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Applied and Environmental Microbiology, September 1998, p. 3264-3269, Vol. 64, No. 9
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Biosensor Determination of the Microscale Distribution of
Nitrate, Nitrate Assimilation, Nitrification, and Denitrification in
a Diatom-Inhabited Freshwater Sediment
Jan
Lorenzen,
Lars Hauer
Larsen,
Thomas
Kjær, and
Niels-Peter
Revsbech*
Department of Microbial Ecology, University
of Aarhus, Ny Munkegade, DK-8000 Aarhus C, Denmark
Received 26 February 1998/Accepted 17 June 1998
 |
ABSTRACT |
High-resolution NO3
profiles in
freshwater sediment covered with benthic diatoms were obtained with a
new microscale NO3 biosensor characterized by
absence of interference from chemical species other than
NO2 and N2O. Analysis of the
microprofiles obtained indicated no nitrification during darkness, high
rates of nitrification and a tight coupling between nitrification and
denitrification during illumination, and substantial rates of
NO3 assimilation during illumination.
Nitrification during darkness could be induced by purging the bulk
water with O2 gas, indicating that the stimulatory effect
on nitrification by illumination was caused by algal production of
O2. NH4+ addition did not stimulate
nitrification during darkness when O2 was restricted to the
upper 1-mm layer, and there was thus a low nitrification potential in
the permanently oxic top 1 mm of the sediment.
| |
INTRODUCTION |
The nitrogen cycle and the bacteria
mediating its various transformations have been studied extensively
since Winogradsky isolated the first nitrifying bacteria in the
late-19th century. Although a tremendous effort has been spent on
quantification of the processes, new findings and developments continue
to give us new and important insights. One such important finding was the recent discovery of extensive nitrogen fixation in the oligotrophic oceans by the filamentous but heterocyst-free cyanobacterium
Trichodesmium sp. (4). An important
methodological development is represented by the isotope pairing method
by Nielsen (17), which compared to previous methods gives us
much better estimates of denitrification in most aquatic environments.
However, there are still problems in the quantification of nitrogen
transformations in a variety of environments. One such environment is
sediments inhabited by benthic microphytes, where assimilation
processes occur simultaneously with nitrification and denitrification.
The influence of microphytobenthos on sediment-water fluxes of
combined, inorganic nitrogen has been investigated by a number of
authors. The presence of benthic microalgae generally reduce the efflux
of NH4+ and NO3 from
the sediment (33), with a minimum efflux during illumination due to algal assimilation (32). Also nitrification and
denitrification have been shown to be affected by light-dark cycles.
Denitrification of NO3 supplied from the
overlying water was reduced during light periods (18, 27),
while denitrification coupled to nitrification in the sediment was
shown to be stimulated during daytime due to a stimulation of
nitrification (27). However, other data indicate lower rates
of potential nitrification in the top 1 cm of sediment inhabited with
microalgae compared to that in sediment without algae (12),
which is to be expected when there is competition for
NH4+. The diurnally integrated rates of
nitrification may thus be lowered by the presence of microalgae even
when oxygen production by the microalgae stimulates nitrification
during illumination compared to dark incubation.
Microsensors for NH4+ and
NO3 based on liquid ion exchangers have been
used to measure microprofiles of NH4+
(7) and NO3 (8, 13) in
sediments, and such sensors could in principle also be used in
photosynthetically active sediments. By modelling based on the measured
microprofiles it would then be possible to calculate vertical profiles
of production and consumption processes for the two chemical species.
The NH4+ consumption rates, however, would not
be very informative for layers with an abundance of microphytes where
assimilation cannot be distinguished from nitrification. The
NO3 microsensors used until now suffer from
bicarbonate interference so that the extremely steep concentration
gradients of bicarbonate in communities of benthic microphytes
(26) result in a substantially inaccurate determinations of
low NO3 concentrations.
