Logo of aemPermissionsJournals.ASM.orgJournalAEM ArticleJournal InfoAuthorsReviewers
Appl Environ Microbiol. Jun 2002; 68(6): 2726–2730.
PMCID: PMC123969

Biodegradation of cis-Dichloroethene as the Sole Carbon Source by a β-Proteobacterium


An aerobic bacterium capable of growth on cis-dichloroethene (cDCE) as a sole carbon and energy source was isolated by enrichment culture. The 16S ribosomal DNA sequence of the isolate (strain JS666) had 97.9% identity to the sequence from Polaromonas vacuolata, indicating that the isolate was a β-proteobacterium. At 20°C, strain JS666 grew on cDCE with a minimum doubling time of 73 ± 7 h and a growth yield of 6.1 g of protein/mol of cDCE. Chloride analysis indicated that complete dechlorination of cDCE occurred during growth. The half-velocity constant for cDCE transformation was 1.6 ± 0.2 μM, and the maximum specific substrate utilization rate ranged from 12.6 to 16.8 nmol/min/mg of protein. Resting cells grown on cDCE could transform cDCE, ethene, vinyl chloride, trans-dichloroethene, trichloroethene, and 1,2-dichloroethane. Epoxyethane was produced from ethene by cDCE-grown cells, suggesting that an epoxidation reaction is the first step in cDCE degradation.

The extensive use of chloroethenes as solvents and synthetic feedstocks has resulted in widespread environmental contamination (22), which is of concern due to the toxicity and carcinogenicity of such compounds. Microbial metabolism is an important factor in determining the fate of chloroethenes in the biosphere (14). Several anaerobic bacteria use tetrachloroethene (perchloroethene) and trichloroethene (TCE) as electron acceptors, producing cis-1,2-dichloroethene (cDCE), vinyl chloride (VC), and ethene as end products (10, 15, 16, 17). Under aerobic conditions, ethene and VC can serve as carbon and energy sources for bacterial growth (3, 8), but thus far, no conclusive evidence exists for aerobic growth on any of the dichloroethenes (cis-dichloroethene, trans-dichloroethene [tDCE], or 1,1-dichloroethene).

Because cDCE accumulation is often a limiting factor in the biodegradation of chloroethenes in subsurface ecosystems, aerobic bacteria capable of growth on cDCE would provide a crucial missing link in the chain of microbial metabolism for this class of compounds. Thermodynamic calculations suggest that cDCE contains sufficient energy to support aerobic growth (4), and enzymes active on cDCE are known in hydrocarbon-oxidizing bacteria (5, 6, 13, 20, 24, 25). In addition, aerobic oxidation of cDCE to CO2 has been observed in microcosm and enrichment culture studies (2, 12). Encouraged by the facts described above, we hypothesized that aerobic growth on cDCE was possible and searched at a variety of contaminated sites for microorganisms able to use this compound as a sole source of carbon and energy.


Chemicals and media.

cis-Dichloroethene (97%), tDCE (98%), TCE (99.5%), and 1,2-dichloroethane (1,2-DCA) (99.8%) were obtained from Sigma-Aldrich. VC (99.5%) was obtained from Fluka, and ethene (99.5%) was obtained from Scott. All other chemicals were reagent grade. A minimal salts medium (MSM) based on that of Hartmans et al. (9) was used for enrichment cultures, with the following modifications: the phosphate concentration was reduced to 20 mM, the ammonium concentration was reduced to 10 mM, and the chloride concentration was reduced to 0.02 mM. The pH of MSM was adjusted to 7.0. Trypticase soy agar in which the nutrient content was one-quarter strength (1/4-TSA) (Difco) was used as a nonselective medium; the agar (Difco) concentration was 20 g/liter.

Enrichment cultures.

Samples of soil (5%, wt/vol), sediment (5%, wt/vol), granular activated carbon (5%, wt/vol), or groundwater (50%, vol/vol) were mixed with MSM to give a total volume of 50 ml in 160-ml serum bottles (headspace, 110 ml of air), which were crimp sealed with Teflon-faced butyl rubber stoppers (Wheaton). cDCE (3 μl of undiluted liquid) was added as the sole carbon source at a concentration of 40 μmol per bottle (initial aqueous concentration, 0.6 mM). Enrichment cultures were incubated at 20°C with shaking at 150 rpm.

