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Degradation of hydrocarbons and alcohols at different temperatures and salinities by Rhodococcus erythropolis DCL14
FEMS Microbiology Ecology 51 (2005) 389–399 www.fems-microbiology.org
Degradation of hydrocarbons and alcohols at different temperatures and salinities by Rhodococcus erythropolis DCL14 Carla C.C.R. de Carvalho *, M. Manuela R. da Fonseca Centro de Engenharia Biolo´gica e Quı´mica, Instituto Superior Te´cnico, Av. Rovisco Pais, 1049-001 Lisboa, Portugal Received 24 March 2004; received in revised form 4 June 2004; accepted 30 September 2004 First published online 27 October 2004
Abstract Rhodococcus erythropolis DCL14 cells were able to metabolise C5–C16 hydrocarbons and C1–C12 alcohols as sole carbon and energy sources, both at 15 and 28 °C. Metabolic activity was also observed at 1.00%, 1.95% and 2.50% sodium chloride. Almost complete degradation of n-, iso- and cyclo-alkanes and aromatic compounds present in fuel oil was achieved after 9 months, 60% being consumed in the first three months. The results from the conditions tested here suggest that this type of bacterium could be involved in bioremediation processes in marine environments such as the Atlantic, Pacific and Indian Ocean. Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords: Bioremediation; Motor oil; Degradation; Fuel oil; Saline conditions
1. Introduction Bioremediation of sites contaminated with spillage of motor oils and fuels is needed where at the same time temperature and salinity may exclude the activity of certain microbes. Temperature has a large impact on both the degradation rates and the toxicity of the hydrocarbons present in the contaminants [12]. Tolerance mechanisms to the hydrocarbons may involve altered compositions of the cytoplasm and outer membrane of microorganisms [20], as well as efflux pumps to remove the solvent from the cells [14]. For instance, Rhodococcus sp. strain Q15 modulates its membrane fluidity in response to low temperature and hydrocarbon toxicity [25]. The prerequisite access to the hydrocarbons is affected by membrane modifications in the lipid composition altering cell surface hydrophobicity [1,25], and by *
Corresponding author. Tel.: +351 21 8417681; fax: +351 21 8419062. E-mail address: [email protected] (C.C.C.R. de Carvalho).
production of extracellular products such as polysaccharides or surfactants [22]. In this study, the strain Rhodococcus erythropolis DCL14, which was isolated from a ditch sediment sample in The Netherlands and has been shown to biodegrade terpenes in organic–aqueous systems [6,7] is used. The rough variant used does not produce extracellular polysaccharides or surfactants, and thus its interactions with the hydrocarbons and the alcohols would be most influenced by membrane properties. The cells are adapted to solvent tolerance, displaying high cell hydrophobicity as shown by migration towards the organic phase in n-dodecane–aqueous systems [10] and cell aggregation in stressful conditions [9,11]. R. erythropolis DCL14 cells also consumed some of the compounds used as organic phase in biotransformation systems [7,8]. Other hydrocarbon-degrading R. erythropolis strains with high hydrophobicity and solvent tolerance were isolated from deep sea [15] and coastal [17] sediments, showing salt tolerance. Consequently, to determine if strain DCL14 would be competent under such natural
0168-6496/$22.00 Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.femsec.2004.09.010
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conditions, one aim of this study was to assess the ability of strain DCL14 to degrade hydrocarbons and alcohols at 15 and 28 °C and in saline conditions. The ability of this strain to degrade both C5–C16 hydrocarbons and C1–C12 alcohols was compared to determine which degradation pathways are present. Two- and four-stroke engine oils and a low sulphur fuel oil were used to assess the potential of this strain to degrade contaminant mixtures that resemble motor oil spills in the environment.
