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In situ detection of starch-hydrolyzing microorganisms in activated sludge

Yun Xia , Yunhong Kong , Per Halkjær Nielsen
DOI: http://dx.doi.org/10.1111/j.1574-6941.2008.00559.x 462-471 First published online: 1 November 2008

Abstract

Polysaccharides constitute a significant part of the organic matter in domestic wastewater and their hydrolysis plays an important role in their transformation and nutrient removal in activated sludge wastewater treatment plants. However, there is no information available about the identity, ecophysiology, and abundance of starch-hydrolyzing organisms (SHOs) in these plants. In this study, fluorescence in situ enzyme staining with BODIPY fluorescein-labeled starch was applied and optimized to label SHOs expressing α-amylase in activated sludge plants. Fluorescence on the surface of bacteria expressing α-amylase activity was clearly visualized. In 11 full-scale nutrient-removing wastewater treatment plants examined, the morphotypes of the dominant SHOs were always cocci in clusters of tetrads, short rods in clusters, and some filamentous organisms. The SHOs were identified by combining in situ enzyme staining and FISH using a range of available oligonucleotide probes. All the SHOs observed were Actinobacteria, and most had the phenotype of polyphosphate-accumulating organisms closely related to the genus Tetrasphaera in the family Intrasporangiaceae. The SHOs were present in most of the wastewater treatment plants examined and comprised, in total, up to 11% of bacterial biovolume and thus formed an important part of the microbial communities.

Keywords
  • starch-hydrolyzing organisms
  • activated sludge
  • BODIPY
  • PAOs

Introduction

Hydrolysis carried out by microorganisms is the first step in the degradation of most organic matter in municipal wastewater treatment plants (WWTPs). The organic substances in domestic wastewater typically consist of 40–60% proteins, 25–50% polysaccharides, and 10–30% lipids (Nielsen et al., 1992), and these macromolecules must be hydrolyzed to their monomers or oligomers before being degraded further. Hydrolysis is also important for the removal of nitrogen (N) and phosphorus (P) in nutrient removal WWTPs because the hydrolysates such as amino acids and their fermentation products (e.g. short-chain fatty acids, SCFAs) are important carbon and nitrogen sources for phosphorus- and nitrogen-removing microorganisms (Rieger et al., 2001). Hydrolysis is a slow process and is usually the rate-controlling step for the phosphorus- and nitrogen-removal processes (Dueholm et al., 2001; Morgenroth et al., 2002). Microbial hydrolysis in activated sludge is carried out by extracellular enzymes (exoenzymes) excreted by microorganisms. These exoenzymes are primarily located on the cell surfaces, where hydrolysis and release of partly degraded macromolecules are repeated until hydrolytic fragments are small enough to be assimilated by the microorganisms (Confer & Logan, 1998; Goel et al., 1998; Kloeke & Geesey, 1999).

Little is known about the identity of the microorganisms involved in hydrolysis in nutrient-removal WWTPs. However, new in situ techniques have been developed recently that can visualize exoenzyme-producing bacteria directly in complex environments. Surface-associated activity of phosphatases, lipases and some other exoenzymes in activated sludge can be detected using the ELF® 97 substrates provided by Molecular Probes (VWR International, Rodovre, Denmark). The principle is that when a nonfluorescing soluble substrate e.g. ELF® 97 palmitate is cleaved by lipases located on the surface of the hydrolytic microorganisms, fluorescent products precipitate on the surface of bacteria, making these visible by epifluorescence microscopy. The ELF approach has been used in combination with FISH to identify phosphatase-excreting microorganisms in activated sludge (Kloeke & Geesey, 1999). Also, lipase-excreting microorganisms have been identified and the filamentous bacteria Candidatus‘Microthrix parvicella’ (Nielsen et al., 2002) and Gordonia amarae-like organisms (Kragelund et al., 2007a) are believed to be among the important bacteria involved in lipid hydrolysis.

