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Applied and Environmental Microbiology, October 1998, p. 3731-3739, Vol. 64, No. 10
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Structural and Functional Dynamics of
Sulfate-Reducing Populations in Bacterial Biofilms
Cecilia M.
Santegoeds,*
Timothy G.
Ferdelman,
Gerard
Muyzer,
and
Dirk
de Beer
Max Planck Institute for Marine Microbiology,
D-28359 Bremen, Germany
Received 26 February 1998/Accepted 31 July 1998
 |
ABSTRACT |
We describe the combined application of microsensors and molecular
techniques to investigate the development of sulfate reduction and of
sulfate-reducing bacterial populations in an aerobic bacterial biofilm.
Microsensor measurements for oxygen showed that anaerobic zones
developed in the biofilm within 1 week and that oxygen was depleted in
the top 200 to 400 µm during all stages of biofilm development.
Sulfate reduction was first detected after 6 weeks of growth, although
favorable conditions for growth of sulfate-reducing bacteria (SRB) were
present from the first week. In situ hybridization with a 16S rRNA
probe for SRB revealed that sulfate reducers were present in high
numbers (approximately 108 SRB/ml) in all stages of
development, both in the oxic and anoxic zones of the biofilm.
Denaturing gradient gel electrophoresis (DGGE) showed that the genetic
diversity of the microbial community increased during the development
of the biofilm. Hybridization analysis of the DGGE profiles with
taxon-specific oligonucleotide probes showed that
Desulfobulbus and Desulfovibrio were the main sulfate-reducing bacteria in all biofilm samples as well as in the bulk
activated sludge. However, different Desulfobulbus and Desulfovibrio species were found in the 6th and 8th weeks
of incubation, respectively, coinciding with the development of sulfate
reduction. Our data indicate that not all SRB detected by molecular
analysis were sulfidogenically active in the biofilm.
| |
INTRODUCTION |
Although sulfate reduction is
thought to be an anaerobic process, sulfate-reducing bacteria (SRB) are
also important in aerobic environments if they can proliferate in
anaerobic zones. For example, in marine sediments (16, 17)
and in aerobic wastewater treatment systems (18, 20),
sulfate reduction accounts for up to 50% of the mineralization of
organic matter. Furthermore, sulfate reduction strongly stimulates
microbially enhanced corrosion of metals (5, 7). Therefore,
the detection of sulfate reducers and sulfate-reducing activity in
sediments, wastewater treatment plants, and fouling biofilms is of
great practical and scientific relevance. Conventional microbial
techniques based on selective culturing are of limited usefulness for
quantification and characterization of environmental populations, as it
is now well recognized that most strains do not grow in vitro, either
because cultivation media poorly resemble natural growth conditions or
because different strains of microorganisms are interdependent (2,
49). Techniques based on the analysis of bacterial DNA and RNA
may complement the conventional microbiological approach and nowadays
are routinely used to determine the presence and distribution of
individual bacterial species, including SRB, in complex communities
such as those in bacterial biofilms (1, 33-35).
So far, these studies of microbial communities, i.e., bacterial
biofilms, have mainly focused on the exploration of bacterial diversity
and on the detection of individual bacterial taxa by molecular
techniques. Studies relating community structure to community function
are scarce, partially because of difficulties in monitoring microbial
activities. Within biofilms, the convection of compounds is hindered,
and consequently mass transfer to the cells often limits conversion
rates. Because of this resistance to mass transfer, biofilms develop
various microenvironments, which differ from the bulk liquid (6,
10, 18). This complicates the interpretation of community
function analysis, because extrapolation of community behavior to that
of individual cells is impossible without knowledge of their
microenvironment.
The most direct way to study different microzonations in biofilms is
with microsensors
needle-shaped devices with a tip measuring from <1
to 100 µm which are sensitive for a specific compound. Due to their
small size, microsensor measurements cause minimal disturbance to the
system. With microsensors, microenvironments can be examined, and
microgradients can be measured. The measured gradients are a function
of local transport and conversion rates. Thus, if the transport process
(usually diffusion) is known, the spatial distribution of microbial
activity can be derived from the substrate profiles (38). An
important advantage of using microsensors is the capability to unravel
closed cycles, such as sulfate reduction coupled with sulfide
oxidation, within a biofilm or sediment (18). With these
systems, measurements of net substrate consumption or product excretion
lead to considerable underestimation of the actual processes within the
biofilms. Internal metabolic cycles can be hidden, although they play a
crucial role in the biofilm and will be reflected in the structure of
the microbial community.