2. Materials and methods 2.1. Microorganism Rhodococcus erythropolis DCL14 was delivered by the Division of Industrial Microbiology of the Wageningen Agricultural University, Wageningen, The Netherlands. The strain was isolated from a sediment sample from a ditch in Reeuwijk, The Netherlands. 2.2. Chemicals The organic solvents used as carbon sources were ethanol (99.8%), butanol (>99.5%), propanol (>99.5%), n-dodecanol, cyclohexane (>99.5%) and toluene (>99.5%) from Merck; n-octane (>99%) from Merck– Schuchardt; methanol (>99.8%), pentanol (>99%), n-hexane (>99%) and iso-octane (>99.5%) from Riedel-de Hae¨n (Seelze, Germany); n-undecane (99%), ntetradecane (99%) and n-hexadecane (99%) from Sigma; cyclohexanol (99%) and n-dodecane (>99%) from Aldrich; pentane (99%) from Fluka; n-heptane (95%) from Lab-Scan and n-nonane (99%) from Acros. The four stroke motor oil tested was Shell Helix Standard 20W50. The two stoke motor oil used was Mobil Super 2T. Fuel oil was from Petrogal, SA (Portugal). 2.3. Growth Cells were grown in cylindrical 100-ml flasks closed with rubber bungs and containing 20 ml of medium (minimal salts medium, pH 7.0) [26], incubated at 15 or 28 °C and 150 rpm on a rotary shaker with an amplitude of 12.5 mm. The organic compounds used as sole carbon sources were present at an initial concentration of 0.125% or 0.25% (v/v). The extent of growth was monitored by measurement of the optical density (OD) at 600 nm and the dry weight, and, in addition, by microscopy and image analysis techniques. The initial OD at 600 nm in the assays carried out at 15 °C was 0.90, which corresponded to a dry weight of 0.30 g and an average of 311 cells per image, while in the experiments at 28 °C the initial OD was 1.24, the dry weight was 0.40 g and an average of 424 cells was observed per image. The percentages of motor oil tested were 0.13%,
0.25%, 0.50%, 1.00% and 2.00% (v/v). Assays were carried out at least in duplicate. 2.4. Carbon source consumption rate The cells were grown in 10-ml test tubes closed with rubber bungs wrapped in aluminium foil, containing 2 ml of minimal salts medium and 0.125% (v/v) of carbon source. The initial OD was 0.12, the dry weight being 0.04 g. The internal diameter of the test tubes was 12.5 mm and the depression caused on the liquid centre by the round bottom of the tubes was 2 mm. The test tubes were incubated at 28 °C and 200 rpm. During growth, at least five samples were taken at different times in order to follow the carbon source consumption. Each sample corresponded to three test tubes: one for biomass measurement and two for determination of the carbon source present. The content of these latter two tubes was extracted with 0.5 ml of ethyl acetate and, after phase separation, the ethyl acetate layer was analysed by gas chromatography (GC). Blank assays, carried out without cells, were also analysed. 2.5. GC analysis The samples were analysed by gas chromatography on a Hewlett–Packard 5890 gas chromatograph with a FID detector, connected to a HP3394 integrator. The capillary column was a SGE HT5, 25 m in length and with internal and external diameters of 0.22 and 0.33 mm, respectively. The oven temperature was 120 °C and that of the injector corresponded to 200 °C. The detector was set at 250 °C. 2.6. Fuel oil consumption Cells were grown in 100-ml flasks, closed with rubber bungs, containing 20 ml of minimal salts medium, R. erythropolis cells and 20, 40, 80, 160 or 320 ll of fuel oil adsorbed on the surface of a plastic tip. Two batches of flasks were used: on one batch, the initial OD600 was 0.78 (0.26 g dry weight); on the other, the initial OD600 was 0.39 ( 0.13 g dry weight). Fuel oil degradation was followed by extraction of the whole volume of 2 flasks with 2 ml of ethyl acetate and by extraction of 1 ml of the aqueous phase of 2 other flasks with 200 ll of ethyl acetate. The ethyl acetate layer was analysed by GC with the following oven temperature program: 50 °C in the first 4 min, followed by an increase of 10 °C min 1 up to 150 and 150 °C in the last 10 min. Several samples were taken in duplicate over a 10-month period, including blank assays without cells. 2.7. Cell hydrophobicity test Cell hydrophobicity tests were carried out according to the ‘‘microbial adhesion to hydrocarbon’’ (MATH)
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test, described by Rosenberg and co-workers [21] as follows: various volumes of n-hexadecane were added in test tubes to 1.2 ml of washed cells (collected at the beginning of the stationary phase) suspended in phosphate buffer (pH 7). After 10 min pre-incubation at 30 °C, the mixtures were agitated at full speed for 120 s on a vortex (Heidolph REAX 2000, Germany). After 15 min, the organic phase was removed and the absorbance of the aqueous phase was measured at 600 nm. Cell hydrophobicity refers to the percentage of cells that migrates towards the organic phase.
