions of CO2 were determined after 0.2, 1, 2, 4, 6, and 14 days. Insulated chambers, equipped as previously described (35), were mounted on top of the permanent frames. Gas samples (13 ml) were taken in preevacuated Exetainers (Labco Ltd., High Wycombe, United Kingdom) after 5, 15, and 45 min and subsequently analyzed for total concentrations and isotopic composition of CO2.

Soil sampling took place after 4 and 14 days. Three soil cores (0 to 20 cm in depth; 2 cm in diameter) were collected and the depth intervals 0 to 5, 5 to 10, and 10 to 20 cm were pooled in separate bags. The samples were stored at 2°C until sieved (<4 mm) and were processed within 2 days. Subsamples were extracted in 1 M KCl and analyzed for urea, NH4+, and NO2 plus NO3 (25, 32). From two of the three blocks, 3- to 3.5-g subsamples from 0- to 5-cm and 10- to 20-cm depths (selected samples) were prepared for PLFA analysis by using a modified Bligh-Dyer single-phase extraction, solid-phase extraction on 100-mg SPE columns (Varian, Harbor City, Calif.), and mild alkaline transesterification as previously described (37).

Soil was dried at 105°C overnight for determination of gravimetric soil moisture. Dried subsamples were analyzed for total C and 13C. The soil bulk density of each depth interval was determined by the end of the experiment.

GC-IRMS analyses.

The concentrations and isotopic composition of CO2 were analyzed with a Europa (Crewe, United Kingdom) Scientific Tracermass isotope ratio mass spectrometer (IRMS) coupled to an automated gas analysis system. The total C content and isotopic composition of soil samples were determined by using an automated combustion elemental analyzer interfaced with a Europa ANCA-SL IRMS system. 13C-labeled PLFA methyl esters were analyzed on a Finnigan Delta Plus XL gas chromatograph (GC)-combustion IRMS (ThermoQuest, Pegnitz, Germany). The gas chromatograph (Hewlett-Packard 6890) was equipped with an HP-5MS column (60 m by 0.25 mm [inner diameter]) and a GC/C III combustion interface. Helium was used as carrier gas.

Fatty acids were tentatively identified from retention times and cross-referencing with samples analyzed by GC-MS. The δ13C values determined by GC-combustion IRMS, based on authentic standards certified relative to PeeDee Belemnite, were corrected for the isotope ratio of the methyl moiety of fatty acid methyl esters (1), as follows: δ13CFA = [(Cn + 1) × δ13CFAME − δ13CMeOH]/Cn, where δ13CFA is the δ13C of the fatty acid, Cn is the number of C atoms in the fatty acid, and δ13CFAME is the δ13C of the fatty acid methyl ester. The fractions of 13C in each fatty acid and the amounts of 13C incorporated were calculated as outlined by Boschker and Middelburg (5).

Statistical analyses.

Concentrations of urea, NH4+, and NO2 plus NO3, respectively, were compared across treatment, sampling day and depth by a linear mixed model, as follows: Y = μ + αS + βT + (αβ)ST + γD + (αγ)SD + (βγ)TD + (αβγ)STD + εB + ETB + ESTB + E′′DTB + E′′′STDB, where Greek letters represent treatment effects, S is sampling day, T is urine treatment, D is depth interval, B is block, and E terms represent random errors. Nitrogen concentrations were log transformed to reduce heteroscedacity.

The PLFA distributions were analyzed by a principal component analysis (PCA) after log(n + 1) transformation of moles percentages and with the covariance matrix. The concentrations of individual fatty acids at the 0- to 5-cm depth were also compared across treatments and sampling days by individual analyses of variance (ANOVAs), using the stepwise Bonferroni procedure to control the overall table-wise error rate (41). Statistical analyses were carried out with SAS 8.2 (SAS Institute, Cary, N.C.).

RESULTS

CO2 emissions and 13C recovery.

