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Applied and Environmental Microbiology, July 2008, p. 4231-4235, Vol. 74, No. 13
0099-2240/08/$08.00+0     doi:10.1128/AEM.02545-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Use of Laser Microdissection for Phylogenetic Characterization of Polyphosphate-Accumulating Bacteria{triangledown}

Stefanie Gloess,1 Hans-Peter Grossart,1 Martin Allgaier,1,{dagger} Stefan Ratering,2* and Michael Hupfer1*

Leibniz Institute of Freshwater Ecology and Inland Fisheries, Alte Fischerhütte 2, D-16775 Stechlin-Neuglobsow, Germany,1 Institute of Applied Microbiology, Justus-Liebig University Giessen, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany2

Received 12 November 2007/ Accepted 22 April 2008


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ABSTRACT
 
Our novel approach for taxonomic identification of uncultured bacteria harboring specific physiological features in complex environmental samples combines cell collection by laser microdissection and subsequent DNA analysis. The newly developed approach was successfully tested for collection and phylogenetic characterization of polyphosphate-accumulating bacteria in activated sludge and lake sediment.


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INTRODUCTION
 
In order to examine the ecological role of bacteria in a (eco)system, it is often necessary to detect and identify these microorganisms in situ, particularly if their specific metabolic capabilities need to be studied. For this purpose, different methods are available; however, currently all of them have distinct disadvantages and limitations. For example, PCR-based techniques using cultivation-independent approaches are very useful for exploring the diversity of microorganisms in environmental samples, but often their metabolic functions remain unknown. More recently, techniques based on fluorescence in situ hybridization (FISH), such as Raman-FISH (17), FISH-secondary-ion mass spectrometry (31), and FISH-microautoradiography (26, 32), have been developed to overcome such problems. These techniques work with substrates labeled with either stable or radioactive isotopes to help identify microbiota and to study their ecophysiology. For the design and application of specific oligonucleotide probes used in FISH, however, knowledge of the target (e.g., 16S rRNA) sequence is essential. The identification of microorganisms involved in specific metabolic processes in a complex environmental sample can also be achieved by additional methods, e.g., separation via stable isotope probing (33) based on the selective recovery of "heavy" 13C-labeled biomarkers (e.g., DNA or RNA). However, this probing technique is restricted by the somewhat limited availability of labeled substrates. In addition to the restriction of limited availability of highly enriched commercial compounds, the actual addition of external substrates and incubation time may affect the activity and composition of the natural bacterial community.

Hence, we aimed to develop an alternative approach for direct phylogenetic characterization of bacteria with specific metabolic functions from various environmental samples, in our case activated sludge and lake sediment. Our novel approach is based on the precise collection of single microorganisms and overcomes all of the abovementioned methodological limitations. In this study, we have separated polyphosphate (poly-P)-accumulating bacteria (PAO) from poly-P-containing sludge and lake sediment. PAO are key players in the phosphorus (P) cycle in sewage treatment plants with enhanced biological P removal. Hence, for optimization of sewage treatment processes, a fundamental understanding of the bacterial sludge community, especially of PAO, is necessary. To date, only a few bacteria of this group have been isolated and characterized; however, they are often of minor relevance for P cycling in full-scale plants (1, 8, 24). Thus, their occurrence in aquatic sediments as well as their role in biotic P transformation is poorly understood and highly controversial (19). In an attempt to specifically characterize PAO in sludge and lake sediment, we used laser microdissection and subsequent phylogenetic characterization to precisely describe members of this bacterial group. Laser microdissection is a powerful technique which has been used previously to analyze eukaryotic cells for a variety of scientific applications (6), e.g., forensics (4), botany (3), pathology (reviewed in reference 7), neurosciences (10), and medical biology (13). It has also been used to analyze pathogenic bacteria in histological tissues, using fluorescently labeled oligonucleotide probes or specific primers (16, 23). To our knowledge, this is the first study using laser microdissection for phylogenetic characterization of specific bacteria in aquatic samples.


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Methodological approach.
 
Activated sludge was obtained from the Wassmannsdorf wastewater treatment plant (near Berlin, Germany) in January 2006. Sediment cores were collected (June 2004) from the deepest point of the oligotrophic Lake Stechlin with a modified Kajak sampler (Uwitech, Mondsee, Austria) equipped with Plexiglas tubes (diameter, 6 cm). The uppermost sediment (ca. 5 mm) was immediately removed using a 50-ml plastic syringe equipped with a flexible tube. To obtain a representative composite sample, six replicate cores were pooled. Poly-P was detected and quantified in both samples by 31P nuclear magnetic resonance analyses according to a standard protocol (18). Poly-P accounted for 0.81 and 2.97 mg P g–1 dry weight in Lake Stechlin and activated sludge, respectively. This high poly-P content provided a good basis for further detection, separation, and identification of potential PAO in these samples.


