Institute of Environmental Engineering, National Central University, Chungli 32054, Taiwan1
Center for Microbial Ecology, Michigan State University, East Lansing, MI 48824, USA2
Department of Urban Engineering, University of Tokyo, Tokyo 113, Japan3
National Institute of Bioscience and Human Technology, Agency of Industrial Science and Technology, 1-1 Tsukuba, Ibaraki, 305-8566 Japan4
Center for Ecological and Evolutionary Studies, University of Groningen, The Netherlands5
Author for correspondence: Wen-Tso Liu. Tel: +886 3422 7151 ext 4683. Fax: +886 3426 9401. e-mail: liuwt{at}cc.ncu.edu.tw
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Keywords: Activated sludge, biological phosphate removal, biomarker, DGGE, 16S rDNA
Abbreviations: DGGE, denaturing gradient gel electrophoresis; EBPR, enhanced biological phosphate removal; PHA, polyhydroxyalkanoate
The GenBank/EMBL/DDBJ accession numbers for the sequences obtained in this report are AF109792 (strain Lpha5), AF109793 (strain Lpha7) and AF124650 to AF124659.
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Methods based on the analysis of various biomarkers and 16S rRNA genes (rDNA) have been used to characterize and monitor microbial communities in various ecological systems. The biomarkers, cellular fatty acids and respiratory quinones, have been used routinely in taxonomy to characterize, differentiate and identify genera, species, and strains of bacteria (Staley et al., 1989 ). Some studies have further shown that the biomarker signature of environmental samples can be statistically analysed and applied to differentiate community profiles (Haack et al., 1994
; Hiraishi et al., 1989
, 1998
). 16S rDNA-based methods can provide more information on the phylogenetic structure of microbial communities than the biomarker method (Bond et al., 1995
; Muyzer et al., 1993
). For example, denaturing gradient gel electrophoresis (DGGE) can be used to resolve PCR-amplified 16S rDNA fragments by electrophoresis through an acrylamide gel that contains an increasing linear gradient of denaturants (Muyzer et al., 1993
). The number and intensity of resolved fragments gives an approximate estimate of the diversity of the predominant species, and further purification of fragments and sequence analysis provides an insight into the phylogenetic affiliation of individual populations (Muyzer et al., 1995
; Nielsen et al., 1999
). Combining the biomarkers and 16S rDNA-based approaches should greatly enhance the characterization of microbial communities found in various systems.
Previously, we varied the phosphorus:carbon (P/C) weight ratio of the feed used for lab-scale EBPR systems and successfully enriched communities that differed in their ability to accumulate phosphate (Liu et al., 1997a ). A gradual increase in the P/C ratio from 20:100 to 2:100 did not affect the carbon uptake and storage under anaerobic conditions, but caused a drop in the sludge P content from ~12% of sludge dry weight (or 33% polyphosphate) to a cellular constituent level (~2% P content or ~0% polyphosphate). As a result, the EBPR activity stopped, and apparent shifts in community structure were reflected by differences in the morphologies of the predominant populations. At a 12% P content the community was predominated by rod-shaped organisms that accumulated both polyphosphate and PHA, whereas at a 2% P content the community was dominated by PHA-accumulating cocci that neither accumulated polyphosphate nor aerobically took up phosphate when provided (Liu et al., 1996
, 1997a
). It was suspected that the loss of polyphosphate-accumulating bacteria upon shifting the feed composition was due to loss of their energy pool (polyphosphate) which they used to transport and store carbon (e.g. PHA) under anaerobic conditions so that it could be subsequently used for growth under aerobic conditions. However, in that study we did not establish whether the apparent shifts in the morphology of the microbial community were indeed caused by changes in the relative abundance of different bacterial populations. Our initial assumption was that polyphosphate-accumulating bacteria would constitute a predominant fraction of the bacterial population in a 12% P-containing sludge, and on altering sludge P content to 2%, could be easily differentiated from non-polyphosphate accumulating bacteria using the biomarker and DGGE approaches. Furthermore, the phylogenetic affiliation of the predominant populations in 2% and 12%-P containing sludge could be easily identifiable from sequence analysis of the predominant 16S rDNA DGGE fragments. This study was aimed at testing the above assumption.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Analyses of chemotaxonomic biomarkers.
