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Phenanthrene degradation by microorganisms isolated from a contaminated stream
ENVIRONMENTAL POLLUTION
Environmental Pollution 101 (1998) 355±359
Phenanthrene degradation by microorganisms isolated from a contaminated stream M.C. Romero*, M.C. Cazau, S. Giorgieri, A.M. Arambarri Instituto Spegazzini, Facultad de Ciencias Naturales, Universidad Nacional de La Plata, calle 53, no 477-1900 La Plata, Argentina Received 22 April 1997; accepted 17 March 1998
Abstract Autochthonous yeast and bacteria strains were isolated from a contaminated stream and studied for their potential to degrade phenanthrene, a three-ring polycyclic aromatic hydrocarbon (PAH), as their only source of carbon and energy. Rhodotorula glutinis and Pseudomonas aeruginosa were the prevailing microorganisms utilizing phenanthrene. Cells of these microorganisms were inoculated in liquid mineral basal medium at 25, 50, 100 and 200 mg phenanthrene literÿ1 in ethanolic solution. To volatilize the ethanol the medium was dried for 40 min at 35 C with UV exposure. Almost complete phenanthrene degradation was observed during the 1-month incubation period. R. glutinis grew exponentially to a maximum of 9.5106 colony-forming units (CFUs) mlÿ1, with a short lag time only at 200 mg literÿ1. The highest density (1.4107 CFU mlÿ1) without a lag time was obtained at 50 mg literÿ1; lower cell numbers were observed at 25 and 100 mg literÿ1. During the logarithmic phase the speci®c growth rate (m) was 0.022 hÿ1 and the corresponding doubling time (td) was 31.5 h (50 mg literÿ1). A reduction to one-tenth of cell yeast numbers was observed at the lowest PAH levels on the 28th incubation day and a decrease of 100-fold was observed at 200 mg literÿ1. Higher m-values (m=d [ln x/x0]/dt), shorter td (td=[ln 2]/m), and a maximum of 7.7±7.4108 CFU mlÿ1 were observed in P. aeruginosa cultures at 50±100 mg literÿ1, respectively, on the 22nd day. At 25 and 200 mg literÿ1 lower bacteria numbers were obtained (2.1±3.0108 CFU mlÿ1) on the 28th day. R. glutinis was as active as P. aeruginosa at growing on phenanthrene; the aromatic hydrocarbon degradation correlated directly to microbial density and biomass increase, the highest biomass reaching 238.0 and 50.0 mg literÿ1 for the yeast and bacteria species, respectively. # 1998 Elsevier Science Ltd. All rights reserved. Keywords: phenanthrene degradation; contaminated steam; R. glutinis; P. aeruginosa
1. Introduction Both bacteria and fungi are plentiful in contaminated soils, and members of both groups contribute to the biodegradation of hydrocarbons. The most abundant fungi in polluted environments are yeasts (Leahy and Colwell, 1990; Berdicevsky et al., 1993); they oxidize polycyclic aromatic hydrocarbon (PAH) with alternative carbon sources (Perry, 1979; Cerniglia, 1992; Sack et al., 1997). Hydrocarbon-degrading bacteria and yeasts have been studied by numerous authors, but in all these studies the potential biodegradation of phenanthrene was determined by co-metabolism and at lower phenanthrene concentrations (Sanseverino et al., 1993; Chaineau et al., 1995). The aims of the present study were to isolate and identify yeast and bacteria strains from a contaminated * Corresponding author. 0269-7491/98/$19.00 # 1998 Elsevier Science Ltd. All rights reserved. PII: S0269 -7 491(98)00056 -6
stream, and to quantify the microbial growth on phenanthrene as a sole source of carbon and energy. 2. Materials and methods 2.1. Isolation of the microorganisms Yeast and bacteria strains were isolated from sediments obtained from a stream heavily contaminated with hydrocarbons, near the YPF-Petroleum Re®nery, La Plata (Argentina). The sediment properties were: pH 6.8; texture: sand 79.9%, lime 9.2%, clay 10.9%; composition: C-organic 4.80%, N-organic 0.15%, P-organic 0.07%. The PAH and phenanthrene concentrations of the studied sediment were 85.0 and 11.0 mg kgÿ1, respectively. The sediment samples were serially diluted and assayed for the presence of phenanthrene-degrading microorganisms by the modi®ed method of Kiyohara et
356
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
al. (1982). The strains were puri®ed by continuously streaking the colonies onto a phenanthrene±agar-plate (as the sole carbon source). Mineral basal salts (MBS) containing (per liter distilled water)/1000 mg (NH4)2SO4, 800 mg K2HPO4, 200 mg KH2PO4, 200 mg MgSO4.7H2O, 100 mg CaCl.2H2O, 5 mg FeSO4.7H2O, and 1.5% agar, were autoclaved. One milliliter of vitamin solution, sterilized by ®ltration, was added to the medium. The pH was adjusted to 6.8±7.0, closer to natural conditions. The surfaces of the sterile agar-plates were coated with 125 ml of ®ltered (0.2-mm pore size) phenanthrene in ethanolic solution (22.5 g literÿ1). To volatilize the solvent the plates were dried for 40 min at 35 C with UV exposure. The cultures were carried out without antibiotic so as not to disturb the microbial response. Sediment dilutions were spread with sterile glass rods onto the agar surface of the Petri dishes and incubated at 251 C for 30 days; but frequently colonies appeared before the ®fth day. Microbial strains were routinely repicked in the presence of phenanthrene to preserve its ability to degrade PAHs. 2.2. Identi®cation and characterization The prevailing yeast strain was identi®ed by colony appearance, cell morphology and