‘Morphological shifts’ in filamentous bacteria isolated from activated sludge processes T.R. Ramothokang*, D. Naidoo and F. Bux Centre for Water and Wastewater Technology, Durban Institute of Technology, PO Box 1334, 4000, Durban, South Africa *Author for correspondence: Tel.: +27-31-204-2346, Fax: +27-31-204-2778, E-mail:
[email protected] Received 16 September 2005; accepted 21 December 2005
Keywords: activated sludge, filamentous bacteria, fluorescent in situ hybridization, microscopic identification, ‘morphological shifts’
Summary This study aimed at isolating filamentous bacteria from full-scale activated sludge processes and studying them in pure culture. Three cultures were isolated using conventional microbiological techniques. The isolates were positively identified as Gordonia amarae, Thiothrix nivea and Type 1863/Acinetobacter spp., using fluorescent in situ hybridization (FISH) with 16S rRNA-targeted oligonucleotide probes. However, a ‘morphological shift’ from filamentous to single-cell form was observed in pure culture. The application of fluorescent in situ hybridization (FISH) showed filamentous bacteria to be much more diverse in their ability to adapt to their changing enviroments. Pure culture studies of filamentous bacteria form the basis for application in full-scale activated sludge plants. It therefore remains important that the taxonomic status of filamentous bacteria be determined.
Introduction Bacterial adaptation to different novel environments usually necessitates morphological changes in response to variations in the environment (Shi & Xia 2003). Variations in temperature, pressure, pH, salinity, concentration of nutrients and other environmental factors have been shown by previously published studies to have the ability to start an intricate series of cellular events that include changes in cellular morphology (Alonso et al. 2002). Filamentous bacteria isolated from activated sludge processes have also been identified in single-cell form (Rosetti et al. 1997). Filamentous bacteria are of interest due to the important roles they play in sludge settleability and filamentous bulking and foaming in activated sludge processes. Bulking sludge settles and compacts slowly, while foaming shows a high level of scum in aeration basins, due to filamentous organisms dominating the floc-formers within the mixed liquor (Richard 1989). Numerous attempts to control bulking and foaming have been made, but they have not been successful, largely due to the failure to identify the causative organisms (Eikelboom 1975). Isolation of filamentous bacteria is a pre-requisite for the investigation of these organisms (Ka¨mpfer 1997). Pure culture studies have shown promise as an essential tool for studying filamentous bacteria on the basis of morphology,
nutritional requirements and physiology. Such knowledge is imperative in order to eventually fully understand activated sludge bulking and foaming phenomena. Isolation of filamentous bacteria is an essential requirement for the investigation of biochemical and metabolic kinetics. However, systematic approaches to the nutritional requirements of different filamentous organisms are limited due to the difficulty of isolating them (Ka¨mpfer et al. 1995; Ka¨mpfer 1997). Procedures for the identification of filamentous bacteria previously relied on morphological characteristics and their response to a number of simple staining techniques (Seviour et al. 1994). The use of morphological criteria for prokaryotes is, however, unreliable as indicators of relatedness, and organisms that look the same, even to a trained eye, may not necessarily be so. It has become clear that organisms like the nocardioform bacteria, which look very similar in mixed liquor or foam samples under the microscope, differ widely in their taxonomy, physiology and biochemistry in pure culture (Seviour et al. 1994). Similarly, some foamproducing organisms like Nocardia pinensis (Soddell & Seviour 1994 as cited by Seviour et al. 1994) may look quite different in foams, and yet very similar in pure culture. Variations in morphology have now also been reported in several other filamentous organisms including Microthrix parvicella (Foot et al. 1992 as cited by
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Seviour et al. 1994), Types 0961 and 0092 (Buali & Horan, 1989 as cited by Seviour et al. 1994) and Type 021N (Ziegler et al. 1990 as cited by Seviour et al. 1994). A chance of incorrect identifications therefore exists, with possibilities that the filaments presently recognized are much more diverse in their properties, and may in fact represent groups of quite different organisms all looking similar (Williams & Unz 1985 as cited by Seviour et al. 1994). Molecular techniques have recently been used in microbial ecology studies in an attempt to overcome the limitations of culture techniques (Van Heerden et al. 2000). The use of gene probes for the in situ identification of filamentous bacteria in activated sludge has been found successful. Oligonucleotides (short strands of nucleic acids, usually 15–30 nucleotides in length), complementary to 16S rRNA and 23S rRNA sequence regions with an intermediate degree of conservation and characteristic for phylogenetic entities like species, genera, families and subclasses, have been used successfully for rapid identification of bacteria. The technique is called fluorescent in situ hybridization (FISH) or whole cell probing (Lindrea et al. 1999). This study aimed at isolating filamentous bacteria from full-scale activated sludge processes and studying them in pure culture. It was with the application of FISH that pure isolates of filamentous bacteria were identified in single-cell form, thereby confirming that filamentous bacteria are indeed much more diverse in their properties than was previously believed.
