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Applied and Environmental Microbiology, March 1999, p. 1289-1297, Vol. 65, No. 3
Lehrstuhl für Mikrobiologie, Technische
Universität München, D-80290 Munich,
Germany,1 and Environmental
Engineering Laboratory,
Received 19 October 1998/Accepted 8 December 1998
A new microscopic method for simultaneously determining in situ the
identities, activities, and specific substrate uptake profiles
of individual bacterial cells within complex microbial communities was
developed by combining fluorescent in situ hybridization (FISH)
performed with rRNA-targeted oligonucleotide probes and microautoradiography. This method was evaluated by using defined artificial mixtures of Escherichia coli and
Herpetosiphon aurantiacus under aerobic incubation
conditions with added [3H]glucose. Subsequently, we were
able to demonstrate the potential of this method by visualizing the
uptake of organic and inorganic radiolabeled substrates
([14C]acetate, [14C]butyrate,
[14C]bicarbonate, and 33Pi) in
probe-defined populations from complex activated sludge microbial
communities by using aerobic incubation conditions and anaerobic
incubation conditions (with and without nitrate). For both defined
cell mixtures and activated sludge, the method proved to be useful for
simultaneous identification and analysis of the uptake of labeled
substrates under the different experimental conditions used. Optimal
results were obtained when fluorescently labeled oligonucleotides
were applied prior to the microautoradiographic developing procedure.
For single-cell resolution of FISH and microautoradiographic signals
within activated sludge flocs, cryosectioned sample material was
examined with a confocal laser scanning microscope. The combination of
in situ rRNA hybridization techniques, cryosectioning,
microautoradiography, and confocal laser scanning microscopy provides a
unique opportunity for obtaining cultivation-independent insights into
the structure and function of bacterial communities.
In most natural and engineered
systems offering a sufficient nutrient supply, microorganisms grow as
spatially organized, matrix-enclosed, multispecies communities in
biofilms, aggregates, or flocs rather than as single planktonic cells
(11). When studying these systems, microbial ecologists
attempt to determine population structure and dynamics, as well as the
spatial distribution of species and the function of species within
communities which are characterized both by various cell-cell
interactions (13) and by pronounced architectural and
chemical heterogeneity (7, 10, 33). Since a rigorous biomass
disaggregation step is a common feature of all traditional
cultivation-based microbiological approaches used for analysis of
complex microbial communities, spatial information is lost. In
addition, biases introduced by conventional cultivation methods result
in pronounced population shifts in the community structure and lead to
a failure to detect dominant bacterial community members (50, 51,
56). Determining the physiological and biochemical properties of
a single species in a laboratory pure culture may also bias the
phenotype since gene expression is strongly influenced by environmental
constraints and the growth mode of the cells (12). Since
isolated strains are known to adapt genetically to the
environmental conditions which they encounter, they may also not be
genetically representative of their environmental counterparts
(8).
With the advent of rRNA in situ hybridization techniques in microbial
ecology, cultivation-independent quantitative examination of the
structure and dynamics of complex microbial communities became possible
(for a review see reference 2). Confocal laser scanning microscopic detection of signals conferred by a variety of
fluorescent strains or fluorescently labeled RNA-targeted nucleic acid
probes also allows analysis of the architecture of bacterial aggregates
(for a recent review see reference 28) and the
spatial arrangement of bacterial species within their habitats (4, 16, 18, 35, 41, 50-54).
Keeping in mind the physiological versatility of many prokaryotes, it
is apparent that identification of a bacterial species in situ does not
provide much information about its function in its habitat, except for
members of a few highly physiologically restricted bacterial lineages
like the ammonia oxidizers belonging to the beta subclass of the
Proteobacteria. While in situ identification of uncultured
microorganisms may allow assumptions concerning possible physiological
characteristics if known physiological properties of the most closely
phylogenetically related species which can be cultivated are taken into
account (26), a more encompassing in situ analytical
approach that yields information about the identity and function of a
cell in its environmental niche would be desirable. Such information
not only would allow a more detailed understanding of the ecology of
complex microbial communities but also might provide information which
could be used for the development of new, more appropriate enrichment
and isolation methods for molecularly identified but uncultured prokaryotes.
Recently, an elegant combination of fluorescent
oligonucleotide probing and microelectrode measurements has
been used for structure-function analysis in microbial ecology. The
relationship between chemical gradients and the stratification of
sulfate-reducing (38) and nitrifying bacteria (41,
42) in trickling filters and a fluidized bed reactor has been
examined by this method. Microelectrodes, however, measure chemical
profiles only at a single point and consequently do not describe
two-dimensional or three-dimensional chemical profiles in heterogeneous
communities. The spatial resolution provided by microsensors is also
above the single-cell level. Other attempts have been made to develop methods for functionally based microbial community analysis on a
single-cell level. One approach has focused on the detection of
specific gene expression in microbial cells by exploiting fusions of a
gene and the coding sequence of a reporter protein (19, 27, 30,
45, 57). While techniques involving reporter proteins provide
valuable information, they are limited in their applicability to
microbial ecology as they require (i) the availability of genetic tools
for the microorganism being studied and (ii) the introduction of
genetically engineered microorganisms into the environment. Other
approaches for demonstrating cell-specific gene expression involve the
use of electron microscopic immunocytochemistry for detection of
specific enzymes (44) or the use of nucleic acid-based techniques for visualization of specific mRNA (17, 20, 21, 23, 48,
55). These methods require either cultivation of the
microorganism being studied for protein purification or detailed information concerning the target mRNA sequences. Use of these methods
has been restricted to pure or simple mixed cultures because of the
requirement for cell permeabilization procedures tailored to the
bacteria being studied.