Ramsing et al. (33) were the first to use both microsensors
and molecular techniques to study sulfate reduction in a
trickling-filter biofilm, which was used for treating municipal
wastewater. Recently, Schramm et al. (42) combined
microsensor and molecular techniques to study nitrification in a
trickling-filter biofilm. In both studies, a good correlation was found
between microbial conversions (sulfate reduction and nitrification) and
microbial population distributions within the biofilms.
The studies described above were performed with well-established and
mature biofilms. In this work, we studied the transients of sulfate
reduction, using microsensors, and followed the successional changes in
the composition of microbial species, using molecular techniques, with
a developing, multispecies bacterial biofilm. This topic is relevant,
as biofilms are often subjected to changing conditions and sloughing is
followed by recolonization. The aim of our study was to assess how
closely species composition reflects activity in a biofilm with
gradually changing microenvironments.
A biofilm developing in an aerobic wastewater treatment plant was used
as a model for biofilms growing in aerobic sulfate-containing waters
with high organic loads. Microsensors with high spatial resolution were
used to measure oxygen and hydrogen sulfide profiles and to infer
aerobic respiration and sulfate-reducing activities. Molecular
techniques for studying SRB were as follows: denaturing gradient gel
electrophoresis (DGGE) analysis of PCR-amplified 16S ribosomal DNA
(rDNA) fragments to determine the complexity of the microbial
community in the biofilm and to monitor its behavior over time,
hybridization analysis of these DGGE profiles with group-specific
oligonucleotide probes to identify SRB, and in situ hybridization with
thin sections of the biofilm to quantify the SRB and determine their
spatial distribution within the biofilm.
| |
MATERIALS AND METHODS |
Biofilm growth and sampling.
Oxygen-impermeable plastic foil
(Ril-O-Ten 80/100X; Otto Nielsen Emballage AS, Lyngby, Denmark) was
submerged as a substratum for biofilm growth in the activated-sludge
basin (first aerated stage) of the Seehausen municipal wastewater
treatment plant in Bremen, Germany, during spring 1996. The temperature
in the basin during the experiment varied between 17 and 20°C. In the
bulk liquid, the concentrations of O2 and
SO42
were 1.8 to 2.4 mg/liter (55 to 75 µM)
and 75 to 115 mg/liter (800 to 1,200 µM), respectively.
NO3 was absent. The biological oxygen demand
was approximately 350 mg/liter (10 mM). Samples of the foil (ca. 6 by
12 cm each) on which the bacterial biofilm was growing were taken
weekly over a period of 12 weeks. While submerged in wastewater,
samples were transferred to the laboratory and used within 1 h for
microelectrode measurements. Replicate samples were frozen at 
20°C
for molecular analysis.
Biofilm fixation and slicing.
Biofilm samples for in situ
hybridization analysis were fixed by incubation in paraformaldehyde
(4% [wt/vol] in phosphate-buffered saline) at 4°C for 1 h and
washed subsequently in phosphate-buffered saline. After fixation,
samples were embedded in embedding medium (OCT compound; Sakura Finetek
USA, Torrance, Calif.) and frozen above evaporating liquid nitrogen.
Thin sectioning of the biofilm was performed with a cryomicrotome
(Microm model HM 505 E) at 18°C. Vertical slices about 10 µm
thick were collected on gelatin-coated microscopic slides. The slides
were air dried and dehydrated in a series of increasing concentrations
of ethanol (50, 80, and 96% [vol/vol]).
Biofilm thickness measurement.
Biofilm thickness was
determined by positioning a thin glass needle mounted on a
micromanipulator on the surface of the biofilm. The needle was moved
down until it touched the substratum, which was detected by bending the
needle, viewed through a dissection microscope. Biofilm thickness was
inferred from micromanipulator readings.
Microsensor measurements.
Microsensor measurements were
performed at room temperature (20 to 23°C) in a flow cell with
aeration and circulation of artificial wastewater containing the
following ingredients: 50 mM KH2PO4, 400 µM
K2HPO4, 760 µM
(NH4)2SO4, 41 µM
MgSO4 · 7H2O, and 200 µM Na-acetate at
pH 7.5 (the last ingredient was added as an additional carbon source to
the bound carbon inside the biofilm to ensure that oxygen remained the
limiting substrate). The oxygen concentration in the flow cell was kept
the same as that in the activated-sludge basin, i.e., approximately 70 µM O2, by bubbling with air and nitrogen. The oxygen
concentration in the flow cell was continuously monitored with an
oxygen microsensor. Microsensors, mounted on micromanipulators, were
positioned on the surface of the biofilm, which was viewed through a
dissection microscope. Profiles were recorded by penetrating the
biofilm with the microsensor in increments of 50 or 100 µm and using
a micromanipulator.
