Blackwell Science, LtdOxford, UKEMIEnvironmental Microbiology 1462-2912Society for Applied Microbiology and Blackwell Publishing Ltd, 20057913921404Original ArticleIn situ activity and assimilation in nitrifying biofilmsA. Gieseke
et al.
Environmental Microbiology (2005) 7(9), 1392–1404
doi:10.1111/j.1462-2920.2005.00826.x
In situ substrate conversion and assimilation by
nitrifying bacteria in a model biofilm
Armin Gieseke,1* Jeppe Lund Nielsen,2
Rudolf Amann,3 Per Halkjær Nielsen2 and
Dirk de Beer1
1
Microsensor Group, Max Planck Institute for Marine
Microbiology, Celsiusstrasse 1, D-28359 Bremen,
Germany.
2
Institute for Life Sciences, Aalborg University,
Sohngaardsholmsvej 57, DK-9000 Aalborg, Denmark.
3
Department of Molecular Ecology, Max Planck Institute
for Marine Microbiology, Celsiusstrasse 1, D-28359
Bremen, Germany.
Summary
Local nitrification and carbon assimilation activities
were studied in situ in a model biofilm to investigate
carbon yields and contribution of distinct populations
to these activities. Immobilized microcolonies
(related
to
Nitrosomonas
europaea/eutropha,
Nitrosomonas oligotropha, Nitrospira sp., and to
other Bacteria) were incubated with [14C]-bicarbonate
under different experimental conditions. Nitrifying
activity was measured concomitantly with microsensors (oxygen, ammonium, nitrite, nitrate). Biofilm thin
sections were subjected to fluorescence in situ
hybridization (FISH), microautoradiography (MAR),
and local quantification of [14C]-bicarbonate uptake
(beta microimaging). Nitrifying activity and tracer
assimilation were restricted to a surface layer of different thickness in the various experiments (substrate
or oxygen limitation). Excess oxygen uptake under all
conditions revealed heterotrophic activity fuelled by
decay or excretion products during active nitrification. Depth limits and intensity of tracer incorporation
profiles were in agreement with ammonia-oxidation
activity (measured with microsensors), and distribution of incorporated tracer (detected with MAR).
Microautoradiography revealed a sharp individual
response of distinct populations in terms of in-/activity depending on the (local) environmental conditions
within the biofilm. Net in situ carbon yields on N,
expressed as e– equivalent ratios, varied between
Received 16 September, 2004; revised 21 January, 2005; accepted
25 January, 2005. *For correspondence. E-mail: [email protected]; Tel. (+49) 421 2028 836; Fax: (+49) 421 2028 690.
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd
0.005 and 0.018, and, thus, were in the lower range of
data reported for pure cultures of nitrifiers.
Introduction
Intensive recent application of in situ techniques like
microsensor measurements or fluorescence in situ hybridization (FISH) has strongly increased our understanding
of the function and structure of microbial communities.
These techniques are of particular use in immobilized
microbial biomass or on organisms that are difficult to
cultivate like nitrifying bacteria. Nitrifiers can cause extensive loss of fertilizer ammonium from soils and subsequent eutrophication of natural waters via leaching of the
product nitrate (Prosser, 1986), but are also responsible
for the first steps in nitrogen removal from aquatic ecosystems (Painter, 1986).
Fluorescence in situ hybridization with oligonucleotide
probes specific for various groups of nitrifying bacteria has
been successfully used for identification, quantification,
and localization of ammonia-oxidizing bacteria (AOB) and
nitrite-oxidizing bacteria (NOB) in sewage treatment systems (e.g. Wagner et al., 1995; Mobarry et al., 1996; Wagner et al., 1996; Juretschko et al., 1998; Schramm et al.,
1998; Schramm et al., 1999). A 16S rRNA-based detection, however, provides only indirect evidence for the physiological potentials and recent activities of the respective
populations. Therefore, this method has been combined
with techniques for in situ analysis of metabolic processes. Several recent studies combined FISH with
microsensor measurements to characterize the microenvironment and N oxidation (i.e. ammonium and nitrite
conversion) activity of nitrifying populations in biofilms
(Schramm et al., 1996; Okabe et al., 1999; Schramm
et al., 1999; Schramm et al., 2000; Gieseke et al., 2001;
Schramm, 2003). Combining both, the measured N oxidation activity and the spatial distribution may allow direct
conclusions with respect to ecology and in situ physiology
of a defined population. In this way, calculation of cellspecific N oxidation rates and in situ substrate affinities
have been achieved.
On the applied technical scale or in multifunctional systems, however, microbial communities in flocs or biofilms
are often complex. The coexistence of several populations
of AOB and NOB, and the lack of a layered architecture
along substrate gradients have been reported for various
In situ activity and assimilation in nitrifying biofilms 1393
treatment systems (Mobarry et al., 1996; Schramm et al.,
1996; Schramm et al., 2000; Gieseke et al., 2001;
Gieseke et al., 2003). Moreover, nitrifying biofilms can
harbour a significant amount of other non-nitrifying organisms (Okabe et al., 1996; Gieseke et al., 2003) contributing to the overall activity. The combined application of
FISH and microsensor measurements in such systems is
not sufficient to resolve the specific contribution of a certain population or individual cells to the overall or local
activity observed in different microenvironments.
The combination of FISH with microautoradiography
(MAR) has been proposed and successfully applied to
investigate whether a specific population or even a single
identified cell actively assimilates a substrate under certain environmental conditions (Nielsen et al., 1998; Lee
et al., 1999; Ouverney and Fuhrman, 1999). Quantification of the assimilation, however, is laborious and time
consuming, because the photographic detection is relatively insensitive and various interfering factors have to be
controlled (Nielsen et al., 2003).
Beta microimaging has been recently suggested in
medical fields as a sensitive technique to measure the
two-dimensional distribution of incorporated radiotracer
with high spatial resolution (10 mm) (Lanièce et al., 1998).