The use of radiolabeled substrates in combination with
microautoradiography does allow analysis of the metabolic activity of
prokaryotes under conditions that approach in situ conditions by direct
visualization of microorganisms with active substrate uptake systems
within a complex community (6). Microautoradiography has been successfully used in many ecological studies to measure activities of members of different autotrophic and heterotrophic prokaryotic groups (9, 37). Microautoradiography has been combined with microelectrodes (37) or simple staining
techniques (3, 34, 36, 47) for characterization of microbial
communities. A major limitation of microautoradiography has been an
inability to correlate the activity detected with identification and
detection of a responsible organism. One attempt to solve this problem
has involved combining microautoradiography with fluorescently labeled antibody (FA) techniques (14, 16). However, considering the low percentage of bacteria successfully cultured from most environments and the enormous structural complexity of many microbial communities, the immunofluorescence approach is limited in its applicability for the
following reasons: first, the FA approach is not completely cultivation
independent due to its requirement for a pure culture of the
microorganism being studied for antibody production; second, the
FA approach can be disturbed by the presence of detritus
particles, fungal spores, and extracellular polymeric substances that
have been reported to cause unspecific binding and to hinder antibody penetration (46); and third, due to the great serological
diversity of complex microbial communities, antibody specificity is for the most part restricted to the species level or below, which implies
that hundreds if not thousands of antibodies must be used for an
encompassing structural analysis of a complex microbial community.
The objective of this study was to develop and evaluate a new method
that involves a direct combination of fluorescent in situ hybridization
(FISH) performed with rRNA-targeted oligonucleotide probes and
microautoradiography for simultaneous in situ identification and
determination of substrate uptake patterns of individual microbial cells within complex microbial consortia. Liquid scintillation counting
of samples and pasteurized control samples was performed in all
experiments in order to confirm that biologically induced uptake of
radiolabeled substrates occurred.
Bacterial strains.
Mixtures of Escherichia coli
ATCC 11775T and Herpetosiphon aurantiacus ATCC
23779T were used in a defined system. E. coli
was cultivated at 37°C on Luria-Bertani agar (Difco), and
H. aurantiacus was cultivated at 30°C on R2A agar (Difco).
Prior to the start of the experiment, both strains were separately
cultivated on R2A broth at 30°C and harvested after incubation for
12 h (E. coli) and 48 h (H. aurantiacus). The cells were washed twice with glucose-free R2A
broth (40) and were diluted to a concentration of
approximately 1 mg (dry weight) ml Activated sludge samples.
Activated sludge samples were
collected from two wastewater treatments plants. Activated sludge
containing high levels of phosphorus-accumulating and nitrifying
bacteria was obtained from the Malmö Water and Sewage Works
(Sweden), a municipal nutrient-removing wastewater pilot plant (50 population equivalents) with high biological phosphorus- and
nitrogen-removing activities. In addition, grab samples of activated
sludge were collected in October 1997 and March 1998 from the
intermittently aerated nitrification-denitrification basin of an
industrial wastewater treatment plant receiving sewage from an animal
carcass-processing facility (Tierkörperbeseitigungsanstalt Kraftisried, Kraftisried, Germany; 6,000 population equivalents). In
March 1998, incomplete nitrification, most probably caused by
nonoptimal pH values (pH >8) and low temperatures, was observed at the
Kraftisried plant. All activated sludge samples were stored at 4°C
before they were used. Samples were incubated with radioactive substrates within 3 to 24 h after collection. Prior to incubation, sludge samples were diluted so that they contained 2 and 1 g of dry matter (suspended solids) per liter with nitrate-free water obtained from filtration (Whatman GF/C glass microfiber filter; pore
size, 1 µm) of activated sludge in order to study heterotrophic activity and autotrophic activity (with
[14C]bicarbonate), respectively. A dry matter content
between 1 and 4 g per liter is typically found in activated sludge
from wastewater treatment plants.
Incubation with radioactive and nonradioactive compounds.
The following substrates, labeled with three different isotopes, were
used: (i) D-[6-3H]glucose (specific activity,
15.6 mCi/mmol), (ii) sodium [14C]bicarbonate (specific
activity, 5.0 mCi/mmol), (iii) [2-14C]acetate (sodium
salt; specific activity, 58.0 mCi/mmol), (iv) [n-14C]butyrate (sodium salt; specific
activity, 10 mCi/mmol), and (v) 33Pi (specific
activity, 3 mCi/mmol). Radioactive chemicals were obtained from
Amersham (Buckinghamshire, United Kingdom), NEN Life Science Products
(Boston, Mass.), and Sigma (Deisenhofen, Germany).
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Combination of Fluorescent In Situ Hybridization
and Microautoradiography
a New Tool for Structure-Function
Analyses in Microbial Ecology
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results AND Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results AND Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results AND Discussion
References
1 in sterile, modified
glucose-free R2A broth.
