Editor: Gilbert Shama
Biofilm formation and interactions of bacterial strains found in wastewater treatment systems
Article first published online: 17 APR 2008
© 2008 Federation of European Microbiological Societies. Published by Blackwell Publishing Ltd. All rights reserved
FEMS Microbiology Letters
Volume 283, Issue 1, pages 83–90, June 2008
How to Cite
Andersson, S., Kuttuva Rajarao, G., Land, C. J. and Dalhammar, G. (2008), Biofilm formation and interactions of bacterial strains found in wastewater treatment systems. FEMS Microbiology Letters, 283: 83–90. doi: 10.1111/j.1574-6968.2008.01149.x
- Issue published online: 17 APR 2008
- Article first published online: 17 APR 2008
- Received 1 February 2008; accepted 28 February 2008.First published online 14 April 2008.
- biofilm formation;
- initial adherence;
- wastewater treatment
Biofilm formation and adherence properties of 13 bacterial strains commonly found in wastewater treatment systems were studied in pure and mixed cultures using a crystal violet microtiter plate assay. Four different culture media were used, wastewater, acetate medium, glucose medium and diluted nutrient broth. The medium composition strongly affected biofilm formation. All strains were able to form pure culture biofilms within 24 h in at least one of the tested culture media and three strains were able to form biofilm in all four culture media, namely Acinetobacter calcoaceticus ATCC 23055, Comamonas denitrificans 123 and Pseudomonas aeruginosa MBL 0199. The adherence properties assessed were initial adherence, cell surface hydrophobicity, and production of amyloid fibers and extracellular polymeric substances. The growth of dual-strain biofilms showed that five organisms formed biofilm with all 13 strains while seven formed no or only weak biofilm when cocultured. In dual-strain cultures, strains with different properties were able to complement each other, giving synergistic effects. Strongest biofilm formation was observed when a mixture of all 13 bacteria were grown together. These results on attachment and biofilm formation can serve as a tool for the design of tailored systems for the degradation of municipal and industrial wastewater.
With increased amounts of wastewater, limited space and stricter regulations and quality controls, the demand for new wastewater treatment processes is growing. This has led to the development of new biofilm-based techniques that have high capacity (Lazarova & Manem, 2000). Increased knowledge of biofilm formation and interspecies interactions has the potential to further develop and refine biofilm processes.
Studies of biofilm development of single-strain microorganisms have focused mainly on clinically relevant bacteria (Costerton et al., 1999; Mack et al., 2007) or strains involved in food spoilage (Midelet & Carpentier, 2004), but areas such as high-grade chemical production (Qureshi et al., 2005; Li et al., 2007), microorganisms in water distribution systems (Simoes et al., 2007) and bioaugmentation through natural genetic transformation (Hendrickx et al., 2000) have also been covered. Dual-species biofilms have been successfully used as model systems to study different types of microbial interactions (Stewart et al., 1997; Christensen et al., 2002; Komlos et al., 2005; Simoes et al., 2007).
The development and persistence of biofilms are affected not only by the surrounding environment but also by the variety of species present (Simoes et al., 2007). Recent observations have shown that factors other than growth rate are important for the relative occurrence and spatial distribution in biofilms (Stewart et al., 1997; Komlos et al., 2005). The mechanisms controlling the microbial interactions in multispecies biofilms are not yet fully understood (Cowan et al., 2000; Komlos et al., 2005).
Microbial ecologists advocate culture-independent, in situ methods to gain further knowledge of individual species in biofilms, their spatial distribution and activity (Daims et al., 2006). This approach is undoubtedly very important but some key properties of the bacteria involved in biofilm formation are possibly more suitable to analysis using pure cultures in vitro. Molin et al. (2000) advocated building up a strong platform of information describing the physiology of organisms using laboratory-based experiments with pure cultures. They further encouraged development of methods for cultivation and characterization of single-species biofilms. It has also been shown that single-species biofilms formed under laboratory conditions exhibit similar overall structural features to those of naturally grown mixed-species biofilms (Davey & O'Toole, 2000).