Isolation and identification of strain JS666.

Repeated isolation attempts with minimal medium plates containing cDCE were unsuccessful, so an approach based on the dilution-to-extinction principle was adopted. Serial 10-fold dilutions of an enrichment culture (10−5 to 10−8) were prepared in triplicate in phosphate buffer, and 100 μl of each dilution was used to inoculate fresh MSM. cDCE was added, and the bottles were incubated as described above for the enrichments. After turbidity appeared in the cultures, samples were spread plated on 1/4-TSA plates, and the purity was evaluated. One resultant isolate (strain JS666) was characterized by standard bacteriological methods (9) and by amplification and sequencing of the 16S rRNA gene (MIDI Labs, Newark, Del.). Clustal-X software (21) was used for sequence alignment and generation of phylogenetic trees.

Growth of strain JS666.

Several colonies from a 1/4-TSA plate were inoculated into 50 ml of MSM and grown on cDCE (four 50-μmol additions) until the late exponential phase. The cells were washed twice in phosphate buffer (20 mM, pH 7.0) and suspended in 700 ml of MSM in a 2,240-ml flask (headspace, 1,540 ml of air) that was crimp sealed as described above. cDCE (60 μl) was added to the culture three to five times, with each addition equivalent to 790 μmol of cDCE/flask (initial aqueous concentration, 0.90 mM). The growth substrate range of strain JS666 was investigated by using various compounds as carbon sources (40 μmol/bottle) in 50 ml of MSM. Growth of cultures and transformation of substrates were monitored as described below. All JS666 cultures were grown at 20°C with shaking at 150 rpm.

Substrate range of resting cells.

Strain JS666 was grown on either succinate (disodium salt, 20 mM) or cDCE in 700 ml of MSM as described above. Cells were harvested in the exponential phase, washed twice in phosphate buffer, and suspended in 0.2 ml of buffer. The cells were transferred to a 10-ml serum vial, and substrate (3 μmol) was added, either dissolved in 0.8 ml of buffer or added as a gas directly to the headspace. In the latter case, the volume of the suspension was adjusted to 1 ml with buffer. Cells were suspended at an optical density at 600 nm (OD600) of 2.5 to 3.0 (1.1 to 1.4 mg of protein/ml) for the substrate range tests and at an OD600 of 15.1 to 15.5 (5.6 to 5.9 mg of protein/ml) for detailed analysis of ethene metabolism. The cell suspension vials were incubated horizontally at 20°C with shaking at 300 rpm. Substrate disappearance and protein concentrations were measured as described below. Abiotic losses (determined with sterile water controls) were subtracted from the observed rates of substrate disappearance before calculation of specific activities.

Kinetic study.

An inoculum (4 ml) from a cDCE-grown JS666 culture was transferred to 68 ml of fresh MSM in a 160-ml serum bottle. The culture was allowed to degrade approximately 100 μmol of cDCE in order to produce a sufficient amount of active biomass for kinetic experiments. Three sequential substrate depletion curves were generated at 20°C for the same serum bottle, which was kept inverted at an angle on a rotary shaker with shaking at 165 rpm between the times when headspace samples were removed. Estimates of the maximum specific substrate utilization rate (k) and the half-velocity constant (Ks) for cDCE transformation were obtained from headspace-based cDCE depletion curves. The data were fitted to the Monod model by using the Aquasim software program, as described previously (18, 26), with a diffusive link (19) included to account for the fact that both the liquid and gaseous phases could act as reservoirs of cDCE. The possibility of mass transfer limitation was addressed by constructing a nonequilibrium model with Aquasim, incorporating a mass transfer coefficient (KLa) measured as described previously (21). The depletion curves predicted by the nonequilibrium model coincided with those of the equilibrium model, which demonstrated the adequacy of our phase equilibrium assumption.

Protein was measured at the beginning of the first depletion curve and at the end of each depletion curve. Because of the relatively large amounts of cDCE (7 to 13 μmol) added at the beginning of each sequential depletion curve, biomass increased significantly during the course of the experiment (from 9.1 to 12.6 mg of protein/liter). However, because most of the batch depletion data were gathered near the end of each curve, the change in biomass within each intensive, data-gathering period was relatively small (<2%, as estimated by using the growth yield coefficient which we report here).