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Detection of PAO.
 
Bacterial cells were extracted from activated sludge and sediment with sodium pyrophosphate (0.1% [wt/vol]) by gentle sonication and shaking in an overhead shaker (level 3) (Reax 2; Heidolph GmbH & Co. KG, Schwabach, Germany) for 30 min. After 30 min of sedimentation, the supernatant was removed and sonicated, and the extracts were fixed in 99% ethanol (1:1 [vol/vol]). PAO were identified by examination of their intracellular poly-P granules, using epifluorescence microscopy (magnification, x400), after extracts were filtered onto a 0.2-µm polycarbonate filter (Nuclepore; Whatman, Germany) and stained with 5 µg ml–1 DAPI (4',6'-diaminido-2-phenylindole dihydrochloride) for 30 min. At this high DAPI concentration, poly-P granules show an intensive yellow fluorescence (36). This characteristic attribute was our criterion for cell separation via laser microdissection, which was performed within 48 h after DAPI staining.


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Collection of PAO by laser microdissection.
 
Laser microdissection and pressure catapulting were performed with a P.A.L.M. MicroBeam system in combination with a P.A.L.M. RoboMover controlled by P.A.L.M. RoboSoftware v2.0 (P.A.L.M. Microlaser Technologies GmbH, Bernried, Germany). The MicroBeam system included an Axiovert 200 M microscope (Carl Zeiss Microimaging GmbH, Germany) equipped with a 100-W mercury lamp. The polycarbonate filters were inspected with an FS18 fluorescence filter (P.A.L.M. Microlaser Technologies GmbH, Bernried, Germany) at a magnification of x400. Single bacterial cells with blue fluorescence and yellow poly-P granules were excised and catapulted into the lid of a 200-µl adhesive-cap tube (P.A.L.M. Microlaser Technologies GmbH, Bernried, Germany) under sterile conditions. For Lake Stechlin sediment, PAO were spread across the whole filter, from which ca. 100 individual cells were selected for phylogenetic analyses. In contrast, sewage sludge PAO often occurred in microcolonies and aggregates, and hence, more cells could be separated (>400 cells). Although single cells were excised, all cells of each sample were pooled in one tube for further phylogenetic analysis. As negative controls, pieces of a sterile polycarbonate filter were used.


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DNA extraction and PCR.
 
All molecular work was performed under sterile conditions in a laminar flow biosafety cabinet. Before and after work, the cabinet and all instruments were irradiated for several hours with UV light and cleaned with DNA decontamination reagent (DNA-ExitusPlus; AppliChem, Darmstadt, Germany). To ensure that no contaminating cells or DNA was present in the Palm adhesive caps or PCR tubes (Biopur Safe-Lock; Eppendorf, Hamburg, Germany), empty tubes were tested by PCR with universal bacterial 16S rRNA gene primers as described below. Since we obtained positive amplification from negative PCR controls for DNA extracted with a Qiagen Micro kit (Qiagen, Hilden, Germany) especially designed for laser microdissection, we extracted DNA from each sample directly in the capture cups. For this purpose, a sterile water-buffer solution was pipetted into the lids of adhesive-cap tubes containing either microdissected cells or negative controls.

With very small amounts of extracted DNA, contaminating DNA can have a large effect on PCR-based analyses. Therefore, in order to avoid this potential problem, DNA extraction and PCR buffers were incubated with DNase I (0.5 to 0.75 U) at room temperature for 12 min, followed by DNase inactivation by incubation at 95°C for 15 min.

In order to directly extract DNA, a DNase I-digested water-buffer solution was pipetted into the lids of adhesive-cap tubes with either microdissected cells or negative controls. The water-buffer solution contained 10 µl water, 1 µl 10x PCR buffer (Qiagen, Hilden, Germany), and 0.75 U DNase I (AppliChem, Darmstadt, Germany) per sample. After pipetting of the DNase I-digested water-buffer solution onto the sample, the adhesive-cap tubes were vortexed with the lid upside down. Further cell-cracking steps were as follows: (i) centrifugation for 2 min at 16,000 x g, (ii) sonication for 2 min, (iii) centrifugation for 5 min at 16,000 x g, and (iv) freeze-thaw. Steps i to iii were performed at room temperature. For the freeze-thaw step, the tubes were incubated three times each for 3 min at 98°C and then in liquid nitrogen.