The activity of surface-associated exoproteases can be detected using 4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacen-3 (BODIPY)-conjugated substrates such as casein and bovine serum albumin (BSA) (Xia et al., 2007). The principle is that the proteins can be conjugated with dye-labeled BODIPY in such a way that almost the entire fluorescence is quenched in the conjugated substrates, and no or little fluorescence can be visualized. Once the conjugated substrates are hydrolyzed by proteases located on the surface of protein-hydrolyzing organisms (PHOs), fluorescence is released, making the PHOs visible by epifluorescence microscopy. FISH analysis showed that the main protein hydrolyzers in activated sludge are epiphytic bacteria (Candidatus genus Epiflobacter) belonging to the family Saprospiraceae of the phylum Bacteroidetes, and some filamentous bacteria belonging to the phylum Chloroflexi, Proteobacteria (related to the genus Aquaspirillum in the class Betaproteobacteria), and the candidate phylum TM7 (Xia et al., 2007, 2008).

No methods are available to detect and identify microorganisms involved in polysaccharide hydrolysis. In activated sludge, polysaccharide mainly includes starch and cellulose (Murakami, 1988), and starch has frequently been used as a model substrate to study the hydrolysis of polysaccharides (Ubukata, 1999; Karahan et al., 2006a, b). Starch is hydrolyzed by exoenzymes of starch-hydrolyzing organisms (SHOs) to a mixture of malto-oligosaccharides and finally to low-molecular-weight sugars such as maltose and glucose (San Pedro et al., 1994; Ubukata, 1999; Karahan et al., 2006a). These hydrolysates may, under anaerobic conditions, be further fermented to SCFAs, which are important carbon and energy sources for polyphosphate-accumulating organisms (PAOs) and denitrifiers (Kong et al., 2004, 2005; Thomsen et al., 2007), contributing to enhanced biological phosphorus removal (EBPR) and nitrogen removal. Starch hydrolysis may, therefore, be an important process and a rate-limiting step for nutrient removal in activated sludge (San Pedro et al., 1994).

Detection of α-amylase activity in the degradation of polymers (starch acetate) has been conducted using BODIPY fluorescein-labeled (FL) DQ starch (Tuovinen et al., 2004). In principle, it is similar to BODIPY FL casein and BSA, and so fluorescence is released if BODIPY FL starch is hydrolyzed by α-amylase (Legaz & Kenny, 1976; Royse & Jensen, 1984). However, it has never been tested on microorganisms whether the fluorescent-labeled hydrolysates actually precipitate on the surface of cells excreting α-amylase and thus can be used to label and identify SHOs in mixed microbial communities. In this study, we have developed a fluorescence in situ enzyme-staining approach using BODIPY FL DQ starch to specifically label SHOs in activated sludge, and have further identified the SHOs by combining the staining method with FISH probing. The distribution and abundance of the SHOs were investigated and their roles in WWTPs are discussed.

Materials and methods

Bacterial strains and culture conditions

Bacillus stearothermophilus (NRRL B-1172) and Bacillus coagulans (NRRL B-14317) were purchased from ARS Culture Collection (USA) (http://nrrl.ncaur.gov). Bacillus strains DSM 2334 and DSM 2349 were purchased from DSMZ Culture Collection (Germany) (http://www.dsmz.de). The medium (DSMZ medium 1) (http://www.dsmz.de) used to cultivate these cultures contained (L−1) peptone 5 g, meat extract 3 g, and, if necessary, agar 15 g. Bacterial cells used in staining tests were prepared in the following way: 50 mL of the sterile medium (in 250-mL Erlenmeyer flasks) was inoculated with 0.1 mL of logarithmically growing cells and incubated at 55 °C on a rotating disk (250 r.p.m.) for 10–15 h (Daron, 1967).

Activated sludge sources

Activated sludge samples were collected from 11 Danish full-scale WWTPs. The influent characteristics and configuration of the plants are described in Table 1. For all experiments, fresh sludge samples were taken from the aerobic tanks and transferred to the laboratory within 0.5–2 h.