and Nonidet, respectively to disperse the floc. After pretreatment, samples were observed under the microscope to assess the success of each pre-treatment procedure (Ramothokang et al. 2003). This was followed by a wash step in Mineral–Salt–Vitamin (MSV) solution and centrifugation at 1898 g and 132 g for 2–5 min respectively in order to separate floc-formers from filamentous bacteria. This facilitated easier isolation of the latter. Pre-treated samples were then serially diluted to 10)1–10)5 in MSV and cultured on a series of media (R2A, CGYA, SCY, I Medium, TGYA, Nocardia Medium, D-Medium, C-Medium and Actinomycete Isolation Agar) employing the spread plate technique, and incubated at 20–25 C for at least 3 weeks. Individual colonies were then plated onto fresh agar plates of the same media components until pure cultures were obtained. Traditional microscopic identification of isolates Isolates were stained and viewed under the microscope according to Jenkins et al. (1986) to select those that exhibited cellular morphology typical of filamentous bacteria. Presumptive filamentous bacteria were selected for further investigations and a more accurate identification by application of the molecular technique, FISH. These isolates were coded HCBCG01 (isolated on CGYA), OSI004b (isolated on I Medium) and SWNCG02 (isolated on CGYA). Oligonucleotide probes
Materials and methods Sampling Grab samples were randomly taken from wastewater treatment plants. The samples were stored half-full in Schott bottles so as to allow for aerobic conditions (Eikelboom 2001). The samples were screened and filamentous bacteria were identified using Gram stain, Neisser stain, PHB (polyhydroxybutyrate) stain, crystal violet sheath stain and cell morphology (Jenkins et al. 1986). Isolation of filamentous bacteria Prior to isolation, activated sludge samples were pretreated by bead-beating, sonication, cellulase hydrolysis
Probes were selected based on the frequency of occurrence of filaments in South Africa. The oligonucleotide probes were synthesized according to specification and were labeled with fluorescein at the 5¢ end (MWG-Biotech, Germany; subsidiary company of Roche´ Products (Pty) Ltd, South Africa). Eight probes (Table 1) were applied to all samples (only positive results are shown) (Table 2). Pure culture fixation The sample was fixed in 4% paraformaldehyde (Amann 1995). Three volumes of paraformaldehyde (PFA) fixative were added to one volume of sample. The samples were then kept at 4 C for 1 min to fix. The normal fixation time of 2–3 h was changed to 1 min. The
Table 1. Oligonucleotide probe sequences and stringencies. Probe
Sequence (5¢–3¢)
Organism
Formamide%
Reference
SNA TNI GA HHY 21N ACA TFR DLP
CATCCCCCTCTACCGTAC CTCCTCTCCCACATTCTA ATGA(CTY)GTCCCCTCTGA GCCTACCTCAACCTGATT TCCCTCTCCCAAATTCTA ATCCTCTCCCATACTCTA CTCCTCTCCACACTCTA CCACCATGCGGCAGGAGCTCA
Sphaerotilus natans Thiothrix nivea Gordonia amarae Haliscomenobacter hydrossis Type 021N Acinetobacter species Thiothrix fructosivorans Nocardioform filamentous bacteria
45 45 45 20 35 35 40 40
Wagner et al. (1994a) Kanagawa et al. (2000) Ka¨mpfer et al. (1996) Wagner et al. (1994a) Wagner et al. (1994a) Wagner et al. (1994b) Kim et al. (2002) Schu¨ppler et al. (1998)
Table 2. Cellular characteristics of the three isolates obtained. Filament strain
HCBCGO1
OSI004b
SWNCGO2
Gram reaction PHB storage Poly-P storage Branched or not Trichome size Sheath Trichome shape
Positive Positive Negative Negative 0.8 lm Negative Smoothly curved
Positive Positive Positive Negative 0.5–0.8 lm Negative Coiled
Positive Positive Negative Negative 1.0 lm Negative Smoothly curved
normal fixation period of 2 h compromised the cell morphology. This was possibly due to the excessive cross-linking of proteins in the cell wall by the PFA molecules (De Los Reyes et al. 1997). Experimental fixation of Gram-positive filamentous bacteria with ethanol was also found to compromise the cell morphology. The cells were pelleted by centrifugation (332g, 4 min) and the supernatant was removed. The cells were then washed in 1 PBS, followed by centrifugation at 332 g for 4 min. The pelleted cells were then resuspended in 1 PBS to 50% of the original sample volume. The cells were then spotted onto slides. In situ hybridization Hybridization was carried out in 50-ml polypropylene centrifuge tubes, which served as a chamber for wholecell hybridization. The tube used for hybridization should be properly sealed to prevent evaporative loss of the hybridization buffer, which might result in nonspecific binding of the probes to the cells (Amann 1995). The concentration of formamide in the hybridization buffers and sodium chloride in the wash buffers was optimized accordingly for each probe. If a high stringency is required formamide (up to 59%) can be added to the hybridization solution (Amann 1995). All the probes were diluted accordingly with the appropriate hybridization buffer. All eight probes were applied to each of the three isolates and to the activated sludge samples. Hybridization was performed according to the methods stated by Amann (1995).
