Ideonella dechloratans gen.nov., sp.nov., a New Bacterium Capable of Growing Anaerobically with Chlorate as an Electron Acceptor

Ideonella dechloratans gen.nov., sp.nov., a New Bacterium Capable of Growing Anaerobically with Chlorate as an Electron Acceptor

System. Appl. Microbiol. 17,58-64 (1994) © Gustav Fischer Verlag, Stuttgart· Jena . New York Ideonella dechloratans gen. nov., sp. nov., a New Bacter...

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System. Appl. Microbiol. 17,58-64 (1994) © Gustav Fischer Verlag, Stuttgart· Jena . New York

Ideonella dechloratans gen. nov., sp. nov., a New Bacterium Capable of Growing Anaerobically with Chlorate as an Electron Acceptor ASA MALMQVIST h , THOMAS WELANDER 1 ,2, EDWARD MOORE 3 , ANDERS TERNSTROM\ GORAN MOLIN4 and INGA-MAJ STENSTROM5 1 2 3 4

5

ANOX AB, Ideon Research Park, 5-223 70 Lund, Sweden Biotechnology, University of Lund, P.O. Box 124,5-221 00 Lund, Sweden Microbiology, G.B.F. - National Research Center for Biotechnology, Mascheroder Weg 1,3300 Braunschweig, FRG Food Technology, University of Lund, P.O. Box 124, S-221 00 Lund, Sweden Applied Microbiology, University of Lund, P.O. Box 124, 5-221 00 Lund, Sweden Received September 23, 1993

Summary The name Ideonella dechloratans is proposed for a new species of Gram-negative, polarly flagellated, chemoorganotrophic, rod shaped bacterium capable of growing anaerobically with chlorate as an electron acceptor. The bacterium is mesophilic, strictly respiratory and has a guanine plus cytosine-content in DNA of 68.1 mol%. Sequence data from 16S ribosomal RNA gene of the type strain, CCUG 30898 T , show Ideonella dechloratans to cluster phylogenetically within the beta subgroup of Proteobacteria. The bacterium was enriched and isolated from activated sludge from a municipal wastewater treatment plant.

Key words: Chlorate Reduction - Anerobic Respiration - Taxonomy - 16S rRNA Sequence - Beta Subgroup of Proteobacteria - Ideonella dechloratans

Introduction The ability to use inorganic electron acceptors other than oxygen, e.g. nitrate, sulfate and carbonate, for energy metabolism is common in the bacterial world. From an energetic point of view, the highly oxidized chlorate ion, CI0 3 -, should be a very favorable electron acceptor. As a matter of fact, calculations of the free energy changes, at standard conditions and pH 7, for the oxidation of organic compounds, e.g. acetate, with oxygen and chlorate, show that chlorate should be an even more efficient electron acceptor than oxygen: CH3 COO- + 4/3C10 3 - + OW - 2HC0 3 - + H 2 0 + 4/3Cl- (1) LlGo, = - 132 kJlmol of electrons exchanged CH3 COO- + 202 + OW - 2HC0 3- + H20 LlGo, - 110 kJlmol of electrons exchanged

(2)

However, chlorate does not occur naturally and has been introduced into the environment only by the activities of humans. This means that time has been short, from an evolutionary point of view, for development of an enzy-

* Corresponding author

matic system needed for the utilization of chlorate as an electron acceptor. Despite this, the ability of several organisms to reduce chlorate to chlorite by means of either of the enzymes nitrate reductase or chlorate reductase has been reported (Azoulay et al. 1971, de Groot and Stouthamer 1969, Oltmann et al. 1976). It has also been suggested that Proteus mirabilis can use chlorate as an electron acceptor in anaerobic electron transport (de Groot and Stouthamer, 1969), but growth of this organism with chlorate as a terminal electron acceptor in an energy-yielding reaction was not demonstrated. In recent years, the interest in microbial reduction of chlorate has increased dramatically due to environmental concerns, since it has been noticed that chlorate in wastewaters being discharged from pulp and paper mills is toxic to aquatic plants, such as bladder wrack (Fucus vesiculosus) (Lehtinen et al., 1988, Rosemarin et al., 1986; 1990). Chlorate is formed and ends up in the wastewater when chlorine dioxide is used for pulp bleaching (Bergnor et al., 1987, Germgard et al., 1981). Several investigators have shown that chlorate can be removed from the waste-