In the present study we used a newly developed microscale
NO3 biosensor (15), characterized
by the absence of interference from species other than
NO2 and N2O, to resolve the
vertical and temporal distribution of NO3 in
diatom-covered sediment during light-dark cycles. By using diffusion
reaction models the vertical distribution of
NO3 assimilation, nitrification, and
denitrification can be calculated from the measured steady-state
NO3 profiles.
| |
MATERIALS AND METHODS |
Sediment sampling.
Sediment cores covered with a thin layer
of pennate diatoms were collected by using 44-mm inside diameter
Plexiglas tubes in the creek inlet of Vilhelmsborg Sø, a small pond
situated 15 km south of Aarhus, Denmark. The pond was created by
damming a creek draining agricultural areas, and the water is therefore rich in NO3 (up to 1 mM) during periods of
high water discharge, but the NO3
concentration may also decline to very low levels during dry summer
periods.
After the return to the laboratory, the sediment cores were manipulated
so that the sediment surface became flush with the upper edge of the
Plexiglas cylinder. The cores were incubated in a constant-temperature
(20°C) water bath containing approximately 43 liters of constantly
aerated tap water (about 5 mM HCO3,
comparable to the lake water) with NO3 and
NH4+ concentrations adjusted to meet the
demands of the specific experiment. No attempt was made to add extra
CO2 during the aeration, and the overlying water therefore
reached a pH of 8.6 compared to a pH of 8.2 in the lake water. Water
movement across the sediment surface was facilitated by directing a
small air jet at the water surface adjacent to the sediment core. The
samples were subjected to a 12-h/12-h light-dark cycle by using a slide
projector giving a light intensity of 110 µmol of photons
m2 s1 in the 400 to 700 nm range to
simulate the low-light conditions of the densely forested sampling
site. The sediment cores were subjected to one or two light-dark cycles
before analysis with microsensors.
Microsensors.
Microprofiles of O2 were obtained
with a Clark-type O2 microsensor (Unisense, Aarhus,
Denmark) mounted on a micromanipulator with computerized depth control
and data acquisition (25). A 2-point linear calibration for
the O2 sensors (23) was obtained by reading the
signal in well-aerated overlying water and anoxic sediment. The
O2 content of air-saturated water was calculated by using
the formula of Garcia and Gordon (10).
Depth profiles of NO
3 were obtained by using
microscale NO
3 biosensors with a sensing tip
of 30 µm in diameter (
15). The
NO
3 sensors were constructed by placing
immobilized, nitrous oxide
reductase-deficient denitrifying bacteria in
front of a N
2O microsensor.
The bacteria reduce
NO
3 and NO
2 to
N
2O, which in turn is detected by the built-in
N
2O sensor.
The signal from the sensor thus represents the
sum of NO
3 and NO
2,
and if any N
2O is present it would result in a signal
corresponding
to 2.4 times the signal of an equivalent concentration of
NO
3 or NO
2
(N
2O contains the N atoms from two
NO
3 or NO
2 ions but
as the diffusivity in water is about 1.2 times that
for the ions the
contribution to the signal is increased by a
factor of 2.4). Readings
performed with a N
2O microsensor (
24)
indicated,
however, that N
2O concentrations were below 1 µM and
therefore no corrections for N
2O were performed. In the
following
we only write NO
3, although the
actual parameter being measured is NO
3 plus
NO
2. The bacterial growth medium within the
sensor contained LiCl
as the dominating salt to ensure a positive tip
potential of the
sensor so that entry of negatively charged ions
including NO
3 and
NO
2 was facilitated.
The NO
3 sensors were mounted on a
computer-controlled system which was similar to that used for the
O
2 microsensors. Before
the experiments the linearity of
the NO
3 sensor signals in the range of 0 to
250 µM was checked by a 5-point
calibration in a test chamber.