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1

Morphotypes of SHOs, stained with BODIPY FL starch, in full-sale WWTPs

BODIPY FL DQ starch staining and incubation conditions

BODIPY FL DQ starch substrate [EnzChek® Ultra Amylase Assay Kit (E33651)] was purchased from Molecular Probes. The working solution was prepared as recommended by the manufacturer. Incubations of pure cultures or activated sludge samples with the BODIPY FL starch were carried out in 10-mL serum bottles, and the final reaction volume was c. 600 μL. Four hundred microliters of fresh-activated sludge biomass [(4 g L−1 mixed liquor suspended solids (MLSS)] or cell suspension (3.2 g dry weight L−1) was transferred into 1.5-mL Eppendorf tubes and spun down (4500 g for 10 min). The supernatant was removed. The BODIPY FL starch was first used at a 2 : 1 ratio, i.e. biomass in 400 μL of cell suspension (3.2 g dry weight cells L−1) was resuspended in 400 μL 1 × reaction buffer +200 μL working solution of BODIPY FL starch, based on our previous experience with staining for PHOs (Xia et al., 2007). The fluorescent signals were examined after 30 min according to the manufacturer's instructions.

To determine the optimal substrate concentrations (in the presence of inhibitors) at which high fluorescent signals on bacterial cells can be observed with a relatively low fluorescent background (signal/noise ratio), the signal/noise ratios for visual determination of SHOs were examined at different BODIPY starch concentrations (3.3, 8.3, 10, 16.7, 23.3, 33.3, 50, 66.7, and 100 mg g−1 MLSS) using sludge samples from AAV and Skagen WWTPs. The optimal incubation time at which the fluorescently stained cells could be observed with a low background under a specific substrate concentration was determined by microscopic observation of the stained samples every 15 min for 180 min. Depending on the final BODIPY FL starch concentration, different amounts of freshly prepared 1 × reaction buffer (1 M MOPS, pH 6.9) were added, and the mixtures were transferred into 10-mL serum bottles and supplemented with the working solution of BODIPY FL starch to a final volume of 600 μL. When necessary, the inhibitors sodium azide and sodium fluoroacetate (Sigma-Aldrich Logistik GmbH, Germany) and sodium iodoacetate (from Merck KGaA Darmstadt, Germany) were added (from a 50 × concentrated stock solution) at a final concentration of 3, 2, and 4 mM, respectively (Xia et al., 2007). The inhibitors were added 20 min before adding the BODIPY FL starch working solution. If the inhibitors were used, the same volume of 1 × reaction buffer was removed to keep the same final reaction volume. Once all the reagents had been added, the serum bottles were immediately wrapped in aluminum paper to exclude light and mixed on a rotating disk (220 r.p.m. at 20±1 °C). Biomass samples were spread on glass slides (see below for details) at different time intervals for microscopic examination. Sludge samples from each plant were stained three times during a 1-year period.

In some experiments, preincubations of activated sludge samples with unlabeled soluble starch were carried out to see whether induction of more SHOs was possible. Fresh sludge (20 mL, 4 g L−1 MLSS) was incubated in 100-mL flasks in the presence of starch (400 mg L−1 final concentration) under aerobic conditions for 1, 6, and 12 h. The incubations were carried out on the same rotating disk mentioned above. The biomass was washed three times with the supernatant from the same plant before incubation with BODIPY FL starch.

Microscopic examination

Activated sludge samples or pure cultures incubated with BODIPY FL DQ starch were spread on gelatin-coated three-well (10–15 μL in each well) Teflon-printed slides (Electron Microscopy Sciences, Hatfield, PA) in a darkroom and allowed to air-dry in a fume hood at room temperature. The slides were mounted with CITI fluor (Citifluor Ltd, London) and examined microscopically. The fluorescent signals of sludge samples stained with BODIPY FL DQ starch (excitation/emission maxima 505/512 nm) were visualized and captured through the FLUOS filter (excitation/emission maxima 450–490/515 nm) using an epifluorescence microscope (Axioskop 2 Plus, Zeiss) equipped with a charge-coupled device (CCD) camera (CoolSNAP HQ, Photometrics, Oberkochen, Germany). The software imagej (http://rsb.info.nih.gov/ij/) was used for image analysis and biovolume determination.