Although its resolution does not allow detection on the
single-cell level, local uptake into tissues can be
quantified.
Microautoradiography together with quantitative beta
microimaging potentially allows determination of local carbon assimilation in biofilms. Related to local substrate
oxidation activities (measured with microsensors), this
would give means to determine in situ carbon yields (i.e.
units C assimilated per unit substrate converted) in immobilized microbial biomass. Until now, growth yields in biofilms have only been determined indirectly, i.e. based on
mass balances of substrates/products and biomass
formed. These methods, however, give overall rather than
local measures and are of limited use for insights into the
ecophysiology of cells immobilized in distinct microenvironments. Furthermore, it is not clear from the published
data whether the yield of cells immobilized in a biofilm is
different from that of suspended cells (van Loosdrecht
et al., 1990; Ellis et al., 2000).
In this study the in situ activity and assimilation of a
complex, nitrifier-dominated community in a model biofilm
was investigated under various conditions. For this purpose, we combined the aforementioned methods, i.e. (i)
FISH for localization of single populations (ii) microsensor
measurements for determination of local N oxidation rates
(iii) MAR to localize carbon incorporation on the singlecell level and (iv) a quantitative beta microimaging technique to quantify local assimilation of inorganic carbon on
a model biofilm. Our hypotheses were (i) that the relative
activity of different populations depends on the respective
microenvironmental conditions and (ii) that the supply of
electron donor (ammonium, nitrite) and acceptor (oxgen)
to the biofilm directly affects the autotrophic carbon fixation (measured as in situ carbon yield) and the relative
extent of concomitant heterotrophic activities.
Results
Microbial populations
The model biofilm was prepared by suspension and
immobilization of flocs from a nitrifying continuous upflow
reactor in an agarose matrix. Fluorescence in situ hybridization of samples from the source reactor with various
oligonucleotide probes revealed the presence of three
main nitrifying populations in the flocs: AOB related to
Nitrosomonas europaea/eutropha (hybridizing with
probes Nso1225, NEU and Nse1472), and to the
Nitrosomonas oligotropha-lineage (positive with probes
Nso1225, NOLI and Nmo218), and one NOB population
affiliated to the genus Nitrospira (hybridization with probes
Ntspa712 and Ntspa662) (Table 1). Both AOB populations
were equally abundant in the source reactor suspension,
and in total ammounted to 1.5 (± 0.51) ¥ 108 cells cm-3
(mean ± SD, n = 5 wells). The abundance of the NOBrelated cells was 4.1 (± 1.2) ¥ 108 cells cm-3. Hybridization
with a mixture of probes EUB338-I to EUB 338-III revealed
8.8 (± 0.64) ¥ 108 cells cm-3 in the suspension. The sum
of AOB and NOB thus accounted for two third of total
bacterial abundance. Most aggregations remained intact
upon preparation of the model biofilm. They consisted of
widely and homogeneously dispersed aggregates of different single phylotype microcolonies, in which individual
cells were easily distinguishable.
Nitrifying activity under different conditions
Local concentrations of oxygen, ammonium and nitrite or
nitrate were measured in the 4-mm thick model biofilm
under different incubation conditions (Table 2). Under
standard conditions (experiment A) profiles of oxygen,
ammonium, and nitrate reflected N oxidation activity. Oxygen penetration depth (defined here as depth where
oxygen < 0.2% air saturation) was about 2.1 mm. Ammonium consumption and nitrate production took place
exclusively within this oxic layer (Fig. 1A, top panel). Local
oxygen consumption rates were similar over large parts
of the oxic layer (Fig. 1A, bottom). Under limitation of
ammonium (experiment B) oxygen was present in all layers of the model biofilm. Within the active surface layer of
1.8 mm thickness the oxygen concentration was
decreased to 70% air saturation, and ammonium was fully
depleted (Fig. 1B, top). Consumption was highest at the
surface (Fig. 1B, bottom). Total (areal) conversion rates of
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
1394 A. Gieseke et al.
Table 1. Probes and hybridization conditions applied in this study.
Probe
Probe Sequence (5¢ to 3¢)
Target sitea
Target organism(s)
FA (%)b
EUB338-I
EUB338-II
EUB338_III
Nso1225
GCTGCCTCCCGTAGGAGT
GCAGCCACCCGTAGGTGT
GCTGCCACCCGTAGGTGT
CGCCATTGTATTACGTGTGA
16S (338–355)
Domain Bacteria
35
80
16S (1224–1243)
35
80
NEUd
CCCCTCTGCTGCACTCTA
16S (653–670)
40
56
(Wagner et al., 1995)
Nse1472e
Nmo218
NOLI191d
ACCCCAGTCATGACCCCC
CGGCCGCTCCAAAAGCAT
CGATCCCCCACTTTCCTC
16S (1472–1489)
16S (218–235)
16S (191–208)
50
35
30
28
80
112
Ntspa712d,f
Ntspa662d,f
CGCCTTCGCCACCGGCCTTCC
GGAATTCCGCGCTCCTCT
16S (712–732)
16S (662–679)
Ammonia-oxidizing
b-proteobacteria
Halophilic/halotolerant
Nitrosomonas spp.
N. europaea
N. oligotropha-lineage
Various members of
the N. oligotrophalineage
Phylum Nitrospira
Genus Nitrospira
(Amann et al., 1990)
(Daims et al., 1999)
(Daims et al., 1999)
(Mobarry et al., 1996)
35
35
80
80
NaCl (mM)c
Reference
(Juretschko et al., 1998)
(Gieseke et al., 2001)
(Gieseke et al., 2001)
(Daims et al., 2000)
(Daims et al., 2000)
a. rRNA position according to E. coli numbering (Brosius et al., 1981).
b. Percentage of formamide in the hybridization buffer.
c. Concentration of sodium chloride in the washing buffer.
d. Applied together with equimolar amount of unlabelled competitor oligonucleotide as indicated in the reference.
e. Referred to as S-*-Nse-1472-a-A-18 in the reference.
f. Referred to as S-*-Ntspa-0712-a-A-21 and S-G-Ntspa-0662-a-A-18, respectively, in the reference.
both, oxygen and ammonium were clearly lower than in
experiment A (Fig. 2). Nitrate release corresponded to the
consumption of ammonium (1:1 ratio), indicating full nitrification in this experiment (Fig. 2). With nitrite as the only
electron donor added (experiment C), oxygen (Fig. 1C,
bottom) and nitrite were consumed at similar rates over
the whole aerated zone, i.e. down to a depth of about
2.6 mm (Fig. 1C, top). The areal oxygen consumption rate
was slightly lower than under standard conditions (Fig. 2).