Calibration of the NO
3 sensors during
incubations was done by linear 2-point calibrations
by using the signal
in the bulk water and the signal at a depth
of 1 cm in the sediment,
where the concentration was assumed to
be zero. Reference
NO
3 and NO
2
concentrations in bulk water samples were determined by high-pressure
liquid chromatography (anion separation column LCA A14; SYKAM,
Gilching, Germany) with 20 mM NaCl as eluent. Peaks were detected
at
220 nm with a SpectroMonitor 3200 spectrometer. In order to
correct for
minor drift in the signal of the NO
3 sensors
we used the average of two sets of calibration points
for the
calibration of each profile, one set obtained just before
and the other
just after profiling. One of the two applied sensors
had a downward
drift in sensitivity of about 4% per hour, while
the other sensor
exhibited a drift of less than 1% per hour. When
signal drift has
resulted in too low a signal, a high sensitivity
can be reestablished
by reversing the polarization of the nitrous
oxide sensor for about 1 min (
15).
Microprofiles were obtained at randomly chosen locations in a single
sediment core. Sediment porosity was determined for the
top 5.5 mm by
weighing and drying four subcores of known volume
taken at the end of
the experiment and was found to be 0.8 ± 0.04
(mean ± standard deviation). As an estimate of the effective sediment
diffusion
coefficients (
Ds) of O
2 and
NO
3, we used the product of the diffusion
coefficient in free water
and the squared porosity (
35).
Diffusion coefficients for O
2 (2.1 × 10
5 cm
2 s
1) and
NO
3 (1.7 × 10
5
cm
2 s
1) in water were found in the literature
(
3,
16). The
Ds estimated
for
O
2 and NO
3 were thus 1.3 × 10
5 cm
2 s
1 and 1.1 × 10
5 cm
2 s
1, respectively.
Estimation of consumption and production profiles.
As the
basis for the calculation of net consumption and production rates of
O2 and NO3 from the measured
microprofiles we used Fick's second law of diffusion including a
production and a consumption term (26):
|
(1)
|
where
C(
z,t) is the concentration at time
t and depth
z,
R is the respiration
rate, and
P is the production rate. Assuming
steady state we
have
|
(2)
|
so Eq.
1 can be reduced to
|
(3)
|
Defining
A(
z) = [
R(
z)
P(
z)]/
Ds and using
Euler's formula for numeric integration we find
|
(4)
|
where
h determines the step size used for numerical
integration. After further integration we have
|
(5)
|
Substituting
C/zn with equation 4 we find
![C<SUB>n+1</SUB>=C<SUB>n</SUB>+h×[∂C/∂z<SUB>n−1</SUB>+h×A<SUB>n−1</SUB>]](/content/vol64/issue9/fulltext/3264/img006.gif)
|
(6)
|
Using equation 6 we can calculate concentration profiles on the
basis of net activities (
An × Ds)
and by altering these net
activities we can minimize the sum of squared
deviations of the
calculated profile from the measured profile. We
chose to use
Microsoft EXCEL Solver to achieve this goal and as a
boundary
condition we introduced a point below the deepest measuring
point
with concentration and activity equal to zero. Starting from this
point we stepwise integrated upwards toward the sediment surface
with
h = 0.1 (the actually used step size during the
measurement
of the profile). Activities were not allowed to vary within
a
measuring step. This procedure on its own often yields oscillating
activities with no biological relevance because of noise in the
measured pore water concentration profiles. To compensate for
this
effect we also minimized the sum of squared first derivatives
of the
guessed activities,
(
A/
z)
2.
These two minima were weighted equally in order to smooth the
oscillations in the activities while still giving a good fit between
the measured and the calculated concentrations. The activities
obtained
this way are per unit volume of pore water, and integrated
activities
expressing the activity per unit of surface area of
sediment must thus
be multiplied with the porosity.
| |
RESULTS |
Profiles and activities in the dark.