Keeping all the microscope setups constant, the exposure time for capturing images showing clearly fluorescing bacterial cells that stained with BODIPY FL starch was 0.03 s for fluorescing bacterial cells with α-amylase activity (B. stearothermophilus and B. coagulans), whereas it was 0.3 s for the bacterial cells without α-amylase activity (Bacillus strains DSM 2349 and DSM 2334). The minimum exposure time observed for the bacterial cells without α-amylase activity did not increase with time (up to 2 h).

BODIPY FL starch staining combined with FISH

BODIPY FL starch staining was combined with FISH probing to identify the SHOs. The fluorescent signals of stained sludge samples were examined as described above. The bacterial cells stained positively for α-amylases were located microscopically, and their positions on the microscopic stage were recorded. CITI fluor on the slides was washed away by gently rinsing (for 1 min) with 70% ethanol before fixation in either 4% paraformaldehyde (for Gram-negative bacteria) or in 50% ethanol (for Gram-positive bacteria) for 2–4 h. For increasing permeability of gene probes, slides for FISH probing of Gram-negative bacteria were also fixed in 50% ethanol for 2–4 h before FISH probing. FISH was carried out according to Amann (1995). Except EUBmix, which was labeled with FLUOS or Cy5, all the oligonucleotide probes used were labeled with Cy3. FISH signals of the bacteria of interest were examined after relocation following the procedure described previously (Xia et al., 2007). In all FISH experiments, probe NONEUB (Wallner et al., 1993) was used as a control for unspecific binding of gene probes. It is not possible to perform FISH before the BODIPY FL starch staining, because the fixation procedure inactivates the exoenzymes.

All the oligonucleotide probes used in this study are listed in Table 2. The specificities, hybridization conditions, use of competitors, and reference information of most of the probes are described in probeBase (Loy et al., 2003). To improve the permeability of probe Myc657, enzymatic pretreatment (Kragelund et al., 2007a) was included.

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2

Oligonucleotide probes used in this study

Biovolume determination

The relative biovolume of all SHOs or the filament-SHOs was determined on a very thin layer of biomass by measuring the percentages of area fluorescing after BODIPY FL starch staining out of the area fluorescing with the EUBmix probe after relocating to the same microscopic field. At least 40 microscopic fields (× 1000) were analyzed for each enumeration (at least 10 for the filament-SHOs). Measurements from each plant were repeated three times during the 1-year experimental period, and the average and SD are reported.

Quantitative FISH was carried out following the procedure described previously (Kong et al., 2006). The percentage of the coccus–SHOs hybridized with Actino-221 was estimated by counting the numbers of cocci labeled by BODIPY FL starch and the cocci hybridized with Actino-221 in 20 fields (in total, c. 300 cells were examined for each sample). The percentage of the rod-SHOs hybridized with Actino-658 was estimated from the fluorescent area of the rod-SHOs labeled by BODIPY FL starch and from the area of the rods hybridized with Actino-658. This was done because the short rods, often forming clusters, were too small to be counted. For each sample at least 20 fields were examined.

Results

BODIPY FL starch staining of pure bacterial cultures

No protocols were available for utilization of the BODIPY FL starch to stain bacterial cultures or activated sludge, and so BODIPY FL starch was first applied on pure cultures with and without α-amylase activity. Bacillus stearothermophilus and B. coagulans have α-amylase activity (Nazina et al., 2001), and strong fluorescent signals on most (>95%) of their vegetative cells were observed against a strong fluorescent background after staining for 30 min, the optimal incubation time according to the manufacturer for measuring pure α-amylase activity. A few endospores were always present, and they were not stained. The fluorescence intensity observed on both bacterial cells and as a background became stronger with staining time (up to 2 h). For the Bacillus strains DSM 2349 and DSM 2334 without α-amylase activity (Daron, 1967; Epstein & Grossowi, 1969), only a weak fluorescence signal was observed on a few (1–3%) cells against a relatively nonfluorescent background, and no fluorescence signal was observed on most cells. Moreover, the fluorescence intensity observed for the cells did not change significantly when the incubation time was extended to 2 h. The weak fluorescence observed on a few cells could be due to autolysis, where BODIPY FL starch reacts with intracellular α-amylase, releasing fluorescence.