Microsensors were used to measure oxygen (37), hydrogen
sulfide (19), and pH (38). The oxygen and pH
electrodes were calibrated as described before (37, 38). The
H2S microsensor was calibrated by measuring the signal in
dilution series of a standard solution (sulfide dissolved in artificial
wastewater with a constant pH, flushed with nitrogen to avoid oxidation
of the sulfide) (18). The concentration of total dissolved
sulfide (H2S plus HS plus S2)
in the dilution series was determined by a spectrophotometric method
(8). The total dissolved sulfide measured in the biofilm was
calculated by using the slope and intercept of the calibration curves.
The sulfide electrode had a linear response to H2S
concentrations of up to 1,000 µM. The detection limit of the
microsensors varied between 1 and 3 µM total sulfide. No pH
correction was necessary, since the sensor was calibrated at the same
pH as the sample medium and the buffer prevented the development of pH
gradients inside the biofilm. The diffusive fluxes were calculated by
using Fick's first law, J = D × (
c/x), where J is flux (in micromoles per square meter
per second), D is diffusion coefficient (in square meters
per second), and c/x is concentration gradient (in micromoles per
cubic meter). Diffusion coefficients inside the biofilm were assumed to
equal the molecular diffusion coefficients (7). Values of
2.12 × 105 cm2/s for oxygen
(3) and 1.39 × 105 cm2/s for
total sulfide (18) at 20°C were used.
Total iron determination.
Ten milliliters of 0.5 M HCl and
0.5 g of dithionite were added to the biofilm sample to extract Fe
oxides, FeS, and FeCO3 (46). The total iron
concentration in this extract was determined spectrophotometrically at
562 nm after reaction of the extract with Ferrozine solution (1%
[wt/vol] hydroxylammoniumchloride and 10% [wt/vol] Ferrozine in 50 mM N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid buffer, pH 7.0) (46). The detection limit was 1 µM
total iron.
Nitrate determination.
The nitrate concentration in the bulk
liquid of the activated-sludge basin was measured spectroscopically
with Spectroquant model 14773 (E. Merck, Darmstadt, Germany). The
detection limit was 16 µM (1 mg/liter)
NO3.
Net sulfate reduction measurement with radiotracers.
Net
sulfate reduction (sulfate reduction minus sulfide oxidation) was
determined by measuring the accumulation of reduced sulfur compounds in
the 7-week-old biofilm. Carrier-free
35SO42 (2.5 MBq; Amersham) was
added to the medium in the microsensor setup after a blank sample of
the biofilm was taken for a reference. During the 30-h incubation
period, samples were taken from the biofilm (3 to 4 cm2
each) and from the medium (4 ml each) at intervals of at least 2.5 h to determine total radioactivity and sulfate concentration. The
entire sample (foil plus adhering biofilm) was preserved in a 20%
(wt/vol) Zn-acetate solution. The fraction of reduced
35SO42 per square centimeter of
plastic foil (i.e., radioactive H2S, FeS, S0,
and FeS2) was determined with the foil and biofilm by the
hot acidic Cr-II procedure as described by Fossing and Jørgensen
(13). The net sulfate reduction was expressed as an areal
rate, i.e., moles of reduced 35S per square meter per
second for comparison with microsensor flux measurements.
Nucleic acid extraction and PCR amplification.
DNA was
extracted from the biofilm samples by a combined freeze-thaw (freezing
three times in liquid nitrogen and thawing at 37°C) and hot
phenol-chloroform-isoamyl alcohol treatment (45). The
ribosomal DNA was enzymatically amplified as described by Muyzer et al.
(29) with either the universal primer 907R and the bacterial
primer GM5F with a GC clamp or the universal primer 907R and the SRB385
primer (1) with a GC clamp, targeting SRB of the delta
subdivision as well as some other bacteria (Table 1). A hot-start, touch-down PCR program
was used for all amplifications to minimize nonspecific amplification
(29). The PCR mixture (100 µl) contained 50 pmol of each
primer, 25 nmol of each of the four deoxynucleoside triphosphates, 300 µg of bovine serum albumin, 10 µl of 10× PCR buffer (HT
Biotechnology Ltd.), and 10 to 20 ng of template DNA.
DGGE analysis of 16S rDNA fragments.
DGGE was performed with
the D-Gene system (Bio-Rad) and the following ingredients and
conditions: 1× TAE (40 mM Tris, 20 mM acetic acid, and 1 mM EDTA at pH
8.3), 1-mm-thick gels, and a denaturant gradient containing 35 to 65%
urea-formamide at 60°C, with 100 V constantly for 17 h version
(29, 30). DGGE gels were photographed on a UV
transillumination table (302 nm) with a Polaroid camera. Photos were
scanned with the software program Fotolook version 2.05 (Agfa), and
inverse images were prepared with Photoshop version 4.0 (Adobe).