The oxygen penetration
into the sediment was only 0.9 mm when the sediment was incubated in
the dark with 27 µM NO3 (actually 25 µM
NO3 and 2 µM NO2)
in the overlying water (Fig. 1A). The
penetration of NO3 extended to a depth of
about 1.5 mm, with some inaccuracy in the determination of
concentrations below 1 µM as evidenced by the scatter of the data
points at these low concentrations. The decrease in
NO3 concentration in the upper part of the
oxic zone was linear and indicated neither
NO3 production (nitrification) nor
consumption (assimilation or denitrification).
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FIG. 1.
Measured O2 ( , mean values with bars
indicating standard deviations, n = 6) and
NO3 ( , mean values with bars indicating
standard deviations, n = 6) concentrations, calculated
profiles (lines) and NO3 assimilation (gray
[top]), nitrification (light grey), and denitrification (gray
[bottom]) rates during darkness (A) and illumination (B). Both the
O2 and the NO3 profiles were
measured at different sites in the core.
|
|
Modelling of the NO
3 profile in the
dark indicated moderate denitrification activities up to 0.03 nmol
cm
3 s
1. The total rate of oxygen
consumption in the dark for the core
analyzed in Fig.
1 amounted to 2.4 mmol m
2 h
1 (Table
1) while the denitrification amounted to
0.08 mmol NO
3 m
2
h
1 (Table
2).
Profiles and activities during illumination.
When the sediment
was illuminated both O2 and NO3
profiles changed dramatically compared to the dark profiles (Fig. 1B).
An O2 peak was created in the upper 1 mm of the sediment
due to a net production of 7.4 mmol of O2 m2
h1 in the upper 0.3 mm while a net O2
consumption of 2.1 mmol m2 h1 below 0.3 mm
in depth decreased the oxygen concentration to zero at a depth of 2.2 mm. The nitrate concentration decreased in the 0.3-mm surface layer
characterized by photosynthetic oxygen production and reached a minimum
of 15 µM before it increased again in the oxic layer below the
photosynthetic zone, where a peak concentration of 28 µM was found at
a depth of 1.8 mm. A concentration of zero was reached at 2.7 mm.
Calculations based on the profile showed an upper net
NO3 consumption zone from 0 to 0.7 mm, a net
production zone from 0.7 to 2.1 mm, and a lower net consumption zone
from 2.1 to 2.7 mm. The integrated rates are listed in Table 2.
Higher NO3 concentration.
Analysis
of another sediment core incubated with 104 µM
NO3 showed comparable data for oxygen
penetration and oxygen consumption in the dark (Table 1), but
denitrification was increased to 0.20 mmol of
NO3 m2 h1 (Table
2). The sediment core was, however, quite different from the core
analyzed in Fig. 1, as the oxygen penetration in the light was about
double, although the rate of net oxygen production in the photic layer
was only 3.9 mmol m2 h1. The values for
nitrate transformations at 104 µM NO3
presented in Table 2 therefore cannot be directly compared with the
values for 27 µM NO3. The general
tendencies are, however, identical between the two experiments,
with a pronounced assimilation zone in the top, a nitrification zone in
the oxic layer below the diatom layer, and a largely anaerobic
denitrification zone. The maximum nitrification activities in both
experiments were found very close to the oxic-anoxic interface where
the nitrifiers can be supplied by ammonium from below.
High O2 concentration in water.
It seemed obvious
that the induction of oxic conditions in deeper layers by
photosynthetic oxygen production caused the relatively high rates of
nitrification during illumination. To verify this assumption an
experiment was conducted where the overlying water was purged with pure
O2. This caused the oxygen penetration to increase from 1.2 to 3.2 mm after purging for 200 min (Fig.
2A). The increased oxygen penetration
caused a gradual change in NO3 profile so
that a situation with a subsurface NO3 peak
was reached after 150 min. Modelling of this nitrate profile resulted
in a distribution of nitrification and denitrification similar to that
observed for the illuminated sediment (Fig.
3), i.e., significant nitrification
activities in the oxic layers next to the oxic-anoxic interface but no
activity in the layer with diatoms.
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|
FIG. 2.