physiological dierences, and by carbohydrate assimilation±fermentation tests, and grown on nitrate as the sole nitrogen source (Pha et al., 1978). Identi®cation was con®rmed by the use of rapid identi®cation methods; API Yeast-Ident and API 20C (Analytab Products, 1992). The dominant bacteria strain was Gram stained by the classical staining method (Murray et al., 1994) and the KOH test (Gregersen, 1978). An oxidase test (Drislide Oxidase; Difco) and microscopy for shape and size were conducted. The classical tests according to `Bergey's Manual of Determinative Bacteriology' (Palleroni, 1984) were performed as was ¯uorescence in UV (Gould et al., 1985). The API 20EN test kit was used (BioMeÂrieux, Marcy-l'Etoile, France) and the manufacturer's instructions were followed. The examination of the trips was realized after 24 and 48 h; the identi®cation was carried out according to the API 20EN identi®cation manual. 2.3. Biotransformation A preculture of the microorganisms was done in 100 ml of sterile MBS amended with 50 mg phenanthrene literÿ1, and incubated at 251 C for 3 days. Then 10 ml of each preculture was utilized as inoculum, containing 1.5102 cells mlÿ1 of the isolated yeast and 2.0102 cells mlÿ1 of the bacterial strain. Phenanthrene biotransformation was determined for the yeast and bacterium in parallel ¯asks by using 2-liter
Erlenmeyers containing 0.5 liters of sterile liquid±MBS plus phenanthrene in four dierent concentrations: 25, 50, 100 and 200 mg literÿ1. After thorough mixing, the ethanol was evaporated exhaustively at 35 C for 40 min. The ¯asks were incubated at 251 C, mixed twice daily, kept in darkness and an initial pH of 6.8±7.0 was maintained during the experiment. Three replicates were done for each treatment. Uninoculated medium ¯asks and autoclaved controls were done in duplicate for each treatment. The growth of both microorganisms on phenanthrene was quanti®ed by the increase of the microbial density during the experiment (colony forming unit [CFU] mlÿ1), the speci®c growth rate (m=d[ln x/x0]/dt), the doubling time (td=[ln 2]/m) and the biomass. Yeast and bacterial densities were obtained by the spread plate method (APHA, 1992); samples were taken aseptically from the cultures, three dilutions in triplicate for each sample were incubated and the CFUs (CFU mlÿ1) were the average of the counted plates. Sterile agar-plate medium was composed of 1.5% agar and MBS spread with ®ltered phenanthrene ethanolic solution. Mineral±agar-plates without phenanthrene as the carbon source were used as a control, as microorganisms can grow on the nutrient agar. Microbial dry biomass was estimated gravimetrically by centrifugating 10 ml of each culture suspension at 8000 rpm for 10 min. Then the supernatant was discarded and the pellet was dried at 90 C for 24 h and weighed (GuÈnther et al., 1995). 2.4. Analytical procedures All chemicals were of analytical grade, naphthalene and phenanthrene were purchased from Aldrich (Milwaukee, USA). The phenanthrene concentrations of the sediments and experimental samples were analyzed by gas chromatography in triplicate. Solid phase microextraction (SPME) technique was used (SPME ®ber assembly, 100 mm polydimethylsiloxane coating for manual holder, SUPELCO (Bellefonte, USA)). The sampling was performed with 4 ml aqueous solution, stirring magnetically for 20 min at room temperature, and using naphthalene as an internal standard, with a response factor of 0.60. A Perkin-Elmer Autosystem Gas Chromatograph equipped with a ¯ame ionization detector was used. Nitrogen was used as a carrier gas (10 psi). The injection port was glass lined and allowed split/splitless injection. The split relation was 1/20, and the temperature 280 C. The detector was maintained at 300 C. The oven temperature was programmed from 50 (initial time 3 min) to 250 C (®nal time 7 min) at a rate of 15 C minÿ1. For the injection with the micro®ber the splitter was closed for 3 min. A fused-silica capillary column, liquid phase (methyl 5% phenyl silicone) permaphase PVMS/54 (25 mm 0.32 mm ID, 1.0-mm ®lm thickness) was used for all
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
determinations. In these conditions the retention times were 10.620 min for naphthalene and 16.512 min for phenanthrene. The per cent recovery of phenanthrene by this technique was calculated by comparing its peak area with the naphthalene standard, and the technique was sensitive to 25 ng phenanthrene mlÿ1. 3. Results and discussion 3.1. Identi®cation and characterization Visible colonies of yeast and bacteria strains occurred on phenanthrene-plates, after 5 and 3 days at 251 C, respectively; but usually the majority of the positive colonies appeared 7 or 10 days later. Based on the characterization described in Table 1, 63% of the yeasts isolated from phenanthrene-plates were identi®ed as Rhodotorula glutinis, and 55% of the bacteria degrading PAH were con®rmed to be Pseudomonas aeruginosa (Table 2). Both microorganism species appeared in the ®rst plating and every further plating with the same morphology, and they always retained the ability to grow on phenanthrene-plates on subsequent platings. As R. glutinis strains losing hydrocarbon transformation capacity had been reported in the literature, some researches controlled possible changes by the induction of yeast cytochrome P-450 (MacGillivray and Shiaris, 1993). R. glutinis showed irregular spheroid cells (2.3±5.0 mm wide, 4.0±10.0 mm long) (cornmeal agar), and budding (yeast extract broth). Colonies on phenanthrene-coated agar were small (0.5±0.9 mm) and slightly pink pigmented. In contrast, on organic-enriched medium