The cumulated emissions of CO2 from soil and vegetation during the 14-day period (average ± standard error) amounted to 19.8 ± 2.9, 25.8 ± 6.0, and 22.6 ± 1.8 g of C m−2 in the CTL, LU, and HU treatments, respectively, and were not statistically different between treatments. In both the LU and HU treatments, the proportion derived from urea constituted around 35% of the total flux at the first sampling after 3 h, a proportion which decreased to <1% by day 4 and to 0.1% by day 14. The cumulated recovery of 13C in CO2 by day 14 was 15 ± 9.2% in the LU treatment and 7.7 ± 2.0% in the HU treatment. The recovery of 13C in the soil was 37 ± 2.6% in the LU treatment and 16 ± 1.0% in the HU treatment. Finally, the recovery of 13C in PLFAs was much less than 1% in both treatments (see below).

Soil nitrogen dynamics.

Background concentrations of extractable N in the pasture soil were very low, and N introduced via simulated urine deposition was therefore readily detected (Table (Table1).1). There were strong vertical gradients, with more than half of the extractable N in the top 5 cm. By day 4, urea concentrations were low, yet they were elevated at the 0- to 5-cm depth in the HU treatment relative to the other treatments. The NH4+ pool remaining in the soil by day 14 was significantly higher in the HU treatment than in the LU treatment at all depths. Accumulation of NO2 plus NO3 occurred at similar rates in the LU and HU treatments and by day 14 corresponded to 23 and 17%, respectively, of the urea N added.

TABLE 1.
Concentrations of extractable nitrogen at three soil depth intervals 4 and 14 days after simulated urine depositiona

PLFA content and composition.

Only the 13 phospholipid fatty acids shown in Table Table22 were consistently detected in all treatments and on both sampling days. Table Table22 presents the moles percent distributions and total yields at the 0- to 5-cm soil depth. Urine deposition resulted in PLFA concentrations increasing from 26.6 to 36 to 39 nmol g−1, an increase that was maintained in the HU treatment, but not in the LU treatment, by day 14. The need to control type 1 errors made the statistical tests of individual PLFAs relatively conservative. Accordingly, only a few significant effects were identified (Table (Table2),2), the most notable being the reduction of cy17:0/16:1ω7 ratios after 4 days. There was a net production of cyclopropane fatty acids (CFAs) in the HU treatment by day 4.

TABLE 2.
Distribution of PLFAs, total yields of PLFAs, and cyclopropane fatty acid-to-precursor ratios at the 0- to 5-cm soil depth in the CTL, LU, and HU treatments

A PCA based on all treatments and both depths (selected samples from 10- to 20-cm depth only) separated the two depth intervals along the first principal component (Fig. (Fig.1A).1A). The samples from the 10- to 20-cm depth were enriched in cy17:0 and cy19:0, in 10Me16:0, and in 18:0 (Fig. (Fig.1B).1B). At the 0- to 5-cm depth, the PLFA profiles of the CTL and HU treatments were similar after 4 days. In contrast, the PLFA composition of the LU treatment had changed dramatically, but it shifted towards the composition of the CTL treatment after 14 days. The PLFA profile of the HU treatment changed in the same direction as that of the LU treatment between day 4 and day 14.

FIG. 1.
A. Score plot of the first two principal components of a PCA based on moles percent distributions of PLFAs in pasture soil from the 0- to 5-cm or 10- to 20-cm depth (selected samples only). The designations indicate treatment (CTL, LU, or HU) and sampling ...

δ13C signature of PLFAs.

The recovery of urea-derived 13C in PLFAs was small, resulting in δ13C changes of not more than 20‰ (Fig. (Fig.2,2, top panels). Incorporation of 13C was observed in branched-chain fatty acids (i15:0, a15:0, and i16:0), in C16 straight-chain fatty acids (mainly 16:0 and 16:1ω7), and in C18 straight-chain fatty acids (18:0, 18:1ω9, and 18:1ω7). Interestingly, a decrease in δ13C was observed with both cy17:0 (LU treatment by day 4) and cy19:0 (LU and HU treatments by day 14). Even though the changes in concentrations of CFAs were moderate, the δ13C depletion thus revealed that these CFAs were characterized by significant turnover. At the 0- to 5-cm soil depth, the δ13Cs of CFAs in the LU and HU treatments were on average 13 to 15‰ lower than the average δ13C for all other PLFAs by day 4 and were 7 to 12‰ lower by day 14. Cyclopropane fatty acids in the CTL treatment were also depleted of 13C, with a δ13C of 2 to 5‰ below the average for all other PLFAs (data not shown).