(A) , measured O2 profile with
atmospheric O2 concentration in the bulk water at time 40
to 5 min (means ± standard deviations [error bars],
n = 6); , measured O2 profile with 930 µM O2 in the bulk water at time 200 to 250 min
(means ± standard deviations [error bars], n = 6). (B) NO3 profiles obtained in the period
between the O2 profiles shown in panel A after oxygenation
for 5 min (), 30 min (), 60 min ( ), 80 min ( ), 100 min
( ), and 150 min (×). Oxygen profiles were measured at the same site
whereas the nitrate profiles were measured at different sites in the
core.
|
|
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|
FIG. 3.
Measured NO3 concentrations
() at time 150 min (cf. Fig. 2), modelled
NO3 profile (line), and nitrification (grey
bars) and denitrification (dark bars) rates with 930 µM
O2 in the bulk water.
|
|
NH4+ addition.
The
NO3 profile in a dark-incubated core did not
change upon addition of 1 mM NH4Cl to the overlying water
(data not shown), indicating that no active nitrifiers were present in
the uppermost 1-mm layer that was oxic during dark incubation.
| |
DISCUSSION |
Nitrification.
The present study shows that nitrification in
sediment covered by benthic microalgae is dependent on the
O2 produced by microphytobenthic photosynthesis. During
darkness nitrification could not be detected in the 1-mm thick oxic
surface layer (Fig. 1A), whereas increased oxygen penetration down to a
depth of 2.2 mm, due to photosynthesis during illumination, resulted in
considerable nitrifying activity in the deeper oxic layers devoid of
microalgae. A high O2 concentration in the overlying water
also resulted in nitrification near a deeply positioned oxic-anoxic
interface but with no activity in the surface zone containing
microalgae. Even addition of 1 mM NH4+ to the
overlying water did not induce nitrifying activity in the surface
layer, and there is thus evidence that nitrifiers were virtually absent
from these layers.
Several reasons might explain the absence or low levels of
nitrification and nitrifying bacteria in the surface layers of
microalgae-inhabited sediments including low
NH
4+ concentration, high O
2
concentration, high pH, low CO
2 concentration,
high light
intensity, and possibly an excretion of chemical agents
with activity
against nitrifiers (
12). To these factors, which
are
discussed below, could be added a high grazing pressure in
the surface
layers where the input of high-quality organic matter
by photosynthesis
is high.
It is evident that the phototrophs with their abundant supply of energy
must be very potent competitors for ammonium. In planktonic
environments microalgae may decrease the NH
4+
concentrations to nanomolar levels, whereas the half-saturation
concentration is reported to be in the range of 5 to 200 µM
NH
4+ for nitrifiers (
12,
31).
Nitrifiers in layers with an abundance
of microalgae should thus be
active only if the supply of ammonium
to the layer is in excess of the
demand for the phototrophs. An
excess of nitrate will not improve the
competitiveness of the
nitrifiers as microalgae generally prefer
NH
4+ over NO
3
(
36). The high O
2 concentrations present in
highly active benthic
communities of microphytes, which may represent
O
2 partial pressures
above 1 atm (
25), may
inhibit nitrifiers (
29), as may the
pH values around 10 often observed in such communities (
25).
High light
intensities are assumed to inhibit nitrification (
11,
34) in
the surface waters of oceanic environments, and the sediment
surface
layers are exposed to similar high light intensities.
Judged from our
data, however, light is not likely to be the main
factor responsible
for low nitrification in the surface layers,
as nitrification was also
absent in the bottom of the diatom mat
(Fig.
1) where the light
intensity must have been very low.
Whatever the reasons are for the inhibition of nitrification in the top
sediment layer such inhibition has some potential
advantages for the
benthic microalgae. The algae do not have to
compete for
NH
4+ with the nitrifying bacteria and nitrogen
loss from the system
is reduced by the lower amount of
NO
3 available for denitrification compared to
a situation with nitrification
going on in the whole oxic layer. It is
thus also possible that
the microalgae produce chemical agents which
inhibit nitrification.