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peated injections, and are quoted for a confidence interval of 95%. Biomass concentration measurements (OD) had an associated error of ±8% based on the standard deviation and sample mean of eight repeated samples, quoted for a confidence interval of 95%. The error associated with the image analysis was ±7% based on the standard deviation and sample mean of twelve repeated images taken from the same sample, quoted for a confidence interval of 95%.
3. Results 2.8. Microscopy and image analysis 3.1. Growth on hydrocarbons at 15 and 20 °C The number of cells per image and the cell viability were measured by fluorescence microscopy, using a LIVE/DEADÒ Bac Lightä Bacterial Viability Kit from Molecular Probes. The microscope was an Olympus CX40, with an Olympus U-RFL-T burner and an UMWB mirror cube unit (excitation filter: BP450-480; barrier filter:BA515). Cell aggregates and cell positioning at the organic-aqueous interface were also observed under brightfield transmitted light. Images were captured using a COHU RGB camera. The acquisition software was Matrox Inspector 2.1. Image analysis was carried out using Visilog 5 for Windows 95 from Noesis SA, Les Ulis, France. Fluorescence microscopy allied with image analysis allowed the rapid quantification of cell growth, positioning and viability in the presence of the tested solvents. 2.9. Error analysis The error associated with the GC quantification of samples was ±6%. The errors were calculated based on the standard deviation and sample mean of seven re-
When cells of R. erythropolis DCL14 were grown on different organic solvents as the sole carbon source, growth was observed 60 h after inoculation on all substrates tested except with 0.25% toluene (data not shown). Clustering of cells was visible with the naked eye from 96 h onwards with n-dodecane and after 120 h with n-tetradecane and n-hexadecane as carbon sources (data not shown). Cell clustering was also considerable when the cells were metabolising the other carbon sources, with the exception of 0.125% toluene and iso-octane, in the presence of which no aggregation was visible. Observation of samples, taken from the different media using fluorescence microscopy, showed that the majority of R. erythropolis cells were either at the surface or inside the solvent droplets (image not shown). Growth was faster at 28 °C than at 15 °C for all the carbon sources tested, the highest growth rates being around 0.1 h 1 with n-dodecane, n-tetradecane and nhexadecane (Fig. 1). At 28 °C, toluene, n-octane and iso-octane appeared to be toxic at concentrations of
Fig. 1. Growth rates of Rhodococcus erythropolis DCL14 during the consumption of hydrocarbons as sole carbon and energy sources.
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0.25%, the growth rate decreasing more than 50% in comparison with that observed at concentrations of 0.125%. At 15 °C, cell growth appeared to be generally higher with 0.25% than with 0.125%, probably due to a lower solubility of the tested solvents in the aqueous phase. Overall, R. erythropolis DCL14 exhibited a nearly 10fold lower growth rate on hydrocarbons with an odd number of carbons than in those with an even number (Fig. 1), and its lag phase was longer on toluene than on the other carbon sources tested. According to the MATH test, the cells hydrophobicity was higher than 80% when growing on cyclohexane, n-hexane, n-octane and iso-octane, at both 15 and 28 °C (Fig. 2). Cells grown on toluene showed values of 54.9% and 92.1% at 15 and 28 °C, respectively. In general, the cell membrane hydrophobicity increased with increasing temperature, except for cells grown on n-hexadecane, which showed a hydrophobicity of 92.1% and 73.7% at 15 and 28 °C, respectively. Fig. 2(I) shows the cell hydrophobicity as a function of the log P of the carbon source, which is a measure of the hydrophobicity and toxicity of the solvent. According to Laane et al. [16], a log P of 4 is the minimum value for an organic solvent to be considered biocompatible. Observation of the results indicates that the data can be grouped in three clusters, corresponding to cells grown on carbon sources with an even (A and B) or odd (C) number of carbon atoms, and with a log P value lower (a) or higher (b) than 6. Cells grown on odd-numbered carbon sources showed lower hydrophobicity than cells grown on carbon sources with an even number of carbons. Considering the same type of carbon source (same cluster in Fig. 2), the cell surface hydrophobicity of strain DCL14 increased with increasing log P values, i.e., with increasing hydrophobicity of the solvent and with an increasing number of carbon atoms on the hydrocarbon chain. At 28 °C, solvents with an even