TABLE 1.
Incubation conditions used
for experimentsa
Sample fixation, washing procedures, and DAPI staining of control samples. After incubation with radioactive and nonradioactive substrates, samples were fixed with 6 ml of 4% paraformaldehyde for 3 h at 4°C. Subsequently, the samples were washed three times with 5 ml of phosphate-buffered saline (PBS) (10 mM sodium phosphate buffer, 130 mM sodium chloride; pH 7.2) in order to remove the excess soluble radioactive substrate and fixative. Alternatively, one of the activated sludge samples incubated with 33Pi was washed twice with 0.1 M sodium citrate-HCl buffer (pH 2) prior to the washing procedure described above for removal of chemically bound phosphorus (29). All washing steps consisted of centrifugation of the sample at 37,100 × g for 5 min, removal of the supernatant, addition of further washing buffer, and mixing. After the washing procedure, the samples used as controls for adsorption and chemography were stained with 4',6-diamidino-2-phenylindole (DAPI) by resuspension of the pellet in 1 ml of a DAPI solution (50 µg/ml in PBS) and incubation for 5 min. Excess DAPI was removed by a subsequent PBS washing step.
Homogenization of activated sludge flocs. Fixed, PBS-washed, radioactive, pure cultures and activated sludge pellets were resuspended in 5 ml of PBS, homogenized with a tissue grinder (Thomas Scientific, Swedesbro, N.J.) for 2 min, and then used for liquid scintillation analysis. For in situ hybridization, aliquots of the homogenized sample material were spotted onto poly-L-lysine (Sigma)-coated slides or coverslips, dried at 46°C, and dehydrated in an ethanol series (50, 80, and 96% ethanol, 3 min each).
Liquid scintillation counting. Liquid scintillation counting of homogenized sample materials and pooled wash solutions was performed in all experiments. A 2-ml portion of a wash solution or dispersed sample material was added to 2 ml of scintillation liquid (Ultima Gold XR; Packard Instrument Co., Meriden, Conn.). After thorough mixing, the sample was allowed to react for at least 3 h at room temperature prior to counting with a Packard model 1600 TR liquid scintillation analyzer as recommended by the manufacturer.
Cryosectioning.
For cryosectioning, nonhomogenized, fixed,
and PBS-washed radioactive activated sludge pellets were resuspended in
500 µl of Tissue-Tek (Microm, Walldorf, Germany) and frozen at
20°C for 1 h. Sections that were 5 to 10 µm thick were cut
with a cryomicrotome (model MIKROM HM500; Microm), transferred onto
poly-L-lysine-coated coverslips, and dehydrated in an
ethanol series (50, 80, and 96% ethanol, 3 min each).
Oligonucleotide probes. The following rRNA-targeting oligonucleotide probes were used for FISH: (i) EUB 338 (5'-GCTGCCTCCCGTAGGAGT-3'), which is complementary to a conserved region of most bacterial 16S rRNA molecules (1); (ii) NEU (5'-CCCCTCTGCTGCACTCTA-3'), which is complementary to a signature region of the 16S rRNA of most halophilic and halotolerant ammonia oxidizers belonging to the beta subclass of the Proteobacteria (54); (iii) CTE (5'-TTCCATCCCCCTCTGCCG-3'), which was used as an unlabeled competitor oligonucleotide probe to ensure specific hybridization of probe NEU (54); and (iv) BET42a (5'-GCCTTCCCACTTCGTTT-3') and GAM42a (5'-GCCTTCCCACATCGTTT-3'), which target 23S rRNA regions conserved in members of the beta and gamma subclasses of the Proteobacteria, respectively (32). Oligonucleotides labeled with 5(6)-carboxyfluorescein-N-hydroxysuccinimide ester (FLUOS) or with the monofunctional, hydrophilic, sulfoindocyanine dyes Cy3 and Cy5 were purchased from Interactiva, Ulm, Germany.
DAPI staining and in situ hybridization. In situ hybridization of the samples was performed as described previously (32). Briefly, 9 µl of hybridization solution containing 50 ng of probe (FLUOS-labeled probes) or 30 ng of probe (Cy3- and Cy5-labeled probes) was applied to a sample immobilized on a slide or coverslip and incubated for 90 min at 46°C in an isotonically equilibrated humid chamber. The probes were removed from the slides or coverslips with 2 ml of washing solution and were immediately immersed in 50 ml of prewarmed (48°C) washing solution and incubated for 15 min. The slides and coverslips were rinsed briefly with ice-cold distilled water and dried with pressurized air. After in situ hybridization, the cells were stained with DAPI (0.33 µg/ml in H2O) for 5 min. The excess DAPI was removed by rinsing the slides or coverslips with ice-cold double-distilled H2O.
