Cadmium Removal by a New Strain of Pseudomonas aeruginosa in ...

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APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Oct. 1997, p. 4075–4078 0099-2240/97/$04.0010 Copyright © 1997, American Society for Microbiology

Vol. 63, No. 10

Cadmium Removal by a New Strain of Pseudomonas aeruginosa in Aerobic Culture CLIFFORD L. WANG,1 PETER C. MICHELS,1 SCOTT C. DAWSON,1 SITSARI KITISAKKUL,1 JOHN A. BAROSS,2 J. D. KEASLING,1 AND DOUGLAS S. CLARK1* School of Oceanography, University of Washington, Seattle, Washington 98195,2 and Department of Chemical Engineering, University of California, Berkeley, California 947201 Received 30 June 1997/Accepted 10 July 1997

A fluorescent pseudomonad (strain CW-96-1) isolated from a deep-sea vent sample grew at 30°C under aerobic conditions in an artificial seawater medium and tolerated cadmium concentrations up to 5 mM. After 140 h, strain CW-96-1 removed >99% of the cadmium from solution. Energy dispersive microanalysis revealed that the cadmium was removed by precipitation on the cell wall; sulfide production was confirmed by growth on Kligler’s agar. Based on 16S ribosomal DNA sequencing and fatty acid analysis, the microorganism is closely related to Pseudomonas aeruginosa. containing citrate. One hundred forty hours after inoculation, strain CW-96-1 removed .99% of the cadmium from solution. Electron micrographs and energy-dispersive microanalysis indicated that cadmium was bound to the cell wall as a sulfur complex with a 1:1 stoichiometry. Fatty acid analysis and 16S ribosomal DNA (rDNA) sequencing identified CW-96-1 as a strain of Pseudomonas aeruginosa. Cell growth and cadmium tolerance. Bacteria were isolated from 2°C samples collected in 1991 from a deep-sea hydrothermal vent plume at the Endeavor Segment of the Juan de Fuca Ridge. Plume water was diluted 10,000-fold in artificial seawater (4), and 10 ml from each dilution was filtered through 0.2-mm-pore-size Nuclepore filters by sterile techniques. The filters were placed on the surface of prechilled FMS agar slants in 50-ml serum bottles (FMS medium is the same as the growth medium described below with the exception that it contains 1.2% purified agar predialyzed in artificial seawater to remove soluble organic material), capped with sterile rubber stoppers and incubated at 5°C with a hydrogen-air (1:9) headspace. Enrichment at 5°C on FMS agar medium yielded a mixed culture, which was then purified at both 5 and 22°C. CW-96-1 was isolated from a 22°C enrichment by plating on Luria agar at 37°C. Growth experiments were performed in artificial seawater medium (pH 7) containing (per liter) 19.6 g of NaCl, 3.3 g of Na2SO4, 0.5 g of KCl, 0.05 g of KBr, 0.02 g of H3BO3, 8.8 g of MgSO4, 1.0 g of NaNO3, 5.0 g of NaS2O3, 2.0 mg of FeSO4 z 7H2O, 0.15 g of MnSO4 z H2O, 0.1 g of CaCl2, 0.43 g of (NH4)2SO4, 35 mg of KH2PO4, 10 mg of yeast extract, and 1.0 g of PIPES [piperazine-N,N9-bis(2-ethanesulfonic acid)]. Ten milliliters of the trace element solution described by Baross (4) was also added. The medium was adjusted to pH 7