FIG. 2.
δ13C values (top panel) and incorporation of urea-derived C (bottom panel) in PLFAs extracted from pasture soil 4 and 14 days after simulated urine deposition (mean ± standard error; n = 2).

Recoveries of urea-derived C in PLFAs after 4 days were comparable in the HU and LU treatments, at 0.001 and 0.0009%, respectively. The concentrations of urea C incorporated in individual PLFAs are presented in Fig. Fig.2,2, bottom panels. These data show a selectivity of 13C incorporation between and within treatments; six fatty acids (i15:0, a15:0, 16:1ω7, 16:0, 18:1ω9, and 18:1ω7) accounted for 85 to 95% of the total 13C incorporation. These fatty acids also accounted for most of the PLFA concentration changes in urine-amended soil relative to the CTL treatment. The LU and HU treatments differed with respect to the incorporation of urea C into the branched-chain fatty acids i15:0 and a15:0, resulting in low i15:0/a15:0 ratios in the HU treatment by day 4 (Table (Table1).1). The absence of excess 13C in the common fatty acid 16:0 in the LU treatment after 4 days is notable, but it cannot be explained at present.

There was a significant (P < 0.01 or better) positive relationship between total concentration changes and the incorporation of 13C in individual PLFAs. In Fig. Fig.3,3, the average fractions of PLFA carbon derived from urea in the LU and HU treatments after 4 and 14 days are plotted against the fractions of CO2 derived from urea. On day 14, the incorporations of urea C in CO2 and PLFAs were relatively similar in the LU and HU treatments (close to the 1:1 line). In contrast, on day 4, the labeling of excess PLFA was greater than the labeling of CO2 emitted, especially in the HU treatment.

FIG. 3.
The fractions of CO2 derived from urea on day 4 and day 14 were plotted against the fractions of urea C in excess PLFA, i.e., PLFA concentration changes in the LU and HU treatments relative to the CTL treatment (mean ± standard error; n = ...

DISCUSSION

Urine patches in grazed pastures constitute a harsh environment that is potentially stressful for soil organisms. Scorching of the vegetation is known to occur, depending on deposition rate and urea concentration (42), and it is a result of osmotic stress, NH3 toxicity, or a combination of both (18, 31, 42). Plants and microbes respond similarly to hyperosmotic conditions (11), and adverse effects on microbial populations can therefore also occur. Polonenko et al. (40) exposed soil columns to osmotic potentials of −0.5 MPa and below and found a significant decrease in the number of viable cells leached from the salt-stressed columns but a significant increase in viable cell numbers following a period of stress relief. In the present study, extractable NH4+ alone corresponded to osmotic potentials of down to −0.19 and −0.33 MPa in the LU and HU treatments, respectively.

The urine patch environment is expected to stimulate nitrification, but high urinary N concentrations may also inhibit nitrifying bacteria (12, 31, 38, 47). Nitrite oxidizers such as Nitrobacter are far more sensitive to adverse environmental conditions than Nitrosomonas, in particular with respect to concentrations of free NH3 (46). Inhibitory NH3 levels of 0.1 to 1 and 10 to 150 mg liter−1 have been reported for Nitrobacter and Nitrosomonas, respectively (3). For the HU and LU treatments, concentrations of free NH3 were estimated from NH4+ concentrations and pH to be 80 to 120 and 40 to 65 mg N liter−1, respectively, suggesting that a selective inhibition of NO2 oxidation was likely to occur, especially in the HU treatment. Monaghan and Barraclough (31) found that maximum NO2 concentrations in urine-amended soil increased progressively from 5 to 160 mg of N kg−1 soil as urine N increased from 3.8 to 25 g of N liter−1. Nitrite accumulation has also been observed at a urea level corresponding to that in the HU treatment in the pasture soil used in the present study (38).

