FEMS Microbiology Ecology 25 (1998) 147^157
Signi¢cance of microbial urea turnover in N cycling of three
Danish agricultural soils
Tommy Harder Nielsen *, Torben Andreas Bonde, Jan SÖrensen
Section of Genetics and Microbiology, Department of Ecology and Molecular Biology, The Royal Veterinary and Agricultural University,
Thorvaldsensvej 40, DK-1871 Frederiksberg C (Copenhagen), Denmark
Received 29 July 1997; revised 17 October 1997; accepted 27 October 1997
Abstract
The importance of microbial urea turnover in N cycling was investigated in three agricultural soils by comparison of gross N
14
C-urea tracer
mineralization determined by the 15 N-NH‡
4 dilution technique and urea turnover determined by a new
technique. Average urea turnover rates were 1.5 to 4.2 Wg N g31 d31 indicating that the soil urea pool was turned over every
9 to 30 min. Urea turnover rates were generally lowest in set-aside soil with increasing activities in bulk and rhizosphere soil
from a barley field. Gross N mineralization and urea turnover rates were correlated (r = 0.79, P 6 0.005) and of similar size in
the three soils. The high urea turnover rates indicated that urea-N was immobilized directly in soil microorganisms, rather than
mineralized to the free NH‡
4 pool. Our study suggests that microbial urea turnover, by-passing the conventional mineralizationimmobilization pathway involving a free NH‡
4 pool, has a significant role in N cycling of agricultural soils. z 1998 Federation of European Microbiological Societies. Published by Elsevier Science B.V.
Keywords : Urea turnover; Gross N mineralization; N cycling; Urease activity
1. Introduction
In soil the production of urea, CO(NH2 )2 , may
arise from bacterial degradation of purines [1] and
of the amino acid arginine; the latter reaction may
involve arginase, arginine decarboxylase and arginine oxidase [2]. A functional ornithine cycle has
also been reported in fungi and a few bacteria [3].
Altogether, little is known about the natural urea
production rates in soil, and virtually nothing is
known about the role of the microorganisms.
Hydrolysis of urea with subsequent release of the
* Corresponding author. Tel.: +45 (35) 28 26 27;
Fax: +45 (35) 28 26 06.
free NH‡
4 (urea ammoni¢cation) has been widely
studied in soils [4^6] and its important role in N
cycling after urea fertilization has been documented
[7]. In contrast, unfertilized agricultural ¢elds typically harbor a very low urea pool [8]; yet, urea ammoni¢cation may still be signi¢cant in the soil N
cycling if urea hydrolysis rates, by the matrix-bound
or microbial enzymes, are high. It is well known that
several bacteria possess a potential for urea hydrolysis [9], including a high-a¤nity urea uptake system,
and subsequent degradation by an `intracellular' urease enzyme [10]. When urea is hydrolyzed intracellularly, the NH‡
4 product may become directly immobilized (assimilated) into new organic material. In
this case, the microbial urea turnover in the soil rep-
0168-6496 / 98 / $19.00 ß 1998 Federation of European Microbiological Societies. Published by Elsevier Science B.V.
PII S 0 1 6 8 - 6 4 9 6 ( 9 7 ) 0 0 0 9 1 - 3
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T. Harder Nielsen et al. / FEMS Microbiology Ecology 25 (1998) 147^157
resents a bypass from the conventional mineralization-immobilization pathway involving a turnover of
‡
the free NH‡
4 pool. Alternatively, if the NH4 product
di¡uses out of the cell [11], the urea turnover may be
considered as part of the gross N mineralization, i.e.
the total NH‡
4 production from degradation of organic N in the soil.
Determinations of urea ammoni¢cation in soil
have typically been based on urea additions in large
excess of the natural pool, thus enabling a direct
quanti¢cation of the process rates by urea disappearance or NH‡
4 production [12,13]. It is clear that this
approach merely provides potential rates of urea ammoni¢cation, rather than in situ rates at the natural
urea concentrations. Only recently has a 14 C tracer
method been developed to determine in situ turnover
rates of urea in soil without enrichment of the natural urea pool [8,14]. Using this method in oak forest
soil, the authors showed activities supporting turnover times of 9^15 min for the indigenous soil urea
pool (29 to 56 ng urea-N g31 ). The data showed that
N cycling through the urea pool was 3 times higher
than the net N mineralization measured by accumu3
lation of NH‡
4 and NO3 in the soil [8]. Gross N
mineralization rates are typically higher than those
of net mineralization [15] and investigations in marine sediments have shown that urea turnover may
constitute up to 80% of the gross N mineralization
[16]. The work by Pedersen et al. [8,14] suggests that
the urea turnover rates in soil may have been comparable to those of gross N mineralization.