number of carbon atoms and with log P values higher than 6 allowed higher growth rates than those with log P values lower than 6, but with the latter, cell hydrophobicity was higher (Fig. 2, II). Cells grown on carbon sources with an odd number of carbons grew at almost the same rate, but cell hydrophobicity varied between 49.1 and 62.6%. 3.2. Growth on alcohols Considering growth of R. erythropolis DCL14 cells on several alcohols, the results from the growth experiments indicated that the cells were able to grow on C1–C12 alcohols as sole carbon and energy source at both 28 °C (Fig. 3) and 15 °C, although at less than half the rate when growing at 15 °C as compared to 28 °C (data not shown). No growth was observed on either of the n-octanol concentrations tested. At 15 °C, the growth rates attained with 0.125% of methanol and ethanol were 0.05 and 0.04 h 1, respectively. At 28 °C and with an initial substrate concentration of 0.125%, the growth rate decreased with increasing number of carbon atoms in the carbon source (Fig. 3). Also, with increasing number of carbon atoms, the growth rates also showed less variance up to concentrations of 1% of each carbon source. The cells were able to grow at concentration up to 1% cyclohexanol, 2% butanol and pentanol, 5% dodecanol, 15% methanol and 20% ethanol. These concentrations were achieved without any strategy to adapt the cells to high concentrations. The growth rates for 15% methanol and ethanol were 0.01 and 0.02 h 1, respectively, and for 20% ethanol it was 0.01 h 1. It was possible to achieve a linear relation between: (i) the highest concentration of the tested alcohols at which the cells were still able to grow completely (comprising lag, exponential and stationary phases) within 30 h and (ii) the difference in the growth rate observed between the highest and the lowest substrate concentration
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Fig. 2. Hydrophobicity of R. erythropolis DCL14 cells as a function of the log P of the carbon source (I) and of the growth rate (II). Carbon sources with: (A) even number of carbon atoms, log P value lower than 6.0; (B) even number of carbons, log P value higher than 6.0; (C) odd number of carbon atoms.
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Carbon source concentration Fig. 3. Growth rates of R. erythropolis DCL14 attained at 28 °C, with several alcohols present at different initial concentrations.
showing a correlation of 0.99 (Fig. 4). Higher differences in growth rates were achieved obviously for the highest concentrations. These results show that there is a relation between the maximum growth rate (which will determine the differences shown in Fig. 4) and the highest substrate concentration at which the cells can still grow. R. erythropolis DCL14 cells were found to be rather hydrophilic when grown on alcohols: only 10% of the cells grown on methanol, around 40% of those grown on ethanol and nearly 60% of cells grown on n-dodecanol migrated towards the n-hexadecane phase during the MATH test. The other alcohols tested as carbon sources produced cells with intermediate values, cell surface hydrophobicity increasing with the number of carbon atoms on the carbon source. Furthermore, no
clusters were observed during growth of cells on these carbon sources, at both 15 and 28 °C, except for the highest concentrations of methanol and ethanol used. The specific carbon source consumption rate was higher for methanol and ethanol than for the hydrocarbons tested (Fig. 5). Ethanol was consumed 30% faster than methanol and its consumption rate was double the average consumption rate of the hydrocarbons with an even number of carbons. 3.3. Growth under saline conditions at 15 °C Since the natural sites at which bioremediation has to take place may present saline conditions to the microorganisms, growth of R. erythropolis DCL14 cells was tested in the presence of different salt concentrations.
Difference in growth rate
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Carbon source consumption (g/h.g d.w. )
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Fig. 5. Carbon source consumption rates attained during growth of R. erythropolis DCL14 cells on alcohols and hydrocarbons as single carbon and energy source.
The lag phase of the cells increased with increasing concentrations of salt: around 50, 100 and 200 h (150 h for ethanol) at an initial NaCl concentration of 1.00%, 1.95% and 2.50%, respectively. At 1.00% NaCl, the highest growth rates (around 0.04 h 1) were observed when the cells were using ethanol, n-hexane and iso-octane as sole carbon sources, whereas the growth rate was almost half for cells growing on methanol and n-octane (Fig. 6). For an initial NaCl concentration of 1.95%, the growth rates attained with all the carbon sources tested only ranged between 0.017 h 1 for cells grown on methanol and 0.026 h 1 for cells grown on iso-octane. Growth rates decreased to nearly 0.014 h 1 at a salt concentration of 2.5% when the cells used methanol, nhexane and n-octane. However, a growth rate almost
as high as that attained when the salt concentration was 1% was observed with cells metabolising ethanol and iso-octane. In these two carbon sources, the cell hydrophobicity also decreased from around 40% at a salt concentration of 2% to less than 10% at a NaCl concentration of 2.5% (data not shown). Fluorescence microscopy and image analysis showed that the cells in contact with the highest salt concentration had decreased their size to about 30% as compared to cells in medium without additional NaCl. 3.4. Degradation of two- and four-stroke motor oils When the cells were incubated at 28 °C with two- and four-stroke motor oils, the optical density of the cultures
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Carbon source Fig. 6. Growth rates of R. erythropolis cells, carrying out the metabolism of alcohols and hydrocarbons as sole carbon and energy sources, observed in salted media at 28 °C.