Procedures for respiratory-quinone analysis were those described by Hiraishi et al. (1989) . Briefly, respiratory quinones were extracted with a mixture of chloroform/methanol (2:1, v/v), purified by TLC and qualitatively analysed by reverse-phase HPLC using a Shimadzu model CTO6-A HPLC equipped with a photodiode array detector. Quinone standards were purchased from Wako Pure Chemicals or extracted from bacterial strains with known quinone compositions (Liu, 1995
). Determination of other quinones, including demethylmenaquinone and rhodoquinone, was not attempted since these quinones were not previously found in EBPR processes (Hiraishi et al., 1989
). Ubiquinones and menaquinones with n isoprene units are abbreviated Q-n and MK-n, respectively. MK-n(Hx) represents a partially hydrogenated menaquinone with x hydrogen atoms on the side chain containing n isoprene units. The total cellular fatty acids were analysed using a protocol described by Rajendran et al. (1992)
.
Cluster analysis (Statistica) was used to statistically discern patterns in the respiratory quinone and total cellular fatty acid data. Dendrograms were constructed by using the single linkage and Euclidean distance rules in the Statistica program. Analysis of variance (ANOVA) was further used to test the change of individual quinone components among sludge samples taken from reactors containing different sludge P contents.
Isolation and PCR amplification of DNA.
DNA from activated sludge was obtained after cell lysis, phenol/chloroform extraction and ethanol precipitation using a previously described protocol (Liu et al., 1997b ). This DNA preparation was used as the template in PCR-reaction mixtures that contained 1x PCR buffer (Gibco-BRL), 200 µM each dNTP, 1·5 mM MgCl2, 0·1 µM each primer, 5% DMSO and 2·5 U Taq DNA polymerase (Gibco-BRL) in a final volume of 100 µl. For amplification of 16S rDNA for DGGE analysis, the 968FGC forward primer with a GC clamp (Heuer et al., 1997
) and the 1392R reverse primer (Ferris et al., 1996
) were used. The PCR was carried out in a Perkin Elmer 9600 thermocycler using a thermal program described previously (Nielsen et al., 1999
). Amplification of DNA was verified by electrophoresis of 2 µl of the PCR product through a 1% agarose gel in 1x TAE buffer (20 mM Tris-acetate, pH 7·4, 10 mM sodium acetate, 0·5 mM EDTA).
DGGE.
DGGE was performed using a D-Gene system (Bio-Rad) according to the manufacturers instructions. PCR products were loaded onto a 6% acrylamide gel (37·5:1, acrylamide:N,N'-methylene-bis-acrylamide) in 1x TAE buffer. The denaturing gradient in the gel was formed by mixing two stock solutions of 6% acrylamide that contained 40% denaturant (2·8 M urea and 18·7% (w/v) formamide; both from Sigma) and 60% denaturant (4·2 M urea, 24% formamide). Denaturants were deionized with AG501-X8 mixed bed resin (Bio-Rad) prior to being used. The DNA fragments were visualized by silver staining as described by Riesner et al. (1989) .
Isolation, cloning and sequencing of DGGE fragments.
The DNA sequences of specific fragments in the DGGE gels were determined. A fragment was excised from the gel using a razor blade and the DNA was eluted overnight in 100 µl TAE buffer. Individual DNA fragments were amplified by PCR with the DGGE primers described above, ligated into the pCRII vector and transformed into competent Escherichia coli cells according to the manufacturers instructions (TA Cloning System; Invitrogen). Clones with the target DNA fragments were identified by amplifying ten randomly chosen clones using the DGGE primers and comparing the electrophoretic mobility of the amplicon with that of the fragments in the original sample. Selected DNA fragments were sequenced at the Michigan State University sequencing facility on an ABI DNA Sequencer model 373 (Applied Biosystems) using the 968FGC (without the GC clamp) and 1392R primers, and the Taq DyeDeoxy Terminator Cycle Sequencing kit (Applied Biosystems).
Phylogenetic analyses.
Partial DNA sequences (~430 bp) obtained in this study were compared to available 16S rRNA sequences in GenBank using the NCBI BLAST program. The most closely related sequences from the NCBI BLAST searches, and important 16S rRNA sequences of environmental clones and bacterial isolates obtained from the EBPR process were retrieved and aligned to those sequenced DGGE bands using the CLUSTAL W program (Thompson et al., 1994 ). A phylogenetic tree was constructed from the evolutionary distance matrix based on the Kimura two-parameter algorithm using the neighbour-joining method (Saitou & Nei, 1987
). The analysis was performed with the MEGA program (Kumar et al., 1993
), and gap sites in the alignment were excluded in the pairwise comparison.