Application and optimization of BODIPY FL starch in activated sludge samples

BODIPY FL DQ starch was applied in activated sludge to stain for SHOs. After 30 min of staining, fluorescence on different cell morphotypes was observed on a fluorescent background. Mainly cocci in clusters of tetrads (Fig. 1a and b), short rods in clusters, and filamentous bacteria fluoresced (Fig. 1c and d). Other morphotypes did not appear during 180 min of staining (microscopic examination was carried out every 15 min). The background fluorescence, however, increased with time.

1

Images of activated sludge after BODIPY FL starch staining and FISH. Image (a) shows cocci in tetrads stained in BODIPY FL starch staining. The squares in image (a) correspond in their positions in image (b) (bright field). Image (c) shows filamentous organisms stained in BODIPY FL starch staining. The arrows in (c) and (d) (bright field) mark the same positions. Images (e) and (f) show the cocci in cluster of tetrads stained with BODIPY FL starch (e) hybridizing with probe Actino-221 [(f), shown as the yellow-colored cells labeled with arrow A], and some Actino-221-positive cells [(f), labeled with arrow B] not stained in BODIPY FL starch staining. The green color indicates hybridization with EUBmix.

BODIPY FL starch staining was also carried out using sludge samples from 10 other WWTPs (Table 1). The SHO morphotypes observed in these WWTPs were the same as those observed in AAV. However, not all the morphotypes were found in each plant. Except in Bjergmarken, Horsens, Aars, Middelfart, and Kerteminde WWTPs, where 0–2 morphotypes were present, all three SHO morphotypes were present in all EBPR and nitrogen-removal WWTPs. No positive signals were observed in sludge samples from Kerteminde WWTP. No fluorescing cells and background were observed in control samples boiled for 15 min, indicating that the fluorescence observed in all these WWTPs was due to the activity of starch-hydrolyzing enzymes excreted by microorganisms.

To specifically label the SHOs in activated sludge and not the consumers of labeled hydrolysate, a set of inhibitors, as used in our previous study on proteases (Xia et al., 2007), was added in the staining incubations. This prevented the consumers of FL starch hydrolysates (e.g. mixture of malto-oligosaccharides, maltose and glucose) from being labeled. Iodoacetate, fluoroacetate, and azide were added to inhibit the glycolysis (Bickis & Quastel, 1965), the TCA cycle (Peters et al., 1953), and the electron transport chain (Myers & Nealson, 1988), respectively. The individual inhibitors were added at concentrations at which the energy metabolisms of all the microorganisms in activated sludge were effectively inhibited (Xia et al., 2007). In the presence of the inhibitors, all the previously observed fluorescing morphotypes were still observed to fluoresce, the only difference being that the number of filamentous morphotypes observed was markedly reduced.

The optimal substrate concentrations and incubation time (in the presence of inhibitors) at which high fluorescent signals on bacterial cells could be observed with a relatively low fluorescent background (signal/noise ratio) were determined. After 30 min of staining, optimal signal/noise ratios were obtained at substrate concentrations between 16.7 and 23.3 mg g−1 MLSS and thus selected as optimal incubation time and substrate concentrations. All the fluorescing morphotypes observed previously were present, and the minimum exposure time for capturing images showing fluorescing bacterial cells was between 0.03 and 0.1 s. At substrate concentrations <16.7 mg g−1 MLSS, the fluorescence signals observed on bacterial cells were weak, and at substrate concentrations >23.3 mg g−1 MLSS, the background fluorescence was too high to distinguish the fluorescent cells from the background. The Bacillus strains with exo-α-amylase activity were also stained under these conditions.