Blotting and hybridization analysis of DGGE gels.
DGGE gels
were blotted onto nylon membranes (Hybond+; Amersham) as described by
Muyzer et al. (29). Hybridization analysis was performed
with probes specific for different groups of sulfate reducers (Table
1). Probe 660 is specific for Desulfobulbus species; probe
687 targets Desulfovibrio species as well as some members of
the genera Geobacter, Desulfomonas,
Desulfuromonas, Desulfomicrobium, Bilophila, and Pelobacter; and probe 804 targets
Desulfobacter, Desulfobacterium,
Desulfosarcina, Desulfococcus, and
Desulfobotulus species (11). The probes were end
labeled with radioactive [
-32P]ATP (New England
Biolabs). The hybridization buffer used was described by
Martinez-Picado and Blanch (25) and contained the following:
10× Denhardt solution, 4× SSC (1× SSC is 0.15 M NaCl and 0.015 M
sodium citrate at pH 7.0), 0.1% (wt/vol) sodium dodecyl sulfate, 2 mM
EDTA, and 50 µg of salmon sperm DNA per ml. Membrane blots were
prehybridized for 3 to 4 h at 40°C before the radioactively labeled probe was added. Hybridization was performed for about 17 h at 40°C. Thereafter, the membranes were washed twice at 40°C for
30 min with 2× SSC-0.1% (wt/vol) sodium dodecyl sulfate. To eliminate nonspecific binding, the membranes were washed two more times
for 15 min at the appropriate dissociation temperature (56°C for
probe 660, 48°C for probe 687, and 52°C for probe 804), determined by the method described by Raskin et al. (36). The
hybridized membranes were dried and exposed for 1 to 2 days to a
PhosphoImager screen. These screens were further analyzed with the
PhosphoImager and the program ImageQuant (Molecular Dynamics, Inc.).
In situ hybridization.
The protocol described by Manz et al.
(22) was used for fluorescence in situ hybridization (FISH)
of the biofilm slices, with probe NON338 as a negative control and
probe SRB385 for the detection of sulfate reducers (Table 1). Probes
NSO1225, NIT3, and NSR1156 were used to estimate the amount of
nitrifying bacteria in the biofilm. The probes were synthesized and
labeled with the fluorochrome CY3 (Interactiva GmbH, Ulm, Germany). The
hybridization buffer contained 0.9 M NaCl, the percentage (vol/vol) of
formamide shown in Table 1, 20 mM Tris-HCl (pH 7.4), and 0.01%
(wt/vol) sodium dodecyl sulfate. The probe concentration was 5 ng/µl.
Hybridization was performed for 1 to 2 h at 46°C. The biofilm
was washed at 48°C for 15 min in a washing buffer containing 20 mM
Tris-HCl (pH 7.4), 5 mM EDTA, the NaCl concentration indicated in Table 1, and 0.01% (wt/vol) sodium dodecyl sulfate. The specimens were examined with an Axioplan epifluorescence microscope (Carl Zeiss, Oberkochen, Germany). CY3-stained cells were counted in 20 to 22 areas of 125 by 625 µm2, both at the bottom and at the
top of the biofilm. The density of the cells in the biofilm was too
high to allow enumeration of the total number stained with
4',6-diamidino-2-phenylindole (DAPI). Therefore, the number of stained
bacteria was expressed per unit of volume of biofilm.
| |
RESULTS |
Biofilm growth and development process.
Within 1 week, a
patchy biofilm ca. 400 µm thick developed on the plastic foil in the
activated-sludge basin (Fig. 1). The thickness of the biofilm increased until the eighth week, after which
it remained more or less constant at 1,000 to 1,200 µm.
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FIG. 1.
Development of biofilm thickness (black bars) and fluxes
of oxygen into the biofilm (gray bars) and sulfide into the oxic zone
(white bars) calculated from the microprofiles.
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Oxygen and sulfide profiles measured at different stages of biofilm
development are shown in Fig.
2. The
graphs show mean
values of four to six profiles measured at different
positions
in the biofilm. pH values measured in the biofilm were within
0.05 U of the pH in the bulk liquid (data not shown) because of
the
buffering capacity of the medium used for measuring the profiles.
Within 1 week, the biofilm developed anaerobic zones, which grew
thicker during the following weeks. Oxygen was depleted within
the top
200 to 400 µm in all stages of biofilm development (except
in some
profiles of the 1-week-old biofilm). Although the anaerobic
environment
in the biofilm would have allowed sulfate reduction,
as there was
enough substrate and sulfate available, no sulfide
production was
measured until the sixth week.