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(Sabouraud dextrose agar), shiny, strongly pink-yellow pigmented, moist colonies and no pseudomycelium (cornmeal agar) were observed. It is conceivable that populations of stressed microorganisms have diculties in forming adequately sized colonies on phenanthrene± mineral plates. The phenanthrene-degrading bacterium (a P. aeruginosa Gram-negative strain) cells were irregularly rod shaped (0.5±2.0 mm) with pyocyanin and pyoverdin production, and without capsula, endospores and motility (ÿ). The characterization of bacteria using the phenotypic test has been subject to debate; the identi®cation by the classical test, API 20EN, Biolog system and REP-PCR methods for environmental PAHdegrading Pseudomonas biovars had been extensively studied (Haack et al., 1995; Johnsen et al., 1996). The conclusion is that the classical test and API 20EN appeared to give the best performance in identifying these bacterial strains. 3.2. Growth on phenanthrene R. glutinis grew exponentially on phenanthrene; the growth curves obtained indicated that yeast reached highest population densities (7.7106±1.4107 CFU mlÿ1) on the 28th day at 25 and 50 mg literÿ1, respectively. At 100±200 mg literÿ1 the maximum cell numbers were obtained on the 22nd incubation day (1.3107± 9.5106 CFU mlÿ1; Fig. 1). The logarithmic phase of growth of R. glutinis slowed down at densities of about 7.7106±1.4107 CFU mlÿ1 at 25±50 mg phenanthrene literÿ1, respectively; during this phase m was 0.022 hÿ1 and the corresponding td was
Table 1 Culture and biochemical characteristics of the isolated Rhodotorula glutinis strain Growth at 37 C (+); pellicle in broth (ÿ); pseodu-true hyphae (ÿ); ascospores (ÿ); chlamydospores (ÿ); germ tubes (ÿ); capsule, India ink (ÿ); urease (+); KNO utilization (+); phenol oxidase (ÿ) Assimilation of:
glucose (+); glycerol (ÿ); gly-2-keto-d-gluconate (+); l-arabinose (ÿ); d-xylose (ÿ); adonitol (+); xylitol (); starch (+); galactose (ÿ); inositol (ÿ); sorbitol (+); methyl-d-glucoside (+); N-acetyl-d-glucosamine (ÿ); cellobiose (+); lactose (ÿ); l-ramnose (ÿ); maltose (+); sucrose (+); trehalose (+); melezitose (+); citric acid (+); ranose ()
Fermentation of:
glucose (ÿ); maltose (ÿ); sucrose (ÿ); lactose (ÿ); galactose (ÿ); trehalose (ÿ)
Table 2 Biochemical characteristics of the isolated Pseudomonas aeruginosa strain Classical test:
(a) consumption of: fructose (+); lactose (ÿ); galactose (+); maltose (ÿ); mannose (+); mannitol (+); gelatine (+); starch (ÿ); sorbitol (+); l-tryptophan (ÿ); meso-inositol (+); cetrimide (+) (b) oxidose (+); catalase (+); urease (+); dihydrolase (+); d-nase (+); ornitine decarboxilase (ÿ); arginine dihydrolase (+) (c) production of indole (ÿ); acetoine (ÿ); nitrite (from nitrate [ÿ]); nitrogen (from nitrate [ÿ]) (d) ¯uorescence on Gould's S1 (+); levan production (+); blue pigment on potato dextrose agar (+); growth on 6.5% NaCl (ÿ); growth at 5, 10 and 42 C (+); hemolisys (+)
API 20EN:
indol production on tryptophan (ÿ); glucose acid (+); arginine dihydrolase (+); urease (ÿ); esculin hydrolysis (ÿ); gelatin hydrolysis (+); b-galactosidase (ÿ); d-glucose (+); l-arabinose (+); d-mannose (+); N-acetyl-d-glucosamine (+); maltose (+); gluconate (+); caprate (+); adipate (ÿ); l-malate (+); citrate (+); phenylacetate (ÿ)
358
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
Fig. 1. Growth curves of Rhodotorula glutinis (&) and Pseudomonas aeruginosa (&) on dierent phenanthrene concentrations (CFU, colonyforming unit).
31.5 h. After 22 days the stationary phase approached at 5.3106±8.7106 CFU mlÿ1, equivalent to a biomass concentration of 89.8±147.9 mg literÿ1. At 100 mg phenanthrene literÿ1 the culture was in logarithmic growth until the 15th day after inoculation (m=0.028 hÿ1; td=24.8 h). A slightly dierent response was obtained with 200 mg phenanthrene literÿ1, with a short lag time at the beginning of the assay and an exponential phase of growth between 7 and 22 days of culture (m=0.018 hÿ1; td=38.5 h). A 10-fold decrease in cell yeast numbers at the 28th culture day (from 1.3107±9.4106 to 1.7106±1.6105) was observed at 100±200 mg phenanthrene literÿ1, respectively. To con®rm that the isolated yeast was capable of degrading PAHs, gas chromatography was used to monitor the disappearance of phenanthrene, and revealed 0.092, 0.147, 0.518 and 0.944 mg literÿ1 of residual substrate at the end of the experiment for each phenanthrene concentration assayed. The compound did not disappear from the controls. P. aeruginosa cultures showed the highest densities (7.7108 CFU mlÿ1) after 22 days on 50 mg literÿ1; after 28 days 2.1, 7.4 and 3.0 CFU mlÿ1 were obtained on 25, 100 and 200 mg literÿ1, respectively, and the maxima were 100-fold higher than the ones obtained for the yeast culture. The m-values were higher (0.026, 0.028, 0.041 and 0.037 hÿ1) and td were shorter (26.7, 24.2, 16.9 and 18.9 h) than the values calculated for