![]() |
RESULTS AND DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
|
|
Phylogeny of predominant populations in EBPR processes
Seven and nine major fragments from sludge samples containing 12·3 and 2% P content, respectively, were excised from the gel and purified. The purified DNA was amplified using the same DGGE primers and electrophoresed in a DGGE gel along with the original sample. Possibly due to the quality of DNA retrieved, only five major fragments from each sample gave PCR products. Since reamplification of a single fragment often resulted in the formation of multiple amplicons, direct sequencing of the reamplified fragment was not possible (data not shown). Instead, the reamplified fragment was cloned and clones with the correct insert were identified by comparing the electrophoretic mobility of the cloned fragments with that of the target fragment found in the original sample. The comparative analysis of these partial 16S rRNA sequences (Fig. 3) revealed the phylogenetic affiliation of the ten sequences retrieved.
|
Another DGGE fragment common to both 2 and 12·3% P sludges (2% P, band c and 12% P band d; Fig. 2) was closely affiliated with the Actinobacteria. The most closely related cultured bacteria were two isolates, strain Lpha5 and Lpha7, from the reactor containing 2% P (Liu, 1995
). These two isolates accumulated PHA but not polyphosphate as granular inclusions and contained the same major menaquinone component [MK-8(H4)] (Liu, 1995
) as observed in the sludge sample. It is likely that these two organisms were not specifically involved in P removal. However, it is also possible that organisms that can accumulate polyphosphate are phylogenetically related to organisms that cannot. A good example is between a phosphate-accumulating bacterium, Microlunatus phosphovorus (Nakamura et al., 1995
), and a non-phosphate-accumulating bacterium, Micropruina glycogenica (Shintani et al., 2000
), that are two phylogentically related genera isolated from EBPR processes. Since different actinobacteria may contain different proportions of nemaquinones (Collins & Jones, 1981
), the change observed in MK-8(H4) content on altering P content could reflect a population shift between non-phosphate-accumulating and phosphate-accumulating actinobacteria with different nemaquinone contents. Nevertheless, these data confirmed that members of this novel bacterial population from the actinobacteria are widely distributed in EBPR systems, and as other reports have suggested (Bond et al., 1999
; Hiraishi et al., 1989
; Nakamura et al., 1995
; Wagner et al., 1994
), were responsible in part for carbon and possibly phosphate metabolism observed in both 2% and 12% P-containing reactors.
DGGE fragments (2%, band a, and 12%, band b; Fig. 2) that were phylogenetically related to species of Caulobacter from the
subclass of the Proteobacteria were observed both in 2% and 12% P-containing systems, even though these fragments did not migrate to the same position on the DGGE gel. It is therefore apparent, as demonstrated here and by other studies (Felske et al., 1998
; Nielsen et al., 1999
), that 16S rDNA fragments that are phylogenetically closely related can have different migration positions in a DGGE gel. Caulobacter spp. are often found in environments with low organic carbon contents (Stahl et al., 1992
), but have been detected in sewage-treatment systems (MacRae & Smit, 1991
) and a full-scale EBPR process (Schuppler et al., 1995
). The presence of Caulobacter may be one reason for the significant amount of Q-10 detected in all sludge samples (Table 1
).
In addition to the common groups, the phylogenetic analysis (Fig. 3) revealed fragments or bacterial populations specific to both 2% and 12% P-containing sludge. Due to the short 16S rRNA sequence obtained and the lack of closely related sequences in the 16S rRNA database, the exact phylogenetic placement of these populations was difficult. Band e (2% P) and band a (12% P) were both associated with the
subclass of the Proteobacteria. Band c (12% P) and band b (2% P) were associated with the high G+C group and unidentified green sulfur bacteria, respectively. These findings were consistent with the result that significant amounts of Q-10 and MK-8(H4) were present in all sludge samples, and a shift in their composition on altering P content was detected. Although those populations unique to the 12% P sludges were possibly the predominant populations performing the EBPR metabolism, their abilities with respect to polyphosphate and PHA metabolism is unknown and warrants further study. Further, in contrast to the respiratory-quinone result, none of the predominant DGGE fragments retrieved and sequenced were associated with the ß subclass of the Proteobacteria. This could be due to our inability to retrieve sequences of some predominant DGGE bands (Fig. 2
) that may have belonged to the ß subclass. The biases associated with community DNA extraction and PCR amplification in some cases (Picard et al., 1992
; Tebbe & Vahjen, 1993
; Wilson, 1997
) could also lead to the misrepresentation of the true community fingerprint. It is also possible that bacteria that produce large quantities of Q-8 but do not belong to the ß-Proteobacteria exist and may be present in activated sludge systems.