A response to urine deposition was detectable in the PLFA profiles of the soil microbial community. Significant growth was indicated by the increase in PLFA yields with both the LU and HU treatments relative to the unamended CTL treatment after 4 days (Table (Table2).2). This was in accordance with results from the related laboratory study with comparable urea amendments to pasture soil, where a twofold increase in potential NH4+ oxidation activity also was observed 3 days after urea amendment (38). In the LU treatment, vigorous growth was further indicated by low CFA-to-precursor ratios (15). In the HU treatment, however, such a reduction in CFA-to-precursor ratios was not observed despite the increase in PLFA yield (Table (Table2).2). Instead, a net production of CFAs occurred in the HU treatment, which could be interpreted as a stress response, since cyclopropane fatty acids have been shown to appear in microbial cell membranes in connection with various stresses, including hyperosmotic conditions (14, 29). There is evidence that lipid extractability or partitioning during sample preparation increases with ionic strength (13), but the importance of this factor is not known.

The LU and HU treatments also differed in the proportions of the branched-chain fatty acids i15:0 and a15:0 after 4 days (Fig. (Fig.2,2, bottom left panel). A similar shift towards production of anteiso fatty acids has been observed in response to salt stress with Listeria monocygotenes and several halotolerant bacteria (reference 10 and references therein); the resulting increase in membrane permeability was explained as a mechanism to facilitate adaptation to hyperosmotic conditions (10). Reinspection of PLFA results from the laboratory study referred to above (38) showed a consistent reduction of i15:0/a15:0 ratios after urea amendment corresponding to the HU treatment but not after urea amendment corresponding to the LU treatment (data available upon request).

As indicated in Fig. Fig.3,3, the incorporation of urea C in PLFA after 4 days was higher than the proportion of urea C in CO2, indicating that part of the CO2 originated from unlabeled substrates in the soil. Stress-induced microbial turnover or degradation of soil organic matter dissolved by the urine could have caused this (24, 38). However, cattle urine contains organic components besides urea, such as hippuric acid (8), and dilution of 13CO2 by degradation of these components could also have occurred.

The observed depletion of 13C in CFAs (Fig. (Fig.2A)2A) was unexpected but testifies to the turnover of these pools during the experiment. Boschker et al. (6) found cy17:0 to be slightly depleted of 13C after [13C]methane amendment to an intertidal sediment, but 13C depletion of CFAs on the order of 10 to 20‰ has not previously been reported for complex microbial communities. Bacterial CFAs are produced from the monoenes 16:1ω7c and 18:1ω7c exclusively by trans-methylation from S-adenosylmethionine (14). Isotope depletion of the active methyl group in S-adenosylmethionine by up to 39‰ has been observed in natural compounds other than fatty acids (19, 49), suggesting that the potential for 13C depletion in CFAs is high.

The source of 13C introduced in the experimental system was a C1 compound, urea. Whereas the isotopic composition of heterotrophs is mostly close to that of the growth substrates (1, 30, 48), the pathways of C assimilation during autotrophic growth can result in 13C depletion of cell material by up to 27‰ (34). Further, lipids are generally depleted in 13C relative to the total biomass (17, 34, 45). Experiments with sulfate-reducing bacteria grown on acetate led to a 13C depletion of 12 and 13‰ in the fatty acid 10Me16:0, whereas growth on CO2 resulted in depletions of 24 and 18‰ (30). Also, as mentioned above, Boschker et al. (6) found a depletion of 13C in cy17:0 isolated from a sediment microbial community after [13C]methane amendment, but this was not the case after [13C]acetate amendment. Finally, discrimination against 13C during algal lipid biosynthesis was shown to increase with CO2 availability (43), suggesting that a high soil CO2 availability in urine patches could also increase isotope fractionation during autotrophic growth.