By convention, gross N mineralization refers to
the total release of free NH‡
4 from organic N degradation. In environmental samples, the process may
therefore be determined by the 15 N dilution technique using a labelling of the NH‡
4 pool. Hence, to
determine if urea turnover was quantitatively signi¢cant relative to gross N mineralization we compared
the in situ urea turnover rate determined by 14 C
tracer technique [8], and the gross N mineralization
determined by 15 N dilution technique [17] in three
Danish agricultural soils. The comparison should
also indicate if in situ urea turnover is merely a
part of gross N mineralization or if urea turnover
should be considered as an internal loop of organic
N transformations in the soils. We ¢nally included a
comparison of the in situ urea turnover rates with
determinations of potential urea ammoni¢cation ac-
tivities, as the latter represents a much adopted
standard assay of urease activity in soils.
2. Materials and methods
2.1. Soil sampling and preparation
All soil samples were collected in the autumns of
1995 or 1996 from a ¢eld site comprising 3.0% clay,
10% silt, 39.5% ¢ne sand and 47.5% course sand (wt/
wt) at the experimental station HÖjbakkegaard
(Taastrup, Denmark). The soil water content was
approximately 12% (wt/dry wt) at all sampling occasions, except one (August 28, 1995) with only approximately 1% water content. Some samples were
collected from the upper 5 cm of soil in a set-aside
plot which had been uncropped for 3 years, with
periodical milling of the soil; these samples are referred to as set-aside bulk soil. Other samples were
collected from the upper 5 cm of bulk soil between
plants in a barley (Hordeum vulgare L.) ¢eld; these
samples are referred to as barley bulk soil. Finally,
soil was collected from the roots of barley stubbles
by gently digging up the roots with the adhering soil.
The stubble was shaken by hand and the soil closely
associated to the barley roots was released and collected; these samples are referred to as barley rhizosphere soil. In all experiments, the soil samples were
sieved through a 2 mm sieve before preincubation in
plastic containers at 20³C for 5 days. All studies of
microbial activities were conducted during the following 1^3 day period. The gravimetric soil water
content was determined after 24 h of incubation at
105³C.
2.2. Gross N mineralization rate
Gross N mineralization was determined by a modi¢cation of the 15 N dilution method described for
sediments by Blackburn [17] and outlined below.
Soils were 15 N-labelled by spraying with a 24 mM
(15 NH4 )2 SO4 solution (98 atom % 15 N; Cambridge
Isotope Laboratories, MA, USA), while the soil was
gently mixed in a polyethylene bag. The water content increased by less than 2% by the 15 NH‡
4 addition
using this procedure. Subsamples of 5 g soil were
weighed into 50 ml polyethylene centrifuge tubes
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with loosely attached screw caps and incubated in a
water bath at 20³C. Three replicate samples were
subsampled 4 or 5 times during the incubation period, adding 24 ml 2 M KCl to each sample and shaking it for 1 h on an orbital shaker (1000 rpm). After
centrifugation at 4500Ug, the supernatant was ¢ltered through KCl-washed GF/C ¢lters and frozen
for later analysis.
The NH‡
4 concentration in the KCl extracts was
analyzed by the method of Verdouw et al. [18] as
modi¢ed by HÖjberg et al. [19]. The 15 N analysis
of the NH‡
4 pool was conducted by the microdi¡usion assay of Risgaard-Petersen et al. [20] and a triple collector mass spectrometer (Tracermass model;
Europa Scienti¢c LTD., Crewe, United Kingdom)
using manual injection of the samples. The 15 N
atom fraction of the NH‡
4 pool was calculated by
the method of Nielsen [21] and the results were
used in the isotope dilution model for NH‡
4 turnover
[17]. The method provides a determination of gross
N mineralization, i.e. total NH‡
4 release (d) and total
consumption of NH‡
4 (i) according to two equations:
P…t† ˆ P0 ‡ …d3i†t;
…1†
d
…d3i†t ‡ P0
ln…R3 n† ˆ ln…R0 3 n†3
Uln
P0
d3i
15
15
…2†
where P0 is the initial 14‡15 N pool and P(t) is the
N pool at time t, 15 n is the natural abundance
15
of N (0.37 atom %) and R0 and R are the relative
abundances of 15 N expressed as 15 N/15 N+14 N at time
0 and time t, respectively. When i and d are constant,
P(t) in Eq. (1) is linear with time. Eq. (2) describes
that ln(R315 n) is linear with ln[((d3i)t+P0 )/P0 ], represented by a slope of 3d/(d3i) and an intercept of
ln(R0 315 n). The gross N mineralization (d) can be
calculated by combination of Eqs. (1) and (2).