R. glutinis. Other authors had reported m-values for growth on phenanthrene uptake even greater than ours (0.058 hÿ1, Weissenfels et al., 1990; 0.069 hÿ1, Boldrin et al., 1993). The gas chromatographic analysis revealed 0.039, 0.083, 0.198 and 0.456 mg literÿ1 of residual phenanthrene at the end of each experiment with P. aeruginosa. Considering the disparity of cell size, calculation on a biomass rather than a per cell basis seems to be the more correct means of comparison for environmental signi®cance. For the isolated microorganisms, 108 cells equated 0.0065 mg (dry wt) of P. aeruginosa strain and 1.70 mg of R. glutinis strain; so the highest biomasses obtained were 238.0 and 50.0 mg literÿ1 for the yeast and bacteria species, respectively. Almost complete phenanthrene transformation was observed in this experiment during the 1-month incubation period, for both microorganisms. However, previous reports showed that 50.9% (Heitkamp and Cerniglia, 1988) and 71% (Heitkamp and Cerniglia, 1989) of this PAH was mineralized by bacteria in the presence of other organic sources; yeasts were only able to co-metabolize phenanthrene at lower rates (MacGillivray and Shiaris, 1994). Partial oxidation of hydrocarbons depending on another C source was found by Ahearn et al. (1971) and Cerniglia and Crow (1981). The lag time preceeding phenanthrene utilization, observed by other authors (Knaebel et al., 1994), was
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
detected in both yeast and bacterial cultures only at 200 mg phenanthrene literÿ1, and once the assimilation was initiated it was similar in all ¯asks. The yeast and bacteria strains were adapted to phenanthrene as the concentration in the sampled sediments was 11 mg kgÿ1, showing less inhibition than those strains which had not undergone pre-exposure. A large number of bacteria from this group, capable of degrading PAH, have been isolated but few authors had reported the growth of yeast with a similar biotransformation potential for phenanthrene. In the present study R. glutinis was able to grow on 200 mg phenanthrene literÿ1, with a short lag time at the beginning of the assay, without co-substrate, with no inhibition response and with similar biomass production; this is an outstanding result and contrasts with other ®ndings. Acknowledgements The authors thank Bact. Mercedes Gatti and Med. Vet. Estela Bruno for the microbiological characterization. This work was supported by grants from the National Council of Scienti®c and Technological Research-CONICET and from CIC (Prov. Bs.As), Argentina. References Ahearn, D.G., Meyers, S.P., Standard, P.G., 1971. The role of yeast in the decomposition of oils in marine environments. Development Industrial Microbiology 12, 126±134. Analytab Products, 1992. API 20EN Analytical Pro®le Index, 5th Edition. `Anal. Proc. 1992'. Analytab Products, Marcy-l'Etoile, France. APHA, 1992. Standard Methods for the Examination of Water and Wastewater, 18th Edition. American Public Health Association, Washington DC. Berdicevsky, I., Duek, L., Merzbach, D., Yannai, S., 1993. Susceptibility of dierent yeast species to environmental toxic metals. Environmental Pollution 80, 41±44. Boldrin, B., Tiehm, A., Fritzsche, C., 1993. Degradation of phenanthrene, ¯uorene, ¯uoranthene and pyrene by a Mycobacterium sp. Applied Environmental Microbiology 59, 1927±1930. Cerniglia, C.E., 1992. Biodegradation of polycyclic aromatic hydrocarbons. Biodegradation 3, 351±368. Cerniglia, C.E., Crow, S.A., 1981. Metabolism of aromatic hydrocarbons by yeast. Archives of Microbiology 129, 9±13. Chaineau, C.H., Morel, J.L., Oudot, J., 1995. Microbial degradation in soil microcosms of fuel oil hydrocarbons from drilling cuttings. Environmental Science and Technology 29, 1615±1621. Gould, W.D., Hagedorn, C., Bardinelli, T.R., Zablotowicz, R.M., 1985. New selective media for enumeration and recovery of ¯uorescent pseudomonads from various habitats. Applied Environmental Microbiology 49, 28±32.
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Gregersen, T., 1978. Rapid method for distinction of Gram-negative from Gram-positive bacteria. European Journal of Applied Microbiology 5, 123±127. GuÈnther, K., Schlosser, D., Fritsche, W., 1995. Phenol and cresol metabolism of Bacillus pumilus isolated from contaminated groundwater. Journal of Basic Microbiology 35, 83±92. Haack, S.K., Garchow, H., Klug, M.J., Forney, L.J., 1995. Analysis of factors aecting the accuracy, reproducibility, and interpretation of microbial community carbon source utilization patterns. Applied Environmental Microbiology 61, 1458±1468. Heitkamp, M.A., Cerniglia, C.E., 1988. Mineralization of polycyclic aromatic hydrocarbons by a bacterium isolated from sediment below an oil ®eld. Applied Environmental Microbiology 54, 1612±1614. Heitkamp, M.A., Cerniglia, C.E., 1989. Polycyclic aromatic hydrocarbon degradation by a Mycobacterium sp. in microcosms containing sediment and water from a pristine ecosystem. Applied Environmental Microbiology 55, 1968±1973. Johnsen, K., Andersen, S., Jacobsen, C.S., 1996. Phenotypic and genotypic characterization of phenanthrene-degrading ¯uorescent Pseudomonas biovars. Applied Environmental Microbiology 62, 3818±3825. Kiyohara, H., Nagao, K., Yana, K., 1982. Rapid screen for bacteria degrading water-insoluble, solid hydrocarbons on agar plates. Applied Environmental Microbiology 43, 454±457. Knaebel, D.B., Federle, T.W., McAvoy, D.C., Vestal, J.R., 1994. Eect of mineral and organic soil constituents on microbial mineralization of organic compounds in a natural soil. Applied