The toxic effects of cadmium on microorganisms are well documented (3, 5, 25) and derive from several mechanisms. Disruption of protein function can occur through binding of cadmium to sulfhydryl groups (7, 12). In addition, cadmium competes with several divalent ions such as Ca21, Zn21, and Mn21 for metal binding sites in biological systems (9, 16). Binding of cadmium to nucleotides leads to single-strand breaks in cellular DNA (9, 16). These potent toxic effects can result in prolonged lag phase, decreased growth rate, lower cell density, or death for bacteria and algae at levels below 1 ppm of cadmium (112.4 ppm 5 1 mM) (2, 14, 16, 20, 22). Microorganisms indigenous to heavy metal-containing environments have evolved several distinct mechanisms of heavy metal tolerance. A common plasmid-encoded mechanism employs heavy metal efflux pumps, which specifically capture and eject undesirable divalent cations through the cell membrane (18, 23). Alternatively, heavy metals can be sequestered by adsorption to the cell wall (17, 21, 29) or by binding to detoxifying ligands, proteins, or polymers (1, 8, 10, 13, 18, 26, 30). Microbially mediated precipitation of heavy metals as insoluble sulfides, carbonates, phosphates, or hydroxides can also reduce the bioavailable concentration of the toxic ions (1, 2, 24). Finally, enzymes can convert metal ions into organometallic compounds, thereby detoxifying or volatilizing the metal ions (18, 24). Hydrothermal vent waters are enriched in heavy metals, including cadmium (27, 28). This habitat might be a rich source of organisms with high tolerance to cadmium and/or efficient mechanisms of cadmium removal from the surrounding solution. Previous investigations of 299 strains isolated from the bodies of hydrothermal vent tube worms indicated that a significant number could tolerate 100-ppm cadmium, although no removal of metal was evident (11, 19). It remained to be seen whether any free-living hydrothermal vent organisms also evolved a high resistance to cadmium, and whether the mechanism of such tolerance might be suitable for the bioremediation of heavy metals. We describe here a microorganism that is able to grow in concentrations of cadmium up to 5 mM. This organism grew rapidly in shake flasks on a defined artificial seawater medium

TABLE 1. Growth and cadmium removal by P. aeruginosa CW-96-1 in artificial seawater medium 50 H Initial Cd (mM)

0 1.0 3.0 5.0

* Corresponding author. Phone: (510) 642-2408. Fax: (510) 6421228. E-mail: [email protected] 4075

OD600

pH

1.37 1.31 1.54 0.45

8.5 8.5 8.3 8.1

140 H Cadmium removed (%)

.99 92 14

OD600

pH

0.43 0.94 1.22 1.30

9.1 9.1 9.1 9.1

Cadmium removed (%)

.99 .99 .99

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cating that at least some cells remained viable after cadmium removal. Sulfide production. Plating strain CW-96-1 on Kligler’s agar yielded black colonies; the appearance of black colonies indicated sulfide production. Electron microscopy and energy-dispersive X-ray spectroscopy (EDXS). In preparation for transmission electron microscopy, cell suspensions were fixed in an equal volume of 1% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.2, for 1 h and postfixed in 0.5% osmium tetroxide for 30 min. The

FIG. 1. Growth and cadmium removal by strain CW-96-1. Open circles are optical density. Filled circles are the cadmium concentration in the culture supernatant. Error bars on the cadmium concentration measurements may not be visible for all data because the error was very small.

with 6 M NaOH and passed through a 0.2-mm-pore-size sterile filter. Cadmium was added from a filter-sterilized stock solution of 1 M CdCl2. The primary carbon source, citrate, was added from a 1 M stock solution to give a final concentration of 30 mM. Cells were grown in 250-ml shake flasks containing 50 ml of cadmium-supplemented medium (0 to 10 mM Cd) inoculated with 0.5 ml of cells (1% inoculum). Flasks were incubated at 30°C and agitated at 200 rpm in a shaker bath. Optical density at 600 nm (OD600) and pH were monitored. Selected samples were transferred to Eppendorf tubes and centrifuged for 3 min at 17,500 3 g. The supernatant was drawn and stored at 220°C for cadmium analysis. The cadmium concentration in the supernatant was determined with a Perkin-Elmer 2380 atomic absorption spectrometer at 228.8 nm with a cadmium lamp. Data for the growth of strain CW-96-1 with and without cadmium are presented in Table 1. After 50 h, growth (as determined by optical density) was not significantly affected by cadmium concentrations up to 3 mM, and optical densities greater than 1.3 were obtained in cultures with and without cadmium. Growth was slower in 5 mM cadmium, achieving an OD600 of only 0.45 after 50 h but reaching an optical density of 1.3 after 140 h. Growth was not observed in 10 mM cadmium. Optical density was affected not only by cell density but also by precipitation of cadmium and flocculation of cells. Suspended precipitate most likely contributed to the high optical density observed after 50 h in 3 mM cadmium. On the other hand, the decrease in optical density after 140 h in 0 to 3 mM cadmium can be explained by settling of flocs from suspension. Increases in pH accompanied growth, with cultures in 0 to 5 mM Cd reaching a pH of ca. 9 by 140 h after inoculation. Cadmium removal. One millimolar soluble cadmium was almost completely removed from solution after 40 h. The majority of the cadmium removal occurred after the exponential growth phase, approximately 30 h after inoculation (Fig. 1). Removal of soluble cadmium was dependent on growth of the cultures, with near-complete removal of up to 5 mM cadmium after 6 days. However, no cadmium was removed in cultures containing 10 mM cadmium due to a lack of growth. Decreases in soluble cadmium were accompanied by the appearance of a bright yellow precipitate, presumably cadmium sulfide. Abiotic precipitation was not a significant contributor to the removal; the soluble cadmium level decreased by less than 10% in cellfree controls. In addition, fresh medium could be inoculated with cultures that had removed .99% of the cadmium, indi-