Ideonella dechloratans gen. nov., sp. nov., a Chlorate Reducing Bacterium

water by biological treatment operated under anaerobic conditions (Axegard et al., 1989, Eckerman, 1987, Germgard and Berglund, 1987, Germgard, 1989, Malmqvist and Welander, 1992, Yu and We/ander, 1990). However, the product formation and microbial mechanisms of the chlorate removal process were unknown until Malmqvist et al. (1991) demonstrated the complete reduction of chlorate to chloride by an enrichment culture growing anaerobically with chlorate as an electron acceptor in an energy-yielding reaction. The enrichment of chlorate reducing microorganisms resulted in a microflora strongly predominated by motile bacteria of a helical morphology. Subsequently, the isolation of chlorate reducing bacteria was reported (Malmqvist and Welander, 1992). However, the characteristics and the taxonomic affiliation of these bacteria were not studied in detail, so that, until now, very little has been known about the microorganisms capable of utilizing chlorate as an electron acceptor. The isolation and characterization of a previously undescribed bacterium, able to grow anaerobically with chlorate as an electron acceptor, is reported in this paper.

Materials and Methods Enrichment of chlorate reducers. The enrichment of chlorate reducing bacteria was carried out in three continuously fed lab scale reactors, one operated as a chemostat and the other two as biofilm processes with reticulated polyurethane foam as support material (biofilm reactors A and B). Initially, the reactors were inoculated with activated sludge from Sjolunda municipal treatment plant, Malmo, Sweden, after which a continuous feed of a defined medium was started. The defined medium contained: 10 mM NaCI0 3 , 25 mM acetic acid, 3 mM NH 3 , 0.25 mM KH 2P0 4, 0.1 mM MgS0 4, 0.1 mM Ca(OHh, 0.05 mM FeS04 " 7 H 2 0, 0.05 mM MnS04 * H 2 0,S 11M NiCI * 6 H 2 0,S 11M CoCh ,c 6 H 2 0,5 11M ZnS04 * 7 H 20, 0.1 11M H 3 B03 , 0.1 11M NazSe04, 0.1 11M Na2 W0 4 and 0.1 11M Na2Mo04. The pH of the medium was adjusted to 7 with NaOH and the temperature in the reactors was controlled to 37°C. The chemostat was operated at a dilution rate of 0.083 h-1 while the dilution rate for the biofilm processes was 1.0 h-1 . The reactors were run under these conditions for several months in order to achieve a strong enrichment of chlorate reducers. A complete reduction of chlorate to chloride was obtained shortly after starting the processes and was maintained throughout the enrichment experiment. Isolation of strains. Chlorate reducing bacteria were isolated from the enrichment cultures using the streak plate method. Samples were taken from the reactors and streaked onto agar plates (1.5% Bactoagar) containing the selective medium described above. The plates were then incubated in an anaerobic jar (GasPak system, BBL, Cockeysville, USA) at 37°C for four days. Well-isolated colonies were picked and restreaked several times to obtain pure cultures of chlorate reducers. Phenotypic characterization. Tests for production of acid from carbohydrates were done in a modified Hugh-Leifson medium containing: 2.0 giL peptone, 5.0 giL NaCl, 0.3 giL K2HP0 4 , 0.06 giL bromocresol purple, and 15.0 giL Bacto-agar. This basal medium, adjusted to pH 7.1, was supplemented with filtersterilized carbohydrate solution to give a final concentration of 1 % (wtlvol) prior to pouring into plates. A yellow zone around the colonies was scored as a positive result. Utilization of individual carbon sources was tested on the basal medium of Palle-