14‡15
2.3. Urea pools
The urea concentration in the soils was determined
by a modi¢cation of the method developed by Pedersen et al. [8]. Soil samples (16 g) were weighed
into 50 ml centrifuge tubes and incubated at 20³C,
in parallel to the incubations for 14 C-urea turnover.
149
Three replicate soil samples were extracted 3 times
during the incubation period by adding 5 ml 1 M
KCl containing 0.5% ZnCl2 to inhibit microbial activity as described by Pedersen et al. [8]. After vortexing the soil, each sample was transferred into a
double-chamber centrifugation unit [22] and centrifuged 10 min at 3³C (6800Ug). The ¢ltrate was immediately frozen for subsequent urea analysis by the
method of Pedersen et al. [8] as modi¢ed from Price
and Harrison [23]. Absorbances were measured at
540 nm in a Microplate reader (Bio-Tek Instruments) and blanks were prepared for each sample
by replacing the diacetylmonooxime reagent with
distilled water.
A recovery experiment to test the urea extraction
procedure was conducted. To avoid hydrolysis of
added urea during this experiment, air-dried bulk
soil from the barley ¢eld was autoclaved three times
for 1.5 h each during a 3 day period [28]. Three
samples of the air-dried soil (each 100 g) were rewetted with 10 ml sterile distilled water or urea solution
containing 4.4 or 8.8 Wg urea-N. The samples were
subsequently incubated for 0.5 h at 20³C before extracting the urea as described above.
2.4. Urea turnover rate
Turnover rates of 14 C-urea were determined in
9 ml polypropylene centrifuge tubes with a butyl
stopper in the lid. Samples of 1.5 g of soil were
weighed into each tube, covered with a perforated
safran ¢lm, and incubated at 20³C for approximately
15 h before use. A sample of 5 Wl 14 C-urea tracer
solution (56 mCi/mmol urea) was then added to
the soil and the tube was sealed with a lid. The tubes
were again incubated at 20³C and subsequently
stopped after 2, 4, 8, 10 and 15 min, respectively.
The urea concentration of the added tracer solution
was adjusted to the indigenous urea concentration in
the soil, since enrichment of the natural urea pool
was shown to in£uence the turnover of 14 C-urea [8].
In the standard protocol, the tracer addition increased the soil water content of the soil sample by
less than 0.5%. One exception to this protocol was
made on August 28, 1995, where the low soil water
content required that the 14 C-urea tracer was added
in a dry form as described by Pedersen et al. [14].
This was accomplished by evaporating an aliquot of
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T. Harder Nielsen et al. / FEMS Microbiology Ecology 25 (1998) 147^157
tracer solution to dryness in a 9 ml polypropylene
centrifuge tube before the soil sample was added.
The dry 14 C-urea was transferred to the soil by
gently shaking the tube, before incubation was continued in a new vial. To determine the transfer of
14
C-urea tracer to the soil, 0.3 ml of the NaOH solution was added directly into the scintillation liquid
(see below). All incubations were stopped by injecting 3 ml NaOH with a hypodermic needle through
the butyl rubber stopper and violently shaking the
tube. The NaOH raised the pH to approximately 14
in the soil slurry, thus terminating all microbial activity and converting the produced 14 CO2 gas to 14 Clabelled carbonate ions in solution. The tubes were
then centrifuged at 15 000Ug and the supernatant
frozen for analysis of 14 C radioactivity in the carbonate pool.
The amount of 14 C trapped in the carbonate pool
was determined by a di¡usion technique. A ¢lter
paper (10 cm2 ) was folded into a 1.5 ml centrifuge
tube and 0.4 ml Carbo-sorb E (Packard Instruments,
Groningen, The Netherlands) was added to the ¢lter.