FIG. 2. Electron micrographs and thin-film EDXS analysis of strain CW96-1. Cells were grown aerobically in citrate medium containing ;1 mM cadmium and harvested in stationary phase. The top picture is an electron micrograph of strain CW-96-1 showing the electron-dense surface layer. The middle and bottom pictures are EDXS pictures showing the distribution of cadmium and sulfur, respectively.

VOL. 63, 1997

CADMIUM REMOVAL BY A NEW STRAIN OF P. AERUGINOSA

FIG. 3. EDXS spectrum of the cell walls shown in Fig. 2. The peaks labeled copper (Cu) and osmium (Os) are due to the copper grid and the osmium tetroxide fixative, respectively.

samples were then dehydrated in a graded acetone series and embedded in Epon-Araldite resin. Sections of 50 nm in thickness were cut with a Reichert Ultracut E microtome and collected on uncoated 300-mesh grids. The transmission electron micrographs of cells in stationary phase showed unusual electron-dense granules, including a dense layer associated with the cell surface (Fig. 2). To determine whether this dense surface layer contained cadmium, EDXS was performed with a JEOL 200CX scanning transmission electron microscope and a Kevex 8000 EDX system (128- by 64-pixel X-ray map; 300-ms dwell time per pixel; TEMSTAR software). EDXS of the cell surface layer revealed the presence of cadmium and sulfur (Fig. 2 and 3), and elemental analysis of this material gave a cadmium-to-sulfur ratio of ca. 1:1. Identification. The microorganism was identified by fatty acid analysis and 16S rDNA sequencing. Fatty acid analysis was carried out by Microbial I.D., Inc. (Newark, Del.), and showed a similarity index of 0.923 to P. aeruginosa. Smallsubunit rDNA of CW-96-1 was amplified in the PCR for automated sequencing. Evolutionary relationships of CW-96-1 to known cultured organisms were determined by the maximum likelihood method with 100 bootstrapped resamplings (Fig. 4).

FIG. 4. Evolutionary relationships of CW-96-1 to known cultured organisms. Numbers at branch points indicate percentages of bootstrap support for the particular branching topology. The scale bar corresponds to 10% nucleotide sequence difference.