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roni and Doudoroff (1972), supplemented with filter-sterilized organic compounds to a concentration of 0.1% (wtlvol). Tests were performed both aerobically and anaerobically, in the latter case the medium was supplemented with 1.06 g NaCI0 3 per liter. Other tests were performed as previously described (Molin and Ternstrom, 1982; 1986). Unless indicated otherwise, all tests were performed at 37°C and read after 4 days. Anaerobic conditions were obtained using an anaerobic jar (GasPak system, BBL). Testing of oxidative capacity for 95 organic compounds on Biolog GN Microplates (Biolog Inc., Hayward, Ca., USA) were done as recommended by the manufacturer. Results were recorded automatically using a MicroStation spectrophotometer (Biolog Inc.). Guanine plus cytosine determination. The mol% G+C in DNA was analyzed at Deutsche Sammlung von Mikroorganismen und Zellkulturen, (DSM) Braunschweig, FRG, by K. D. Janke und S. Kirschner using a HPLC-technique described by Mesbach et al. (1989). Sequencing of PCR-amplified 16 S ribosomal RNA gene and data analysis. Approximately 0.1 g of cells were washed and resuspended with 560 III TE buffer (10 mM Tris-Cl, 1 mM EDTA, pH 8.0). The cells were lysed by incubating for 1 hour with 60 Ilg Proteinase K (Sigma Chemie GmbH, Deisenhofen, Germany) and 30 III SDS (10%). DNA was prepared by the CT AB (hexadecyltrimethyl ammonium bromide) (Sigma Chemie GmbH, Deisenhofen, Germany) miniprep protocol for the preparation of bacterial genomic DNA (Wilson, 1987). The 16S ribosomal DNA (rDNA) was amplified by polymerase chain reaction (PCR) using a standard protocol (Saiki et al., 1988) with a Techne PHC-2 thermal cycler (Techne Corp., Cambridge, U.K.). Overlapping PCR-DNA fragments were generated in two separate reactions of 10 mM Tris-Cl (pH 8.3), 50 mM KCI, 1.5 mM MgCI 2, 200 11M of each dNTP, 60 pmol of each amplification primer, 1.0 Ilg chromosomal DNA, and 2.5 U Taq DNA polymerase (Boehringer, Mannheim, Germany) in 100 Itl reaction volumes overlaid with sterile mineral oil. Conserved priming sites; nucleotide positions 8-27 and the complement of positions 1488-1511 of the 16S rRNA gene (E. coli numbering) allowed the production of the first fragment and positions 1099-1113 of the 16S rRNA gene and the complement of positions 456-473 of the 23S rRNA gene (located approximately 0.7 kb downstream from the 3'-terminus of the 16S rRNA gene) enabled the production of the second, overlapping, fragment. The PCR used 30 cycles of the following profile: 1 minute denaturation at 94°C; 1 minute primer annealing at 52 °C; and 2 minutes primer extension at 72°C. Reaction products were elongated with a final incubation of 10 minutes at 72 0C. Amplified PCRDNA was extracted with chloroform: isoamyl alcohol (24: 1) and visualized, by agarose gel (1 %) electrophoresis and EtBr-staining, as single bands. DNA was excised from the gel and eluted using the Gene-Clean (Bio 101 Inc., La Jolla, Ca., U.S.A.) protocol. The isolated DNA was resuspended to a final volume of 40 III prior to sequencing. The 16S rDNA was sequenced directly by a modified Sequenase (United States Biochemical GmbH, Bad Homburg, Germany) procedure described by Rogall et al. (1990), using 8-10 III of the purified PCR-amplified DNA and 5 pmol of sequencing primer. The primers used for the 16S rRNA gene sequence determination have been described previously (Lane, 1991). Sequence data were aligned manually with known 16S rRNA sequences using conserved primary sequence and secondary structure characteristics as references (Woese et al., 1983, Gutell et al., 1985), and compared with more than 1200 bacterial and archael16S rRNA sequences (Olsen et aI., 1991). Sequence similarity and evolutionary distance values, incorporating the Jukes and Cantor (1969) correction factor for reverse mutations, were calculated for sequence-pair comparisons using only unambigu-

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A. Malmqvist, T. Welander, E. Moore,

A. Ternsrrom. G. Molin, and L-M. Stenstrom

ous, homologous nucleotide positions. A sequence "mask" (Lane, 1991) that allowed the comparison of 1323 positions was used. Phylogenetic trees were generated using a pairwise, weighted, least squares distance method (Olsen, 1987).

Results Four isolates able to reduce chlorate to chloride were obtained, one from the chemostat (isolate 1), one from biofilm reactor A (isolate 2) and two from biofilm reactor B (isolates 3 and 4). All four isolates were subjected to

phenotypic characterization and showed exactly the same results in all tests. Biolog GN microplates were not suitable for identification of the chlorate degrading isolates, possibly because of interference with the redox indicator used in the system. In spite of several attemps to modify inoculation strength and incubation conditions, it was not possible to obtain reproducable results. The G+C content was then determined for isolates 1 and 2, which showed identical G+C contents. Finally, ribosomal RNA sequencing was done on isolate 1, which was chosen as a type strain. An estimated 98%

Fig. 1. Electron micrograph of Ideonella dechloratans showing the cell wall structure. Bar, 0.5 !lm.