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FIG. 2.
Oxygen ( ) and total sulfide ( ) microprofiles in
1-, 5-, 6-, 8-, 10- and 12-week-old biofilms. Biofilm thickness is
indicated by the shaded areas. The biofilm surface is at a depth of 0 mm, the water phase is indicated by negative values, and the biofilm is
indicated by positive values.
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In the sixth week, a total sulfide concentration of 40 µM was
detected, which increased to approximately 100 µM in the 10-
and
12-week-old biofilms (Fig.
2). No sulfide diffused out of
the biofilm,
and oxygen and sulfide profiles overlapped, indicating
that all sulfide
produced by sulfate reduction was oxidized in
the aerobic part of the
biofilm.
Iron, which could have bound the sulfide, was not detected either in
biofilm samples (measured at weeks 5 and 11) or in the
activated sludge
(data not shown).
Sulfate reduction rates measured by a radiotracer method with
35SO
42 showed much lower values
(approximately 0.017 µmol/m
2 · s
1)
than the rates calculated with microsensor data (Fig.
1). Also,
the
radioactive tracer technique showed that no reduced S compounds
accumulated inside the biofilm or bulk water.
Community development.
To determine how the structure of the
community developed with time and to identify the presence of SRB, DGGE
of PCR-amplified DNA fragments and hybridization analysis of these DGGE
profiles with group-specific probes were performed. Figure
3 shows the DGGE pattern of 16S rRNA gene
fragments obtained after PCR amplification with a primer pair which
amplifies members of the domain Bacteria. Figure
4 shows a graph of the total number of
bands as well as new bands appearing relative to the activated sludge
and the total number of bands remaining from the activated sludge.
Nearly all bands obtained from the activated sludge remained in the
biofilm samples. However, new bands also appeared during the maturation of the biofilm.
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FIG. 3.
DGGE analysis of 16S rDNA fragments obtained after PCR
amplification with primer pair GM5F-907R and genomic DNA from bacterial
biofilm samples taken at different time points. From left to right:
sample taken directly from the activated sludge (t = 0 weeks) and samples taken from 1-, 3-, 5-, 6-, 8-, 10-, and 12-week-old
biofilms (as indicated above the lanes).
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FIG. 4.
Graphical representation of the number of bands in the
denaturing gradient gel shown in Fig. 3. Total number of DGGE bands,
black bars; total bands remaining from week 0 (activated sludge), gray
bars; new bands after week 0, white bars.
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Detection of SRB in DGGE profiles.
DGGE gels with DNA
fragments amplified with primers for members of the domain
Bacteria were blotted and hybridized with probes specific
for different groups of SRB (probes 660, 687, and 804). No
hybridization was observed, probably due to the small ratio of each SRB
strain to the total number of bacteria in the biofilm.
To increase the sensitivity of SRB detection in our samples, we used
the SRB385 sequence with a GC clamp as a forward primer,
which targets
sulfate reducers as well as some other delta proteobacteria
and
gram-positive bacteria, and the 907R sequence as the reverse
primer. In
this way, we excluded the amplification of 16S rDNA
fragments of most
of the
Bacteria and enriched PCR products from
SRB. These
DGGE profiles were subsequently blotted and hybridized
with
radioactively labeled oligonucleotide probes specific for
different
groups of SRB.
In these SRB-enriched DGGE gels, clear hybridization was obtained with
probes 660 and 687 (Fig.
5B and
6B) but not with 804
(data not shown).
This result indicates the presence of
Desulfobulbus (probe
660) and possibly
Desulfovibrio (probe 687, which also
targets members of the genera
Geobacter,
Desulfomonas,
Desulfuromonas,
Desulfomicrobium,
Bilophila, and
Pelobacter) and the absence of
all species targeted by probe
804 (
Desulfobacter,
Desulfobacterium,
Desulfosarcina,
Desulfococcus, and
Desulfobotulus) in all stages
of biofilm development and in
activated sludge. Furthermore, hybridization
analysis of the DGGE
profiles with probe 660 showed the initial
presence of two
Desulfobulbus strains and the appearance of another
strain
after week 6 (compare lanes 2 to 5, Fig.
5B, with lanes
6 to 9),
coinciding with the first detected sulfide production
by microsensors.
The result for probe 687 was similar; initially
two
Desulfovibrio strains were present, and a third appeared
during
week 8 (compare lanes 2 to 6, Fig.
6B, with lanes 7 to 9).
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FIG. 5.