Analytical methods.

cDCE was analyzed in headspace samples (100 or 250 μl) by gas chromatography with flame ionization detection, and either a capillary column (GSQ; Agilent) or a packed column (10% SP-1000 on 100/120 Supelcoport [Supelco]). Both columns separated cDCE from tDCE, which was present as an impurity in the cDCE source at a concentration of approximately 2%. cDCE was quantified (micromoles per bottle) by comparison to a standard curve derived from known quantities of cDCE in serum bottles with the same gas and liquid volumes as the experimental bottles. For the kinetic studies, cDCE concentrations were converted to micromolar concentrations by methods described previously (7) by using a dimensionless Henry's constant of 0.1232 for cDCE at 20°C.

Protein concentrations were measured with the Micro-BCA reagent (Pierce Chemical Company). Culture fluid (0.9 ml) was mixed with 0.1 ml of 10 M NaOH, heated to 90°C for 10 min to effect cell lysis, and cooled to room temperature, and 1 ml of Micro-BCA working reagent was added. Samples were then incubated at 60°C for 60 min and cooled to room temperature, and the absorbance at 562 nm was read with a spectrophotometer (Hewlett Packard 8452A or Varian Cary 3E). Absorbance values were compared to the values for bovine serum albumin standards treated identically. The growth rate was calculated by fitting an exponential function to the plot of protein versus time. The growth yield was calculated from a linear regression of protein versus amount of cDCE consumed.

Chloride was quantified by the colorimetric method of Bergmann and Sanik (1), which we modified by using 1-ml samples, 0.2 ml of iron reagent, and 0.4 ml of thiocyanate reagent. Cells were removed by centrifugation (16,000 × g, 2 min) before the supernatant was used in the chloride assay.


Despite extended incubation times and the use of inocula (soil, water, sediments) from chloroethene-contaminated sites, only two active cultures were obtained from 18 aerobic enrichments (20°C) initiated with cDCE as the sole carbon source. One of the active enrichments was chosen for further study. The inoculum for the culture was granular activated carbon from a pump-and-treat plant (Dortmund, Germany) processing groundwater contaminated with perchloroethene, TCE, and cDCE. Biodegradation of cDCE in the enrichment began after 50 days, and cDCE-degrading activity was subsequently maintained for 7 months in the absence of other carbon sources through 11 transfers (5%, vol/vol) transfers in minimal medium.

A pure culture that grew on cDCE was obtained from serial dilutions of the Dortmund enrichment culture. The cDCE-degrading isolate, strain JS666, is a yellow-pigmented, gram-negative, nonmotile, oxidase-positive, catalase-negative rod. Phylogenetic analysis of the 16S rRNA gene (GenBank accession no. AF408397) indicated that JS666 is a member of the family Comamonadaceae in the β-proteobacteria, with 97.9% sequence identity to the Antarctic isolate Polaromonas vacuolata (Fig. (Fig.1).1). It is unclear whether JS666 is truly a Polaromonas strain due to many phenotypic differences, including differences in motility, the catalase reaction, pigmentation, and the optimum temperature (11). Assignment of strain JS666 to a genus is difficult at present due to the lack of taxonomic data for the isolate and to the uncertain phylogeny of some members of the Comamonadaceae (28).

FIG. 1.
Phylogeny of strain JS666 based on 16S rRNA gene comparison. GenBank accession numbers are given below the species names. Positions containing gaps or ambiguous nucleotides were removed, leaving sequences consisting of 1,434 bases for analysis. Bootstrap ...

Strain JS666 grew on cDCE as the sole carbon and energy source (Fig. (Fig.2).2). Batch cultures grown on cDCE as the sole carbon source did not reach high optical densities, but protein measurements (Fig. (Fig.2,2, inset) confirmed that growth was coupled to cDCE degradation, with a calculated growth yield of 6.1 ± 0.4 g of protein/mol of cDCE. The doubling time on cDCE calculated from the protein data was 74 ± 8 h at 20°C (data not shown). The growth rate and yield with cDCE are both lower than corresponding values for growth on 1,2-DCA (23) or VC (8), probably due to the lower incubation temperature used in this study and the lower energy content of cDCE (4).

FIG. 2.
Growth of strain JS666 on cDCE as the sole carbon and energy source. Symbols: [open triangle], cDCE content; □, cumulative amount of cDCE consumed; ○, biomass expressed as OD600; [open diamond], chloride content. Due to partitioning between the headspace ...