Environmental Microbiology 60, 4500±4508. Leahy, J.G., Colwell, R.R., 1990. Microbial degradation of hydrocarbons in the environment. Microbiology Review 54, 305±315. McGillivray, A.R., Shiaris, M.P., 1993. Biotransformation of polycyclic aromatic hydrocarbons by yeast isolated from coastal sediments. Applied Environmental Microbiology 59, 1613± 1618. MacGillivray, A.R., Shiaris, M.P., 1994. Relative role of eukaryotic and prokaryotic microorganisms in phenanthrene transformation in coastal sediments. Applied Environmental Microbiology 60, 1154±1159. Murray, R.G., Doetsch, R.N., Robinow, C.F., 1994. Determinative and cytological light microscopy. In: Gerhardt, P., Murray, R.G.E., Wood W.A., Krieg, N.R. (Eds.), Methods for General and Molecular Bacteriology. American Society for Microbiology, Washington DC, pp. 21±41. Palleroni, N.J., 1984. Pseudomonas. In: Tansill, B. (Ed.), Bergey's Manual of Determinative Bacteriology. Williams and Wilkins, Baltimore, pp. 141±199. Perry, J.J., 1979. Microbial cooxidations involving hydrocarbons. Microbiology Review 43, 59±72. Pha, H.J., Miller, M.W., Mrak, E.M., 1978. The Life of Yeast, 2nd Edition. Harvard University Press, Cambridge. Sack, U., Heinze, T.M., Deck, J., Cerniglia, C.E., Martens, R., Zadrazil, F., Fritsche, W., 1997. Comparison of phenanthrene and pyrene degradation by dierent wood-decaying fungi. Applied Environmental Microbiology 63, 3919±3925. Sanseverino, J., Applegeta, B.M., King, J.M.H., Sayler, G.S., 1993. Plasmid-mediated mineralization of naphthalene, phenanthrene and anthracene. Applied Environmental Microbiology 59, 1931± 1937. Weissenfels, W.D., Beyer, M., Klein, J., 1990. Degradation of phenanthrene, ¯uorene, and ¯uoranthene by pure bacterial cultures. Applied Microbiology and Biotechnology 32, 479±484.
Environmental Pollution 101 (1998) 355±359
Phenanthrene degradation by microorganisms isolated from a contaminated stream M.C. Romero*, M.C. Cazau, S. Giorgieri, A.M. Arambarri Instituto Spegazzini, Facultad de Ciencias Naturales, Universidad Nacional de La Plata, calle 53, no 477-1900 La Plata, Argentina Received 22 April 1997; accepted 17 March 1998
Abstract Autochthonous yeast and bacteria strains were isolated from a contaminated stream and studied for their potential to degrade phenanthrene, a three-ring polycyclic aromatic hydrocarbon (PAH), as their only source of carbon and energy. Rhodotorula glutinis and Pseudomonas aeruginosa were the prevailing microorganisms utilizing phenanthrene. Cells of these microorganisms were inoculated in liquid mineral basal medium at 25, 50, 100 and 200 mg phenanthrene literÿ1 in ethanolic solution. To volatilize the ethanol the medium was dried for 40 min at 35 C with UV exposure. Almost complete phenanthrene degradation was observed during the 1-month incubation period. R. glutinis grew exponentially to a maximum of 9.5106 colony-forming units (CFUs) mlÿ1, with a short lag time only at 200 mg literÿ1. The highest density (1.4107 CFU mlÿ1) without a lag time was obtained at 50 mg literÿ1; lower cell numbers were observed at 25 and 100 mg literÿ1. During the logarithmic phase the speci®c growth rate (m) was 0.022 hÿ1 and the corresponding doubling time (td) was 31.5 h (50 mg literÿ1). A reduction to one-tenth of cell yeast numbers was observed at the lowest PAH levels on the 28th incubation day and a decrease of 100-fold was observed at 200 mg literÿ1. Higher m-values (m=d [ln x/x0]/dt), shorter td (td=[ln 2]/m), and a maximum of 7.7±7.4108 CFU mlÿ1 were observed in P. aeruginosa cultures at 50±100 mg literÿ1, respectively, on the 22nd day. At 25 and 200 mg literÿ1 lower bacteria numbers were obtained (2.1±3.0108 CFU mlÿ1) on the 28th day. R. glutinis was as active as P. aeruginosa at growing on phenanthrene; the aromatic hydrocarbon degradation correlated directly to microbial density and biomass increase, the highest biomass reaching 238.0 and 50.0 mg literÿ1 for the yeast and bacteria species, respectively. # 1998 Elsevier Science Ltd. All rights reserved. Keywords: phenanthrene degradation; contaminated steam; R. glutinis; P. aeruginosa
1. Introduction Both bacteria and fungi are plentiful in contaminated soils, and members of both groups contribute to the biodegradation of hydrocarbons. The most abundant fungi in polluted environments are yeasts (Leahy and Colwell, 1990; Berdicevsky et al., 1993); they oxidize polycyclic aromatic hydrocarbon (PAH) with alternative carbon sources (Perry, 1979; Cerniglia, 1992; Sack et al., 1997). Hydrocarbon-degrading bacteria and yeasts have been studied by numerous authors, but in all these studies the potential biodegradation of phenanthrene was determined by co-metabolism and at lower phenanthrene concentrations (Sanseverino et al., 1993; Chaineau et al., 1995). The aims of the present study were to isolate and identify yeast and bacteria strains from a contaminated * Corresponding author. 0269-7491/98/$19.00 # 1998 Elsevier Science Ltd. All rights reserved. PII: S0269 -7 491(98)00056 -6
stream, and to quantify the microbial growth on phenanthrene as a sole source of carbon and energy. 2. Materials and methods 2.1. Isolation of the microorganisms Yeast and bacteria strains were isolated from sediments obtained from a stream heavily contaminated with hydrocarbons, near the YPF-Petroleum Re®nery, La Plata (Argentina). The sediment properties were: pH 6.8; texture: sand 79.9%, lime 9.2%, clay 10.9%; composition: C-organic 4.80%, N-organic 0.15%, P-organic 0.07%. The PAH and phenanthrene concentrations of the studied sediment were 85.0 and 11.0 mg kgÿ1, respectively. The sediment samples were serially diluted and assayed for the presence of phenanthrene-degrading microorganisms by the modi®ed method of Kiyohara et