Autoradiographic procedure and combination with in situ hybridization. An autoradiographic procedure in which autoradiographic emulsion LM-1 (Amersham International) was used was performed as described by Andreasen and Nielsen (3). The optimum exposure time for the samples was between 3 and 7 days. Kodak D19 developer was used as the developer (2 min), and after each preparation was washed in demineralized water for 1 min, it was fixed in 30% (wt/vol) thiosulfate for 4 min. The slides and/or coverslips were washed for 10 min in gently running tap water and then were washed twice (2 min each) in demineralized water. The slides and coverslips were allowed to air dry. Application of the in situ hybridization protocol before and after the autoradiographic procedure was tested. A summary of the method used is presented as flow scheme in Fig. 1.
|
Microscopy. A model LSM 510 scanning confocal microscope (Carl Zeiss, Oberkochen, Germany) equipped with a UV laser (351 and 364 nm), an Ar ion laser (450 to 514 nm), and two HeNe lasers (543 and 633 nm) was used to record optical sections. The formation of silver grains in the autoradiographic film covering a sample was observed by using the transmission mode of the instrument. Image processing was performed with the standard software package delivered with the instrument (version 1.6). Reconstructed and processed images were printed by using the software package Microsoft Power Point (version 7.0; Microsoft, Redmond, Wash.) and a Kodak model 8650 PS printer.
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RESULTS AND Discussion |
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Top
Abstract
Introduction
Materials and Methods
Results AND Discussion
References
|
|---|
Combination of microautoradiography and FISH: method development and evaluation of pure cultures. Separate cultures of E. coli and H. aurantiacus, a nonphotosynthetic member of the green nonsulfur bacteria, were incubated aerobically with a mixture of unlabeled glucose and [3H]glucose (Table 1). Uptake of radioactive glucose was initially monitored after homogenization by liquid scintillation counting. After 3 h of incubation the E. coli and H. aurantiacus cells contained 15.2 and 1.1% of the added radioactively labeled glucose, respectively (Fig. 2A). After 24 h of incubation, a significant increase in glucose uptake by H. aurantiacus (7.9%) (Fig. 2A) was observed. Differences in growth rate, glucose uptake kinetics, and glucose degradation kinetics between E. coli and H. aurantiacus might have contributed to the differences detected in the amounts of radioactive glucose per gram of biomass of the two species. It should be noted that liquid scintillation counting underestimates the actual amount of glucose taken up since a fraction of the radioactive label is excreted as 3H2O due to metabolic glucose transformations. In control experiments performed with pasteurized E. coli and H. aurantiacus cells, only traces of radioactively labeled glucose were detected (E. coli, 0.1%; H. aurantiacus, 0.7%) (Fig. 2A), which demonstrated that there was no adsorption.
|
|
Combination of microautoradiography and FISH: use with complex microbial communities. After we developed the protocol for combining FISH and microautoradiography, this technique was used to document in situ substrate uptake in several probe-defined microbial populations within different complex activated sludge microbial consortia by using different types of radioactively labeled substrates.
[14C]bicarbonate. Comparative experiments were performed by using [14C]bicarbonate and probe NEU targeting halophilic and halotolerant members of the beta subclass ammonia oxidizers (54) with activated sludge with high nitrifying activity (Kraftisried, October 1997; more than 80% N removal) and low nitrifying activity (Kraftisried, March 1998; approximately 10% N removal). Experimental details are given in Table 1 (experiments 2 and 3). After FISH and autoradiographic development, most silver grains accumulated close to clusters of ammonia oxidizers detectable with probe NEU in the homogenized sample obtained in October 1997 (Fig. 4A and B), reflecting the autotrophic activity of the ammonia oxidizers. While all of the NEU-positive cell clusters were labeled with radioactive bicarbonate, DAPI-positive cell clusters that were labeled with radioactive bicarbonate but were not stained with probe NEU were also detected (data not shown). These clusters most likely represented additional autotrophic nitrifying bacteria (e.g., ammonia oxidizers which were not detectable with probe NEU, as well as nitrite-oxidizing bacteria belonging to the genus Nitrospira [25]). In the sample obtained in March 1998, which displayed low nitrifying activity, very few NEU-positive and/or radioactively labeled cells were visualized. In pasteurized controls for both samples no uptake of radioactively labeled bicarbonate was observed. These results were confirmed by liquid scintillation counting of subsamples in both experiments (Fig. 4I).
|
33Pi. Phosphate removal from wastewater is a key feature of modern sewage treatment plants. Traditionally, this removal was achieved by chemical phosphate precipitation with metal salts. Over the last few decades chemical phosphate precipitation has been replaced or supplemented by the so-called enhanced biological phosphorus removal (EBPR) process in many sewage treatment plants (24). Despite the importance of the EBPR process, the microorganism(s) responsible for it has still not been identified (49, 51). Here we investigated the possibility of identifying phosphorus-accumulating microorganisms in situ by a combination of FISH and microautoradiography. Activated sludge obtained from the nutrient removal plant at Malmö was analyzed by anaerobic-aerobic incubation with 33Pi (Table 1, experiment 4) and subsequent in situ hybridization with probes BET42a and GAM42a which are specific for the members of the beta and gamma subclasses of the Proteobacteria, respectively. When these procedures were used, sludge flocs labeled with radioactive phosphate were also detected in the pasteurized negative control, indicating that chemical phosphate precipitation occurred (data not shown). Consequently, an additional wash step performed at pH 2 was included to remove chemically bound phosphorus prior to FISH and the autoradiographic development procedure. Following application of the modified protocol, radioactive phosphate was no longer microscopically detected in the pasteurized control sludge. The influence of the pH of the wash buffers on the amount and nature of the radioactive phosphate detected in the activated sludge was also demonstrated by the results of the liquid scintillation counting procedure. While significant relative amounts of radioactive phosphate (6.9%) were detected in the pasteurized control after the standard pH 7 washing procedure, negligible relative amounts of radioactive phosphate (0.2%) remained after the pH 2 washing step (Fig. 4II). These results demonstrate that proper negative control reactions are important to differentiate between chemical adsorption and biological uptake of the labeled substrate. Once chemical adsorption has been observed for a specific substrate, adequate additional washing procedures must be used in the protocol.