The centrifuge tube was then installed in a string
hanging from the butyl rubber lid of a 100 ml infusion bottle. A 0.5 ml sample was placed in the infusion bottle and acidi¢ed by injecting 1.5 ml 5 N
H2 SO4 with a hypodermic needle through the lid to
release CO2 from the NaOH solution. The released
CO2 was allowed to absorb to the Carbo-sorb overnight before the CO2 trap was transferred to a 20 ml
scintillation vial and ¢lled with scintillation liquid
(Ecoscient A, National Diagnostics, Atlanta, GA).
The 14 C activity was measured in a Beckman LS
1801 scintillation counter with an automatic quench
correction software program. Radioactivity (dpm
g31 soil) was calculated using the soil water content
and the amount of NaOH added. The 14 CO2 background in the 14 C-urea tracer solution was subtracted from all results. The radioactivity of added
tracer was determined by adding 5 Wl 14 C-urea tracer
solution directly into the scintillation liquid.
The 14 C-urea turnover rate was calculated using
the steady-state model described by Lund and Blackburn [24]. This model is valid if the urea pool size
remains constant during the incubation period and
the 14 C-urea pool decreases exponentially with time.
The model determines the turnover rate constant (k)
as the slope of the natural logarithm (ln) of 14 C-urea
content against time, and the turnover rate is determined as k times the urea pool. The 14 C-urea content
(%) left in the soil during the incubation was calculated knowing the amount of added 14 C-urea tracer
and the amount of produced 14 CO2 . As suggested by
Pedersen et al. [8] the turnover rate constant was
calculated from data points where 6 90% of the
added 14 C-urea was hydrolyzed. The standard deviation on the turnover rate was determined using the
standard deviation on the urea pool, whereas no
statistical analysis was performed on the urea turnover rate constant.
2.5. Potential urea ammoni¢cation
Potential urea ammoni¢cation activity was quanti¢ed as NH‡
4 production from added urea. From
each soil 4 replicates of 0.1 g soil samples were
weighed into 1.5 ml Eppendorf tubes and amended
with 0.2 ml non-bu¡ered substrate solution of 5 mM
urea. The soil slurries were shaken and incubated at
20³C for 0.5 h. The incubations were stopped by the
addition of 0.8 ml ice-cold 2 M KCl solution and
shaken for 1 min before centrifugating at 15 000Ug
for 5 min. The supernatant was collected and frozen
for later NH‡
4 analysis. Control samples (3 replicates)
were prepared as described above, but incubation
was stopped immediately after substrate addition.
2.6. Statistics
The statistical analysis of correlations between the
urea turnover rate, gross N mineralization and urea
ammoni¢cation rates was conducted by using a simple correlation analysis [25].
3. Results
3.1. Assay of gross N mineralization
Gross N mineralization rates were determined by
N isotope dilution technique during a 2^4 day soil
15
N
incubation, in which the NH‡
4 concentration and
‡
atom % in the NH4 pool were followed. This is
exempli¢ed in Fig. 1, showing progress curves (A
and B) for the analyzed parameters and a plot (C)
of the slope of 3ln[((d3i)t+P0 )/P0 ] versus ln(%
15
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151
process [27]. However, the consumption activity (i)
was clearly stimulated by the initial 15 NH‡
4 addition
leading to a steady decrease of the soil NH‡
4 pool
during the incubation period. Short incubation periods (24 h) were therefore necessary when analysing
gross N mineralization rates in the barley rhizosphere soil samples, in which NH‡
4 consumption
was rapid (data not shown).
3.2. Assay of urea pools and urea turnover rates
15
Fig. 1. Measurements of NH‡
N atom
4 concentration (A) and
% (B) during the gross N mineralization assay in set-aside bulk
soil sampled on October 23, 1995. Part C shows parameters used
for the calculation of gross N mineralization rate (see text).
15
N30.37). The latter was used for calculation of
gross N mineralization rates (d) [17]. This regression
was chosen since the multiple determination of NH‡
4
concentration and 15 N-atom % during the same incubation period gave us several data points based on
steady mineralization rates rather than a two-point
determination of gross mineralization rates [26,27].