Results and discussion Following are the images of pure cultures of filamentous bacteria demonstrating morphological changes from filaments to single cells in laboratory growth conditions and, their positive identification in their single cell form as filamentous bacteria using FISH. The isolate, HCBCGO1, was identified as Gordonia amarae using the species-specific probe GA (Figure 1). The culture displayed characteristics similar to the species Nocardia amarae. Nocardia amarae was renamed Gordonia amarae (De Los Reyes et al. 1998). Gordonia species form part of the mycolic acid-containing actinomycetes group (Davenport et al. 2000). This group of organisms has exhibited filamentous, coccoidal and rod-shaped forms in mixed liquor and foaming samples (Davenport et al. 2000). The culture identified as Gordonia amarae had single coccoidal and rod-shaped cells. HCBCGO1 was identified as a Grampositive isolate, which also demonstrated a positive result for PHB storage. However, the same isolate had presented a negative result for poly-P accumulation which may be due to the fact that it was grown in pure culture and might have therefore lost its polymer storage capabilities or shown this property due to the change in environment. The comparative analyses of the Gram and the PHB staining reactions with FISH results coincide with that of Gordonia amarae. The culture OSIOO4b was identified as Thiothrix nivea. Thiothrix species belong to the filamentous sulphur bacterial group (Jenkins et al. 1993). The TNI
Figure 1. Micrographs of the same isolate HCBCG0 1(1000 immersion oil). (a) Gram-positive HCBCGO1 3 weeks after isolation in the filamentous form. (b) HCBCGO1 hybridized with fluorescein (green)-labeled GA probe, 6 weeks after isolation.
Figure 2. Micrographs of the same isolate OSI004b (1000 immersion oil). (a) Gram-positive OSI004b 3 weeks after isolation in the filamentous form. (b) OSI004b hybridized with fluorescein (green)-labeled TNI probe, 6 weeks after isolation.
probe had bound to coccoidal cells (Figure 2b). A previous study of a plant in Høgeland, Germany, found the TNI probe had bound to coccoidal cells. It was hypothesized that the single cells represented gonidia, which is a non-filamentous form of Thiothrix (Nielsen et al. 1998). According to the criteria routinely used for morphological identification of Thiothrix spp., the cells may accumulate sulphur and are either square or longer than they are wide and are sometimes present as rosettes or gonidia (Pernelle et al. 1998). Thiothrix nivea cells are Gram variable when significant amounts of intracellular sulphur granules are present (Seviour et al. 1999). OSIOO4b was identified as being positive for the Gram reaction, PHB storage and poly-P accumulation. These phenotypic characteristics provide verification that the culture, OSIOO4b, is Thiothrix nivea. The culture SWNCGO2 showed positive hybridization for the ACA probe only (Figure 3b). The morphology of the hybridized cells was oval shaped. The morphology of this culture is analogous to that of Acinetobacter cells, which are characterized by oval shaped cells commonly referred to as cocco-bacilli (Blackall et al. 1998). Acinetobacter cells are Gram negative and they are Neisser positive for polyphosphate (Blackall et al. 1998). The probe ACA has also been reported to show positive hybridization signals for the filament Type 1863. Type 1863 has demonstrated variable morphology ranging from cocco-bacilli to rods in pure culture (Seviour et al. 1999). The cells are generally Gram negative but some cells within the filament have stained positive (Rosetti et al. 1997). The culture SWNCGO2
presented a positive reaction for PHB storage and for the Gram stain, but a negative reaction for poly-P accumulation which may, yet again, be due to the change in growth conditions in pure culture. It has been suggested that Acinetobacter and Type 1863 share a very close relationship (Wagner et al. 1994b). An Eikelboom Type 1863 strain RT2 was isolated, identified and phylogenetically placed in the gamma sub-class of the proteobacterium. Acinetobacter johnsonii, based on its 16S rDNA sequence (Rosetti et al. 1997). Based on the above results and comparison with literature it can be deduced that SWNCGO2 is Type 1863. Previously published research has also identified an Australian Type 1863 as Acinetobacter using the Biolog System (Seviour & Blackall 1999).