The actual contribution of nitrifiers to 13C incorporation in PLFAs is not known. The greatest amounts of 13C were found in the common fatty acids 16:0, 16:1ω7c, and 18:1ω7c, which predominate in Nitrosomonas and Nitrobacter cell membranes (4, 27, 28) but are also present in many other organisms. Fixation of CO2 by heterotrophic bacteria may have accounted for some incorporation of 13C into PLFAs, although a more uniform distribution of label within a range of microbial PLFAs would have been expected if this had been the main mechanism for urea C incorporation (44). Both cy17:0 and cy19:0 can be synthesized by autotrophic NO2 oxidizers, including Nitrobacter winogradski (27, 28). If Nitrobacter was the main source of de novo CFA synthesis, then this might explain why 13C depletion was so extreme in these compounds. Future work should address the response of Nitrobacter to urine deposition in more detail.

In summary, carbon and nitrogen transformations are intense in pasture soil affected by urine. The response of the pasture soil microbial community was complex. The HU treatment was characterized by osmotic pressures and free NH3 concentrations which probably caused some stress-induced metabolism and a partial inhibition of nitrification activity during the first few days after deposition (12, 38, 40, 47). The low recovery of urea-derived C in PLFAs did not suggest intracellular urea hydrolysis as a major mechanism for turnover of urinary urea, but still some information about the microbial response to urine deposition was obtained that was not revealed by overall PLFA dynamics. A stress response of nitrifiers to urine deposition was indicated, which differed between the two levels of urinary urea applied.

Acknowledgments

This study was supported by the EU Framework Programme 5 project MIDAIR (EVK2 CT-2000-00096) and by the Danish Research Centre for Organic Farming.