Gross N mineralization rates were not expected to
be a¡ected by the 15 NH‡
4 isotope addition, since
NH‡
4 is itself a product rather than substrate of the
The urea extraction procedure modi¢ed from Pedersen et al. [8] was tested by adding a known
amount of urea to autoclaved soil samples and measuring the recovery of extractable urea. As seen in
Table 1 there was a slight increase in KCl-extractable
urea when the air-dried soil was rewetted, but otherwise the urea content in the extracts was most similar
to the expected values after addition of 44 ng N g31
(21% increase) or 88 ng N g31 (43% increase) urea to
the soil. The 100% extraction e¤ciency indicated
that no binding of the added urea to soil matrix
took place, and an e¤cient extraction of the native
urea pool was therefore likely.
We further tested the extraction e¤ciency for
14
CO2 produced in the soil after a complete turnover
of the added 14 C-urea tracer. Fig. 2A shows the rapid accumulation of 14 CO2 during incubation of a setaside bulk soil sample with 14 C-urea tracer; 90% of
the added 14 C-urea was hydrolyzed to 14 CO2 within
the ¢rst 30 min, indicating a high turnover rate constant. A 85^95% recovery was found in all incubations suggesting that most of the added tracer was
accessible for hydrolysis. Subsequent incubations
were made with incubation periods of only 15 min.
Fig. 2B shows the % of added 14 C-urea, which was
left in the three di¡erent soils during 15 min of incubation. The results gave a linear progress curve on
Table 1
Recovery of urea added to sterilized set-aside bulk soila
Urea pool size
31
Measured (ng N g )
Expected rangeb (ng N g31 )
Dry soil
Dry soil+water
Dry soil+1Uureac
Dry soil+2Uuread
134 þ 3
206 þ 3
255 þ 6
244^256
306 þ 6
288^300
a
Values are means (n = 3) þ standard deviation.
Expected concentration range is calculated as: Urea pool in dry soil+water þ 2Ustd+urea pool added.
c
1Uurea is 44 ng N g31 .
d
2Uurea is 88 ng N g31 .
b
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were reporting a steady-state activity, a series of urea
hydrolysis assays were conducted for di¡erent periods (17^21, 22^26 and 41^45 h) during the long-term
assay for gross N mineralization. The results showed
that there were no signi¢cant changes in average
urea concentrations or turnover rate constants during the 2 day incubation period (Table 2).
3.3. Comparisons of process rates in three agricultural
soils
Fig. 2. Part A shows measurements of % 14 CO2 production originating from 14 C-urea hydrolysis in a set-aside bulk soil sampled
on October 23, 1995. Part B shows a logarithmic plot of % remaining 14 C-urea versus incubation time, used for determining
the turnover rate constant (see text) in the set-aside bulk, barley
bulk and barley rhizosphere soil sampled on October 23, 1995.
the log-scale, which allowed the turnover rate constant to be determined from the slope of a linear
regression. Finally, to determine if the short-term
incubations (15 min) for determining turnover rates
As seen in Table 3, the average water content was
between 9 and 14% in the three soils during the
sampling period in September^December of 1995^
1996. This gave a comparable water content between
these samples, which in turn allowed a direct comparison of their process rates at an incubation temperature of 20³C. The data from the dry soil collected on August 28, 1995 are not included in
Table 3, but reported separately below. During the
sampling period, urea concentrations were constant
at approximately 20 ng g31 in the set-aside bulk soil.
Similar concentrations were found in the barley bulk
soil, except at one occasion when a higher level of 34
ng g31 was reached. Finally, the barley rhizosphere
soil had a concentration level between 25 and 36 ng
g31 . The urea turnover rate constant was highly variable within each soil, although the rate constants
were generally similar in the three soils.
Table 3 also shows the calculated urea turnover
rates and the gross N mineralization rates determined by 15 N isotope dilution technique. During
the autumn, the urea turnover rate in the set-aside
bulk soil varied from 1.5 to 3.3 (mean 2.1) Wg N g31
d31 while rates in the barley bulk and rhizosphere
soils were higher, ranging from 2.1 to 2.2 (mean 2.2)
and 1.6 to 4.2 (mean 2.9) Wg N g31 d31 , respectively.