Experiment D was planned to subject AOB and NOB
populations to competition for oxygen. Under conditions
where the oxygen concentration was reduced to 20% and
both, ammonium and nitrite were present in high concentrations only a weak oxidation of ammonium could be
observed. Oxygen penetrated less deep, i.e. to 1.1 mm,
and the areal oxygen consumption rate was the lowest of
all experiments (except negative control, N) (Fig. 2). Highest local oxygen consumption within the model biofilm
took place about 0.5 mm below the surface (Fig. 1D,
bottom).
In all experiments the areal oxygen consumption rates
were stoichiometrically too high for the observed N con-
version (Fig. 2). The different incubation conditions
involved different stoichiometries. Therefore, a mass balance was calculated based on electron equivalents (e–
eqs) from acceptor to donor (8 e– eqs per mol ammonium
and 2 e– eqs per mol nitrite oxidized; 4 e– eqs per mol
oxygen reduced). This resulted in a ratio of e– eqs to
acceptor (oxygen) divided by e– eqs from donor (ammonium and/or nitrogen) greater than one in all cases
(Table 3). In experiments A–D the excess oxygen consumption not explained by nitrification (i.e. the e– eqs to
acceptor not originating from ammonium and/or nitrite)
was between 15% and 69% of total oxygen consumption
(Table 3). Only a small consumption of oxygen in the order
of 11% ± 3% of that under standard conditions was
observed when neither ammonium nor nitrite were
present (negative control N; Fig. 2).
Assimilation activity on single-cell level
The combination of MAR and FISH was used to relate the
assimilation of inorganic carbon within the model biofilm
(visualized by MAR signals) to individual cells or single
Table 2. Incubation conditions in the experiments.
Designation
Experimental conditions
Ammonium concentration
(mM)
Nitrite concentration
(mM)
Oxygen concentration
(% air saturation)
A
B
C
D
N
Standard
Ammonium limitation
Nitrite oxidation
Competition
Negative control
0.7
0.02
0.07
0.3
–
–
–
0.6
0.3
–
100
100
100
20
100
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
In situ activity and assimilation in nitrifying biofilms 1395
Ammonium, Nitrate (mM)
Ammonium, Nitrate (mM)
500
600
700
0
0
100
200
0
10
25
Ammonium, Nitrite (mM)
20
30
50
75
Nitrite (mM)
0
200
400
0
100
200
300
200
300
400
500
Depth (mm)
–1
0
1
2
A
3
0
25
50
75
B
100 0
25
50
75
C
100 0
25
50
75
D
100 0
25
50
75
100
Oxygen (% air)
Depth (mm)
0
1
2
A
0.0
–0.4
–0.8
B
0.0
–0.4
–0.8
C
0.0
–0.4
Local oxygen consumption (mmol cm
–0.8
D
0.0
–0.4
–0.8
–3 h–1)
Fig. 1. Mean concentration profiles of oxygen (), ammonium (), nitrite () and nitrate () (top), and distribution of local oxygen consumption
rates with depth (bottom) in the model biofilm. Designations A–D refer to the incubation conditions (Table 2). Note the different scales. The top
offset axis refers to the concentration of ammonium, the top axis to either nitrate (A and B) or nitrite (C and D) respectively. (Error bars: SE;
number of profiles: see Fig. 2).
phylotype microcolonies identified by FISH. A dark layer
of silver grains covered cells with an active uptake of [14C]labelled inorganic carbon (Fig. 3).
Results of MAR are summarized in Table 4. Under aeration and presence of 0.7 mM ammonium (experiment A),
all three nitrifier populations were the MAR-positive
throughout the model biofilm, with only slight differences
in silver grain density along depth. Under ammonium limitation (experiment B) the MAR pattern was changed:
Only cells close to the surface were MAR-positive. A few
AOB cells hybridizing with probe Nmo218, but the majority
of AOB hybridizing with probe Nso1225 only, and half of
the Ntspa712-positive microcolonies (NOB) incorporated
the label in this experiment. When a high concentration of
nitrite was added to the incubation (experiment C), cells
related to either of the two AOB populations were almost
completely MAR negative. NOB hybridizing with Ntspa712
were found to be strongly MAR-positive throughout the
model biofilm. However, a few colonies near the surface
were negative. A few algal contaminants present in the
model biofilm were MAR-positive. Presence of ammonium
and nitrite in similar high concentrations, and limited
amounts of oxygen (experiment D), caused a few N. oligotropha-related cells at the surface to be covered with silver
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
1396 A. Gieseke et al.
Fig. 2. Areal conversion rates of oxygen (grey), ammonium (hatched),
nitrite (white) and nitrate (cross-hatched) under different experimental
conditions. Values were calculated based on the slope of the concentration gradients through the diffusive boundary layer. Positive data
correspond to a release, negative data to a consumption of the
respective solute. Designations A–D and N refer to incubation conditions as described in Table 2. (Error bars: 95% confidence limits;
numbers of profiles indicated in diagram).
grains. The majority of N. europaea/eutropha-affiliated
cells at the surface was also weakly MAR-positive. In
deeper layers, however, most cells were MAR-negative.
NOB (hybridizing with the probe Ntspa712) were MARnegative only near the bottom.