In summary, our current understanding of the diversity of microbial populations in EBPR processes has primarily come from studies using fluorescent in situ hybridization (Bond et al., 1999 ; Kämpfer et al., 1996
; Schuppler et al., 1998
; Wagner et al., 1994
) and analysis of environmental 16S rDNA clone libraries (Bond et al., 1995
; Schuppler et al., 1995
). All studies suggest a high degree of phylogenetic diversity, including at least 30 different phylotypes from major phyla of the domain Bacteria. Nevertheless, no study to date has firmly identified a specific phylotype directly associated with the accumulation of polyphosphate, PHA or both. We initially suspected that the inconclusive findings were attributed to the use of sludge samples from full-scale EBPR processes, in which the population diversity is influenced by a variety of environmental factors (e.g. multiple substrates, electron acceptors and constant changes in the P/C feeding ratio) (Cech & Hartman, 1993
; Kuba et al., 1993
; Liu et al., 1997a
), and phosphate-accumulating bacteria were not necessarily the major population. Our study indicated that even under well-controlled and enriched EBPR systems, the detectable microbial populations were phylogenetically diverse. While a shift of microbial community structure on altering sludge P content was observed from the biomarker and DGGE fingerprints, our results further found that microbial communities in 2% and 12% P-containing sludge not only included predominant bacterial populations specific to each sludge, but also shared phylogenetically closely related populations. It was further suspected that specific phylogenetic groups might include both non-phosphate-accumulating and phosphate-accumulating populations. Thus, the combined use of biomarkers and DGGE methods was insufficient to identify organisms that accumulate phosphate. A promising approach will be to combine microautoradioagraphy with fluorescent in situ hybridization to link functional traits to a phylogenetic population in activated sludge processes (Lee et al., 1999
). This approach with a refined hierarchical set of probes (e.g. Mobarry et al., 1996
; Raskin et al., 1994
) should provide a better understanding of the organisms responsible for EBPR.
![]() |
ACKNOWLEDGEMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Bond, P., Erhart, E., Wagner, M., Keller, J. & Blackall, L. (1999). Identification of some of the major groups of bacteria in efficient and nonefficient biological phosphorus removal activated sludge systems.Appl Environ Microbiol 65, 4077-4084.
Cech, J. S. & Hartman, P. (1993). Competition between polyphosphate and polysaccharide accumulating bacteria in enhanced biological phosphate removal system.Water Res 27, 1219-1225.
Collins, M. D. & Jones, D. (1981). Distribution of isoprenoid quinone structural types in bacteria and their taxonomic implications.Microbiol Rev 45, 316-354.
Comeau, Y., Hall, K. J., Hancock, R. E. W. & Oldham, W. K. (1986). Biochemical model for enhanced biological phosphorus removal.Water Res 20, 1511-1521.
Dawes, E. A. & Senior, P. J. (1973). Energy reserve polymers in microorganisms.Adv Microbiol Physiol 10, 135-266.[Medline]
Felsenstein, J. (1985). Confidence limits of phylogenies: an approach using the bootstrap.Evolution 39, 783-791.
Felske, A., Wolterink, A., Van Lis, R. & Akkermans, A. D. (1998). Phylogeny of the main bacterial 16S rRNA sequences in Drentse A grassland soils (The Netherlands).Appl Environ Microbiol 64, 871-879.
Ferris, M. J., Muyzer, G. & Ward, D. M. (1996). Denaturing gradient gel electrophoresis profiles of 16S rRNA-defined populations inhabiting a hot spring microbial mat community.Appl Environ Microbiol 62, 340-346.[Abstract]
Fuhs, G. W. & Chen, M. (1975). Microbiological basis of phosphate removal in the activated sludge process for the treatment of wastewater.Microb Ecol 22, 119-138.