Table 2
Urea concentrations, urea turnover rate constants and urea turnover rates in the set-aside bulk soil sampled on December 6, 1995 and
incubated over a 46 h period (see text)a
Period of
determination (h)
Water contentb
Urea pool size
(ng N g31 )
Turnover rate
constant (k)
Urea turnover
(Wg N g31 d31 )
17^21
22^26
41^45
12.1 þ 0.2
12.1 þ 0.2
12.1 þ 0.2
19.4 þ 8.5
22.9 þ 8.1
19.6 þ 8.3
0.050
0.048
0.054
1.4 þ 0.6
1.6 þ 0.6
1.5 þ 0.6
a
b
Values are means (n = 3) þ standard deviation.
Water content is indicated as % (wt/dry wt).
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153
set-aside bulk, barley bulk and rhizosphere soils. The
potential urea ammoni¢cation activities were from
8.5 to 52.6 (mean 30.6), 47.7 to 56.4 (mean 52.4)
and 117.6 to 169.2 (mean 138.5) Wg N g31 d31 , respectively.
4. Discussion
4.1. Urea turnover in soil
Fig. 3. Comparison of urea turnover rates and gross N mineralization rates in the set-aside bulk, barley bulk and barley rhizosphere soil sampled during autumn 1995 and 1996, including `noactivity' data points observed in dry soils sampled on August 28,
1995.
By comparison, the set-aside bulk soil showed gross
N mineralization rates varying from 0.7 to 1.5 (mean
1.0), whereas the rates in the barley bulk and rhizosphere soils were again higher, from 1.5 to 2.0 (mean
1.9) Wg N g31 d31 and from 2.6 to 4.8 (mean 3.8) Wg
N g31 d31 , respectively.
Finally, Table 3 shows that the potential rates of
urea ammoni¢cation also measured when comparing
The urea concentrations of 20^35 ng N g31 in the
three agricultural soils were comparable, yet slightly
lower than those of 30^90 ng N g31 reported in a
grassland soil by Pedersen et al. [14]. Quantitative
extraction of the small urea pools from the soil
was important for determination of in situ urea turnover rates. Test experiments with added urea pools
showed extraction e¤ciencies close to 100% (Table
1), which suggested an e¤cient extraction of the
small, native urea pool. The linearity of 14 C-urea
hydrolysis, as expressed on a log-scale (Fig. 2B) during the initial 15 min of incubation, indicated that
the urea existed as a single pool in the soil. Similar
results have been found in 14 C-urea tracer experiments in forest soil [8], whereas two urea pools
Table 3
Soil water contents, urea concentrations, urea turnover rate constants, urea turnover rates, gross N mineralization rates, and potential
urea ammoni¢cation rates in the set-aside bulk, barley bulk and barley rhizosphere soilsa
Soil
Water
contentb
Urea pool size
(ng N g31 )
Turnover rate
constant (k)
Urea turnover rate
(Wg N g31 d31 )
Gross N mineral. rate
(Wg N g31 d31 )
Potential urea ammon. rate
(Wg N g31 d31 )
B
9.4 þ 0.1
C
12.6 þ 0.0
D
12.1 þ 0.2
Barley bulk
21.0 þ 5.4
17.1 þ 7.8
20.7 þ 8.1
0.110
0.061
0.051
3.3 þ 0.9
1.5 þ 0.7
1.5 þ 0.6
1.5 þ 0.2
0.7
0.8
52.6 þ 5.5
8.5
ND
A
12.0 þ 0.0
B
10.4 þ 0.1
C
13.0 þ 0.1
Barley rhizosphere
20.1 þ 3.2
16.0 þ 2.1
33.6 þ 7.5
0.078
0.090
0.044
2.2 þ 0.4
2.1 þ 0.3
2.2 þ 0.5
2.0
1.5 þ 0.2
2.1
53.0 þ 6.0
56.4 þ 5.3
47.7
A
B
C
35.7 þ 3.5
25.0 þ 8.0
31.5 þ 7.6
0.082
0.082
0.035
4.2 þ 0.8
2.9 þ 0.9
1.6 þ 0.4
4.0
4.8 þ 0.6
2.6
169.2 þ 24.5
117.6 þ 14.4
128.6
Set-aside bulk
13.1 þ 0.1
13.0 þ 0.1
13.7 þ 0.1
a
Values are means (n = 3) þ standard deviation.
Water content is indicated as % (wt/dry weight).
ND, not determined.
Sampling times were : A: September 13, 1995; B: October 23, 1995; C: November 6, 1996; D: December 6, 1995. Measurements on
December 6, 1995 are average values of data from Table 2.
b
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with distinctively di¡erent turnover rates seemed to
occur in grassland soil [14].