(A) DGGE analysis of PCR-amplified 16S rDNA fragments
obtained with primers SRB385 and 907R and bacterial genomic DNA from
biofilm samples taken at different time points. From left to right:
marker 1 (M1), Desulfonema limicola (Dnl) and
Desulfococcus multivorans (Dcm); sample taken from the
activated sludge (t = 0 weeks); samples from 1-, 3-, 5-, 6-, 8-, 10-, and 12-week-old biofilms (as indicated above the
lanes); marker 2 (M2), Desulfobulbus propionicus (Dbp,
positive control for probe 660), Desulfomicrobium baculatum
and Desulfovibrio salexigens (Dmb and Dvs, positive controls
for probe 687). (B) Hybridization results of the same DGGE pattern
hybridized with probe 660, which is specific for
Desulfobulbus species. The bands in the lower parts of the
gels are single-stranded DNA and should be disregarded.
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FIG. 6.
(A) DGGE analysis of PCR-amplified 16S rDNA fragments
obtained with primer set SRB385 and 907R. From left to right: marker 1 (M1), Desulfonema limicola (Dnl) and Desulfococcus
multivorans (Dcm); sample taken from the activated sludge
(t = 0 weeks); samples from 1-, 3-, 5-, 6-, 8-, 10-, and 12-week-old biofilms (as indicated above the lanes); marker 2 (M2),
Desulfobulbus propionicus (Dbp, positive control for probe
660), Desulfomicrobium baculatum, and Desulfovibrio
salexigens (Dmb and Dvs, positive controls for probe 687). (B)
Hybridization results of the same DGGE pattern hybridized with probe
687, which targets Desulfovibrio species. The bands in the
lower parts of the gels are single-stranded DNA and should be
disregarded.
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The DGGE patterns obtained with primers targeting sulfate reducers
(Fig.
5 and
6) show less variation with time than those
obtained with
bacterial primers (Fig.
3). This trend can be seen
more clearly in Fig.
7, which contains a graph of the number
of
total, new, and remaining DGGE bands. The number of bands
representing
SRB varies only between 14 and 21, whereas Fig.
4 shows a
variation
of 14 to 37 bands from all bacteria during biofilm
development.
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FIG. 7.
Graphical representation of the number of bands in the
denaturing gel shown in Fig. 5A and 6A. Total number of DGGE bands,
black bars; total bands remaining from week 0 (activated sludge), gray
bars; new bands, white bars.
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In situ hybridization of sulfate-reducing and nitrifying
bacteria.
In situ hybridization with the SRB385 probe of thin
sections of biofilms taken at weeks 2 and 11 stained
Vibrio-shaped bacteria and short fat rods. Both morphotypes,
resembling Desulfovibrio and Desulfobulbus cells,
were present over the whole thickness of the biofilm. In the 2-week-old
biofilm, the number of SRB was (64 ± 23 [mean ± standard
deviation]) × 106 SRB/ml on the surface and 114 ± 33 × 106 SRB/ml at the bottom of the biofilm. In the
11-week-old biofilm, the SRB varied from 201 ± 78 × 106 SRB/ml to (358 ± 127) × 106
SRB/ml on the surface and at the bottom of the biofilm,
respectively.
In situ hybridization with probes specific for nitrifying bacteria
revealed the presence of ammonia-oxidizing bacteria (30
× 10
6 to 90 × 10
6 cells per ml) and
nitrite-oxidizing bacteria,
Nitrobacter species
(0.6 × 10
6 to 1 × 10
6 cells per ml)
and
Nitrospira species (50 × 10
6 to
100 × 10
6 cells per ml), respectively, in the 2- and
6-week-old biofilms.
| |
DISCUSSION |
Process and biofilm development.
The thickness and community
characteristics of the biofilm developed gradually during 12 weeks
without large-scale sloughing. The biofilm had a heterogeneous
structure during the whole experimental period, as was reported in
other studies (10, 26), explaining the variations in the
amounts of oxygen and sulfide (Fig. 1). Heterogeneity may increase
surface area exchange, therefore allowing more efficient transport of
substrates and products. In determining the fluxes, we assumed that
diffusion was the only transport mechanism; thus, the fluxes might be
underestimated. However, the fluxes measured in this study are close to
those calculated by Kühl and Jørgensen (18) for a
trickling-filter biofilm.
The microprofiles (Fig.
2) showed that no sulfide diffused out of the
biofilm. Since iron was absent, iron sulfide precipitation
can be
excluded. The low value for the sulfate reduction rate,
determined with
radiotracers, compared to the rates calculated
from microsensor data,
can be explained only by cycling between
reduced and oxidized
35S compounds. Most of the
35S
2
formed is rapidly oxidized to
35SO
42 or other oxidized
35S compounds, which are not detectable by our radiotracer
method
(H
2S, S
0, FeS
2, and FeS).