Nitrification is easy to inhibit as the ammonium
monooxygenase
is sensitive to a wide range of chemical species
(
20).
Denitrification.
The NO3 profiles
obtained during darkness indicate that denitrification occurs at
O2 concentrations of up to 20 µM (Fig. 1A). During
illumination, on the other hand, net denitrification was not detected
until O2 concentrations had dropped below 1 µM, above which nitrification dominated. It is, however, likely that the zones of
nitrification and denitrification overlapped during the day, so that
the low denitrification rates at O2 concentrations below 20 µM were masked by nitrification. Aerobic denitrification has been
observed in pure cultures of denitrifying bacteria (28), and
it is possible that the bacteria mediating the nitrification also
participate in an aerobic denitrification as denitrification has been
observed in both Nitrosomonas spp. (2) and
Nitrobacter spp. (9). Repeated analysis of
denitrification in a biofilm by use of N2O microsensors
also indicated denitrification when the O2 concentration
was <20 µM (6). Denitrification in sediments in the
presence of up to 10 µM O2 has been suggested by
Blackburn et al. (1), who used a computer model to simulate
NO3 profiles obtained with liquid ion
exchanger-type NO3 microelectrodes
(14).
Comparison of data on nitrogen cycling obtained by the
NO3 biosensor and other techniques.
The
depth-integrated rate of denitrification was highest in the illuminated
sediments (Table 2). This is contradictory to early reports on
denitrification in illuminated sediments (5), where it was
found that photosynthesis increased the O2 penetration and
thereby lowered denitrification by causing a reduced flux of nitrate
from the overlying water to the denitrifying layers. This earlier work
was based on acetylene inhibition of N2O reductase, but the
addition of acetylene also blocked nitrification and the method thus
only gave reliable results in environments where tightly coupled
nitrification-denitrification did not occur. Later work based on the
isotope pairing method (17) showed, however, that coupled
nitrification-denitrification could be stimulated by light-induced oxygen production (27) and that this coupled process was
almost independent of the NO3 concentration
in the overlying water. The data presented here thus support the
findings obtained by the isotope pairing method. However, in contrast
to the isotope pairing method the NO3
microprofiles also yield information about the exact location of the
NO3-transforming processes. Analysis of
NO3 microprofiles may also result in good
estimates of nitrifying activity, even when the isotope pairing method
cannot yield such information due to high rates of
NO3 assimilation, but the zones of
NO3 assimilation and nitrification should
then be well separated as they were in this study. An overlap between
the nitrifying and denitrifying layers may also result in minor
underestimations of both nitrification and denitrification.
By use of the microscale NO
3 biosensors it is
possible to get very detailed information about the processes producing
or consuming
NO
3. Due to the absence of
interference it is now possible to analyze
such processes in all
environments with sufficient water content.
Coupled
nitrification-denitrification in some environments such
as the plant
rhizosphere (
22) and manure-soil interface (
19)
is difficult to study by conventional techniques, and the microsensors
may prove useful for the study of these ecologically very important
microenvironments. With the rise of molecular microbial ecology
any new
microsensor for an ecologically important chemical species
also
improves our possibility of linking species distribution
and in situ
gene expression, etc., with microscale chemical transformations,
such
as has already been done in a few studies (
21,
30).
| |
ACKNOWLEDGMENTS |
This study was supported by the Commission of the European
Community under the Mast III Programme MICROMARE, project no. 950029, and by the Danish Biotechnology Programme.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Microbial Ecology, University of Aarhus, Ny Munkegade, Building 540, DK-8000, Aarhus C, Denmark. Phone: 45 89 42 32 44. Fax: 45 86 12 71 91. E-mail: revsbech{at}biology.aau.dk.
| |
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Applied and Environmental Microbiology, September 1998, p. 3264-3269, Vol. 64, No. 9
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