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doubled during the first 20 h at initial oil fractions lower than 0.5% Shell Helix standard 20W-50 and for all Mobil Super 2T concentrations tested. With increasing oil concentrations, however, the growth rate decreased (Fig. 7). Significant cell aggregation was observed after 15 h with Shell Helix oil fractions higher than 0.5% and after 30 h with lower oil fractions. In the presence of Mobil Super 2T, cell clustering was observed after 40 h of cell growth. Thus quantification of growth by measurement of the optical density became impossible, and the results were calculated using the data obtained via fluorescence microscopy and image analysis and also by dry weight measurements. At 15 °C, the growth rate increased with increasing concentrations of Mobil Super 2T up to a fraction of 0.5% and remained almost constant with higher oil fractions at a value of nearly 0.04 h 1 (Fig. 7). When the cells were using Shell Helix, a sharp linear decrease of
the growth rate with increasing oil fractions was observed. At lower oil fractions, cell growth started at a higher rate when the cells were using Shell Helix but at higher fractions, cell growth was faster when cells were using Mobil 2T. Very small clusters were visible after 50 h in all growth experiments with both oils. The higher the oil fraction, the larger the cell clusters were. Images taken with a microscope showed cells gathered around small oil droplets of both oils (data not shown). The droplet diameter ranged between 0.02 and 0.09 mm. The MATH hydrophobicity test showed that, for both oils, cells were less hydrophobic when grown on 0.5% oil fraction (data not shown). The most hydrophobic cells were those grown on either the lowest (0.125%) or the highest (2%) initial oil fraction. Furthermore, cells grown on Mobil Super 2T were more hydrophobic than those consuming Shell Helix, at identical initial fractions.
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Concentration (%, v/v) Fig. 7. Growth rates attained with R. erythropolis DCL14 cells growing on motor oils at 15 and 28 °C.
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Time (month) Fig. 8. Percentage of fuel oil remaining at different times during its biodegradation by R. erythropolis cells.
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In the presence of 2.5% NaCl, the growth curves followed similar trends to those shown in Fig. 7, but the growth rates were half of those achieved in media without salt (data not shown). 3.5. Degradation of fuel oil Fuel oil is composed of saturated hydrocarbons, aromatics, resins and asphaltenes. GC analysis only allowed the detection of alkanes and aromatics in the fuel oil used. The asphaltenes and resins were probably retained at the injector. The first compounds to be degraded were n-hexane, cyclohexane and iso-octane, followed by ndodecane. Almost complete degradation of the n-, isoand cyclo-alkanes, as well as aromatics present in fuel oil, was achieved by R. erythropolis DCL14 after 9 months of biodegradation (Fig. 8). After 3 months, more than 60% of the hydrocarbons had already been consumed. The percentage of oil remaining decreased with time according to a second order polynomial curve with a correlation of 0.99.