Fig.2. Electron micrograph of Ideonella dechloratans negatively stained, showing the flagella. Bar, 1 !lm.

Ideonella dechloratans gen. nov., sp. nov., a Chlorate Reducing Bacterium (1500 nucleotide positions) of the complete 16S rRNA gene sequence of I. dechloratans was determined. The sequence has been deposited with the EMBL data library under accession number X72724. As a first analysis, the 16S rRNA sequence of I. dechloratans was aligned with those of representative organisms from all known taxa of procaryotes for which there is sequence data. This comparison determined that I. dechloratans clusters phylogenetically within the beta-subgroup of the Proteobacteria. A subsequent comparison was done against the sequences of organisms of the Proteobacteria beta subgroup (25 organisms). This comparison determined that I. dechloratans was most similar to the organisms of the evolutionary branch that includes Comamonas testosteroni, Alcaligenes eutrophus and Burkholderia cepacia (Table 1). Evolutionary distances were calculated from masked sequence comparisons (Table 1) and a rooted dendrogram was generated to show the estimated phylogenetic relationship of I. dechloratans within the Proteo bacteria beta subgroup (Fig. 3). Table 1. Similarity and evolutionary distance calculations between the 16S rDNA sequence of ldeonella dechloratans and species of the Proteobacteria beta-subgroup and other reference orgamsms Organisms compared with Ideo nella dechloratans

% Sequence similarity"

Evolutionary distance* ".

Comamonas testosteroni Alcaligenes eutrophus Burkholderia cepacia Nitrosomonas europaea Nitrosolobus multiformis Spirillum volutans Achromobacter xylosoxidans Alcaligenes faecalis Chromobacterium fluviatile Methylophilus methylotrophus Chromobacterium violaceum Vitreoscilla stercoraria Neisseria gonorrhoeae Kingella kingae Simonsiella muelleri Eikenella corrodens Neisseria denitrificans Kingella denitrificans Neisseria elongata Escherichia coli Pseudomonas aeruginosa Agrobacterium tumefaciens Desulfovibrio desulfuricans Bacillus subtilis

90 90 89

0.06

'f

85

87 87 89 87 88

81

88

86 84 84 84 84 85 85 85 79 82 77

76 74

0.08 0.08 0.11 0.10 0.12

0.09

0.10 0.11 0.16 0.10 0.12 0.13 0.12 0.13 0.12 0.12 0.12 0.12 0.19 0.16 0.22 0.24 0.28

* Determined from unmasked sequence comparisons. * Determined from masked sequence comparisons.

Discussion The reactors used in the enrichment procedure were all inoculated from the same source and they were all fed the same medium. In view of this, it is not surprising that the

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results of the phenotypic characterization clearly showed the four isolates studied to be representatives of the same bacterial species. This chlorate reducing bacterium has a typically "pseudomonad" phenotype, i.e. it is a Gram negative, respiring, cytochrome C oxidase positive, motile rod capable of utilizing a wide spectrum of different classes of small organic molecules. However, defined as such, the genus Pseudomonas has for long been a "dumping ground" for bacteria now known to be phylogenetically unrelated. De Vos et al. (1989) determined that [Pseudomonas] organisms are distributed throughout the alpha (e.g. [Pseudomonas] diminuta, [Pseudomonas] azotocolligans, [Pseudomonas] echinoides, [Pseudomonas] paucimobilis), beta (e.g. Burkholderia cepacia, [Pseudomonas] acidovorans, [Pseudomonas] solanacearum) and gamma (e.g. Pseudomonas fluorescens and other "true" Pseudomonas spp., and [Pseudomonas] spp. belonging to the Alteromonas branch, the Vibrionaceae branch, the Shewanella branch and the Aeromonadaceae branch) subgroups of the Proteobacteria. Many of the [Pseudomonas] spp. listed in Bergey's Manual of Systematic Bacteriology, Volume 1 (Palleroni, 1984) have been renamed or transferred to other genera, such as Comamonas, Acidovorax, and Hydrogenophaga, etc. Most of these reclassified bacteria have G+C contents of 65-70 mol%, while the "true" Pseudomonas spp. have G+C contents of 57-64 mol%. Within the beta subgroup, at least four evolutionary branches have been recognized (De Vos et al., 1989). I. dechloratans belongs to a line comprising very different phenotypes: the very specialized plant pathogenic bacteria centered around Burkholderia cepacia, the Comamonas group which, despite a vigorous attack on most organic compounds seldom utilize carbohydrates, and the Hydrogenophaga group consisting mostly of hydrogen oxidizing bacteria ([Alcaligenes] eutrophus belongs to this group). Neither group has a phenotype corresponding to the chlorate reducing bacterium. The highest rRNA (unmasked) sequence similarity values obtained between the chlorate reducing bacterium and members of the beta subgroup were approximately 90% (Table 1). Such a 16S rRNA sequence similarity value (between 90-93%) is often noticed between members of different but related genera (e.g., Comamonas, Burkholderia cepacia and Alcaligenes). It is clear that I. dechloratans clusters within the "acidovorans complex". However sequence comparison of the 16S rRNA gene of I. dechloratans is limited due to the limited amount of sequence data currently available for organisms of the beta subgroup. Except for very few complete or nearly complete sequences, the only additonal 16S rRNA sequence analyses of organisms from the beta subgroup has come from the work of Busse et al. (1992), who analyzed xenobioticdegrading isolates and some reference organisms by comparison of sequence positions 1220 to 1377 (E. coli numbering). When the same sequence region of I. dechloratans was compared with the sequence data from the study of Busse et al., (1992) the closest similarity was found to Comamonas acidovorans (data not shown). There is a significant error in the comparison of sequence data of only 157 nucleotide positions, but this evidence helps con-