Conclusions The cultures had undergone a ‘morphological shift’, from a filamentous form to single cocci or rod shaped cells. The morphological changes had occurred after they had been isolated, and sub-cultured. The change in morphology had taken place while the cultures where stored at 4 C. All growth is normally stopped when they are stored at this temperature, however this was not the case. This change in morphology could have been a response to the change in the culture environment. Many bacteria form spores during adverse conditions and when favorable conditions prevail they revert to their normal morphological form or to vegetative cells.
Figure 3. Micrographs of the same isolate SWNCGO2 (1000 immersion oil). (a) Gram-positive SWNCGO2 3 weeks after isolation in the filamentous form. (b) SWNCGO2 hybridized with fluorescein (green)-labeled ACA probe, 6 weeks after isolation.
When they were isolated onto solid media there was a change in the supply of nutrients and pH, amongst other factors. All these factors could contribute to a stressful environment, which could trigger a morphological change, as a survival mechanism. To substantiate the findings of this study further, more pure culture work needs to be done. ‘Morphological shifts’ need to be examined in more detail to provide more definitive answers to the following questions: 1. Why do ‘morphological shifts’ occur? 2. What triggers the change in morphology? 3. Do these changes occur in pure culture only or in activated sludge as well and do they play the same role in the process regardless of their morphology? 4. Is there any link between the changes in filamentous bacterial morphology and process performance? In light of the findings of this study it is imperative that the extent of diversity and the current taxonomic status of filamentous bacteria be reviewed. The result of pure culture studies forms the basis for applications in full-scale activated sludge plants and to improve our understanding of bulking and foaming problems. Thus it is vital that for future work, conditions prevalent in the system are mimicked as closely as possible during pure culture studies. Acknowledgments The authors would like to thank the National Research Foundation of South Africa for their financial support and the eThekwini Municipality (Durban, South Africa) for their assistance. References Alonso, J.L, Mascellaro, S., Moreno, Y., Ferru˘s, M.A. & Herna´ndez, J. 2002 Double-staining method for differentiation of morphological changes and membrane integrity of Campylobacter coli cells. Applied and Environmental Microbiology 68, 5151–5154. Amann, R.I. 1995 In situ identification of microorganisms by whole cell hybridization with rRNA-targeted nucleic acid probes. In Molecular Microbial Ecology Manual, eds. Akkermans A.D.L., Elsas J.D. & Bruij F.J. Vol. 3.3.6., London: Kluwer. 0792334116. Blackall, L.L., Bond, P.L., Crocetti, G.C. & Christensson, M. 1998 The microbial diversity of complex consortia in waste treatment processes: correlation between process performance and microbial composition. In: Microbial Community and Functions in Wastewater Treatment Processes. Proceedings of the 2nd International Symposium of the Center For Excellence University of Tokyo. 94–101. De Los Reyes, F.L., Ritter, W. & Raskin, L. 1997 Group-specific small unit rRNA hybridization probes to characterize filamentous foaming bacteria in activated sludge. Applied and Environmental Microbiology 63, 1107–1117. De Los Reyes, F.L., De Los Reyes, F.L. II, Hernandez, M. & Raskin, L. 1998 Quantification of Gordona Amarae strains in foaming activated sludge and anaerobic digester systems with oligonucleotide hybridization probes. Applied and Environmental Microbiology 64, 2503–2512.
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