Quantification of carbon uptake
Beta microimaging was used to detect the distribution and
quantity of incorporated [14C]-labelled inorganic carbon in
the model biofilm. The distribution of [14C] in vertical thin
sections showed a similar heterogeneity of signals as the
distribution of microcolonies after hybridization. Figure 4
visualizes examples of tracer uptake from all experiments
as colour-coded quantitative 2D distribution within thin
sections of the model biofilm. Differences to MAR results
can be attributed to the higher sensitivity of beta microimaging. Whereas the uptake of [14C] was high under standard conditions (experiment A), only a low amount of
tracer was incorporated in homogeneously distributed
patches along depth in the negative control (N). In experiment C distinct tracer uptake also occurred deeper in the
model biofilm. In contrast, this took place mostly at the
surface in experiments B and D, with the exception of a
few isolated local spots with tracer situated in deeper
layers. Examples in Fig. 4 Demonstrate that similar signal
intensities were obtained in experiments A and C, but
lower ones in experiments B and D.
Beta microimaging allowed quantitative analysis of local
inorganic carbon assimilation. For that purpose [14C]
incorporation, representing assimilation of total inorganic
carbon present in the medium, was analysed along vertical profiles (Fig. 5). The depth, to which incorporation of
inorganic carbon was observed, was closely similar to the
penetration depth of the limiting component, i.e. oxygen
(experiments A and D), oxygen and nitrite (experiment C),
or ammonium (experiment B) respectively (Fig. 5). The
local assimilation rate under standard conditions was
highest at the surface of the model biofilm (upper 400 mm)
and decreased with depth under standard conditions
(experiment A, Fig. 5). In experiments B and D we
observed the same pattern, although rates were generally
lower and decreased more sharply with depth. In experiment C a more homogeneous distribution of the [14C]
incorporation rates was found in the upper 2.5 mm. In
general, the vertical decrease of local rates of carbon
uptake (Fig. 5) was much sharper than that of oxygen
conversion (Fig. 1, bottom). Total (areal) uptake rates of
inorganic carbon under the different experimental
conditions were: 0.78 nmol cm-2 h-1 (experiment A),
0.29 nmol cm-2 h-1 (experiment B), 0.45 nmol cm-2 h-1
(experiment C; with a substantial incorporation in layers
below 1 mm) and 0.18 nmol cm-2 h-1 (experiment D)
respectively.
Table 3 summarizes the assimilation of carbon (from
beta microimaging) and the conversion of nitrogen and
oxygen (from microsensor measurements) for all experimental conditions. The data are areal rates (calculated
from depth-integration of local volumetric rates) presented
as electron equivalents (Rittmann and McCarty, 2001) to
allow comparison of the different conditions with different
electron donors added. In experiments A and B ammonium was fully oxidized (release of 8 e– eqs per mol),
whereas in experiment C nitrite oxidation was dominant
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
In situ activity and assimilation in nitrifying biofilms 1397
Table 3. Acceptor and donor conversions (expressed as electron equivalents), and biomass and energy yields under different incubation
conditions.
Name
Experiment
Nitrogena
(me– eqs cm-2 h-1)
Oxygena
(me– eqs cm-2 h-1)
Carbona
(me– eqs cm-2 h-1)
Other donorsb
(%)
fs,meas.c
(–)
fe,meas.d
(–)
fe,max.e
(–)
A
B
C
D
Standard
Ammonium limitation
Nitrite oxidation
Competition
-0.35
-0.08
-0.33f
-0.04g
0.54
0.25
0.39
0.13
3.1 ¥ 10-3
1.2 ¥ 10-3
1.8 ¥ 10-3
0.7 ¥ 10-3
35
66
15
69
0.009
0.014
0.005
0.018
1.5
3.0
1.2
3.3
0.991
0.986
0.995
0.982
a. Data represent rates (from depth integration of local volumetric rates) multiplied by no. of electron equivalents produced/consumed per mol
[i.e. carbon reduction: 4; ammonia-oxidation (to nitrate): 8; nitrite oxidation (to nitrate): 2; oxygen reduction: 4] (Rittmann and McCarty, 2001). A
negative sign indicates e– eqs from donor, a positive e– eqs to acceptor.
b. Data based on amount of e– equivalents consumed by reduction of oxygen (col. 4) not originating from nitrification (col. 3).
c. Ratio fs (amount of e– eqs to carbon divided by e– eqs from donor) represents measured carbon yield.
d. Ratio fe (amount of e– eqs to oxygen divided by e– eqs from donor) represents measured energy yield.
e. Ratio fe,max. (equals 1-fs) represents maximum energy yield based on stoichiometric conversion of nitrogen and oxygen.
f. Data based on nitrite consumption rates.
g. Data based on ammonia-oxidation rates to nitrite (6 e– eqs mol-1) and to nitrate (8 e– eqs mol-1).
Table 4. Results of MAR-FISH for the different incubations.
Experiment
Population affiliated to
A
B
C
D
N. oligotropha
Topa
Middle
Bottom
+++b
++
++
++c
–
–
+
–
–
++c
–
–
N. europaea/eutropha
Top
Middle
Bottom
+++
++
++
++d
–
–
–
–
–
+d
–
–
Nitrospira sp.
Top
Middle
Bottom
+++
++
++
++e
–
–
+++d
+++
+++
+++d
++
–
a. Refers to depth of the investigated layer as follows: top 0–0.2 mm,
middle 0.2–1.5 mm, bottom > 1.5 mm.
b. Signs refer to estimated silver grain density around probe positive
cells as follows: +++ very high density, ++ high density, + low density,
– no silver grains.
c. MAR signal associated only with a few probe-positive cells.
d. MAR signal associated with the majority of all probe-positive
colonies.
e. MAR signal associated with about half of all probe-positive
colonies.