Haack, S. K., Garchow, H., Odelson, D., Forney, L. J. & Klug, M. J. (1994). Accuracy, reproducibility, and interpretation of fatty acid methyl ester profiles of model bacterial communities.Appl Environ Microbiol 60, 2483-2493.[Abstract]
Heuer, H., Krsek, M., Baker, P., Smalla, K. & Wellington, E. M. (1997). Analysis of actinomycete communities by specific amplification of genes encoding 16S rRNA and gel-electrophoretic separation in denaturing gradients.Appl Environ Microbiol 63, 3233-3241.[Abstract]
Hiraishi, A., Masamune, K. & Kitamura, H. (1989). Characterization of the bacterial population structure in an anaerobic-aerobic activated sludge system on the basis of respiratory quinone profiles.Appl Environ Microbiol 55, 897-901.[Medline]
Hiraishi, A., Ueda, Y. & Ishihara, J. (1998). Quinone profiling of bacterial communities in natural and synthetic sewage activated sludge for enhanced phosphate removal.Appl Environ Microbiol 64, 992-998.
Jenkins, D. & Tandoi, V. (1991). The applied microbiology of enhanced biological phosphate removal accomplishments and needs.Water Res 25, 1471-1478.
Kämpfer, P., Erhart, R., Beimfohr, C., Böhringer, J., Wagner, M. & Amann, R. (1996). Characterization of bacterial communities from activated sludge: culture dependent numerical identification versus in situ identification using group- and genus-specific rRNA-targeted oligonucleotide probes.Microb Ecol 322, 101-121.
Kuba, T., Smolders, G., van Loosdrecht, M. C. M. & Heijnen, J. J. (1993). Biological phosphorus removal from wastewater by anaerobic-anoxic sequencing batch reactor.Water Sci Technol 27, 241-252.
Kumar, S., Tamura, K. & Nei, M. (1993). MEGA: molecular evolutionary genetics analysis, version 1.0. University Park, PA: Pennsylvania State University.
Lee, N., Nielsen, P. H., Andreasen, K. H., Juretschko, S., Nielsen, J. L., Schleifer, K.-H. & Wagner, M. (1999). Combination of fluorescent in situ hybridization and microsutoradiography new tool for structure-function analyses in microbial ecology.Appl Environ Microbiol 65, 1289-1297.
Liu, W.-T. (1995). Function, dynamics, and diversity of microbial population in anaerobic aerobic activated sludge processes for biological phosphate removal. PhD thesis, University of Tokyo.
Liu, W.-T., Mino, T., Nakamura, K. & Matsuo, T. (1994). Role of glycogen in acetate uptake and polyhydroxyalkanoate synthesis in anaerobicaerobic activated sludge with a minimized polyphosphate content.J Ferment Biotechnol 77, 535-540.
Liu, W.-T., Mino, T., Matsuo, T. & Nakamura, K. (1996). Glycogen accumulating population and its anaerobic substrate uptake in anaerobicaerobic activated sludge without biological phosphate removal.Water Res 30, 75-82.
Liu, W.-T., Nakamura, K., Matsuo, T. & Mino, T. (1997a). Internal energy-based competition between polyphosphate- and glycogen-accumulating bacteria in biological phosphorus removal reactor effect of the P/C feeding ratio.Water Res 31, 1430-1438.
Liu, W.-T., Marsh, T. L., Chen, H. & Forney, L. J. (1997b). Characterization of microbial diversity by determining terminal restriction fragment length polymorphisms of gene encoding 16S rRNA.Appl Environ Microbiol 63, 4516-4522.[Abstract]
MacRae, J. D. & Smit, J. (1991). Characterization of caulobacters isolated from wastewater treatment systems.Appl Environ Microbiol 573, 751-758.
Marais, G. v. R., Lowenthal, R. E. & Siebritz, I. P. (1983). Observations supporting phosphate removal by biological excess uptake a review.Water Sci Technol 15, 15-42.
Mino, T., Arun, V., Tsuzuki, Y. & Matsuo, T. (1987). Effect of phosphorus accumulation on acetate metabolism in the biological phosphorus removal process. In Advances in Water Pollution Control: Biological Phosphate Removal from Wastewaters, pp. 27-38. Edited by R. Ramadori. Oxford: Pergamon Press.
Mino, T., van Loosdrecht, M. C. M. & Heijnen, J. J. (1998). Microbiology and biochemistry of the enhanced biological phosphate removal process.Water Res 32, 3193-3207.