The aim of the present study was to provide a platform of biofilm-related information on single and mixed bacterial strains with qualities important for wastewater treatment. The research presented here includes a study of (1) the ability to form biofilm of 13 bacterial strains commonly found in wastewater treatment environments, (2) the occurrence of four adherence factors in each bacterial strain and (3) coadhesion and competition in mixed cultures. This information can serve to support further development, design and operation of biofilm processes.
Materials and methods
Bacteria and culture media
Thirteen bacterial strains commonly found in wastewater treatment plants were included in this study (Table 1). All experiments were performed in sterile-filtered municipal wastewater from Käppala wastewater treatment plant, Sweden (126 mg Chemical Oxygen Demand L−1, 45 mg N L−1, 7 mg P L−1, pH 7.5). In addition, tenfold diluted nutrient broth (dNB) and two defined minimal media (Mbwele, 2006) prepared with either 1% (w/v) acetate or glucose (sodium acetate trihydrate, Merck; d-glucose, Sigma) as carbon source, 0.24% (w/v) HEPES (Sigma), 0.21% (w/v) NaHCO3 (Merck), 0.0033% (w/v) Bushnell–Haas Broth (Difco), 0.05% (w/v) yeast extract (Scharlau) and 0.05% (w/v) Casamino acids (Difco) were used for the biofilm formation assay of pure cultures.
|Acinetobacter calcoaceticus||ACA652*||ATCC 23055|
|Aeromonas hydrophila L6||GAM42a†||Mbwele (2006)|
|Bacillus cereus SJV||LGC‡||Labisolate§|
|Brachymonas denitrificans B79||OTU6-178¶||Leta et al. (2003)|
|Brachymonas denitrificans SJV||OTU6-178¶||Labisolate§|
|Comamonas denitrificans 110||COM1424∥||Gumaelius et al. (2001)|
|Comamonas denitrificans 123||COM1424∥||Gumaelius et al. (2001)|
|Delftia sp. SJV||BET42a†||Labisolate§|
|Escherichia coli AF1000||ECO1167**||Sandén et al. (2003)|
|Escherichia coli K-12||ECO1167**||ATCC 10798|
|Klebsiella pneumoniae SJV||GAM42a†||Labisolate§|
|Pseudomonas aeruginosa||Pae997∥||MicroBioLogics 0199 (ATCC 19429)|
|Zoogloea ramigera||BET42a†||CCUG 35504T (ATCC 19544)|
Biofilm formation assay
Each microorganism was cultured in shake flasks overnight for 18 h at 30 °C (180 r.p.m.) in all four media and then diluted to an OD of 0.1 at 620 nm (OD620 nm) in the same medium. Next, 100 μL diluted culture was transferred to each of eight sterile polystyrene microtiter plate wells (Sarstedt 96-well). Sterile culture medium was used as a control. The plates were prepared in duplicate and incubated at 30 °C, with shaking at 100 r.p.m., for 24 and 48 h, respectively. Planktonic growth was measured spectrophotometrically before and after incubation. Biofilm formation was quantified according to Djordjevic et al. (2002), using a crystal violet method. This assay was repeated three times, using fresh overnight cultures. A classification of the strains into nonbiofilm formers, Abs≤Abscontrol, and strong biofilm formers, Abs≥4 × Abscontrol, were made (Stepanovic et al., 2000). Statistical analyses were performed using a paired Student's t-test with a 95% confidence interval.
In addition, biofilm formation of dual-strain mixtures of the taxa listed in Table 1 was examined, as well as a choice of multistrain mixtures: all organisms (A), the six strongest single-strain biofilm formers (B), the three strongest single-strain biofilm formers (C), the three strongest combined with the three weakest single-strain biofilm formers (D), and the strongest dual-strain biofilm formers (H–J) were used in three different combinations (E–G). The mixed cultures were prepared by combining equal volumes of the diluted (OD620 nm=0.1) overnight pure cultures. The biofilm assay was performed as described above using wastewater medium.