(release of 2 e– eqs per mol). Both, ammonium and nitrite
oxidation contributed to the rate of e- eqs from donor in
experiment D. Yields are expressed as ratios of e– eqs to
acceptor for energy generation (fe) or to carbon (fs) divided
by e– eqs from donor. These ratios reflect relative substrate partitioning and sum up to unity. The fs based on
measured carbon assimilation and ammonium (or nitrite)
conversion rates was between 0.005 (experiment C) and
0.018 (experiment D). The latter result in a calculated
maximum fe between 0.982 and 0.995, which represents
the ratio of e– eqs to the acceptor divided by e– eqs from
nitrification.
Discussion
Activity on the community level
Our experiments aimed at studying in situ the effects of
different (autotrophic) incubation conditions on (i) the distribution of activity among AOB and NOB microcolonies
(as detected with MAR-FISH) and on (ii) the carbon
assimilation and concomitant heterotrophic activities in a
complex, nitrifier-dominated community in a model biofilm.
Fig. 3. Example of a MAR-FISH micrograph
(from experiment D). Microautoradiographic
image (A) and overlay of MAR image with fluorescence micrograph (B) after hybridization
with probes Ntspa712 (red) and EUB338
(green). Cells appearing yellow hybridized with
both probes. Scale bar: 20 mm.
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
1398 A. Gieseke et al.
Fig. 4. Two-dimensional distribution of [14C]bicarbonate incorporation in vertical sections of
the model biofilm as visualized in high resolution by beta microimaging. Designations A–D
and N refer to the incubation conditions
(Table 2). Signal intensity per pixel
(10 mm ¥ 10 mm) is shown by colour-coding
(identical scale for all images), arrows indicate
surface and base of the model biofilm. Scale
bar: 1 mm; cpm, counts per minute.
Fig. 5. Vertical distribution of local inorganic carbon assimilation rates under different experimental conditions determined from beta microimaging
data (see Fig. 4). The shaded areas show the penetration depth of the limiting substrate, i.e. either oxygen (experiments A, C and D) or ammonium
(B) in the respective experiment (see also Fig. 1 and Table 2). (Error bars: SE; n = 6 thin sections).
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
In situ activity and assimilation in nitrifying biofilms 1399
The measured distributions of oxygen, ammonium,
nitrite and nitrate indicated mass transport limitations similar to that of a grown biofilm: The community was limited
by oxygen (experiments A, C and D) or ammonium (experiment B). Depth limits of the N oxidation activity, as followed with microsensors, correlated well with that of
actively assimilating microcolonies (i.e. covered with silver
grains after MAR-FISH) and local assimilation rates (as
detected by beta microimaging, see below).
Areal conversion rates of oxygen, ammonium and
nitrite, however, were 3–12 times lower than in naturally
grown nitrifying biofilms (Okabe et al., 1999; Schramm
et al., 1999; Gieseke et al., 2001). This was resulting from
the low cell density in our model biofilm (5.6 ¥ 108 nitrifier
cells cm-3) compared with grown biofilms (≥ 1 ¥ 1011 nitrifier cells cm-3) (Schramm et al., 1999; Gieseke et al.,
2001). The prime purpose of using a model system was
to create a transport-limited situation (and subsequent
formation of gradients and diverse microenvironments)
that is typical for biofilms and other aggregated microbial
biomass, while avoiding the structural heterogeneity and
high cell density of a grown biofilm. The abundances,
although artificially low, were sufficient to support strong
gradients over small distances that lead to differential
activity of distinct populations (see below).
Non-nitrifying Bacteria were present in significant
amounts. Relative abundances are comparable to those
reported for grown nitrifying biofilms (Okabe et al., 1996;
Gieseke et al., 2003). The oxygen uptake in excess to
nitrification demands under all experimental conditions
(fe > 1, see Table 3) suggests heterotrophic activity of
these populations with local carbon sources as donors
(see also below). Interestingly, heterotrophic oxygen consumption in absolute numbers is rather low when nitrification is absent (negative control) but about 3 times higher
when nitrification is active (experiments A and B). Donor
limitation and an additional supply of electron donors,
potentially provided by an active nitrifying guild, might
explain this increased activity. This ‘fuelling’ has been
observed earlier in chemostat experiments (Rittmann
et al., 1994).
Experiment D was designed to promote competition
between AOB and NOB, and the potential activity of a
‘nitrifier denitrification’ (Jetten et al., 1997). The relevance
of the latter process has been suggested for activated
sludge (Muller et al., 1995) and biofilms (Siegrist et al.,
1998; Helmer et al., 1999; Schmid et al., 2000). While the
appropriate population was present in our system, the
results do not show evidence for this type of metabolism:
Neither consumption of ammonium and/or nitrite nor
assimilation of inorganic carbon was observed in anoxic
layers. Colocalized oxidation of ammonium and nitrite prevailed in this experiment, as indicated by microsensor
data (Fig. 1) and MAR results (see below).
Contribution of the individual populations
Microautoradiography was used to detect how much the
distinct populations (detected by FISH) were involved in
the N oxidation and C assimilation activity. The activity of
single populations in the experiments was regulated primarily by the quality of the local microenvironment, i.e. the
(vertical) position of a cell or microcolony in the model
biofilm. The activity of all three nitrifying populations under
standard conditions (experiment A) suggests that there is
no cut-off competition for oxygen or substrate. Microautoradiography results (and also beta microimaging, see
below) showed that in layers void of oxygen or substrate
no carbon fixation occurred in any of the populations
including the N. europaea/eutropha-like cells (Table 4).
Forced substrate limitation (e.g. experiments B and D)
caused surface-bound assimilation activities without pronounced differences in activities among cells from one
distinct population. This indicates threshold levels, under
which a whole population is inactive (e.g. N. oligotropha
in experiments B and D) and above which fully active (N.
europaea/eutropha in experiments B and D, Nitrospira sp.
in experiments C and D).