PCR amplification of 16S rRNA genes was performed using the bacterial primer pair 341f and 907r (30) (MWG Biotech AG, Ebersberg, Germany), as this primer set allows for a rough but rapid phylogenetic analysis. The reaction mixture contained 250 nM of each primer, 0.5 µg of bovine serum albumin (Roth, Karlsruhe, Germany), a 200 µM concentration of each deoxynucleoside triphosphate (Bioline GmbH, Luckenwalde, Germany), 2 mM MgCl2, 1 µl of 10x PCR buffer, and 0.2 U HotStarTaq DNA polymerase (Qiagen, Hilden, Germany). Ultrapure water (Bioline GmbH, Luckenwalde, Germany) was used to bring the reaction volume up to 11 µl. Note that to avoid contamination by traces of Escherichia coli DNA, the PCR master mix was incubated with DNase I (final concentration, 0.51 U) at room temperature for 12 min, followed by a DNase inactivation and polymerase activation step at 95°C for 15 min.

Four microliters of the cell-cracked sample and 11 µl of the DNase-digested PCR master mix were transferred into a new sterile Biopur Eppendorf cap. Both the DNase-digested water-buffer solution and PCR master mix were checked for PCR amplification without the addition of template. Solutions showing DNA amplification were discarded. PCR amplification was performed in a Gradient Cycler PT200 instrument (MJ Research), using the following thermal profile: (i) initial denaturation at 95°C for 2 min, (ii) denaturation at 95°C for 1 min, (iii) annealing at 55°C for 40 s, (iv) extension at 72°C for 1 min 30 s (48 cycles of steps 2 to 4 were performed), and (v) final extension at 72°C for 10 min. Cooling down at 4°C completed the reaction.

The greater number and increased cell size of separated sewage sludge bacteria led to more extracted and amplified DNA than that obtained from Lake Stechlin sediments.

We also tested primer pair 8f and 907r to try to maximize PCR product length; however, this primer pair did not amplify Lake Stechlin sediment-derived DNA. For sewage sludge DNA, both primer pairs revealed PCR products but gave different results for phylogenetic community composition. Whereas members of the Alphaproteobacteria, Actinobacteria, and Firmicutes were detected with both primer sets, the primer pair 8f-907r additionally revealed the occurrence of members of the Betaproteobacteria and Bacteroidetes. This bias might be caused by differences in primer selectivity of the forward primer 8f (14), although detection of similar clones in Lake Stechlin sediment suggests that they may have a role in poly-P accumulation.

Other sources of error may have been (i) the undefined efficiency of the above-described DNA extraction procedure for environmental samples and (ii) the small amounts of PCR products obtained, especially for sediment samples, despite optimizing the PCR cycle number. Nevertheless, the protocol described above proved to be sufficient for cloning and subsequent phylogenetic analysis of microdissected cells, whether they were derived from sludge or sediment.


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Clone libraries.
 
For construction of clone libraries, the 16S rRNA gene PCR products obtained using the primer pair 341f and 907r were purified using SureClean (Bioline GmbH, Luckenwalde, Germany) and then cloned into competent E. coli cells, using pGEM-T Easy vector system II (Promega GmbH, Mannheim, Germany) according to the manufacturer's protocol. For each clone library, over 100 clones were picked and the plasmids purified with the Wizard Plus Minipreps DNA purification system (Promega GmbH, Mannheim, Germany). Clones with an incomplete insert and sequences of poor quality were excluded from further analyses. Between 72 and 84 clones of each clone library were processed for sequencing.


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Sequencing and phylogenetic classification of PAO.
 
For sequencing, a Big Dye Terminator v3.1 cycle sequencing kit and an ABI Prism 3100-Avant genetic analyzer (Applied Biosystems, Darmstadt, Germany) were used. The cloned 16S rRNA gene fragments were partially sequenced (about 400 to 550 bp for most and 250 to 300 bp for some) with primer M13f (29) or 341f. Phylogenetic classifications were determined by BLAST searches (http://www.ncbi.nlm.nih.gov/BLAST) and the sequence classification tool of the Ribosomal Database Project RDP II (http://rdp.cme.msu.edu/). Clone libraries constructed for PAO found in both sewage sludge and Lake Stechlin sediment were dominated by members of the Alphaproteobacteria and Actinobacteria (Fig. 1; Table 1). In Lake Stechlin sediment, a more diverse population of PAO was found, which is in accordance with the fact that sediments consist of different microniches and zones (37). In contrast, sewage sludge is more adept at harboring specific microbial communities due to its more uniform composition and conditions. In both samples, relatively large numbers of Propionibacterium-related sequences were found. Poly-P storage has been described previously for some members of the genus Propionibacterium and for the genus Paracoccus, which dominated the sludge clone library. Other known poly-P bacteria detected belonged to the genera Micrococcus, Rhodococcus, Corynebacterium, and Staphylococcus (present in the sewage sludge), as well as Pseudomonas (present in Lake Stechlin sediment). Since the full genome of "Candidatus Accumulibacter phosphatis" was recently studied by metagenomic analysis of an enrichment culture from enhanced biological phosphorus removal sludge (27), it was slightly surprising that no trace of this bacterial group was found.