Cells were grown overnight (18 h, 30 °C, 180 r.p.m.) in wastewater. Ten milliliters of diluted cell suspension [OD620 nm=0.1 in sterile phosphate-buffered saline (PBS), pH 7.2] was added to polystyrene Petri dishes (Sarstedt), which were incubated for 30 min at 30 °C. The number of cells in each diluted suspension was determined by cell count using a Bürker-chamber. After washing five times with 5 mL PBS the adhered cells were stained for 5 min with 1% crystal violet. Cells were counted under light microscopy and the number of adhered cells per Petri dish area was determined. Results are presented as percentage of adhered cells out of the total number of added cells.
The cell-surface hydrophobicity of cells cultured overnight (18 h, 30 °C, 180 r.p.m.) in wastewater was investigated using the water–xylene method described by Heilmann et al. (1996). Results are presented as the percentage of cells in the hydrophobic phase.
Congo red agar method
The presence of amyloid fibers on the cell surface was investigated by culturing the strains on Congo red (CR) agar. The protocol described by Freeman et al. (1989) was used. The plates were incubated aerobically at 30 °C for 24 h. Amyloid-positive organisms produced red colonies and amyloid-negative organisms produced white colonies (Castonguay et al., 2006).
Visualization of biofilms was performed with fluorescence in situ hybridization (FISH) and confocal laser scanning microscopy (CLSM). Cells were grown in 96-well cell culture plates (Greiner) for 24 h in wastewater as described above. After incubation the wells were washed five times in Milli-Q water. The biofilm was fixed with 4% paraformaldehyde according to a previously described procedure (Amann, 1995). Subsequent hybridization followed Manz's protocol and was performed at 46 °C for 60 min (Manz et al., 1992). The universal oligonucleotide probe EUB338 (Amann et al., 1990) was used for all samples. In addition, more specific probes were used according to Table 1. Following hybridization, the wells were subjected to 20 min washing at 48 °C. The β-glucan (an extracellular polymeric substance, EPS) binding dye calcofluor white was applied (50 μL, 45 min, 50 mg L−1, Sigma F3543). Excess dye was removed by washing with PBS.
Pure culture biofilm formation
The ability to form biofilm was investigated for the bacteria presented in Table 1 using a crystal violet microtiter plate assay. Three taxa were able to form biofilm in all four culture media, namely Acinetobacter calcoaceticus, Comamonas denitrificans 123 and Pseudomonas aeruginosa (Fig. 1). All 13 organisms were able to form biofilm in at least one medium. Biofilm formation was most consistently obtained in acetate medium, where eight strains formed strong biofilm. Wastewater and glucose gave statistically equal results (P=1), whereas dNB resulted in poorer biofilm formation (P<0.05). Strongest planktonic growth was observed in glucose medium, followed by acetate. In wastewater and dNB growth was significantly lower (P=0.001, data not shown). Culture time (24 or 48 h) did not affect biofilm formation (P=0.8), with the exception for Zoogloea ramigera in acetate medium, where biofilm formation decreased from strong at 24 h to none at 48 h.
The importance of initial adherence, hydrophobicity, amyloid production and EPS production for biofilm formation was investigated for the bacteria listed in Table 1. Comamonas denitrificans 110 and Brachymonas denitrificans B79 most readily adhered to polystyrene surfaces (15.1% and 13.5%, respectively) according to the initial adherence test (Table 2). Bacillus cereus SJV, Z. ramigera, B. denitrificans SJV and Klebsiella pneumoniae SJV showed poor adherence properties (<1%). Only Acinetobacter calcoaceticus and B. denitrificans B79 possessed hydrophobic cell-surface properties (>70%) and no more than two organisms, B. cereus SJV and B. denitrificans B79, produced Congo red binding amyloid adhesins (Table 2). FISH-CLSM showed that biofilms of C. denitrificans strains 110 and 123 and B. cereus SJV formed scattered cell clusters surrounded by EPS while A. calcoaceticus produced a uniform cell mat with little or no visible EPS. Brachymonas denitrificans B79 and P. aeruginosa also formed uniform cell mats but with moderate presence of EPS in the multilayer sections of the biofilm. Figure 2 shows the different biofilm morphologies of C. denitrificans 110 and A. calcoaceticus. Biofilm thickness of pure cultures was 9–21 μm for all organisms, corresponding to a few cell layers.