In order to ensure labeling of all SHOs potentially active in activated sludge, preincubations of activated sludge samples with unlabeled soluble starch (without the inhibitors) were carried out in some experiments before staining with a BODIPY FL starch to induce or enhance the α-amylase activities. This was done to rule out the possibility that the α-amylase activity of some SHOs might require a longer (>2 h) induction time or that their in situα-amylase activity was too weak to be detected within 30 min of enzyme staining. The preincubations were carried out under aerobic conditions for 1, 6, or 12 h. The same fluorescing morphotypes were still observed in sludge samples from all WWTPs investigated, indicating that the main SHOs were labeled after 30 min staining. Therefore, no preincubation was included in the subsequent experiments. Interestingly, this also implies that other microorganisms in activated sludge could not be induced to produce α-amylases within 12 h.

Identification of SHOs using enzyme staining combined with FISH

FISH probing using different oligonucleotide probes was used to identify the SHOs labeled with BODIPY FL starch staining. In the WWTPs examined, all the SHO morphotypes (cocci in clusters of tetrads, short rods in clusters, and filaments) hybridized only with probes HGC69a and HGC236 designed for Actinobacteria. They did not hybridize with ALF968, BET42a, GAM42a, and SRB385+385Db designed for the classes Alpha-, Beta-, Gamma-, and Deltaproteobacteria, respectively, with probe mixtures LGCmix designed for Firmicutes, with Bac111 designed for sludge clones in Saprospiraceae in the phylum Bacteroidetes, with GNSB941 and CFX1223 designed for most members in the phylum Chloroflexi, or with probe TM7-905 designed for the candidate phylum TM7. For all these probes tested, bacteria were present in the WWTPs without being stained with BODIPY FL starch staining.

Further FISH probing using more specific probes targeting Actinobacteria showed that, in all the WWTPs examined, most (>80%) of the coccus-SHOs hybridized with probe Actino-221 designed for the uncultured Actinobacteria related to genus Tetrasphaera (Fig. 1e and f). A varying fraction of the rod-SHOs hybridized with Actino-658 (Table 2). For example, rods hybridized with Actino-658 accounted for 6% of the total bacterial biovolume in Aars WWTP (Table 3), but none of these stained for α-amylase (Table 1). In other WWTPs examined, 10–70% (naked eye estimation) stained for α-amylase. Interestingly, some rod-SHOs also hybridized with probe NLI-265 designed for the filamentous Candidatus‘Nostocoida limicola’. A small fraction of the coccus-SHOs and some rod-SHOs did not hybridize with probes Actino-221 and Actino-658. Only a few filament-SHOs in Hjorring WWTP hybridized with probe Myb736a designed for the Mycobacterium complex. All the other actinobacterial filament-SHOs (Table 3) did not hybridize with any other probe tested, including Myc657 targeting the Mycobacterium subdivision (mycolic acid-containing actinomycetes), NLIMII-175 targeting Nostocoida limicola II strains, and MPA-645 targeting Candidatus‘Microthrix parvicella’. Their identity, therefore, is still not clear.

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Biovolume of total amount of SHOs and probe-defined potential SHOs in full-scale WWTPs

Distribution and abundance of the SHOs in wastewater treatment plants

The distribution and abundance of the total SHOs and the probe-defined SHO groups in 11 full-scale WWTPs are listed in Table 3. The biovolume of all SHOs measured ranged from none to 11% of the total bacterial biovolume. Middelfart and Kerteminde, both nitrogen-removal plants with chemical phosphorus precipitation, had none or very few SHOs. No significant difference in the total biovolume of SHOs was found in the other plants despite their differences in influent characteristics (municipal or industrial) and configurations (EBPR or nitrogen-removal with chemical phosphorus precipitation). The groups defined by probes Actino-221 and Actino-658 in all plants except Middelfart and Kerteminde accounted for 6±2% and 7±2% of the total bacterial biovolume, respectively. The actinobacterial filament-SHOs comprised <1% of the total bacterial biovolume in all plants investigated.