The stoichiometry of chloride production (Fig. (Fig.2)2) indicates that cDCE was completely dechlorinated (1.94 mol of Cl produced/mol of cDCE degraded). There was no detectable growth in JS666 cultures without cDCE, and there was no significant disappearance of cDCE in flasks inoculated with autoclaved cells (data not shown). The pH optimum for growth of strain JS666 on cDCE was 7.2. Growth on cDCE was optimal at temperatures between 20 and 25°C and was not detectable at 30°C. tDCE was present as an impurity (approximately 2%) in the cDCE used in this work, but tDCE was not metabolized during growth on cDCE. In other experiments (data not shown), some of the tDCE impurity was transformed after cDCE was depleted.

Strain JS666 did not grow on tDCE, TCE, VC, 1,2-DCA, or ethene as a carbon source, but cells could transform all these compounds after growth on cDCE (Table (Table1).1). The enzymes involved are inducible, as indicated by the lower activity in succinate-grown cells. The ability of cDCE-grown JS666 cells to transform other chloroethenes may prove to be very useful at contaminated sites, where mixtures of pollutants may be encountered (22). It is surprising that strain JS666 did not grow on ethene, which seems to be the most likely natural substrate of the cDCE-degrading enzymes, particularly considering the fact that the VC-assimilating bacteria isolated to date also use ethene as a carbon source (8, 26) and at least in one case appear to have evolved directly from ethene-degrading bacteria (27).

Activity of cDCE-grown and succinate-grown JS666 cells with chloroethenes, ethene, and 1,2-DCA as substrates

Dense suspensions of cDCE-grown cells oxidized ethene stoichiometrically to epoxyethane (Fig. (Fig.3)3) and also oxidized propene to epoxypropane (data not shown). Succinate-grown cells also catalyzed ethene transformation, but the rate was only 40% of the rate observed with cells grown on cDCE (data not shown). At this time we cannot explain the observed differences in the activity of succinate-grown cells in the epoxyethane accumulation experiment and the substrate range assays (Table (Table1),1), although it should be noted that the former experiment was performed with much denser cell suspensions, which may have affected activity.

FIG. 3.
Production of epoxyethane ([open triangle]) from ethene (○) by cDCE-grown JS666 cells. Due to partitioning of both compounds between the headspace and the liquid phase, the concentrations are expressed in micromoles per bottle. The data points are averages ...

The ability of JS666 cells to convert ethene to epoxyethane suggests that oxidation of cDCE to the corresponding epoxide is the first step in cDCE biodegradation in strain JS666. cDCE epoxide was not detected when cells were incubated with cDCE, however, which could have been due to further metabolism of the epoxide by the cells. Monooxygenase-catalyzed epoxidations appear to be a universal initial step in the aerobic metabolism of chloroethenes in bacteria (5, 6, 8, 24, 25). Further work will be required to determine rigorously whether strain JS666 uses the same strategy.

The kinetics of cDCE metabolism in strain JS666 at 20°C with continuous agitation were studied by simultaneously fitting depletion curves to three sets of data (Fig. (Fig.4).4). A k of 12.6 ± 0.3 nmol/min/mg of protein and a Ks of 1.6 ± 0.2 μM best fit all three data sets. The k value calculated from depletion curves (Fig. (Fig.4)4) agreed fairly well with the cDCE utilization rate seen in substrate range assays (Table (Table1).1). However, by using the k value and growth yield (Y) (Fig. (Fig.2),2), a doubling time (ln2/Yk) of 150 h was estimated, which is at odds with the doubling time determined directly from protein measurements during growth (74 h). The discrepancy suggests that there was underestimation of k in the substrate depletion experiments, probably due to a lower active fraction of protein (i.e., protein measurements probably overestimated the active biomass) under the conditions of the substrate depletion assay compared to cells in exponential-phase cultures. Note that differences in the active fraction of protein would not have affected estimates of Ks.

FIG. 4.
Three sequential depletion curves for cDCE at 20°C obtained for JS666 (value ± 95% confidence interval). Symbols: ○, experiment 1 measured values; □, experiment 2 measured values; [open triangle], experiment 3 measured values. ...