In the homogenized unpasteurized sludge, several morphotypes belonging to the beta subclass of the Proteobacteria labeled with radioactive phosphate were detected, while no representatives of the gamma subclass of the Proteobacteria were found to contain radioactive phosphate (Fig. 4C and D). These preliminary findings confirmed the results of previous molecular studies which indicated that members of the beta subclass of the Proteobacteria (as well as gram-positive bacteria with a high DNA G+C content) are more important than Acinetobacter spp. and other members of the gamma subclass of the Proteobacteria in the EPBR process (5, 51). We are currently intensifying the search for the phosphorus-accumulating organism(s) by (i) establishing phylogenetic 16S rRNA-based inventories of microbial communities involved in EBPR, (ii) designing a probe set for the bacteria, and (iii) combining FISH performed with these probes and microautoradiography, using 33Pi as the substrate.14C-labeled fatty acids. Activated sludge obtained from the Kraftisried plant in October 1997 was incubated with 14C-labeled acetate and 14C-labeled butyrate. These two fatty acids were selected because they comprise a main fraction of the organic load of the influent wastewater of the Kraftisried plant. Uptake of both fatty acids under aerobic incubation conditions and anaerobic incubation conditions (with and without nitrate) was studied in order to determine the ability of the combined technique to monitor differences in the uptake of substrates by the microbial consortium under different metabolic conditions. Experimental details are given in Table 1 (experiments 5 to 10). Subsequent in situ hybridization was performed with probes NEU and BET42a, which are specific for halophilic and halotolerant beta subclass ammonia oxidizers and members of the beta subclass of the Proteobacteria, respectively. Ammonia oxidizers stained during in situ hybridization with probe NEU failed to take up acetate or butyrate under the different conditions (Fig. 4E and F), suggesting that they were not able to grow mixotrophically. After aerobic incubation with radioactively labeled acetate or butyrate, most sludge flocs, which were composed mainly of members of the beta subclass of the Proteobacteria, were heavily coated with silver grains (Fig. 4E and F), while under anaerobic conditions only insignificant uptake of these fatty acids, comparable to the uptake observed in pasteurized control samples, was detected. Interestingly, under denitrifying conditions (anaerobic conditions with nitrate) the microbial activated sludge population took up significantly more butyrate than acetate (data not shown). The substrate uptake patterns of the activated sludge under different conditions observed microscopically were confirmed by parallel liquid scintillation counting of subsamples (Fig. 4III).
As observed in the [14C]bicarbonate and 33Pi experiments (Fig. 4A and C), insufficient dispersion of cell aggregates and activated sludge flocs by the homogenization procedure hampered single-cell resolution by microautoradiography (Fig. 4E). To improve the spatial resolution, we replaced the homogenization step in the protocol by using cryosections (thickness, 5 to 10 µm) of the radioactively labeled activated sludge flocs for in situ hybridization and subsequent autoradiographic development. This modification allowed us to correlate silver grain formation with individual probe-stained microbial cells, even within voluminous sludge flocs (Fig. 4G, H, and J). However, it should be noted that the native architecture of the sludge flocs might still have been disrupted to some extent by sample handling (e.g., fixation, centrifugation, and resuspension) prior to the cryosectioning step.Conclusions. Over the last decade molecular methods have revealed new and fascinating insights into the microbial diversity present in environmental samples (22, 31, 43, 56). Probe-based techniques have also allowed detailed quantitative studies of environmental microbial community structures and dynamics to be performed (1, 38, 39, 53). Probe-based techniques have also allowed detailed quantitative studies of environmental microbial community structures and dynamics to be performed (1, 38, 39, 50, 51). While they have provided important information, such studies have failed to reveal the functions of the microorganisms within their ecological niches. The combination of FISH and microautoradiography described in this paper allowed us to directly analyze the in vivo uptake of substrates by probe-identified microorganisms under different environmental conditions. While microautoradiography does not provide quantitative data, the information obtained from experiments which combined the two techniques should help identify key functional microbial groups that are active in the environment. The possible applications range from fundamental studies in microbial ecology to identifying bacteria involved in biotechnological processes, such as sewage treatment and bioremediation. In addition, information concerning the in vivo physiology of novel bacteria taxa identified by molecular techniques should facilitate the design of appropriate isolation techniques for many of these hitherto uncultured microorganisms.