356
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
al. (1982). The strains were puri®ed by continuously streaking the colonies onto a phenanthrene±agar-plate (as the sole carbon source). Mineral basal salts (MBS) containing (per liter distilled water)/1000 mg (NH4)2SO4, 800 mg K2HPO4, 200 mg KH2PO4, 200 mg MgSO4.7H2O, 100 mg CaCl.2H2O, 5 mg FeSO4.7H2O, and 1.5% agar, were autoclaved. One milliliter of vitamin solution, sterilized by ®ltration, was added to the medium. The pH was adjusted to 6.8±7.0, closer to natural conditions. The surfaces of the sterile agar-plates were coated with 125 ml of ®ltered (0.2-mm pore size) phenanthrene in ethanolic solution (22.5 g literÿ1). To volatilize the solvent the plates were dried for 40 min at 35 C with UV exposure. The cultures were carried out without antibiotic so as not to disturb the microbial response. Sediment dilutions were spread with sterile glass rods onto the agar surface of the Petri dishes and incubated at 251 C for 30 days; but frequently colonies appeared before the ®fth day. Microbial strains were routinely repicked in the presence of phenanthrene to preserve its ability to degrade PAHs. 2.2. Identi®cation and characterization The prevailing yeast strain was identi®ed by colony appearance, cell morphology and physiological dierences, and by carbohydrate assimilation±fermentation tests, and grown on nitrate as the sole nitrogen source (Pha et al., 1978). Identi®cation was con®rmed by the use of rapid identi®cation methods; API Yeast-Ident and API 20C (Analytab Products, 1992). The dominant bacteria strain was Gram stained by the classical staining method (Murray et al., 1994) and the KOH test (Gregersen, 1978). An oxidase test (Drislide Oxidase; Difco) and microscopy for shape and size were conducted. The classical tests according to `Bergey's Manual of Determinative Bacteriology' (Palleroni, 1984) were performed as was ¯uorescence in UV (Gould et al., 1985). The API 20EN test kit was used (BioMeÂrieux, Marcy-l'Etoile, France) and the manufacturer's instructions were followed. The examination of the trips was realized after 24 and 48 h; the identi®cation was carried out according to the API 20EN identi®cation manual. 2.3. Biotransformation A preculture of the microorganisms was done in 100 ml of sterile MBS amended with 50 mg phenanthrene literÿ1, and incubated at 251 C for 3 days. Then 10 ml of each preculture was utilized as inoculum, containing 1.5102 cells mlÿ1 of the isolated yeast and 2.0102 cells mlÿ1 of the bacterial strain. Phenanthrene biotransformation was determined for the yeast and bacterium in parallel ¯asks by using 2-liter
Erlenmeyers containing 0.5 liters of sterile liquid±MBS plus phenanthrene in four dierent concentrations: 25, 50, 100 and 200 mg literÿ1. After thorough mixing, the ethanol was evaporated exhaustively at 35 C for 40 min. The ¯asks were incubated at 251 C, mixed twice daily, kept in darkness and an initial pH of 6.8±7.0 was maintained during the experiment. Three replicates were done for each treatment. Uninoculated medium ¯asks and autoclaved controls were done in duplicate for each treatment. The growth of both microorganisms on phenanthrene was quanti®ed by the increase of the microbial density during the experiment (colony forming unit [CFU] mlÿ1), the speci®c growth rate (m=d[ln x/x0]/dt), the doubling time (td=[ln 2]/m) and the biomass. Yeast and bacterial densities were obtained by the spread plate method (APHA, 1992); samples were taken aseptically from the cultures, three dilutions in triplicate for each sample were incubated and the CFUs (CFU mlÿ1) were the average of the counted plates. Sterile agar-plate medium was composed of 1.5% agar and MBS spread with ®ltered phenanthrene ethanolic solution. Mineral±agar-plates without phenanthrene as the carbon source were used as a control, as microorganisms can grow on the nutrient agar. Microbial dry biomass was estimated gravimetrically by centrifugating 10 ml of each culture suspension at 8000 rpm for 10 min. Then the supernatant was discarded and the pellet was dried at 90 C for 24 h and weighed (GuÈnther et al., 1995). 2.4. Analytical procedures All chemicals were of analytical grade, naphthalene and phenanthrene were purchased from Aldrich (Milwaukee, USA). The phenanthrene concentrations of the sediments and experimental samples were analyzed by gas chromatography in triplicate. Solid phase microextraction (SPME) technique was used (SPME ®ber assembly, 100 mm polydimethylsiloxane coating for manual holder, SUPELCO (Bellefonte, USA)). The sampling was performed with 4 ml aqueous solution, stirring magnetically for 20 min at room temperature, and using naphthalene as an internal standard, with a response factor of 0.60. A Perkin-Elmer Autosystem Gas Chromatograph equipped with a ¯ame ionization detector was used. Nitrogen was used as a carrier gas (10 psi). The injection port was glass lined and allowed split/splitless injection. The split relation was 1/20, and the temperature 280 C. The detector was maintained at 300 C. The oven temperature was programmed from 50 (initial time 3 min) to 250 C (®nal time 7 min) at a rate of 15 C minÿ1. For the injection with the micro®ber the splitter was closed for 3 min. A fused-silica capillary column, liquid phase (methyl 5% phenyl silicone) permaphase PVMS/54 (25 mm 0.32 mm ID, 1.0-mm ®lm thickness) was used for all