REFERENCES

1. Abraham, W.-R., C. Hesse, and O. Pelz. 1998. Ratios of carbon isotopes in microbial lipids as an indicator of substrate usage. Appl. Environ. Microbiol. 64:4202-4209. [PMC free article] [PubMed]
2. Allison, S. M., and J. I. Prosser. 1991. Urease activity in neutrophilic autotrophic ammonia-oxidizing bacteria isolated from acid soils. Soil Biol. Biochem. 23:45-51.
3. Anthoniesen, A. C., R. C., Loehr, T. B. S. Prakasam, and E. G. Srinath. 1976. Inhibition of nitrification by ammonia and nitrous acid. J. Water Pollut. Contol F 48:835-852. [PubMed]
4. Blumer, M., T. Chase, and S. W. Watson. 1969. Fatty acids in the lipids of marine and terrestrial nitrifying bacteria. J. Bacteriol. 99:366-370. [PMC free article] [PubMed]
5. Boschker, H. T. S., and J. J. Middelburg. 2002. Stable isotopes and biomarkers in microbial ecology. FEMS Microbiol. Ecol. 40:85-95. [PubMed]
6. Boschker, H. T. S., S. C. Nold, P. Wellsbury, D. Bos, W. de Graaf, R. Pel, R. J. Parkes, and T. E. Cappenberg. 1998. Direct linking of microbial populations to specific biogeochemical processes by 13C-labelling of biomarkers. Nature 392:801-805.
7. Boschker, H. T. S., J. F. C. de Brouwer, and T. E. Cappenberg. 1999. The contribution of macophyte-derived organic matter to microbial biomass in salt-marsh sediments: stable carbon isotope analysis of microbial biomarkers. Limnol. Oceanogr. 44:309-319.
8. Bristow, A. W., D. C. Whitehead, and J. E. Cockburn. 1992. Nitrogenous constituents in the urine of cattle, sheep and goats. J. Sci. Food Agric. 59:387-394.
9. Butler, J. L., M. A. Williams, P. J. Bottomley, and D. D. Myrold. 2003. Microbial community dynamics associated with rhizosphere carbon flow. Appl. Environ. Microbiol. 69:6793-6800. [PMC free article] [PubMed]
10. Chihib, N.-E., M. R. da Silva, G. Delattre, M. Laroche, and M. Federighi. 2003. Different cellular fatty acid pattern behaviours of two strains of Listeria monocytogenes, Scott A and CNL 895807, under different temperature and salinity conditions. FEMS Microbiol. Lett. 218:155-160. [PubMed]
11. Csonka, L. N. 1989. Physiological and genetic responses of bacteria to osmotic stress. Microbiol. Rev. 53:121-147. [PMC free article] [PubMed]
12. Darrah, P. R., P. H. Nye, and R. E. White. 1987. The effect of high solute concentrations on nitrification rates in soil. Plant Soil 97:37-45.
13. Frostegård, Å., A. Tunlid, and E. Bååth. 1991. Microbial biomass measured as total lipid phosphate in soils of different organic content. J. Microbiol. Methods 14:151-163.
14. Grogan, D. W., and J. E. Cronan, Jr. 1997. Cyclopropane ring formation in membrane lipids of bacteria. Microbiol. Mol. Biol. Rev. 61:429-441. [PMC free article] [PubMed]
15. Guckert, J. B., M. A. Hood, and D. C. White. 1986. Phospholipid ester-linked fatty acid profile changes during nutrient deprivation of Vibrio cholerae: increases in the trans/cis ratio and proportions of cyclopropyl fatty acids. Appl. Environ. Microbiol. 52:794-801. [PMC free article] [PubMed]
16. Hanson, J. R., J. L. Macalady, D. Harris, and K. M. Scow. 1999. Linking toluene degradation with specific microbial populations in soil. Appl. Environ. Microbiol. 65:5403-5408. [PMC free article] [PubMed]
17. Hayes, J. M. 2001. Fractionation of carbon and hydrogen isotopes in biosynthetic processes. Rev. Min. Geochem. 43:225-278.
18. Haynes, R. J., and P. H. Williams. 1992. Changes in soil solution composition and pH in urine-affected areas of pasture. J. Soil Sci. 43:323-334.
19. Hegazi, M. F., R. T. Borchardt, and R. L. Schowen. 1979. α-Deuterium and carbon-13 isotope effects for methyl transfer catalyzed by catechol-O-methyltransferase. SN2-like transition state. J. Am. Chem. Soc. 101:4359-4365.
20. Hoult, E. H., and J. W. McGarity. 1986. The measurement and distribution of urease activity in a pasture soil. Plant Soil 93:359-366.
21. Jahns, T., A. Zobel, D. Kleiner, and H. Kaltwasser. 1988. Evidence for carrier-mediated, energy-dependent uptake of urea in some bacteria. Arch. Microbiol. 149:377-383.
22. Johnsen, A. R., A. Winding, U. Karlson, and P. Roslev. 2002. Linking of microorganisms to phenanthrene metabolism in soil by analysis of 13C-labelled cell-lipids. Appl. Environ. Microbiol. 68:6106-6113. [PMC free article] [PubMed]
23. Kebreab, E., J. France, D. E. Beever, and A. R. Castillo. 2001. Nitrogen pollution by dairy cows and its mitigation by dietary manipulation. Nutr. Cycl. Agrecosys. 60:275-285.
24. Keeney, D. R., and A. N. MacGregor. 1978. Short-term cycling of 15N-urea in a ryegrass-white clover pasture. N. Zealand J. Agric. Res. 21:443-448.