Even traces of ammonium, however, were sufficient to
support some carbon fixation in both AOB populations
(MAR results of experiments B and C). The N. europaea/
eutropha-related cells showed assimilation activity only in
experiment B (low ammonium, nitrite absent), the N. oligotropha-affiliated population was more active in experiment C (traces of ammonium, nitrite present). Earlier
studies suggest that the latter group is adapted to low
substrate concentrations (Bollmann and Laanbroek,
2001), whereas N. europaea/eutropha-like populations
are more competitive under high concentrations of substrate (Schramm et al., 2000). It is critical to extrapolate
physiological characteristics of a taxonomic group from
information of a limited number of strains. Furthermore,
the structure (and potential function) of the community
and the coexistence of both AOB populations is determined by niche differentiation under the conditions in the
source reactor. The community architecture in the model
biofilms, however, is random and not formed by ecological
opportunities or constraints. Under suitable microenvironmental conditions both populations can be active during
the short-term incubations. Therefore, no conclusions on
adaptation can be drawn from the observed activity patterns. Possible explanations for the observed differential
activity under similar microenvironmental conditions, thus,
have to be attributed to potential physiological constraints
of the individual populations, e.g. (i) a differential effect of
low oxygen concentrations (Gieseke et al., 2001), and/or
(ii) a toxicity of the nitrite/nitrous acid (Anthonisen et al.,
1976) in experiments C and D. The activity of the N.
europaea/eutropha-like population is obviously affected to
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
1400 A. Gieseke et al.
a higher extent by either or both factors in experiments C
and D. Under some conditions (experiments B to D)
assimilation occurred only in some of the – sometimes
even neighbouring – microcolonies. This observation indicates the existence of additional mechanisms of local
activity control within a more or less similar microenvironment. In future studies it might be worthwhile to consider,
besides ammonia availability, also competition for oxygen
and product toxicity to explain the ecophysiology of different AOB strains.
The finding of MAR-positive Nitrospira sp. down to the
penetration limit of oxygen in experiments C and D supports the hypothesis of adaptation of Nitrospira sp. to low
oxygen levels (Schramm et al., 1999; Schramm et al.,
2000; Gieseke et al., 2001; Gieseke et al., 2003). An inhibition of the activity of Nitrospira sp. under high oxygen
concentration as suggested earlier to explain its predominant restriction to deeper biofilm layers (Okabe et al.,
1996) is not confirmed by our results. In all experiments
oxygen concentration was high at the surface, and MARpositive Nitrospira were found under these conditions.
In situ assimilation
Carbon assimilation rates and nitrification activities measured in situ allow the calculation of in situ yields (Table 3).
Conversion rates obtained from microprofiles, however,
are inherently sensitive to noise, and, thus, are prone to
substantial error. Adding the uncertainty of carbon assimilation rates, and the obvious contribution of heterotrophic
processes to the overall activities, the resulting data
should only be considered as estimates. Consideration of
stoichiometry and mass balances helps to provide a solid
description in this case. This can be reached by describing
the mass balance in terms of e– eqs (Table 3) transferred
from donor to acceptor and to biomass (Rittmann and
McCarty, 2001).
The net yields, expressed as fs (i.e. the ratio of e– eqs
to carbon divided by e– eqs from donor, Table 3), are 0.005
(experiment C) to 0.018 (experiment D). Robust gross
yields (fs) of 0.028 and 0.02 (i.e. 0.14 and 0.10 e– eqs
biomass, C5H7O2N, per e– donor) have been assigned
earlier to AOB and NOB respectively (Rittmann and
McCarty, 2001). Typical carbon yields (i.e. mol C assimilated per mol N oxidized) reported for pure cultures are in
a range of 0.014–0.096 mol C (mol N)-1 oxidized for various AOB, and 0.0125–0.031 mol C (mol N)-1 for Nitrobacter sp. and Nitrococcus mobilis (Prosser, 1989;
Poughon et al., 2001). These values correspond to fs
between 0.009 and 0.064 for AOB, and 0.025–0.062 for
NOB respectively. The fs determined here are in the lower
range of these data. This is first of all because of the fact
that the net yields determined in this study reflect the sum
of assimilation, maintenance and decay, and, thus, are to
be lower than the gross yield fs. Moreover, the data represent in situ yields for mixed communities in a transportlimited environment, which have not been measured previously. Effects like mass transfer limitations of oxygen or
substrate, substrate partitioning between populations, and
product toxicity (see above) might have lowered the yield
in situ compared with those obtained with suspensions of
pure cultures. Interestingly, a comparison of all experiments shows that the relative total amount of excess
oxygen consumed increases with the net carbon yield of
the nitrifiers (Table 3). The inverse is observed with the
carbon assimilation rate (excluding experiment C). A
higher net yield, observed under more restrictive conditions here (ammonium limitation, experiment B, or competition, experiment D), thus goes along with a stronger
relative contribution of heterotrophic organisms to oxygen
consumption. This relation potentially indicates an
increasing discrepancy between net and gross yields of
the nitrifying guild and an intensified ‘fuelling’ of the heterotrophic members of the community. More detailed
investigations on grown nitrifying biofilms might help to
explain this observation.
The net yield in experiment C (nitrite oxidation) of 0.005
is mainly associated with the activity of Nitrospira sp., and
is remarkably low compared with those obtained in experiments A or B (full nitrification or predominant ammoniaoxidation). Moreover, the abundance of Nitrospira sp. was
about 4 times higher than that of the AOB populations in
our model biofilm, and the thickness of the active layer
was larger. Thus, the cell-specific yield is even lower. This
result clearly supports earlier hypotheses that members
of the genus Nitrospira sp. are typical K strategists
(Schramm et al., 2000), characterized by a low maximum
substrate conversion rate, a low growth rate and yield
(Watson et al., 1986; Ehrich et al., 1995; Rittmann and
McCarty, 2001). Similar suggestions were inferred earlier
from in situ studies on nitrifying biofilms (Okabe et al.,
1999; Schramm et al., 1999).