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ACKNOWLEDGMENTS |
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This work was supported by postdoctoral grant 97-256, from the Swedish Foundation for International Cooperation in Research and Higher Education to N.L., by the Körber Award, by the Danish Technical Research Council (Activity and Diversity in Complex Microbial Systems framework program), and by Sonderforschungsbereich SFB411 from the Deutsche Forschungsgemeinschaft (Research Center for Fundamental Studies of Aerobic Biological Wastewater Treatment). K.H.A. was supported by an industrial Ph.D. fellowship.
We thank Thomas Fritsche for critically reading the manuscript. We gratefully acknowledge the contribution of David Stahl in formulating concepts used in this study.
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FOOTNOTES |
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* Corresponding author. Mailing address: Lehrstuhl für Mikrobiologie, Technische Universität München, D-80290 Munich, Germany. Phone: 49 89 2892 2373. Fax: 49 89 2892 2360. E-mail: wagner{at}mikro.biologie.tu-muenchen.de.
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REFERENCES |
|---|
|
Top
Abstract
Introduction
Materials and Methods
Results AND Discussion
References
|
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| 1. |
Amann, R. I.,
L. Krumholz, and D. A. Stahl.
1990.
Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology.
J. Bacteriol.
172:762-770 |
| 2. |
Amann, R. I.,
W. Ludwig, and K. H. Schleifer.
1995.
Phylogenetic identification and in situ detection of individual microbial cells without cultivation.
Microbiol. Rev.
59:143-169 |
| 3. | Andreasen, K., and P. H. Nielsen. 1997. Application of microautoradiography for the study of substrate uptake by filamentous microorganisms in activated sludge. Appl. Environ. Microbiol. 63:3662-3668[Abstract]. |
| 4. | Assmus, B., P. Hutzler, G. Kirchhof, R. Amann, J. R. Lawrence, and A. Hartmann. 1995. In situ localization of Azospirillum brasiliense in the rhizosphere of wheat with fluorescently labeled rRNA-targeted oligonucleotide probes and scanning confocal laser microscopy. Appl. Environ. Microbiol. 61:1013-1019[Abstract]. |
| 5. | Bond, P. L., P. Hugenholtz, J. Keller, and L. L. Blackall. 1995. Bacterial community structures of phosphate-removing and non-phosphate-removing activated sludges from sequencing batch reactors. Appl. Environ. Microbiol. 61:1910-1916[Abstract]. |
| 6. | Brock, T. D., and M. L. Brock. 1968. Autoradiography as a tool in microbial ecology. Nature 209:734-736. |
| 7. | Caldwell, D. E., D. R. Korber, and J. R. Lawrence. 1992. Confocal laser microscopy and digital image analysis in microbial ecology. Adv. Microb. Ecol. 12:1-67. |
| 8. | Caldwell, D. E., G. M. Wolfaardt, and D. R. Korber. 1997. Do bacterial communities transcend Darwinism? Adv. Microb. Ecol. 15:105-191. |
| 9. | Carman, K. R. 1990. Radioactive labeling of a natural assemblage of marine sedimentary bacteria and microalgae for the trophic studies: an autoradiographic study. Microb. Ecol. 19:279-290. |
| 10. |
Costerton, J. W.,
Z. Lewandowski,
D. de Beer,
D. E. Caldwell,
D. R. Korber, and G. A. James.
1994.
Biofilms: the customized microniche.
J. Bacteriol.
176:2137-2142 |
| 11. | Costerton, J. W., Z. Lewandowski, D. E. Caldwell, D. R. Korber, and H. M. Lappin-Scott. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49:711-745[Medline]. |
| 12. |
Deretic, V.,
M. J. Schurr,
J. C. Boucher, and D. W. Martin.
1994.
Conversion of Pseudomonas aeruginosa to mucoidy in cystic fibrosis: environmental stress and regulation of bacterial virulence by alternative sigma factors.
J. Bacteriol.
176:2773-2780 |
| 13. | Dworkin, M. 1991. Introduction, p. 1-6. In M. Dworkin (ed.), Microbial cell-cell interactions. American Society for Microbiology, Washington, D.C. |
| 14. | Fliermans, C. B., and E. L. Schmidt. 1975. Autoradiography and immunofluorescence combined for autoecological study of single cell activity with Nitrobacter as a model system. Appl. Microbiol. 30:676-684[Medline]. |
| 15. | Fuhrmann, J. A., and F. Azam. 1982. Thymidine incorporation as a measure of heterotrophic bacterioplankton production in marine surface waters: evaluation and field results. Mar. Biol. 66:109-120. |
| 16. | Ghiorse, W. C., D. N. Miller, R. L. Sandoli, and P. L. Siering. 1996. Applications of laser scanning microscopy for analysis of aquatic microhabitats. Microsc. Res. Tech. 33:73-86[Medline]. |
| 17. |
Hahn, D.,
R. I. Amann, and J. Zeyer.
1993.
Detection of mRNA in Streptomyces cells by whole-cell hybridization with digoxigenin-labeled probes.
Appl. Environ. Microbiol.
59:2753-2757 |
| 18. | Harmsen, H. J. M., A. D. L. Akkermans, A. J. M. Stams, and W. M. De Vos. 1996. Population dynamics of propionate-oxidizing bacteria under methanogenic and sulfidogenic conditions in anaerobic granular sludge. Appl. Environ. Microbiol. 62:2163-2168[Abstract]. |
| 19. |
Harry, E. J.,
K. Pogliano, and R. Losick.
1995.