Discussion

In situ detection of the SHOs with exo-α-amylase activity using the BODIPY FL starch staining

The experiments with pure cultures expressing exo-α-amylase and several activated sludge samples clearly showed that a part of the hydrolyzed BODIPY FL starch precipitated on the surface of the bacteria, making a visual detection of SHOs possible. Only a part of the fluorescent hydrolysates precipitated onto the bacteria as fluorescence increased with time both on the bacterial surface and in the solution. Therefore, it was important to inhibit other bacteria from taking up some of the fluorescently labeled hydrolysates by various metabolic inhibitors as was also necessary for detection of protein hydrolyzers (Xia et al., 2007). The negative controls showed that the fluorescent intensity was insignificant in non-SHOs, indicating that SHOs with exo-α-amylase activity present in full-scale WWTPs were reliably labeled using the BODIPY FL starch staining method. The strong fluorescence observed on the surface of the SHOs confirms that most exoenzymes in microbial aggregates are mainly associated with the cell surface (Confer & Logan, 1998; Goel et al., 1998; Kloeke & Geesey, 1999; Nielsen et al., 2002; Xia et al., 2007).

The use of inhibitors to exclude consumers of starch hydrolysates had no effect on the level of fluorescence of the cocci in clusters of tetrads, of short rods in clusters, and of many filamentous organisms. The number of fluorescent filaments was, however, significantly reduced, indicating that some filaments were most probably consumers of starch hydrolysates and did not specifically produce exo-α-amylase. This observation makes sense as a number of filamentous organisms, e.g., most members of Chloroflexi (Kragelund et al., 2007b), and Type 021N (Williams & Unz, 1985) can take up glucose and other sugars in activated sludge while very few other probe-defined species in activated sludge can do so (Kragelund et al., 2007b; Thomsen et al., 2007).

The possible unspecific labeling of bacteria undergoing autolysis was overcome by adopting short exposure time to capture only images of the fluorescing morphotypes (putative SHOs), followed by their identification using FISH probing. Intracellular α-amylase as well as other enzymes from cell autolysis are present in the extracellular polymeric substances (EPS) matrix in activated sludge (Rudd et al., 1983; Urbain et al., 1993), but they are insignificant compared with the amount of exoenzymes (Frolund et al., 1995). This observation is also supported by a pure culture study of B. subtilis, where the intracellular α-amylase activity was insignificant compared with the activity of exoenzymes (Coleman & Elliott, 1962). However, we cannot completely rule out the presence of bacteria producing low levels of α-amylase, and so the method primarily provides information about the presence of highly active α-amylase producers. Likewise, the possibility that some bacteria primarily excrete α-amylase into bulk liquid cannot be ruled out, and these bacteria would be missed in our SHOs screening.

The preincubation with unlabeled starch before incubation with BODIPY FL starch failed to reveal more types of SHOs. Therefore, the SHOs identified using BODIPY FL starch staining combined with FISH were the SHOs that were active under in situ conditions. Moreover, although α-amylase is the main amylolytic enzyme, the presence of other amylolytic enzymes e.g. β-amylase, α-glucosidase, pullulanase and glucoamylase could not be ruled out. Clostridium thermosulfurogenes, an anaerobic bacterium secreting β-amylase and glucoamylase, can hydrolyze starch in the absence of significant pullulanase or α-amylase activity (Hyun & Zeikus, 1985). Thus, SHOs having amylolytic activity by enzymes other than α-amylase or a weak α-amylase activity may also be missed in our screening. This possibility could not be confirmed because the α-amylase-specific BODIPY FL starch is the only substrate available among this group of substrates.

Identity of SHOs in full-scale WWTPs

Interestingly, all the SHOs in 11 WWTPs examined were Actinobacteria. The dominant SHO morphotypes were cocci in clusters of tetrads and short rods in clusters hybridized with probes Actino-221 and Actino-658, respectively. Phylogenetically, they are related to uncultured Actinobacteria with a PAO phenotype relatively close to the genus Tetrasphaera in the family Intrasporangiaceae. The hybridization of the rod-SHOs with probe NIL-265 also supports their affiliation to the genus Tetrasphaera because, although designed for the Candidatus‘Nostocoida limicola’, probe NIL-265 also perfectly matches some of the Tetrasphaera-related clones targeted by probe Actino-658.