|Species||Planktonic growth, 24 h (OD620 nm)*||Initial adherence (%)||Hydro- phobicity (%)||CRA method†||FISH-CLSM‡ (cells/EPS)|
|A. calcoaceticus||0.09 ± 0.019||7.6||79||Pink||++/−|
|A. hydrophila L6||0.15 ± 0.034||1.2||30||White||+/−|
|B. cereus SJV||0.18 ± 0.026||0.1||0||Red||+/++|
|B. denitrificans B79||0.21 ± 0.025||13.5||76||Red||++/+|
|B. denitrificans SJV||0.28 ± 0.041||1.0||9||White||+/+|
|C. denitrificans 110||0.14 ± 0.023||15.1||0||Pink||+/++|
|C. denitrificans 123||0.22 ± 0.044||1.7||3||Pink||++/++|
|Delftia sp. SJV||0.29 ± 0.035||5.4||5||White||++/+|
|E. coli AF1000||0.12 ± 0.019||1.6||8||White||−/−|
|E. coli K-12||0.17 ± 0.032||2.3||6||White||−/−|
|K. pneumoniae SJV||0.14 ± 0.026||1.0||14||White||−/−|
|P. aeruginosa||0.07 ± 0.026||9.0||8||Pink||+/+|
|Z. ramigera||0.25 ± 0.022||0.7||10||Pink||+/+|
Dual-strain biofilm formation
Interspecies interactions were studied in dual-strain biofilms of the bacteria in Table 1 using the crystal violet microtiter plate assay. Figure 3 illustrates the results from the biofilm formation assay performed on the dual-strain mixtures. Five organisms were determined to be strong coadherers. Acinetobacter calcoaceticus, Delftia sp. SJV, B. denitrificans B79, P. aeruginosa and C. denitrificans 123 all formed biofilm when cocultured with any of the other 12 organisms. One strain, Aeromonas hydrophila L6, formed biofilm with nine of the 12 organisms. The remaining seven organisms were considered to be weak coadherers and did not form a detectable biofilm when cocultured with each other, except for the combination of C. denitrificans 110 and Escherichia coli AF1000, which formed a weak biofilm. Synergistic interactions, meaning that the dual-strain biofilm value exceeded both single-strain biofilm values (Simoes et al., 2007), were observed in 14 of the 78 dual biofilm combinations, while antagonistic interactions were found in 22 combinations (Fig. 3). Delftia sp. SJV and P. aeruginosa were most often involved in synergistic biofilm interactions (six combinations each) and B. cereus SJV in antagonistic interactions (nine of 12 combinations). FISH-CLSM confirmed the presence of both strains involved in the dual-strain biofilms. Pseudomonas aeruginosa, A. calcoaceticus and C. denitrificans 110 dominated the mixtures while E. coli AF1000 and K-12 appeared in low numbers. Dual-strain biofilm thickness varied between 10 and 25 μm.
Multistrain biofilm formation
Formation of multistrain biofilms of a selection of the bacteria in Table 1 was investigated using the crystal violet microtiter plate assay. The results are presented in Fig. 4. Overall strongest biofilm formation was obtained when all microorganisms were cultured together (A). The biofilms formed by the six (B) and the three (C) strongest biofilm formers were equally strong (P=1), although significantly weaker than (A). When the three weakest biofilm formers were added to the three strongest (D), biofilm formation was reduced. The strongest coadherers, selected from Fig. 3 and combined together (E–G), did not form stronger biofilms than corresponding dual-strain mixtures (H–J; Fig. 4). FISH-CLSM of biofilms formed by the mixture of all organisms (A) showed that P. aeruginosa and C. denitrificans strains 110 and 123 were the dominant organisms, while the very strong biofilm coformer A. calcoaceticus was present only in small numbers. The probe ECO1167 gave no visible signal, indicating low/undetectable cell numbers for the two E. coli strains. Owing to the lack of species-specific probes for A. hydrophila, K. pneumoniae, Delftia sp. and Z. ramigera, their presence could not be adequately confirmed.