The relatively low measured Ks value, considered in conjunction with the relatively high k value, is significant considering the possible participation of this organism in natural attenuation of cDCE. If JS666 were present and active at a cDCE-contaminated site, the cDCE utilization rate would be one-half the maximum rate at a cDCE concentration of 160 μg/liter. The Environmental Protection Agency-mandated maximum contaminant level for cDCE in drinking water is 70 μg/liter (http://www.epa.gov). Therefore, under appropriate conditions in the field, JS666 should easily be able to oxidize cDCE to obtain levels below drinking water standard levels. Perhaps more relevant is the observation that in the experiments described here, cDCE was degraded routinely to concentrations below 0.03 μg/liter.

The discovery of a bacterium able to grow on cDCE shows that aerobic biodegradation of cDCE in the absence of other carbon substrates is possible. Our results with enrichment cultures indicate that such bacteria appear to be rare and may exist only in highly selective artificial environments, such as the activated carbon filter that was the source of strain JS666. The existence of cDCE-assimilating bacteria suggests that there is potential for bioaugmentation, which could lead to a self-sustaining, low-cost bioremediation strategy at sites where cDCE is a problem contaminant. Our results indicate that growth on cDCE as a carbon source could be a previously unrecognized factor in determining the environmental fate of this compound. Further characterization of JS666 should facilitate the search for similar strains and allow evaluation of the role of such strains in the natural attenuation of cDCE and other chlorinated ethenes.


We thank Petra Koziollek for providing the activated carbon sample. Mike Allen and John Dunlap helped with phenotypic characterization of JS666.

This work was funded by the U.S. Strategic Environmental Research and Development Program. N.V.C. was supported by a postdoctoral fellowship from the Oak Ridge Institute for Science and Education (U.S. Department of Energy).