Examination of the specific activity of the
reduced sulfur pool during
the course of the experiment showed
that the pool rapidly became
equilibrated (within 3 h) with the
initially labeled sulfate. This
result indicates that a small
reduced sulfur pool (sulfide) with a high
turnover rate is present
in the biofilm.
We concluded that all sulfide, produced by sulfate reduction in the
anoxic zone, was oxidized in the oxic layer in the biofilm.
Thus, at
the point of the steepest gradient, the sulfide flux
in the zone
adjacent to the aerobic zone can be used as a conservative
measure of
SRB activity. Diffusive fluxes of oxygen into the biofilm
and sulfide
from the anaerobic zone into the aerobic zone, calculated
from the
microprofiles, are given in Fig.
1. Sulfide fluxes ranged
from 15% in
week 6 to 35% in week 11 of the oxygen fluxes (Fig.
1). Since
oxidation of 1 mol of sulfide to sulfate requires 2
mol of oxygen, a
large part (up to 70%) of the oxygen diffusing
into the biofilm was
used for the reoxidation of sulfide. Thus,
sulfate reduction may be as
important as aerobic mineralization
in the biofilm. Since the biomass
yield by sulfate reduction is
much lower than by aerobic mineralization
(
21), rates of biomass
production in wastewater treatment
plants based on biofilms may
be affected significantly. The turnover of
the sulfide pool in
the biofilm, calculated by dividing the pool size
by the fluxes
(both determined from microsensor profiles), lasted only
minutes.
Consequently, sulfate reduction rates in biofilms can be
reliably
measured in situ only by microsensors.
The development of anaerobic zones in the biofilm created an optimal
environment for sulfate reducers. However, sulfide production
could not
be measured during the first 6 weeks of biofilm development.
In theory,
sulfide could have been oxidized anaerobically, e.g.,
by
Fe
3+ or NO
3 as an electron
acceptor. However, neither Fe
3+ nor
NO
3 was present in the bulk liquid of the
activated-sludge basin.
In situ hybridization showed the presence of
ammonia and nitrite
oxidizers in the 2- and 6-week-old biofilms;
consequently, some
nitrate could have been formed by nitrification
inside the biofilm.
However, even if sulfide reduction was coupled with
anaerobic
sulfide oxidation in the initial stages of biofilm
development,
some sulfide should have been detected by the sensitive
sulfide
microsensor. We therefore conclude that no sulfate reduction
occurred
in the biofilm during the first 6 weeks. Similar lag phases in
developing biofilms have been detected for sulfate-reducing
(
48)
and other slow-growing bacteria, such as nitrifiers
(
47) and
methanogens (
14). Slow adaptation of the
sulfate-reducing population
to the biofilm microenvironment might
explain the lag phase in
sulfate reduction that was found in our study.
Community development.
Comparative DGGE analysis of PCR
products obtained with primers for the 16S rDNA of all bacteria showed
that the species composition of the biofilm changed within 1 week from
that of the activated sludge (Fig. 3). The original strains remained,
but new bands appeared each week (Fig. 4), leading to a more complex
bacterial community. Exceptions were week 8 and, to a lesser degree,
week 3, when fewer bands were detected. These exceptions might have been caused by a lower amount of DNA being loaded on the gel. Our
results show that biofilms can harbor a much wider range of microbial
species than bulk liquid. The increase in bacterial diversity, shown by
the increasing number of DGGE bands, was the result of the development
of different microhabitats inside the biofilm. Due to resistance to
mass transfer and conversions, microzonations develop within the
biofilm, providing a broader range of niches for bacteria with
different physiological characteristics.
DGGE analysis of PCR products obtained with primers specific for
sulfate reducers, i.e., primer pair SRB385F/907R, showed
that the
community of delta proteobacteria, including the sulfate
reducers, was
more stable during biofilm development than the
total community
(compare Fig.
4 and
7).
Desulfobulbus and
Desulfovibrio species were present from the beginning.
Additional populations
of
Desulfobulbus and of
Desulfovibrio appeared, respectively,
after 6 and 8 weeks of
incubation. We speculate that the additional
strains from weeks 6 and 8 are responsible for the sulfide production
in the biofilm.