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A. Malmqvist, T. Welander, E. Moore, A. Ternstrom. G. Molin, and I.-M. Stenstrom

I

-

I

Bar represents approximately 0.01 evolutionary distance unit

Ideonella dechloratans Comamonas testosteroni Alcaligenes eutrophus Burkholderia cepacia

I I

Alcaligenes xylosoxidans Alcaligenes faecalis Nitroso monas europaea Spirillum volutans

r--

Chromoba cterium fluviatile Chromobacterium violaceum

'--

-

-

---1

Vitreoscilla stercora ria Eikenella corrodens Methylophilus methylotrophus Escherichia coli

firm the phylogenetic position of I. dechloratans within the "acidovorans" branch of the beta subgroup. Thus, it is apparent from the phylogenetic analysis, as well as from phenotypic characterization, that the chlorate reducing bacterium merits a separate genus, and we therefore propose the name Ideonella dechloratans for these orgalllsms. Although the isolation and characterization of Ideonella dechloratans definitely shed some light on the previously unknown bacteria able to reduce chlorate to chloride in the absence of oxygen, many questions concerning the microbiology of chlorate reduction remain to be answered. It is yet not known whether the ability to grow with chlorate as an electron acceptor is limited to a very specialized group of bacteria within the beta subgroup of the Proteobacteria or if it is widespread amongst different groups of heterotrophic bacteria, as is the ability to grow with nitrate as terminal electron acceptor. The earlier observations of enriched chlorate reducing bacteria with a morphology very different from that of the bacterium described in this paper (Malmqvist et al., 1991) indicate the existence of chlorate reducers other than ldeonella dechloratans. However, it is not possible to draw any conclusions about the phylogenetic relatedness of the spiralshaped bacteria and ldeonella dechloratans, based on morphological observations only.

Fig. 3. The estimated phylogenetic position of Ideonella dechloratans among other representative organisms belonging to the beta subgroup of the Proteobacteria. Sequence data used for comparison were obtained through the Ribosomal Database Project (Olsen et al., 1991).

The evolutionary as well as the ecological and biochemical aspects of the existence of microorganisms with the ability to grow with chlorate as an electron acceptor are intriguing. An interesting question is whether the chlorate reducing bacteria posess a unique electron transport chain, or if it is basically the same as for denitrifying bacteria. The resemblance of the chlorate ion to nitrate, which perhaps explains the previously reported reduction of chlorate to chlorite by the enzyme nitrate reductase, could mean that the chlorate reduction to chloride is carried out by a modified nitrate reducing enzyme system. This would explain the occurrence of chlorate reducers in nature in spite of the short time that has passed since chlorate was introduced into the environment by humans. It could also be an explanation to the inhibition of chlorate reduction by oxygen, though the energy yield with chlorate as an electron acceptor should be, at least theoretically, higher than the yield with oxygen as terminal electron acceptor. It must be stressed that for most organisms, despite the similarity between chlorate and nitrate, these compounds are not interchangeable as electron acceptors. The vast majority of denitrifiers seem to be incapable of growing with chlorate as an electron acceptor and Ideonella dechloratans lost its ability to utilize nitrate for respiration after several sub cultivations on chlorate. This may be explained by significant differences between the respiratory