The sharp decrease of local carbon uptake rates with
depth is explained by two facts. First, nitrifying bacteria
have a high maintenance energy demand (75–76% for
AOB and 53–81% for NOB respectively) (Prosser, 1989;
Poughon et al., 2001). Subsequently, gross yield
decreases with depth, i.e. the amount of energy for maintenance increases relative to the amount left for assimilation. Second, nitrifiers have a relatively high KM(O2) and
therefore are relatively poor competitors for oxygen
(Prosser, 1989). With increasing depth (or decreasing
oxygen concentration, respectively), the relative amount
of oxygen consumed by heterotrophs might increase.
In conclusion, the different methods produced consistent results. Nitrification as followed by the formation of
nitrite and nitrate and the consumption of oxygen with
microsensors was mostly limited to a certain depth in the
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
In situ activity and assimilation in nitrifying biofilms 1401
model biofilm. Correspondingly, this depth limit of activity
was reflected by the distribution of MAR-positive cells, and
by the depth distribution of uptake rates of [14C], as measured by beta microimaging. The integrated approach
allowed the calculation of in situ carbon yields and mass
balances, revealing effects like a ‘fuelling’ of coexisting
heterotrophic populations under autotrophic incubation
conditions. It facilitated the investigation of activity on the
community, population and single-cell level. Together with
FISH its application to environmental biofilms opens new
possibilities in studying the ecophysiology of single populations in situ and their impact on the overall function of
complex microbial communities.
Samples from experimental incubations were carefully
removed from the support and fixed with PFA accordingly
directly after incubation. Thereafter, the samples were transferred to cryoembedding medium (Jung tissue embedding
medium, Leica GmbH, Nussloch, Germany), incubated for
24 h at 4∞C, frozen on dry ice, and then sliced vertically on
a cryomicrotom (HM505E, Microm, Walldorf, Germany). The
sections of 20 mm thickness were immobilized on gelatincoated cover glasses (Lee et al., 1999). After drying the
embedding compound and traces of precipitated [14C]-bicarbonate within the sections were removed by gentle washing
with a 0.1-M glycine buffer solution, pH 3, followed by rinsing
in distilled water. After drying samples were stored at -18∞C
until further processing.
Experimental set-up
Experimental procedures
Nitrifying fluidized bed reactor
A conical 360-ml continuous upflow reactor, operated as
described previously (de Beer et al., 1993), served as a
source for nitrifying flocs. The reactor was fed with a medium
composed of NH4Cl (5.7 mM), Na2SO4 (0.5 mM), K2HPO4
(0.25 mM) and trace elements (MgCl2, 2.9 ¥ 10-2 mM; FeCl3,
1.8 ¥ 10-2 mM; MnCl2, 3.5 ¥ 10-3 mM; MoO42–, 1.4 ¥ 10-3 mM;
in 0.2 M HCl). The medium was recirculated at a rate of
1 ml s-1. Na2CO3 (0.6 M) was dosed for maintenance of a
constant pH (8.0), leading to a total carbonate concentration
of 2 mM. The system had been operated under stable conditions for more than 16 months before sampling.
Preparation of model biofilm
For every experiment fresh samples were taken from the
reactor by introducing a long tube from above. This ensured
a mixed vertical sample from the whole reactor column. Flocs
were broken down into smaller aggregates by vortexing, and
750 ml of the suspension were mixed with an equal amount
of low melting agarose (melting temperature 26∞C to 30∞C)
kept at 30∞C. The mixture was portioned in sterile cut-off
8 ¥ 4 mm lids (d ¥ h) from 2-ml Eppendorf tubes using a
syringe with an injection needle, and the suspension was
immediately solidified on ice. The model biofilm was kept
covered with liquid medium until experimental incubation.
Fluorescence in situ hybridization
Samples from the source reactor were fixed directly with
fresh paraformaldehyde (PFA) for 4 h as described earlier
(Amann et al., 1990). After washing in PBS and transfer to
PBS/EtOH, the samples were ultrasonicated with an amplitude of 126 mm for 3 ¥ 20 s. Aliquots of 6 ml per well were
spread on gelatin-coated microscopic slides with wells, dried
at room temperature, and subjected to FISH according to
earlier descriptions (Manz et al., 1992). Probes and
hybridization conditions are listed in Table 1. Populations
were quantified by microscopic counting on an epifluorescence microscope (Zeiss Axiophot II, Carl Zeiss, Jena,
Germany).
Model biofilms were attached by their support to the bottom
of a 100-ml wide-neck glass bottle with silicone grease. A
volume of 12 ml of the respective medium was added, and
the incubation chamber was closed by a lid with two holes.
A small curved glass capillary for aeration and convection of
the medium was inserted through the peripheral hole. The
opening of the capillary was placed directly above the
medium surface to avoid aerosol formation. A second central
opening in the lid was used to access the sample with
microsensors during incubation. The liquid medium was similar to the reactor feed, except that K2HPO4 was replaced by
Na2HPO4, and fixed amounts of NH4Cl, NaNO2 (Table 2) and
Na2CO3 were added. The buffer modification was necessary
because of cross-sensitivity of the NH 4+ microsensor to K+.
The pH was adjusted to 7.8 by addition of either acid or base.
Exact total carbonate concentration in the medium was determined with a total organic carbon analyser (TOC 5050 A,
Shimadzu Europe, Duisburg, Germany), and was
1.49 ± 0.08 mM (mean ± SD, n = 6 aliquots) in all incubations. Several experiments with different conditions were performed as shown in Table 2. In experiment D (decreased
oxygen concentration) argon was used for aeration.
Radiotracer incubation and microsensor measurements
Each experiment was started by adding 5.9 MBq of [14C]bicarbonate per 12 ml of medium. Incubation was performed
for a period of 8.8 h. Microprofiles were measured concomitantly with amperometric oxygen microsensors (Revsbech,
1989) and ion-selective microelectrodes for ammonium, and
either nitrite or nitrate (de Beer et al., 1997). Measurements
were started after 1 h of incubation to allow development of
a steady state (no change in the shape of microprofiles after
this time). Between 14 and 20 microprofiles were recorded
in each individual experiment at different lateral positions in
the model biofilm sample.