Use of immunofluorescence to visualize cell-specific gene expression during sporulation in Bacillus subtilis.
J. Bacteriol.
177:3386-3393 |
| 20. | Hodson, R. E., W. A. Dustman, R. P. Garg, and M. A. Moran. 1995. In situ PCR for visualization of microscale distribution of specific genes and gene products in prokaryotic communities. Appl. Environ. Microbiol. 61:4074-4082[Abstract]. |
| 21. | Hönerlage, W., D. Hahn, and J. Zeyer. 1995. Detection of mRNA of nprM in Bacillus megaterium ATCC 14581 grown in soil by whole-cell hybridization. Arch. Microbiol. 163:235-241. |
| 22. |
Hugenholtz, P.,
C. Pitulle,
K. L. Hershberger, and N. R. Pace.
1998.
Novel division level bacterial diversity in a Yellowstone hot spring.
J. Bacteriol.
180:366-376 |
| 23. |
Hurek, T.,
T. Egener, and B. Reinhold-Hurek.
1997.
Divergence in nitrogenases of Azoarcus spp., Proteobacteria of the beta subclass.
J. Bacteriol.
179:4172-4178 |
| 24. |
Jenkins, D., and V. Tandoi.
1991.
The applied microbiology of enhanced biological phosphate removal accomplishments and needs.
Wat. Res.
25:1471-1478.
|
| 25. |
Juretschko, S.,
G. Timmerman,
M. Schmid,
K.-H. Schleifer,
A. Pommerening-Röser,
H.-P. Koops, and M. Wagner.
1998.
Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations.
Appl. Environ. Microbiol.
64:3042-3051 |
| 26. |
Kane, M. D.,
L. K. Poulsen, and D. A. Stahl.
1993.
Monitoring of the enrichment and isolation of sulfate-reducing bacteria by using oligonucleotide hybridization probes designed from environmentally derived 16S rRNA sequences.
Appl. Environ. Microbiol.
59:682-686 |
| 27. | Kremer, L., A. Baulard, J. Estaquier, O. Poulain-Godefroy, and C. Locht. 1995. Green fluorescent protein as a new expression marker in mycobacteria. Mol. Microbiol. 17:913-922[Medline]. |
| 28. |
Lawrence, J. R.,
G. M. Wolfaardt, and T. R. Neu.
1998.
The study of biofilms using confocal laser scanning microscopy, p. 431-465.
In
M. H. F. Wilkinson, and F. Schut (ed.), Digital image analysis of microbesimaging, morphometry, fluorometry and motility techniques and applications. John Wiley & Sons, Chichester, United Kingdom.
|
| 29. | Lee, N., M. Wagner, and P. H. Nielsen. Unpublished results. |
| 30. | Lewis, P. J., C. E. Nwoguh, M. R. Barer, C. R. Harwood, and J. Errington. 1994. Use of digitized video microscopy with a fluorogenic enzyme substrate to demonstrate cell- and compartment-specific gene expression in Salmonella enteritidis and Bacillus subtilis. Mol. Microbiol. 13:655-662[Medline]. |
| 31. |
Liesack, W., and E. Stackebrandt.
1992.
Occurrence of novel groups of the domain Bacteria as revealed by analysis of genetic material isolated from an Australian terrestrial environment.
J. Bacteriol.
174:5072-5078 |
| 32. | Manz, W., R. Amann, W. Ludwig, M. Wagner, and K.-H. Schleifer. 1992. Phylogenetic oligodeoxynucleotide probes for the major subclasses of proteobacteria: problems and solutions. Syst. Appl. Microbiol. 15:593-600. |
| 33. | Massol-Deyá, A. A., J. Whallon, R. F. Hickey, and J. M. Tiedje. 1995. Channel structures in aerobic biofilms of fixed-film reactors treating contaminated groundwater. Appl. Environ. Microbiol. 61:769-777[Abstract]. |
| 34. |
Meyer-Reil, L. A.
1978.
Autoradiography and epifluorescence microscopy combined for the determination of number and spectrum of actively metabolizing bacteria in natural waters.
Appl. Environ. Microbiol.
36:506-512 |
| 35. | Møller, S., A. R. Pedersen, L. K. Poulsen, E. Arvin, and S. Molin. 1996. Activity and three-dimensional distribution of toluene-degrading Pseudomonas putida in a multispecies biofilm assessed by quantitative in situ hybridization and scanning confocal laser microscopy. Appl. Environ. Microbiol. 62:4632-4640[Abstract]. |
| 36. | Nielsen, P. H., K. Andreasen, M. Wagner, L. L. Blackall, H. Lemmer, and R. J. Seviour. 1998. Variability of type 021N in activated sludge as determined by in situ substrate uptake pattern and in situ hybridization with fluorescent rRNA targeted probes. Wat. Sci. Technol. 37:423-430. |
| 37. | Paerl, H. W. 1984. Alteration of microbial metabolic activities in association with detritus. Bull. Mar. Sci. 35:393-408. |
| 38. | Ramsing, N. B., M. Kühl, and B. B. Jörgensen. 1993. Distribution of sulfate-reducing bacteria, O2 and H2S in photosynthetic biofilms determined by oligonucleotide probes and microelectrodes. Appl. Environ. Microbiol. 59:3820-3849. |
| 39. |
Raskin, L.,
J. M. Stromley,
B. E. Rittmann, and D. A. Stahl.
1994.