1. Bergmann, J. G., and J. Sanik. 1957. Determination of trace amounts of chlorine in naphtha. Anal. Chem. 29:241-243.
2. Bradley, P. M., and F. H. Chapelle. 2000. Aerobic microbial mineralization of dichloroethene as sole carbon source. Environ. Sci. Technol. 34:221-223.
3. de Bont, J. A. M., and W. Harder. 1978. Metabolism of ethylene by Mycobacterium E 20. FEMS Microbiol. Lett. 3:89-93.
4. Dolfing, J., A. J. van den Wijngaard, and D. B. Janssen. 1993. Microbiological aspects of the removal of chlorinated hydrocarbons from air. Biodegradation 4:261-282. [PubMed]
5. Ensign, S. A., M. R Hyman, and D. J. Arp. 1992. Cometabolic degradation of chlorinated alkenes by alkene monooxygenase in a propylene-grown Xanthobacter strain. Appl. Environ. Microbiol. 58:3038-3046. [PMC free article] [PubMed]
6. Fox, B. G., J. G. Bornemann, L. P. Wackett, and J. D. Lipscomb. 1990. Haloalkene oxidation by the soluble methane monooxygenase from Methylosinus trichosporium OB3b: mechanistic and environmental implications. Biochemistry 29:6419-6427. [PubMed]
7. Gossett, J. M. 1987. Measurement of Henry's Law constants for C1 and C2 chlorinated hydrocarbons. Environ. Sci. Technol. 21:202-208.
8. Hartmans, S., and J. A. M de Bont. 1992. Aerobic vinyl chloride metabolism in Mycobacterium aurum L1. Appl. Environ. Microbiol. 58:1220-1226. [PMC free article] [PubMed]
9. Hartmans, S., A. Kaptein, J. Tramper, and J. A. M. de Bont. 1992. Characterization of a Mycobacterium sp. and a Xanthobacter sp. for the removal of vinyl chloride and 1,2-dichloroethane from waste gases. Appl. Microbiol. Biotechnol. 37:796-801.
10. Holliger, C., D. Hahn, H. Harmsen, W. Ludwig, W. Schumacher, B. Tindall, F. Vazquez, N. Weiss, and A. J. Zehnder. 1998. Dehalobacter restrictus gen. nov. and sp. nov., a strictly anaerobic bacterium that reductively dechlorinates tetra- and trichloroethene in an anaerobic respiration. Arch. Microbiol. 169:313-321. [PubMed]
11. Irgens, R. L., J. J. Gosink, and J. T. Staley. 1996. Polaromonas vacuolata gen. nov., sp. nov., a psychrophilic, marine, gas-vacuolate bacterium from Antarctica. Int. J. Syst. Bacteriol. 46:822-826. [PubMed]
12. Klier, N. J., R. J. West, and P. A. Donberg. 1999. Aerobic biodegradation of dichloroethylenes in surface and subsurface soils. Chemosphere 38:1175-1188. [PubMed]
13. Koziollek, P., D. Bryniok, and H.-J. Knackmuss. 1999. Ethene as an auxiliary substrate for the cooxidation of cis-dichloroethene and vinyl chloride. Arch. Microbiol. 172:240-246. [PubMed]
14. Lee, M. D., J. M. Odom, and R. J. Buchanan, Jr. 1998. New perspectives on microbial dehalogenation of chlorinated solvents: insights from the field. Annu. Rev. Microbiol. 52:423-452. [PubMed]
15. Maymó-Gatell, X., Y. Chien, J. M. Gossett, and S. H. Zinder. 1997. Isolation of a bacterium that reductively dechlorinates tetrachloroethene to ethene. Science 276:1568-1571. [PubMed]
16. Maymó-Gatell, X., I. Nijenhuis, and S. H. Zinder. 2001. Reductive dechlorination of cis-1,2-dichloroethene and vinyl chloride by “Dehalococcoides ethenogenes.” Environ. Sci. Technol. 35:516-521. [PubMed]
17. Neumann, A., H. Scholz-Murumatsu, and G. Diekert. 1994. Tetrachloroethene metabolism of Dehalospirillum multivorans. Arch. Microbiol. 162:295-301. [PubMed]
18. Reichert, P. R. 1994. AQUASIM-a tool for simulation and data analysis of aquatic systems. Water Sci. Technol. 2:21-30.
19. Reichert, P. R. 1998. AQUASIM 2.0-user manual. Swiss Federal Institute for Environmental Science and Technology. Dubendorf, Switzerland.
20. Semprini, L. 1997. Strategies for the aerobic co-metabolism of chlorinated solvents. Curr. Opin. Biotechnol. 8:296-308. [PubMed]
21. Smatlak, C. R. 1995. Comparative kinetics of H2 utilization for dechlorination of tetrachloroethene and methanogenesis by an anaerobic enrichment culture. M.S. thesis. Cornell University, Ithaca, N.Y.
22. Squillace, P. J., M. J. Moran, W. W. Lapham, C. V. Price, R. M. Clawges, and J. S. Zogorski. 1999. Volatile organic compounds in untreated ambient groundwater of the United States, 1985-1995. Environ. Sci. Technol. 33:4176-4187.
23. van den Wijngaard, A. J., R. D. Wind, and D. B. Janssen. 1993. Kinetics of bacterial growth on chlorinated aliphatic compounds. Appl. Environ. Microbiol. 59:2041-2048. [PMC free article] [PubMed]
24. van Hylckama Vlieg, J. E. T., J. Kingma, A. J. van den Wijngaard, and D. B. Janssen. 1998. A glutathione S-transferase with activity towards cis-dichloroepoxyethane is involved in isoprene utilization by Rhodococcus sp. strain AD45. Appl. Environ. Microbiol. 64:2800-2805. [PMC free article] [PubMed]
25. van Hylckama Vlieg, J. E. T., W. de Koning, and D. B. Janssen. 1996. Transformation kinetics of chlorinated ethenes by Methylosinus trichosporium OB3b and detection of unstable epoxides by on-line gas chromatography. Appl. Environ. Microbiol. 62:3304-3312. [PMC free article] [PubMed]
26. Verce, M. F., R. L. Ulrich, and D. L. Freedman. 2000. Characterization of an isolate that uses vinyl chloride as a growth substrate under aerobic conditions. Appl. Environ. Microbiol. 66:3535-3542. [PMC free article] [PubMed]
27. Verce, M. F., R. L. Ulrich, and D. L. Freedman. 2001. Transition from cometabolic to growth-linked biodegradation of vinyl chloride by a Pseudomonas sp. isolated on ethene. Environ. Sci. Technol. 35:4242-4251. [PubMed]
28. Wen, A., M. Fegan, C. Hayward, S. Chakraborty, and L. I. Sly. 1999. Phylogenetic relationships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong 1926 and Tamaoka et al. 1987) gen. nov., comb. nov. Int. J. Syst. Bacteriol. 49:567-576. [PubMed]

Articles from Applied and Environmental Microbiology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...