It is widely believed that most bacterial species can be incorporated in biofilms but it is not known if all can form single-strain biofilms without the aid of coadhering strains (Li et al., 2007). The present results show that all of the bacterial strains investigated form single-strain biofilm in 24 h in at least one of the tested culture media (Fig. 1), confirming the importance of carbon sources and nutrient supply for biofilm growth (Li et al., 2007; Wijman et al., 2007). The carbon sources present influence quorum sensing signals, regulating swarming motility within some species, which has a strong effect on biofilm formation (Shrout et al., 2006). Biosynthesis of other signal substances, cell-surface appendages and EPS might also be affected by the presence or absence of certain carbon sources. The present results show further that biofilm growth did not depend on planktonic growth. Similar observations were made by Wijman et al. (2007) and von Canstein et al. (2002).
The number of initially adhered cells was not critical for biofilm formation, as can be seen for C. denitrificans 123 and Delftia sp. SJV (Table 2, Fig. 1). As the initial adherence test allowed a short cell–substratum contact time (30 min), strains with slow adherence mechanisms gave a low adherence percentage but were still able to form biofilms. Even though the initial adherence number did not seem to affect the amount of biofilm produced, the morphological patterns of the initially adhered cells were still present in the 24-h-old biofilms (Fig. 2).
Hydrophobic cells readily adhere to hydrophobic surfaces through hydrophobic interactions (Donlan & Costerton, 2002; Qureshi et al., 2005). Thus, we expected the hydrophobic strains to adhere more rapidly and to a larger extent to polystyrene than the hydrophilic strains. In the same way we expected the amyloid-producing strains to show strong initial adherence. Amyloid adhesins have been found in 5–40% of prokaryotes in natural biofilms (Larsen et al., 2007) and are suggested to be an important adhesion factor among environmental isolates (Castonguay et al., 2006). However, no clear correlation between these properties and the initial adherence was observed (Table 2), leading to the conclusion that predictions of an organism's adherence capacity cannot be estimated by determining its hydrophobicity or amyloid production. The strongest initial adherence was observed for C. denitrificans 110 (15.1%), a completely hydrophilic strain with no amyloid production. The adherence factors studied were selected based on previous research (Zogaj et al., 2003, Denkhaus et al., 2007, Larsen et al., 2007). However, a large number of factors, besides cell-surface hydrophobicity and amyloid production, have been proposed to affect the initial adherence stage of biofilm formation, such as nutrient supply, hydrodynamic forces, surface charges and presence of other adhesion proteins such as pili or flagella (Denkhaus et al., 2007). When attachment of different bacterial strains is examined, the range of adherence mechanisms is wide, which complicates simple cause–effect conclusions.
The strong biofilm formers here, B. cereus SJV and C. denitrificans 123, produce large amounts of EPS, building up a bulky matrix surrounding the cells (Table 2, Fig. 1). The non- or weak biofilm-forming organisms, K. pneumoniae SJV and E. coli strains K-12 and AF1000, did not produce detectable quantities of EPS under our experimental conditions. However, A. calcoaceticus formed the strongest biofilm of all pure-strains in wastewater, but did not produce any detectable EPS. The biofilm structure and the hydrophobicity of A. calcoaceticus indicate that hydrophobic or electrostatic forces hold the biofilm together.
Strains unable to form single-strain biofilm in wastewater medium were able to form strong biofilm when cocultured with other species. The dual-strain system used to investigate biofilms made it feasible to distinguish the role of each individual organism, as also pointed out by Stewart et al. (1997). According to the present results, the choice of organisms to coculture to attain synergistic effects must be carefully made. It is not possible to predict performance in mixed cultures based on results from pure cultures. One hypothesis is that the clearest synergistic effects appeared when two organisms with rather different properties are mixed. An organism that rapidly adheres to polystyrene surfaces may create a new cell-based surface onto which organisms with slow or deficient initial adherence could attach. Similar conclusions were previously drawn by Stewart et al. (1997). The important role of EPS in biofilm formation was acknowledged by our results. As single strains, E. coli AF1000 and Delftia sp. SJV, a non- and a moderate EPS producer, respectively, together formed a strong biofilm comprising clearly visible EPS, indicating synergistic effects on metabolism and biosynthesis. Pseudomonas aeruginosa formed biofilm in all dual cultures and was found to be one of the dominating species in group A (Fig. 4), but barely formed biofilm in pure culture. These results indicate that P. aeruginosa possesses favorable characteristics in competitive biofilm situations. Acinetobacter calcoaceticus, by contrast, formed strong biofilms in all single- and dual-strain cultures but was not one of the dominating organisms in the mixed-strain biofilms. These characteristics make it a potential coadherer, inducing biofilm formation of other bacteria without out-competing them. Organisms with such characteristics could be used as anchors for attachment of adherence-deficit strains, with potential applications in several industries.