The constant urea concentrations observed during
each of the standard 14 C-urea incubation periods
were also maintained during longer incubations of
2^4 days, corresponding to the long-term incubation
assay for gross N mineralization (Table 2). The
small, but constant urea pools in the soil implied
that the urea hydrolysis rate was balanced by a similar rate of urea production. Direct measurements of
turnover rate constants (Table 3) by the 14 C assay
suggested a turnover time of 9 to 30 min for the urea
pool. Purine catabolism [1] and amino acid hydrolysis via arginine degradation [2] are likely bacterial
sources of urea in the soil. Indigenous urea production by bacteria containing the complete ornithine
cycle has also been described [3], but so far only a
few bacteria have been demonstrated to harbor this
trait. It may be added, however, that much attention
has recently been paid to urea production by several
groups of bacteria, including denitrifying, sulfate
reducing or fermenting bacteria [29]. Signi¢cant
bacterial production of urea has further been demonstrated in marine sediments [30]. In soil microfungi, the role of urea production is yet unknown but
urea production and metabolism have been demonstrated in fungal species [31,32]. In contrast, protozoa and nematodes have not yet been shown to produce urea in soil (Bryan Gri¤ths, personal
communication).
In the present study, the soil water content was
approximately 12%, which corresponded to a concentration range for urea in the soil water of 10^20
WM. The low urea concentration in the three soils
suggested that the urea hydrolysis was controlled
by microbial activity, since microbial urease enzymes
are reported to have low Km values [33]. Urea hydrolysis may take place intracellularly in microorganisms [4,9], and a urea uptake system has been
observed in both Klebsiella pneumoniae and Alcaligenes eutrophus, with low Km values of 13 and 38
WM, respectively [10]. These values are one order of
magnitude lower than the Km values of 280 and 650
WM, respectively, for the urease enzyme per se [10].
Hence, urea hydrolysis rather than urea uptake may
regulate the overall rate of urea turnover at the very
low concentrations in soils. By comparison, reported
Km values for total urease activity, including both
intracellular and matrix-bound enzymes, in various
soils are as high as 1.3 to 62.5 mM [34].
In the assay of potential urea ammoni¢cation (urease activity), the addition of 5 mM urea resulted in
rates which were approx. 80% of Vmax values in the
three soils; apparent Km values were approx. 2 mM
(data not shown). Hence, the apparent Km value for
total urease activity in the soils was higher than reported values for microbial uptake or hydrolysis of
urea, but at the low end of values reported for whole
soil samples. This suggests that both microbial (intracellular) and matrix-bound (extracellular) components are involved in urease activity of the soils. This
is con¢rmed by the results of Pettit et al. [4], suggesting that the matrix-bound urease activity in soil may
sometimes be up to 60% of the total activity. The
potential urea ammoni¢cation rates (Table 3)
showed an increasing level of activity from the setaside bulk soil to the barley bulk and rhizosphere
soils (in this order), which could re£ect a general
increase of organic content [5] or microbial biomass
and activity [35]. However, the absence of correlation between potential ammoni¢cation rates (urease
activity) and microbial urea turnover rates (P 6 0.2,
Table 3) con¢rmed that the former gave no speci¢c
or detailed information on microbial urea transformations in the soils, possibly because of a variable
matrix-bound component.
4.2. Comparison of urea turnover and gross N
mineralization
Urea turnover rates of 1.5 to 4.2 Wg N g31 d31
observed in the three soils (Table 3) were comparable
to turnover rates of 3.7 Wg N g31 d31 in an oak
forest soil [8] and 0.3 Wg N g31 d31 to 13.0 Wg N
g31 d31 in a grassland soil [14]. Gross N mineralization rates of 0.7 to 4.8 Wg N g31 d31 in the present
study (Table 3) were also similar to results from
grassland soils (1.4 Wg N g31 d31 ) reported by Davidson et al. [27] and from control soil (0.72 Wg N
g31 d31 ), oil-seed rape residues (0.87 to 2.2 Wg N g31
d31 ) and winter wheat residues (0.92 Wg N g31 d31 )
reported by Watkins and Barraclough [36].