Mobarry, B. K., Wagner, M., Urbain, V., Rittmann, B. E. & Stahl, D. A. (1996). Phylogenetic probes for analyzing abundance and spatial organization of nitrifying bacteria.Appl Environ Microbiol 62, 2156-2162.[Abstract]
Muyzer, G., de Waal, E. C. & Uitterlinden, A. G. (1993). Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA.Appl Environ Microbiol 59, 695-700.[Abstract]
Muyzer, G., Teske, A. & Wirsen, C. O. (1995). Phylogenetic relationships of Thiomicrospira species and their identification in deep-sea hydrothermal vent samples by denaturing gradient gel electrophoresis of 16S rDNA fragments.Arch Microbiol 164, 165-172.[Medline]
Nakamura, K., Hiraishi, A., Yoshimi, Y., Kawaharasaki, M., Masuda, K. & Kamagata, Y. (1995). Microlunatus phosphovorus gen. nov., sp. nov., a new gram-positive polyphosphate-accumulating bacterium isolated from activated sludge.Int J Syst Bacteriol 45, 17-22.[Abstract]
Nielsen, A. T., Liu, W.-T., Philips, C., Grady, L.Jr, Molin, S. & Stahl, D. A. (1999). Identification of a novel group of bacteria in sludge from a deteriorated biological phosphorus removal process.Appl Environ Microbiol 65, 1251-1258.
Picard, C., Ponsonnet, C., Paget, E., Nesme, X. & Simonet, P. (1992). Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction.Appl Environ Microbiol 58, 2717-2722.[Abstract]
Rajendran, N., Matsuda, O., Imamura, N. & Urushigawa, Y. (1992). Variation in microbial biomass and community structure in sediments of eutrophic bays as determined by phospholipid ester-linked fatty acids.Appl Environ Microbiol 58, 562-571.[Abstract]
Raskin, L., Stromley, J. M., Rittmann, B. E. & Stahl, D. A. (1994). Group-specific 16S rRNA hybridization probes to describe natural communities of methanogens.Appl Environ Microbiol 60, 1232-1240.[Abstract]
Riesner, D., Steger, G., Zimmat, R., Owens, R. A., Wagenhofer, M., Hillen, W., Vollbach, S. & Henco, K. (1989). Temperature-gradient gel electrophoresis of nucleic acids: analysis of conformational transitions, sequence variations, and proteinnucleic acid interactions.Electrophoresis 10, 377-389.[Medline]
Saitou, N. & Nei, M. (1987). The neighbor-joining method: a new method for reconstructing phylogenetic trees.Mol Biol Evol 4, 406-425.[Abstract]
Schuppler, M., Mertens, F., Schön, G. & Göbel, U. B. (1995). Molecular characterization of nocardioform actinomycetes in activated sludge by 16S rRNA analysis.Microbiology 141, 513-521.[Abstract]
Schuppler, M., Wagner, M., Schön, G. & Göbel, U. B. (1998). In situ identification of nocardioform actinomycetes in activated sludge using fluorescent rRNA-targeted oligonucleotide probes.Microbiology 144, 249-259.[Abstract]
Shintani, T., Liu, W.-T., Hanada, S., Kamagata, Y., Miyaoka, S., Suzuki, T. & Nakamura, K. (2000). Micropruina glycogenica gen. nov., sp. nov., a new Gram-positive glycogen-accumulating bacterium isolated from activated sludge.Int J Syst Evol Microbiol 50, 201-207.[Abstract]
Stahl, D. A., Key, R., Flesher, B. & Smit, J. (1992). The phylogeny of marine and freshwater caulobacters reflects their habitat.J Bacteriol 174, 2193-2198.[Abstract]
Staley, J. T., Bryant, M. P., Pfennig, N. & Holt, J. G. (editors) (1989). Bergeys Manual of Systematic Bacteriology, vol. 3. Baltimore: Williams & Wilkins.
Tebbe, C. C. & Vahjen, W. (1993). Interference of humic acids and DNA extracted directly from soil in detection and transformation of recombinant DNA from bacteria and a yeast.Appl Environ Microbiol 59, 2657-2665.[Abstract]
Thompson, J. D., Higgins, D. G. & Gibson, T. J. (1994). CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice.Nucleic Acids Res 22, 4673-4680.[Abstract]
Wagner, M., Erhart, R., Manz, W., Amann, R., Lemmer, H., Wedi, D. & Schleifer, K. H. (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.[Abstract]
Wilson, I. G. (1997). Inhibition and facilitation of nucleic acid amplification.Appl Environ Microbiol 63, 3741-3751.
Received 4 October 1999;
revised 17 December 1999;
accepted 19 January 2000.