Ideonella dechloratans gen. nov., sp. nov., a Chlorate Reducing Bacterium

enzymes of chlorate reducers and denitrifiers, but this observation may also be explained by differences in the ability of the cell to transport chlorate and nitrate across the cell membrane. In any case, the enzymes involved in the reduction of chlorite to chloride have, so far, not been identified. Studies of the enzymology of chlorate reduction, e.g. in cell free extracts, together with microbiological investigations focused on the taxonomy and ecology of chlorate reducing bacteria are interesting tasks for further research in the field of microbial chlorate reduction. Hopefully, such studies will lead to a better understanding of the biochemistry, the evolution and the ecology of chlorate reducing bacteria in the near future. Description of Ideonella gen. nov. Ideonella (Ide.on.el'. lao M.L. fem.n. derived from Ideon, the research centre where the bacterium was first described, and -ella. M.L. dim. ending). Gram-negative, mesophilic, straight or slightly curved asporogenous rods, 0.7 to 1.0 by 2.5 to 5 !lm in size. Motile by two or several polar or subpolar flagella. Do not produce prosthecae. No resting stages are known. Aerobic, having a strictly respiratory type of metabolism, with oxygen as terminal electron acceptor; chlorate but not sulphate can be used as an alternative electron acceptor, allowing growth to occur anaerobically. Chlorate is reduced to chloride. Cytochrome C-oxidase positive. Catalase weakly positive. Chemoorganotrophic, utilizing organic acids, amino acids and carbohydrates as sole carbon sources. Lipolytic and proteinolytic. The DNA base composition is 68 mol% G+C, as determined by HPLC. Member of the Comamonas group of the Bsubclass of Proteobacteria. The type species is Ideonella dechloratans. Description of Ideonella dechloratans sp. nov. Ideonella dechloratans (de.chlor.at'ans. M.L. adj; de L. pref. from; chloratans derived from chlorate; the chlorate reducing bacterium). Cells are 0.7 to 1.0 by 2.5 to 5!lm, typically 0.8 by 3 !lm, straight or slightly curved rods; and occurs as singles, in pairs and sometimes as short filaments with 4-5 cells. When occurring in pairs or short filaments, individual cells have a fusiform appearence with pointed ends. Cells are Gram-negative and contain several inclusions of unknown composition. Motile by two or several polar or subpolar flagella. Do not produce prosthecae. Do not produce endospores. No resting stages are known. Growth occurs at 12 to 42°C. No growth occurs at 10 or 46 0c. No growth occurs in broth with 3% NaCl. Colonies are circular, smooth and nonpigmented. Cytochrome C-oxidase positive. Catalase is produced, but in low quantities; conventional agar colony catalase test may be interpretated as negative. Metabolism strictly respiratory. Oxygen, nitrate and chlorate serve as terminal electron acceptors. The ability of using nitrate as an electron acceptor may be lost after several subcultivations on chlorate. Chlorate is reduced to chloride, nitrate to nitrite. Nitrite and sulphate do not serve as electron acceptors. Acetate, alanine, asparagine, butyrat, fructose, glucose, lactate, propionate, pyruvat, and succinate are utilized as sole carbon sources in mineral medium; both aerobically and anaerobically, in the latter case with chlorate as electron acceptor. With chlorate but not with oxygen, adipate

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and fumarate is also utilized. Aminobenzoate, phenol, and phenylalanine is not utilized. No acid is produced from cellobiose, glucose, or maltose in modified Hugh-Leifson medium. Proteinolytic against casein but not against egg yolk. Lipolytic against egg yolk, Tween 20, Tween 40, Tween 60 and Tween 80, but not against Tween 85. Isolated from chlorate enriched sewage water. The DNA base composition of two studied isolates was 68.1 mol% G+C, as determined by HPLC. Type strain is CCUG 30898 T . Acknowledgements. We thank Professor Birger Bergh, Department of Classical Studies, Lund University, for his kind help with the latinization of the name. We are also very grateful to Professor Claes Weibull, Department of Microbiology, Lund University, for the electron microscopy.

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Asa Malmqvist, ANOX AB, Ideon Research Park, S-22370 Lund, Sweden