25. Keeney, D. R., and D. R. Nelson. 1982. Nitrogen—inorganic forms. Agron. Monogr. 9:643-693.
26. Klose, S., and M. A. Tabatabai. 2000. Urease activity of microbial biomass in soils as affected by cropping system. Biol. Fertil. Soils 31:191-199.
27. Knief, C., K. Altendorf, and A. Lipski. 2003. Linking autotrophic activity in environmental samples with specific bacterial taxa by detection of 13C-labelled fatty acids. Environ. Microbiol. 5:1155-1167. [PubMed]
28. Lipski, A., E. Spieck, A. Makolla, and K. Altendorf. 2001. Fatty acid profiles of nitrite-oxidizing bacteria reflect their phylogenetic heterogeneity. Syst. Appl. Microbiol. 24:377-384. [PubMed]
29. Loffeld, B., and H. Keweloh. 1996. cis/trans isomerization of unsaturated fatty acids as possible control mechanism of membrane fluidity in Pseudomonas putida P8. Lipids 31:811-815. [PubMed]
30. Londry, K. L., L. L. Jahnke, and D. J. Des Marais. 2004. Stable isotope ratios of lipid biomarkers of sulfate-reducing bacteria. Appl. Environ. Microbiol. 70:745-751. [PMC free article] [PubMed]
31. Monaghan, R. M., and D. Barraclough. 1992. Some chemical and physical factors affecting the rate and dynamics of nitrification in urine-affected soil. Plant Soil 143:11-18.
32. Mulvaney, R. L., and J. M. Bremner. 1979. A modified diacetyl monoxime method for colorimetric determination of urea in soil extracts. Commun. Soil Sci. Plant Anal. 10:1163-1170.
33. Nielsen, T. H., T. A. Bonde, and J. Sørensen. 1998. Significance of microbial urea turnover in N cycling of three Danish agricultural soils. FEMS Microbiol. Ecol. 25:147-157.
34. Pancost, R. D., and J. S. Sinninghe Damsté. 2003. Carbon isotope compositions of prokaryotic lipids as tracers of carbon cycling in diverse settings. Chem. Geol. 195:29-58.
35. Petersen, S. O. 1999. Nitrous oxide emissions from manure and inorganic fertilizers applied to spring barley. J. Environ. Qual. 28:1610-1618.
36. Petersen, S. O., S. G. Sommer, O. Aaes, and K. Søegaard. 1998. Ammonia losses from urine and dung of grazing cattle: effect of N intake. Atmos. Environ. 32:295-300.
37. Petersen, S. O., P. S. Frohne, and A. C. Kennedy. 2002. Dynamics of a soil microbial community under spring wheat. Soil Sci. Soc. Am. J. 66:826-833.
38. Petersen, S. O., S. Stamatiadis, and C. Christofides. Short-term nitrous oxide emissions from pasture soil as influenced by urea level and soil nitrate. Plant Soil, in press.
39. Pettit, N. M., A. R. J. Smith, R. B. Freedman, and R. G. Burns. 1976. Soil urease: activity, stability and kinetic properties. Soil Biol. Biochem. 8:479-484.
40. Polonenko, D. R., C. I. Mayfield, and E. B. Dumbroff. 1986. Microbial responses to salt-induced osmotic stress. Plant Soil 92:417-425.
41. Rice, W. R. 1989. Analyzing tables of statistical tests. Evolution 43:223-225.
42. Richards, I. R., and K. M. Wolton. 1975. A note on urine scorch caused by grazing animals. J. Br. Grassland Soc. 30:187-188.
43. Riebesell, U., A. T. Revill, D. G. Holdsworth, and J. K. Volkman. 2000. The effects of varying CO2 concentration on lipid composition and carbon isotope fractionation in Emiliana huxleyi. Geochim. Cosmochim. Acta 64:4179-4192.
44. Roslev, P., M. B. Larsen. D. Jørgensen, and M. Hesselsøe.Submitted for publication.
45. Schouten, S., M. Strous, M. M. M. Kuypers, W. I. C. Rijpstra, M. Baas, C. J. Schubert, M. S. M. Jetten, and J. S. Sinninghe Damsté. 2004. Stable carbon isotopic fractionations associated with inorganic carbon fixation by anaerobic ammonium-oxidizing bacteria. Appl. Environ. Microbiol. 70:3785-3788. [PMC free article] [PubMed]
46. Smith, R. V., L. C. Burns, R. M. Doyle, S. D. Lennox, B. H. L. Kelso, R. H. Foy, and R. J. Stevens. 1997. Free ammonia inhibition of nitrification in river sediments leading to nitrite accumulation. J. Environ. Qual. 26:1049-1055.
47. Stark, J. M., and M. K. Firestone. 1995. Mechanisms for soil moisture effects on activity of nitrifying bacteria. Appl. Environ. Microbiol. 61:218-221. [PMC free article] [PubMed]
48. Teece, M. A., M. L. Fogel, M. E. Dollhopf, and K. H. Nealson. 1999. Isotope fractionation associated with biosynthesis of fatty acids by a marine bacterium under oxic and anoxic conditions. Org. Geochem. 30:571-1579.
49. Weilacher, T., G. Gleixner, and H.-L. Schmidt. 1996. Carbon isotope pattern in purine alkaloids. A key to isotope discriminations in C1 compounds. Phytochemistry 41:1073-1077.

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