Antagonistic interactions observed in dual-strain biofilms with B. cereus SJV and B. denitrificans B79 disappeared when the two strains were mixed with A. calcoaceticus. Nevertheless, a more diverse mixture did not necessarily lead to a stronger biofilm formation. This can be clearly seen when comparing mixtures E–G in Fig. 4 and corresponding dual-strain mixtures H–J. The complexity of multistrain biofilm interactions makes it difficult to explain this. Cometabolism, secretion of polymers, communication through quorum sensing, which regulates gene expression, and a possible production of bacteriocins all affect the bacterial community (Davey & O'Toole, 2000).
Naturally occurring biofilms usually consist of a variety of species living in a state of symbiosis. Further research is needed before a complete understanding of the mechanisms underlying the formation and maintenance of bacterial biofilms are fully understood. In vitro investigation of biofilm behavior of pure or mixed cultures might lead us one step towards this goal. The results presented here may prove useful for the design of and for choice of inoculum for biofilm processes in wastewater treatment. By applying our knowledge of the synergistic and antagonistic effects of bacterial interactions on attachment and biofilm formation, tailored systems for the degradation of municipal and industrial wastewater could be designed.
We would like to acknowledge Laurent Barbe, PhD, for CLSM expertise, Johan Lindberg for assistance with statistical analyses and Anna Norström for critical review.
- 1995) In Situ Identification of Micro-Organisms by Whole Cell Hybridisation with rRNA-Targeted Nucleic Acid Probes. Kluwer Academic Publishers, Dordrecht. , (
- 1996) rRNA-targeted oligonucleotide probes for the identification of genuine and former pseudomonads. Syst Appl Microbiol 19: 501–509. , , , , & (
- 1990) Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl Environ Microbiol 56: 1919–1925. , , , , & (
- 2006) Biofilm formation by Escherichia coli is stimulated by synergistic interactions and co-adhesion mechanisms with adherence-proficient bacteria. Res Microbiol 157: 471–478. , , , , , & (
- 2002) Metabolic commensalism and competition in a two-species microbial consortium. Appl Environ Microbiol 68: 2495–2502. , , & (
- 1999) Bacterial biofilms: a common cause of persistent infections. Science 284: 1318–1322. , & (
- 2000) Commensal interactions in a dual-species biofilm exposed to mixed organic compounds. Appl Environ Microbiol 66: 4481–4485. , , & (
- 2006) Wastewater treatment: a model system for microbial ecology. Trends Biotechnol 24: 483–489. , & (
- 2000) Microbial biofilms: from ecology to molecular genetics. Microbiol Mol Biol Rev 64: 847–867. & (
- 2007) Chemical and physical methods for characterisation of biofilms. Microchimica Acta 158: 1–27. , , & (
- 2002) Microtiter plate assay for assessment of Listeria monocytogenes biofilm formation. Appl Environ Microbiol 68: 2950–2958. , & (
- 2002) Biofilms: survival mechanisms of clinically relevant microorganisms. Clin Microbiol Rev 15: 167–193. & (
- 1989) New method for detecting slime production by coagulase negative staphylococci. J Clin Pathol 42: 872–874. , & (
- 2001) Comamonas denitrificans sp. nov., an efficient denitrifying bacterium isolated from activated sludge. Int J Syst Evol Microbiol 51: 999–1006. , , & (
- 1996) Characterization of Tn917 insertion mutants of Staphylococcus epidermidis affected in biofilm formation. Infect Immun 64: 277–282. , , & (