Microautoradiography and FISH
Vertical cryosections of the model biofilm from each experiment were hybridized with the different probes (Table 1) as
described above. The sections were subsequently covered
with a photographic emulsion for microautoradiography and
© 2005 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 7, 1392–1404
1402 A. Gieseke et al.
allowed to expose for approximately 7 days before development as described earlier (Andreasen and Nielsen, 1997).
The developed slides were dried and immediately inspected
under the microscope. Triplicates were analysed for each
incubation condition and probe.
Beta microimaging
Incorporation of [14C] was quantified by use of a beta emission microimager (Micro Imager, Biospace Mesures, Paris,
France). For that purpose, washed and dried cryosections
were covered with a scintillation foil and mounted in the
device. The working principle of the beta microimager is
described elsewhere (Lanièce et al., 1998). Shortly, scintillation signals are enhanced in an array of microphotomultipliers
with an individual diameter of 10 mm. Each signal event is
retranslated into an optical signal, detected via a CCD chip
and recorded on a PC. For each experimental condition 6
randomly chosen thin sections were scanned for a time interval of 45 min each. Within the counting interval, between
7500 and 17500 counts were obtained for each individual thin
section, with the exception of the negative control, where a
maximum of 2200 counts was obtained in 45 min. For determination of counting efficiency, a set of thin sections was first
scanned on the beta microimager. Then, the sections were
washed off by incubation in trichloroacetic acid at 95∞C for
30 min. Radioactivity in the liquid phase was subsequently
quantified by scintillation counting and compared with the
results from beta microimaging.
Rates of nitrification and [14C] incorporation
The microprofiles were used to calculate total areal conversion rates of oxygen, ammonium, nitrite and nitrate for the
whole model biofilm, and local volumetric consumption rates
of oxygen in different layers. For the areal conversion rates,
fluxes through the diffusive boundary layer (DBL) were calculated according to earlier descriptions (Schramm et al.,
1996). The molecular diffusion coefficients in water of
2.12 ¥ 10-5 cm2 s-1, 1.76 ¥ 10-5 cm2 s-1, 1.65 ¥ 10-5 cm2 s-1
and 1.71 ¥ 10-5 cm2 s-1 for oxygen, ammonium, nitrite and
nitrate, respectively, were taken from literature data (Broecker
and Peng, 1974; Li and Gregory, 1974), adjusted to the
incubation temperature of 20∞C. Local volumetric conversion
rates were obtained by a step-size calculation procedure for
each profile as previously described (de Beer and Stoodley,
1999). In case of local volumetric rate calculation within the
model biofilm, the effective diffusion coefficient in the matrix,
i.e. 0.75% agarose, was assumed to be 0.96 ¥ D0 (Beuling,
1998). The resulting mean rates were smoothed by adjacent
averaging to reduce noise effects.
Inorganic carbon incorporation data as measured by beta
microimaging was analysed with the analytical software
package provided by the manufacturer (Betavision, Micro
Imager, Biospace Mesures, Paris, France). After determination of the original surface in the 2D visualized data, counts
were obtained (as counts per minute – cpm) from individual
fields (50 mm height) along a vertical column with a width of
1.5–2.5 mm and a total length (along depth) of 4 mm (i.e. 80
fields). Counts along depth from randomly chosen thin sec-
tions (n = 6) were horizontally averaged for each depth interval (i.e. 50 mm). The resulting mean profiles of counts were
used to quantify the vertical distribution of [14C] incorporation
from inorganic carbon according to:
C = z · (d · v · A · t)-1
with C: [14C] incorporation rate (nmol cm-3 h-1); z: measured
radioactivity (cps); d: counting efficiency (cps Bq-1); v: voxel
size (cm3), given by field dimensions (5 ¥ 10-3 cm ¥ 0.15–
0.25 ¥ 10-1 cm), and section thickness (2 ¥ 10-3 cm); A:
molar radioactivity (Bq nmol-1); t: apparent local incubation
time in the experiment (h). The latter was calculated by subtracting the time needed for diffusional transport of tracer to
a specific depth of the model biofilm from total incubation
time. The time interval for a 50% tracer saturation for each
layer (compared with the medium) was calculated (Crank,
1975) with a diffusion coefficient for bicarbonate of 1.06 ¥
10-5 cm2 s-1 in freshwater at 20∞C (Li and Gregory, 1974).
Rates were corrected for unspecific uptake and bicarbonate
precipitates by subtracting results of the negative control from
each profile. Areal uptake rates were calculated by integration of local uptake rates over the whole depth of activity. For
the reason of comparability between experiments, we integrated the rates of the upper 2.5 mm in each section.
Yields and substrate partitioning were calculated based on
mass balances of e– eqs (Rittmann and McCarty, 2001).
Shortly, 8 e– eqs are released per 1 mol of the donor ammonium, or 6 per mol ammonium, and 2 per mol nitrite oxidized
respectively. These can be partitioned between the acceptor
for energy generation (oxygen), and the biomass (carbon).
The partitioning is reflected by the ratios fe and fs, referring
to the relative amounts of e– eqs from donor used for energy
generation (fe) and for assimilation of carbon (fs), that sum up
to unity. For carbon assimilation we assumed 4 e– eqs to be
necessary per mol carbon fixed as biomass (Rittmann and
McCarty, 2001).
Acknowledgements
The authors would like to thank Anja Eggers, Gaby Eickert
and Ines Schröder for preparation of oxygen microelectrodes,
and Tim Ferdelman for his help with the isotopic work. Henk
Jonkers is acknowledged for assistance in the carbonate
determination, and Peter Stief and three anonymous reviewers for a critical review of the manuscript.
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