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Combined with a high percentage similarity (97% of 1,331 homologous nucleotides), molecular phylogeny clearly indicates that CW-96-1 is a strain of P. aeruginosa. The results obtained for strain CW-96-1 are exceptional in many respects. It appears that a primary mechanism of cadmium removal is precipitation as CdS. However, most sulfideproducing organisms capable of metal precipitation do so only under anaerobic conditions (6, 15). In contrast, strain CW-96-1 grows and removes cadmium under aerobic conditions. This mechanism of cadmium resistance was observed previously for two strains of Klebsiella aerogenes (1, 2) but only at cadmium concentrations up to 60 ppm (0.54 mM). By comparison, strain CW-96-1 grows in medium containing 5 mM cadmium, though the bioavailability of Cd21 may be affected by formation of cadmium-citrate complexes. Efficient metal removal and growth over a range of metal concentrations under aerobic conditions are advantages that may offer unique opportunities to employ this organism for metal remediation in simple reactors or even in situ. We are grateful to Sarah Perkins for assistance with the experiments and to Andrew Magyarosy for many helpful discussions. This research was funded by the Biotechnology Research and Education Program at the University of California and by a Department of Energy grant to N. Pace (DE-FG03-96ER62293). REFERENCES 1. Aiking, H., A. Stijnman, C. V. Garderen, H. V. Heerikhuizen, and J. V. T. Riet. 1984. Inorganic phosphate accumulation and cadmium detoxification in Klebsiella aerogenes NCTC 418 growing in continuous culture. Appl. Environ. Microbiol. 47:374–377. 2. Aiking, H., K. Kok, H. V. Heerikhuizen, and J. V. T. Riet. 1982. Adaptation to cadmium by Klebsiella aerogenes growing in continuous culture proceeds mainly via formation of cadmium sulfide. Appl. Environ. Microbiol. 44:938– 944. 3. Babich, H., and G. Stotzky. 1977. Sensitivity of various bacteria, including Actinomycetes, and fungi to cadmium and the influence of pH on sensitivity. Appl. Environ. Microbiol. 33:681–695. 4. Baross, J. A. 1993. Isolation and cultivation of hyperthermophilic bacteria from marine and freshwater habitats, p. 21–30. In P. F. Kemp, B. F. Sherr, E. B. Sherr, and J. J. Cole (ed.), Handbook of methods in aquatic microbiology. Lewis Publishers, Boca Raton, Fla. 5. Bowman, J. P., L. I. Sly, and A. C. Hayward. 1990. Patterns of tolerance to heavy metals among methane-utilizing bacteria. Lett. Appl. Microbiol. 10: 85–87. 6. Brown, D. E., G. R. Groves, and J. D. A. Miller. 1973. pH and eH control of cultures of sulphate-reducing bacteria. J. Appl. Chem. Biotechnol. 23:141– 149. 7. Cunningham, D. P., and L. L. Lundie. 1993. Precipitation of cadmium by Clostridium thermoaceticum. Appl. Environ. Microbiol. 59:7–14. 8. Gekeler, W., E. Grill, E. Winnacker, and M. H. Zenk. 1988. Algae sequester heavy metals via synthesis of phytochelatin complexes. Arch. Microbiol. 150:197–202. 9. Hughes, M. N., and R. K. Poole. 1989. Metals and microorganisms, p. 290. Chapman and Hall, New York, N.Y. 10. Inouhe, M., M. Hiyama, H. Tohoyama, M. Joho, and T. Murayama. 1989. Cadmium-binding protein in a cadmium-resistant strain of Saccharomyces cerevisiae. Biochim. Biophys. Acta 993:51–55. 11. Jeanthon, C., and D. Prieur. 1990. Susceptibility to heavy metals and characterization of heterotrophic bacteria isolated from two hydrothermal vent polychate annelids, Alvinella pompejana and Alvinella caudata. Appl. Environ. Microbiol. 56:3308–3314. 12. Jungmann, J., H.-A. Reins, C. Schobert, and S. Jentsch. 1993. Resistance to cadmium mediated by ubiquitin-dependent proteolysis. Nature 361:369–371. 13. Kurek, E., A. J. Francis, and J.-M. Bollag. 1991. Immobilization of cadmium by microbial extracellular products. Arch. Environ. Contam. Toxicol. 20:106– 111. 14. Les, A., and R. W. Walker. 1984. Toxicity and binding of copper, zinc, and cadmium by the blue-green alga, Chroosoccus paris. Water Air Soil Pollut. 23:129–139. 15. Lovley, D. R. 1993. Anaerobes into heavy metal: dissimilatory metal reduction in anoxic environments. Tree 8:213–217. 16. Mitra, R. S., and I. A. Bernstein. 1978. Single-strand breakage in DNA of Escherichia coli exposed to Cd21. J. Bacteriol. 133:75–80. 17. Mullen, M. D., D. C. Wolf, F. G. Ferris, T. J. Beveridge, C. A. Flemming, and G. W. Bailey. 1989. Bacterial sorption of heavy metals. Appl. Environ. Microbiol. 55:3143–3149.

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