- 2000) In situ monitoring of natural genetic transformation of Acinetobacter calcoaceticus BD413 in monoculture biofilms. Water Sci Technol 41: 155–158. , & (
- 2002) The microbial community composition of a nitrifying-denitrifying activated sludge from an industrial sewage treatment plant analyzed by the full-cycle rRNA approach. Syst Appl Microbiol 25: 84–99. , , & (
- 2005) Interaction of Klebsiella oxytoca and Burkholderia cepacia in dual-species batch cultures and biofilms as a function of growth rate and substrate concentration. Microb Ecol 49: 114–125. , , & (
- 1999) Acetogenic and sulfate-reducing bacteria inhabiting the rhizoplane and deep cortex cells of the sea grass Halodule wrightii. Appl Environ Microbiol 65: 5117–5123. , , & (
- 2007) Amyloid adhesins are abundant in natural biofilms. Environ Microbiol 9: 3077–3090. , , , , & (
- 2000) Innovative biofilm treatment technologies for water and wastewater treatment. Biofilms II: Process Analysis and Applications (BryersJD, ed), pp. 159–206. Wiley-Liss, New York. & (
- 2003) Identification of efficient denitrifying bacteria from tannery wastewaters in Ethiopia and a study of the effects of chromium III and sulfide on their denitrification rate. World J Microbiol Biotechnol 20: 405–411. , , & (
- 2007) Single-species microbial biofilm screening for industrial applications. Appl Microbiol Biotechnol 76: 1255–1262. , & (
- 2007) Microbial interactions in Staphylococcus epidermidis biofilms. Anal Bioanal Chem 387: 399–408. , , , , & (
- 1992) Phylogenetic oligodeoxynucleotide probes for the major subclasses of Proteobacteria: problems and solutions. Syst Appl Microbiol 15: 593–600. , , , & (
- 2006). Microbial phosphorus removal in waste stabilisation pond wastewater treatment systems. Thesis, Royal Institute of Technology: Stockholm. (
- 2004) Impact of cleaning and disinfection agents on biofilm structure and on microbial transfer to a solid model food. J Appl Microbiol 97: 262–270. & (
- 2000) Molecular ecology of biofilms. Biofilms II: Process Analysis and Applications (BryersJD, ed), pp. 89–120. Wiley-Liss, New York. , , , et al. (
- 1995) Detection of microbial cells in aerosols using nucleic acid probes. Syst Appl Microbiol 18: 113–122. , & (
- 2005) Biofilm reactors for industrial bioconversion processes: employing potential of enhanced reaction rates. Microb Cell Factories 4: 24. , , , & (
- 2003) Limiting factors in Escherichia coli fed-batch production of recombinant proteins. Biotechnol Bioeng 81: 158–166. , , et al. (
- 2006) The impact of quorum sensing and swarming motility on Pseudomonas aeruginosa biofilm formation is nutritionally conditional. Mol Microbiol 62: 1264–1277. , , , , & (
- 2007) Biofilm interactions between distinct bacterial genera isolated from drinking water. Appl Environ Microbiol 73: 6192–6200. , & (
- 2000) A modified microtiter-plate test for quantification of staphylococcal biofilm formation. J Microbiol Methods 40: 175–179. , , , & (
- 1997) Spatial distribution and coexistence of Klebsiella pneumoniae and Pseudomonas aeruginosa in biofilms. Microb Ecol 33: 2–10. , , , & (
- 2002) Species diversity improves the efficiency of mercury-reducing biofilms under changing environmental conditions. Appl Environ Microbiol 68: 2829–2837. , , & (
- 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. , , , , , & (
- 2007) Air-liquid interface biofilms of Bacillus cereus: formation, sporulation, and dispersion. Appl Environ Microbiol 73: 1481–1488. , , , & (
- 2003) Production of cellulose and curli fimbriae by members of the family Enterobacteriaceae isolated from the human gastrointestinal tract. Infect Immun 71: 4151–4158. , , & (