The phylogeny of the SHOs not hybridizing with Actino-221 and Actino-658 is still not clear and has to be investigated further. The identity of the filamentous-SHOs was not investigated further as only a few specific probes targeting filamentous Actinobacteria are available.

Some ecophysiological aspects of SHOs in full-scale WWTPs

The SHOs identified in this study were widespread in full-scale WWTPs and present in almost all the WWTPs examined (except one), constituting up to 11% of the total bacterial biovolume. The abundance of the SHOs (Table 3) might have been slightly overestimated because the fluorescent areas of the individual SHO cells resulting from the surface-associated BODIPY FL starch staining are slightly larger than the areas resulting from FISH probing targeting intracellular RNA, as can be seen in Fig. 1e and f. The abundance varied in the WWTPs. This might have be due to different levels of starch in the influent but this could not have been related to the available information about influent characteristics or plant configuration. The presence and relative abundance of the SHOs in most WWTPs did not change significantly (naked eye observation) during the 1 year experimental period, and so the SHOs seem to be common members of the microbial communities in nutrient-removing WWTPs.

The biovolume of SHOs measured by FISH using probes Actino-221 and Actino-658 is only an approximate estimation of all coccus and rod-SHOs in WWTPs. Not all the bacteria hybridizing with these probes were SHOs, and not all the coccus-SHOs and rod-SHOs hybridized with these two probes. The biovolumes measured with probes Actino-221 and Actino-658 in WWTPs were three to four times higher than the total biovolume of SHOs in most plants. For example, the Actino-658 defined group accounted for a significant fraction (6±2%) (mean value±SD) of the microbial community in Aars, but none of them behaved as active SHOs. Most probably, some hidden biodiversity exists among the SHO groups.

Bacteria defined by probes Actino-221 and Actino-658 are considered to be PAOs involved in EBPR and this study shows that they are also main SHOs. Very interestingly, recent results have shown that the Actino-221 defined cocci are also involved in fermentation of glucose (Kong et al., 2007, 2008), a behavior agreeing well with hydrolysis of starch. It seems that they are multifunctional bacteria involved in hydrolysis, fermentation, and phosphorus-removal, which, to our knowledge, has never been reported before for any bacterial group. However, how these functions are regulated is still unclear. Moreover, as discussed earlier, hidden biodiversity, i.e. the presence of different species or different ecotypes of a probe-defined group, could not be ruled out. This is especially true for the Actino-658 defined group as their percentages of being active SHOs varied greatly in different WWTPs.

In this study the focus was on degradation of starch but other polysaccharides may also be degraded in activated sludge. We also attempted to investigate the cellulose degradation using EnzChek® Cellulase substrates [EnzChek® Cellulase Substrate blue fluorescent, 339/452 (E33953)] from Molecular Probes. The cellulase substrate, prepared as recommended by the manufacturer, was applied in sludge samples from 11 WWTPs, but no or very little positive signals were detected (data not shown). Perhaps, the cellulase activity was too weak to be detected because of its high resistance to enzyme hydrolysis (Beguin & Aubert, 1994), resulting in the presence of high levels of cellulose in the excess sludge (Honda, 2002). Also, surface-associated activities of β-d-galactosidase and β-d-glucuronidase, which are involved in hydrolysis of oligosaccharides (lactose and β-d-glucopyranosiduronic derivatives, respectively), were determined using ELF® 97 β-d-galactopyranoside and ELF® 97 β-d-glucuronide, following the procedure by Kragelund (2005). Only some unidentified filaments were positive. Future studies are needed to show whether other poly- and oligosaccharides are degraded by specialized populations similar to those degrading starch and protein.

Acknowledgements

This study was supported by the Danish Technical Research Council in the program ‘Identification and characterization of uncultured bacteria involved in hydrolysis and fermentation in nutrient removal plants’ (grant number 26-04-0115). We are grateful to L.H. Pedersen for providing pure culture of B. stearothermophilus and B. coagulans.

Footnotes

  • Editor: Michael Wagner

References

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