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
determinations. In these conditions the retention times were 10.620 min for naphthalene and 16.512 min for phenanthrene. The per cent recovery of phenanthrene by this technique was calculated by comparing its peak area with the naphthalene standard, and the technique was sensitive to 25 ng phenanthrene mlÿ1. 3. Results and discussion 3.1. Identi®cation and characterization Visible colonies of yeast and bacteria strains occurred on phenanthrene-plates, after 5 and 3 days at 251 C, respectively; but usually the majority of the positive colonies appeared 7 or 10 days later. Based on the characterization described in Table 1, 63% of the yeasts isolated from phenanthrene-plates were identi®ed as Rhodotorula glutinis, and 55% of the bacteria degrading PAH were con®rmed to be Pseudomonas aeruginosa (Table 2). Both microorganism species appeared in the ®rst plating and every further plating with the same morphology, and they always retained the ability to grow on phenanthrene-plates on subsequent platings. As R. glutinis strains losing hydrocarbon transformation capacity had been reported in the literature, some researches controlled possible changes by the induction of yeast cytochrome P-450 (MacGillivray and Shiaris, 1993). R. glutinis showed irregular spheroid cells (2.3±5.0 mm wide, 4.0±10.0 mm long) (cornmeal agar), and budding (yeast extract broth). Colonies on phenanthrene-coated agar were small (0.5±0.9 mm) and slightly pink pigmented. In contrast, on organic-enriched medium
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(Sabouraud dextrose agar), shiny, strongly pink-yellow pigmented, moist colonies and no pseudomycelium (cornmeal agar) were observed. It is conceivable that populations of stressed microorganisms have diculties in forming adequately sized colonies on phenanthrene± mineral plates. The phenanthrene-degrading bacterium (a P. aeruginosa Gram-negative strain) cells were irregularly rod shaped (0.5±2.0 mm) with pyocyanin and pyoverdin production, and without capsula, endospores and motility (ÿ). The characterization of bacteria using the phenotypic test has been subject to debate; the identi®cation by the classical test, API 20EN, Biolog system and REP-PCR methods for environmental PAHdegrading Pseudomonas biovars had been extensively studied (Haack et al., 1995; Johnsen et al., 1996). The conclusion is that the classical test and API 20EN appeared to give the best performance in identifying these bacterial strains. 3.2. Growth on phenanthrene R. glutinis grew exponentially on phenanthrene; the growth curves obtained indicated that yeast reached highest population densities (7.7106±1.4107 CFU mlÿ1) on the 28th day at 25 and 50 mg literÿ1, respectively. At 100±200 mg literÿ1 the maximum cell numbers were obtained on the 22nd incubation day (1.3107± 9.5106 CFU mlÿ1; Fig. 1). The logarithmic phase of growth of R. glutinis slowed down at densities of about 7.7106±1.4107 CFU mlÿ1 at 25±50 mg phenanthrene literÿ1, respectively; during this phase m was 0.022 hÿ1 and the corresponding td was
Table 1 Culture and biochemical characteristics of the isolated Rhodotorula glutinis strain Growth at 37 C (+); pellicle in broth (ÿ); pseodu-true hyphae (ÿ); ascospores (ÿ); chlamydospores (ÿ); germ tubes (ÿ); capsule, India ink (ÿ); urease (+); KNO utilization (+); phenol oxidase (ÿ) Assimilation of:
glucose (+); glycerol (ÿ); gly-2-keto-d-gluconate (+); l-arabinose (ÿ); d-xylose (ÿ); adonitol (+); xylitol (); starch (+); galactose (ÿ); inositol (ÿ); sorbitol (+); methyl-d-glucoside (+); N-acetyl-d-glucosamine (ÿ); cellobiose (+); lactose (ÿ); l-ramnose (ÿ); maltose (+); sucrose (+); trehalose (+); melezitose (+); citric acid (+); ranose ()
Fermentation of:
glucose (ÿ); maltose (ÿ); sucrose (ÿ); lactose (ÿ); galactose (ÿ); trehalose (ÿ)
Table 2 Biochemical characteristics of the isolated Pseudomonas aeruginosa strain Classical test:
(a) consumption of: fructose (+); lactose (ÿ); galactose (+); maltose (ÿ); mannose (+); mannitol (+); gelatine (+); starch (ÿ); sorbitol (+); l-tryptophan (ÿ); meso-inositol (+); cetrimide (+) (b) oxidose (+); catalase (+); urease (+); dihydrolase (+); d-nase (+); ornitine decarboxilase (ÿ); arginine dihydrolase (+) (c) production of indole (ÿ); acetoine (ÿ); nitrite (from nitrate [ÿ]); nitrogen (from nitrate [ÿ]) (d) ¯uorescence on Gould's S1 (+); levan production (+); blue pigment on potato dextrose agar (+); growth on 6.5% NaCl (ÿ); growth at 5, 10 and 42 C (+); hemolisys (+)
API 20EN:
indol production on tryptophan (ÿ); glucose acid (+); arginine dihydrolase (+); urease (ÿ); esculin hydrolysis (ÿ); gelatin hydrolysis (+); b-galactosidase (ÿ); d-glucose (+); l-arabinose (+); d-mannose (+); N-acetyl-d-glucosamine (+); maltose (+); gluconate (+); caprate (+); adipate (ÿ); l-malate (+); citrate (+); phenylacetate (ÿ)
358
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
Fig. 1. Growth curves of Rhodotorula glutinis (&) and Pseudomonas aeruginosa (&) on dierent phenanthrene concentrations (CFU, colonyforming unit).