Group-specific 16S rRNA hybridization probes to describe natural communities of methanogens.
Appl. Environ. Microbiol.
60:1232-1240 |
| 40. |
Reasoner, D. J., and E. E. Geldreich.
1985.
A new medium for the enumeration and subculture of bacteria from potable water.
Appl. Environ. Microbiol.
49:1-7 |
| 41. | Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K.-H. Schleifer. 1996. Structure and function of a nitrifying biofilm as determined by in situ hybridization and microelectrodes. Appl. Environ. Microbiol. 62:4641-4647[Abstract]. |
| 42. |
Schramm, A.,
D. DeBeer,
M. Wagner, and R. Amann.
1998.
Identification and activities in situ of Nitrosospira and Nitrospira spp. as dominant populations in a nitrifying fluidized bed reactor.
Appl. Environ. Microbiol.
64:3480-3485 |
| 43. | Snaidr, J., R. Amann, I. Huber, W. Ludwig, and K.-H. Schleifer. 1997. Phylogenetic analysis and in situ identification of bacteria in activated sludge. Appl. Environ. Microbiol. 63:2884-2896[Abstract]. |
| 44. | Spieck, E., S. Ehrich, J. Ammand, and E. Bock. 1998. Isolation and immunocytochemical location of the nitrite-oxidizing system in Nitrospira moscoviensis. Arch. Microbiol. 169:225-230[Medline]. |
| 45. | Stewart, G. S., and P. Williams. 1992. lux genes and the applications of bacterial bioluminescence. J. Gen. Microbiol. 138:1289-1300[Medline]. |
| 46. | Szwerinski, H., S. Gaiser, and D. Bardtke. 1985. Immunofluorescence for the quantitative determination of nitrifying bacteria: interference of the test in biofilm reactors. Appl. Microbiol. Biotechnol. 21:125-128. |
| 47. |
Tabor, P. S., and R. A. Neihof.
1984.
Direct determination of activities for microorganisms of Chesapake Bay populations.
Appl. Environ. Microbiol.
48:1012-1019 |
| 48. | Tolker-Nielsen, T., K. Holmström, and S. Molin. 1997. Visualization of specific gene expression in individual Salmonella typhimurium cells by in situ PCR. Appl. Environ. Microbiol. 63:4196-4203[Abstract]. |
| 49. | van Loosdrecht, M. C. M., G. Smolders, T. Kuba, and J. J. Heijnen. 1997. Metabolism of micro-organisms responsible for enhanced biological phosphorus removal from wastewater. Use of dynamic enrichment cultures. Antonie Leeuwenhoek 71:109-116. |
| 50. |
Wagner, M.,
R. Amann,
H. Lemmer, and K.-H. Schleifer.
1993.
Probing activated sludge with oligonucleotides specific for proteobacteria: inadequacy of culture-dependent methods for describing microbial community structure.
Appl. Environ. Microbiol.
59:1520-1525 |
| 51. |
Wagner, M.,
R. Erhart,
W. Manz,
R. Amann,
H. Lemmer,
D. Wedi, and K.-H. Schleifer.
1994.
Development of an rRNA-targeted oligonucleotide probe specific for the genus Acinetobacter and its application for in situ monitoring in activated sludge.
Appl. Environ. Microbiol.
60:792-800 |
| 52. | Wagner, M., R. Amann, P. Kämpfer, B. Assmus, A. Hartmann, P. Hutzler, N. Springer, and K.-H. Schleifer. 1994. Identification and in situ detection of Gram-negative filamentous bacteria in activated sludge. Syst. Appl. Microbiol. 17:405-417. |
| 53. | Wagner, M., B. Assmus, A. Hartmann, P. Hutzler, and R. Amann. 1994. In situ analysis of microbial consortia in activated sludge using fluorescently labelled, rRNA-targeted oligonucleotide probes and confocal laser scanning microscopy. J. Microsc. 176:181-187[Medline]. |
| 54. | Wagner, M., G. Rath, R. Amann, H.-P. Koops, and K.-H. Schleifer. 1995. In situ identification of ammonia-oxidizing bacteria. Syst. Appl. Microbiol. 17:251-264. |
| 55. | Wagner, M., M. Schmid, S. Juretschko, K. H. Trebesius, A. Bubert, W. Goebel, and K.-H. Schleifer. 1998. In situ detection of a virulence factor mRNA and 16S rRNA in Listeria monocytogenes. FEMS Microbiol. Lett. 160:159-168[Medline]. |
| 56. | Ward, D. M., M. M. Bateson, R. Weller, and A. L. Ruff-Roberts. 1992. Ribosomal RNA analysis of microorganisms as they occur in nature. Adv. Microb. Ecol. 12:219-286. |
| 57. |
Webb, C. D.,
A. Decatur,
A. Teleman, and R. Losick.
1995.
Use of green fluorescent protein for visualization of cell-specific gene expression and subcellular protein localization during sporulation in Bacillus subtilis.
J. Bacteriol.
177:5906-5911 |
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