Figure 1
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  		FIG. 1. Neighbor-joining phylogenetic tree of representative sequences derived from the 16S rRNA gene clone libraries for sewage sludge and Lake Stechlin sediment, using the primer pair 341f-907r. Sequences from this study are shown in bold. Sequences from sewage sludge are additionally marked by asterisks. GenBank accession numbers are given. The scale bar corresponds to 10 base substitutions per 100 nucleotide positions.


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  		TABLE 1. Phylogenetic classification of clones (obtained with primer pair 341f-907r) from sewage sludge and Lake Stechlin sedimenta

In summary, our study shows that PAO could be identified in both samples by use of laser microdissection and subsequent phylogenetic analyses of the excised cells. In addition to already known PAO, several new bacterial taxa for which poly-P storage has not yet been described were identified in the lake sediment clone library (e.g., the genera Hyphomicrobium and Microbacterium). However, the phylogenetic characterization of PAO is an indication of the potential ability to store poly-P and cannot be taken as evidence of a real physiological role. In this study, intracellular poly-P was visualized by DAPI staining only. Therefore, additional studies, preferably with cultures, are necessary to verify the intracellular presence and formation of poly-P in these bacterial groups. Furthermore, the development and use of highly specific oligonucleotide probes (e.g., for FISH) would reveal more detailed information about the relative abundances of PAO in natural bacterial communities and their contribution to the actual poly-P content measured. Nevertheless, our clone libraries suggest that poly-P accumulation by sewage sludge and sediment bacteria is a common feature to both bacterial communities.

The Palm MicroBeam technology coupled with molecular analyses is an excellent approach for the phylogenetic classification of hitherto unknown putative PAO from environmental samples.

Our results show that this approach is a powerful tool for direct identification of uncultured sediment and other aquatic bacteria with microscopically detectable specific features. This approach has the potential to select different bacterial groups from a variety of environments by using markers other than intracellular poly-P granules, e.g., intracellular polyhydroxybutyrate, FISH probes (medical microbiology [23]), or specific radiolabeled substrates (microautoradiography).


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Nucleotide sequence accession numbers.
 
The partial sequences of 16S rRNA genes obtained in this study were deposited in GenBank under accession numbers EU152875 to EU152914, EU196056 to EU196127, and EU515684 to EU515781.


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ACKNOWLEDGMENTS
 
We thank Peter Schmieder for his assistance with the nuclear magnetic resonance investigations carried out at the Research Institute of Molecular Pharmacology, Berlin, Germany. We also thank Christiane Herzog for her valuable help with chemical-analytical work and Monika Degebrodt for running the sequencer. Sylvia Schnell and Udo Jäckel (Institute of Applied Microbiology, University of Giessen) contributed to this study by their inspiring discussions. Gabriele Friedemann (Palm Microlaser Technologies GmbH, Bernried, Germany) is thanked for her introduction to P.A.L.M. MicroBeam technology and for usage of P.A.L.M. instruments.


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FOOTNOTES
 
* Corresponding author. Mailing address for Stefan Ratering (microdissection): Institute of Applied Microbiology, Justus-Liebig University Giessen, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany. Phone: 49 641 9937354. Fax: 49 641 9937359. E-mail: Stefan.Ratering{at}agrar.uni-giessen.de. Mailing address for Michael Hupfer (poly-P bacteria): Leibniz Institute of Freshwater Ecology and Inland Fisheries, Müggelseedamm 310, D-12587 Berlin, Germany. Phone: 49 30 64181 605. Fax: 49 30 64181682. E-mail: hupfer{at}igb-berlin.de Back

{triangledown} Published ahead of print on 2 May 2008. Back

{dagger} Present address: DOE Joint Genome Institute, 2800 Mitchell Dr., Walnut Creek, CA 94598. Back


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Applied and Environmental Microbiology, July 2008, p. 4231-4235, Vol. 74, No. 13
0099-2240/08/$08.00+0     doi:10.1128/AEM.02545-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.




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