The question of the metabolic activity of the species identified as
SRB, present in the first 5 weeks, is unresolved. Our
FISH data
indicate that a significant portion of the SRB are present
and
proliferate in the oxic part of the biofilm. The ability of
Desulfobulbus and
Desulfovibrio species to
respire oxygen (
12,
24) could explain their presence in
activated sludge and the
initial biofilm. As the activated-sludge basin
is aerated, other
nonrespiring sulfate reducers may be unable to
survive in this
aerobic environment. Also, the finding of Teske et al.
(
45),
that
Desulfobulbus and
Desulfovibrio species are the main SRB
in the aerobic layers
of a stratified fjord, underlines their
ability to survive (and maybe
thrive) in the presence of oxygen.
Alternatively, SRB could use a
different metabolic pathway. Some
Desulfobulbus and
Desulfovibrio species can utilize nitrate instead
of sulfate
as a terminal electron acceptor (
27,
43,
52).
There is
conflicting evidence about whether or not nitrate is
used in the
presence of sulfate. It has been reported that nitrate
can suppress
sulfate reduction (
43) and, conversely, that sulfate
inhibits the use of nitrate as a terminal electron acceptor
(
9).
However, no nitrate was found in the bulk liquid of the
activated-sludge
basin. Another metabolic role of SRB could be the
fermentation
of organics, such as fumarate or malate (
51).
With in situ hybridization, we found 10
7 to 10
8
SRB/ml of biofilm volume. Considering the total cell density of
10
10 to 10
11 cells per ml of biofilm reported
by others (
7,
31), we estimate
that the relative percentage
of bacteria that are SRB is less
than 1%. This, however, contradicts
our finding that a large part
of the oxygen diffusing into the biofilm
is used for oxidation
of the formed sulfide and that sulfate reduction,
therefore, plays
an important role in the biofilm. An explanation for
this discrepancy
is that either the SRB are extremely active or our
FISH technique
underestimates the number of SRB. From the
H
2S fluxes and the
number of SRB (determined by FISH), the
specific sulfate reduction
rate (moles of
SO
42 cell
1 day
1)
was calculated to be 70 × 10
15 mol of
SO
42 cell
1 day
1
(at week 11). This value is in the upper range of the specific
sulfate
reduction rates reported by Jørgensen (
15).
By using the molecular techniques described in this study, we were able
to follow community development and to detect different
groups of SRB
in complex biofilms with many species. This would
be difficult to
achieve with conventional cultivation techniques.
However, molecular
techniques also have biases and limitations.
First, PCR amplification
is not quantitative, as preferential
amplification can occur (
39,
44). Therefore, band intensities
cannot be extrapolated to
indicate the abundance of a particular
bacterial population. Second,
the number of DGGE bands is dependent
on the amount of DNA that is
applied to the gel as well as on
the number of different DNA fragments.
Third, some of the oligonucleotides,
i.e., SRB385 and probe 687, which
were used in this study, are
not as specific as originally described.
However, we used only
primer SRB385 in our PCRs to exclude most non-SRB
and to enrich
the SRB populations. Probe SRB385 was also used for in
situ hybridization
analysis. For this application, the specificity of
the probe is
increased, as the gram-positive cells, targeted by SRB385,
are
not accessible to the oligonucleotide when paraformaldehyde is
used
as the fixative (
40). These biases demonstrate the
importance
of combining different molecular methods and comparing them
with
activity measurements.
Relating community analysis obtained by molecular techniques to
processes occurring within microbial consortia is of great
practical
relevance. However, the molecular techniques currently
available, and
used in this study, seem insufficient to accurately
predict the
behavior of a gradually changing but complex microbial
community. More
specific techniques, perhaps involving the detection
or expression of
specific genes, are needed so that molecular
techniques can be used as
reliable diagnostic tools. Further analysis
of the biofilm samples,
nitrate microsensor measurements, and
in situ hybridization with more
specific probes (
23) might provide
a more detailed picture
of the abundance, distribution, and functioning
of the SRB groups.
| |
ACKNOWLEDGMENTS |
This work was financially supported by the Körber
Foundation (Hamburg, Germany) and the Max Planck Society
(München, Germany).
We thank Dror Minz and Kerstin Sahm for their help with the
hybridization analysis and Td determination;
Ronnie Glud for his help with the radiotracer experiment; Susanne
Klöser for the total iron detection; and Gaby Eickert, Anja
Eggers, and Vera Hübner for technical assistance with the
microsensors.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Max Planck
Institute for Marine Microbiology, Celsiusstr. 1, D-28359 Bremen,
Germany. Phone: 49-421-2028-838. Fax: 49-421-2028-580. E-mail: ssantego{at}mpi-bremen.de.
Present address: Netherlands Institute for Sea Research (NIOZ),
1790 AB Den Burg, The Netherlands.
| |
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Abstract
Introduction