31.5 h. After 22 days the stationary phase approached at 5.3106±8.7106 CFU mlÿ1, equivalent to a biomass concentration of 89.8±147.9 mg literÿ1. At 100 mg phenanthrene literÿ1 the culture was in logarithmic growth until the 15th day after inoculation (m=0.028 hÿ1; td=24.8 h). A slightly dierent response was obtained with 200 mg phenanthrene literÿ1, with a short lag time at the beginning of the assay and an exponential phase of growth between 7 and 22 days of culture (m=0.018 hÿ1; td=38.5 h). A 10-fold decrease in cell yeast numbers at the 28th culture day (from 1.3107±9.4106 to 1.7106±1.6105) was observed at 100±200 mg phenanthrene literÿ1, respectively. To con®rm that the isolated yeast was capable of degrading PAHs, gas chromatography was used to monitor the disappearance of phenanthrene, and revealed 0.092, 0.147, 0.518 and 0.944 mg literÿ1 of residual substrate at the end of the experiment for each phenanthrene concentration assayed. The compound did not disappear from the controls. P. aeruginosa cultures showed the highest densities (7.7108 CFU mlÿ1) after 22 days on 50 mg literÿ1; after 28 days 2.1, 7.4 and 3.0 CFU mlÿ1 were obtained on 25, 100 and 200 mg literÿ1, respectively, and the maxima were 100-fold higher than the ones obtained for the yeast culture. The m-values were higher (0.026, 0.028, 0.041 and 0.037 hÿ1) and td were shorter (26.7, 24.2, 16.9 and 18.9 h) than the values calculated for
R. glutinis. Other authors had reported m-values for growth on phenanthrene uptake even greater than ours (0.058 hÿ1, Weissenfels et al., 1990; 0.069 hÿ1, Boldrin et al., 1993). The gas chromatographic analysis revealed 0.039, 0.083, 0.198 and 0.456 mg literÿ1 of residual phenanthrene at the end of each experiment with P. aeruginosa. Considering the disparity of cell size, calculation on a biomass rather than a per cell basis seems to be the more correct means of comparison for environmental signi®cance. For the isolated microorganisms, 108 cells equated 0.0065 mg (dry wt) of P. aeruginosa strain and 1.70 mg of R. glutinis strain; so the highest biomasses obtained were 238.0 and 50.0 mg literÿ1 for the yeast and bacteria species, respectively. Almost complete phenanthrene transformation was observed in this experiment during the 1-month incubation period, for both microorganisms. However, previous reports showed that 50.9% (Heitkamp and Cerniglia, 1988) and 71% (Heitkamp and Cerniglia, 1989) of this PAH was mineralized by bacteria in the presence of other organic sources; yeasts were only able to co-metabolize phenanthrene at lower rates (MacGillivray and Shiaris, 1994). Partial oxidation of hydrocarbons depending on another C source was found by Ahearn et al. (1971) and Cerniglia and Crow (1981). The lag time preceeding phenanthrene utilization, observed by other authors (Knaebel et al., 1994), was
M.C. Romero et al./Environmental Pollution 101 (1998) 355±359
detected in both yeast and bacterial cultures only at 200 mg phenanthrene literÿ1, and once the assimilation was initiated it was similar in all ¯asks. The yeast and bacteria strains were adapted to phenanthrene as the concentration in the sampled sediments was 11 mg kgÿ1, showing less inhibition than those strains which had not undergone pre-exposure. A large number of bacteria from this group, capable of degrading PAH, have been isolated but few authors had reported the growth of yeast with a similar biotransformation potential for phenanthrene. In the present study R. glutinis was able to grow on 200 mg phenanthrene literÿ1, with a short lag time at the beginning of the assay, without co-substrate, with no inhibition response and with similar biomass production; this is an outstanding result and contrasts with other ®ndings. Acknowledgements The authors thank Bact. Mercedes Gatti and Med. Vet. Estela Bruno for the microbiological characterization. This work was supported by grants from the National Council of Scienti®c and Technological Research-CONICET and from CIC (Prov. Bs.As), Argentina. References Ahearn, D.G., Meyers, S.P., Standard, P.G., 1971. The role of yeast in the decomposition of oils in marine environments. Development Industrial Microbiology 12, 126±134. Analytab Products, 1992. API 20EN Analytical Pro®le Index, 5th Edition. `Anal. Proc. 1992'. Analytab Products, Marcy-l'Etoile, France. APHA, 1992. Standard Methods for the Examination of Water and Wastewater, 18th Edition. American Public Health Association, Washington DC. Berdicevsky, I., Duek, L., Merzbach, D., Yannai, S., 1993. Susceptibility of dierent yeast species to environmental toxic metals. Environmental Pollution 80, 41±44. Boldrin, B., Tiehm, A., Fritzsche, C., 1993. Degradation of phenanthrene, ¯uorene, ¯uoranthene and pyrene by a Mycobacterium sp. Applied Environmental Microbiology 59, 1927±1930. Cerniglia, C.E., 1992. Biodegradation of polycyclic aromatic hydrocarbons. Biodegradation 3, 351±368. Cerniglia, C.E., Crow, S.A., 1981. Metabolism of aromatic hydrocarbons by yeast. Archives of Microbiology 129, 9±13. Chaineau, C.H., Morel, J.L., Oudot, J., 1995. Microbial degradation in soil microcosms of fuel oil hydrocarbons from drilling cuttings. Environmental Science and Technology 29, 1615±1621. Gould, W.D., Hagedorn, C., Bardinelli, T.R., Zablotowicz, R.M., 1985. New selective media for enumeration and recovery of ¯uorescent pseudomonads from various habitats. Applied Environmental Microbiology 49, 28±32.
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