Our direct measurements of both urea turnover
rates and gross N mineralization rates by tracer techniques make it possible to compare the two processes
in the soils. In Fig. 3, all measurements of urea turn-
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T. Harder Nielsen et al. / FEMS Microbiology Ecology 25 (1998) 147^157
over and gross N mineralization rates of the present
study are shown, including those of the dry barley
bulk and rhizosphere soils on August 28, 1995,
where none of the processes seemed to be active. A
most important observation was that in situ urea
turnover and gross N mineralization rates were comparable in absolute numbers. To our knowledge,
however, the high signi¢cance of urea turnover in
relation to gross N mineralization has not previously
been demonstrated directly in soils.
The results in Fig. 3 show that the process rates
were well correlated (r = 0.79, P 6 0.005) using data
from all soils, but further analysis of the data indicates that the signi¢cance of urea turnover relative to
gross N mineralization is di¡erent in the three soils.
The rhizosphere soil showed urea hydrolysis activities which were similar to or only 60% of the gross
N mineralization rates. In the latter case, at least
part of the organic N was degraded to NH‡
4 without
involvement of the urea pool. In the barley bulk soil,
the urea turnover was approximately 100% of the
gross N mineralization, which indicated that all N
mineralizations could, in theory, have passed via the
urea pool. Finally, in the set-aside soil, the urea hydrolysis was always 200% of the gross N mineralization, suggesting that urea hydrolysis did not result in
concomitant NH‡
4 release, but was assimilated directly into the microbial biomass.
In the set-aside bulk soil, the recorded di¡erence
between urea turnover and total NH‡
4 release by
gross N mineralization suggested that 0.7^1.8 Wg N
g31 d31 could have been directly assimilated into
microbial cells. Assuming a C/N ratio of 4.2 as derived from data of Bakken [37] in the microbial biomass the N immobilization would require a C assimilation of 2.5^6.4 Wg C g31 d31 corresponding to 0.1^
0.3 Wg C g31 h31 . To test whether this C assimilation
is likely, data from measurements of bacterial
growth in soils using the [3 H]thymidine incorporation technique may be useful. Reported C assimilation rates using this technique are 0.1^ 4 Wg C g31
h31 in bulk sand [38], 0.7^ 1.0 Wg C g31 h31 in
unplanted soil, and 0.6^0.8 Wg C g31 h31 in bulk
soil [39]. These numbers indicate that the above hypothesis of direct immobilization of N released by
urea hydrolysis is indeed a plausible explanation
for the urea turnover exceeding the gross N mineralization in the set-aside bulk soil.
155
4.3. Signi¢cance of urea turnover in soil N cycling
It has previously been stated that organic N compounds in soils may be mineralized via two pathways
or a combination of both of them: (1) In the conventional mineralization-immobilization turnover (MIT)
pathway, organic N compounds are mineralized by
extracellular soil enzymes and released to the free
NH‡
4 pool before immobilization into microorganisms takes place [40]. (2) In the `direct' hypothesis,
organic compounds are taken into the microorganisms, deaminated, and only the surplus N is released
(i.e. mineralized) into the free NH‡
4 pool [40,41]. Assimilation of organic N directly into microorganisms
has been reported in soils incubated with glycine and
leucine [41]. (3) Finally, in a `parallel' hypothesis suggested by Barraclough [43], both of the
two above models are operative simultaneously
in the soils. Findings by Hadas et al. [42] also
suggested both pathways operated concurrently. Finally, glutamic acid, leucine and NH‡
4 were also
shown to be immobilized together in soil bacteria
[44].
When urea turnover is sometimes higher than the
gross N mineralization as seen in the set-aside soil,
we propose that this could be due to an intracellular
urea hydrolysis and immobilization of the NH‡
4,
without a release (i.e. mineralization) into the free
NH‡
4 pool (`direct' hypothesis). Such a reaction
thus by-passes the conventional mineralization-immobilization turnover (MIT model), proceeding via
the free extracellular NH‡
4 pool. Since the urea turnover rates are high, and sometimes higher than the
measured gross N mineralization rates, we further
suggest that the urea cycle in the soils has a profound signi¢cance in soil N cycling.
Acknowledgments
The authors thank Niels O.G. JÖrgensen for his
assistance in 14 CO2 analysis, Henning Pedersen for
critically reading the manuscript and Bryan Gri¤ths
for commenting on urea production in soil protozoa
and nematodes. This work was supported by the
Danish Strategic Environmental Research Programme 1992^1996.
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