4. Discussion Nowadays, all kinds of motors are widely present in our environment and accidental oil spills are frequent. The bioremediation of sites contaminated with hydrocarbons, resulting from e.g. spillage of motor oils, diesel or jet fuels, may have to be carried out in the natural environment and thus at temperatures considerably below the optimum and/or under saline conditions. Microorganisms that present the ability to degrade these contaminants under different environmental conditions can represent a useful tool for the remediation of contaminated soil or for dedicated effluent treatment. Bacteria and fungi initiate the aerobic metabolism of alkanes using mono-oxygenases, which by a radical mechanism introduce a hydroxyl group in the alkane molecule through a partial reduction of O2. The resulting alkanol is further oxidised and metabolised in the b-oxidation pathway. In actinomycetes (including Rhodococcus, Mycobacterium and Nocardia), the b-oxidation pathway can be both a catabolic pathway and a source of fatty acids for the production of a large variety of lipids needed for survival and growth under adverse conditions [1]. Comparing the results obtained in this study for the degradation of the alkane with those of the corresponding alcohol, it is observed that on pentanol R. erythropolis DCL14 cells grew at a rate 5-fold higher than on n-pentane, whilst on cyclohexanol and dodecanol the DCL14 cells grew at a third of the rate obtained with cyclohexane and dodecane, respectively. No growth was observed on n-octanol, even at an initial concentration of 0.125%. These results indicate that during pentane metabolism in the tested strain, the forma-
tion of pentanol from pentane must be the limiting step. During the consumption of other hydrocarbons, the degradation of the corresponding alcohol, which should be more toxic according to the log P scale, is the limiting step since rather low concentrations of these compounds may damage the cells. However, at 28 °C, the cells were able to grow up to concentrations of 1% cyclohexanol, 2% butanol and pentanol, 5% dodecanol, 15% methanol and 20% ethanol. Growth on these high concentrations was achieved without any strategy to adapt the cells to increasing concentrations of either carbon source. Studies carried out by Solano-Serena et al. [22] showed that growth on cyclohexane and iso-octane may be difficult: most of the populations able to grow on solid medium using cyclohexane as substrate could not grow in liquid medium on this substrate; the only strain that was able to grow on iso-octane solely was isolated from a gasoline-polluted sample. Degradation of cyclohexane was achieved by co-metabolism [2] and by a mixed microbial population, which by several unknown interactions, was able to perform an accordant attack on the substrate [18]. R. erythropolis DCL14 was able to degrade these two recalcitrant compounds and to grow in medium with only these substrates as carbon and energy sources, showing the great potential of this strain for bioremediation. Furthermore, toluene, another toxic compound, was degraded at an initial concentration of 0.125%, although the lag phase was longer than with the other carbon sources. In a previous study, de Carvalho et al. [11] observed that when R. erythropolis DCL14 cells were incubated with toluene, non-viable cells were twice as large as the viable ones, indicating that toluene should increase the fluidity of the cellular membrane. After incubation with 2.5% toluene, the colonies growing on agar plates were yellow, i.e., the cells that survived produced phenotypically different colonies. Sokolovska´ et al. [23] found a clear correlation between the carbon source and the mycolic acid profiles of R. erythropolis E1: there was a lack of odd-numbered carbon chains in cells growing on linear alkanes with an even number of carbon atoms; mycolic acids were present with both even and odd-numbered carbon chains in cells growing on compounds with an odd number of carbon atoms, branched alkanes or mixtures of these compounds. These results, and the available literature, led the authors to conclude that there are two different pathways for the synthesis of mycolic acids in this strain: one by addition of acetyl coenzyme A and the other from addition of propionyl coenzyme A. According to Alvarez [1], during cultivation of the strain R. erythropolis 17 on n-alkanes, the cells produced fatty acids with a chain length associated with the chain length of the growth substrate, as well as with other fatty acids derived from the b-oxidation pathway. The strain used
C.C.C.R. de Carvalho, M. Manuela R. da Fonseca / FEMS Microbiology Ecology 51 (2005) 389–399
by this author [1] showed a more efficient mechanism for the production of the intermediate propionyl-CoA, precursor of fatty acids containing an odd-number of carbon atoms. However, in our case, the results indicate that R. erythropolis DCL14 cells have a less efficient mechanism for producing the precursor propionylCoA than other strains of the Rhodococcus genus, as the growth rates obtained with carbon sources with an odd number of carbon atoms were 10-fold lower than those obtained with an even number of carbons. The absence of a clear and linear relation between the hydrophobicity of the carbon source used and the observed cell hydrophobicity, suggests that the latter is the result of modifications in the membrane composition and not of direct incorporation of solvent molecules at the cellular membrane. Since a rough variant was chosen, i.e., the strain did not produce extracellular polymeric substances, modifications at the cell surface hydrophobicity by changes of the lipid composition of the membrane were probably the only available mechanism for protection from the toxic effects of the solvents. The maximum growth rate attained with a certain carbon source tested was related to the highest substrate concentration at which R. erythropolis DCL14 cells could still grow. Thus, the results clearly indicate that there is a relation between the DCL14 ability to degrade a certain compound and the cellÕs resistance to that compound, which may be crucial for the degradation of toxic substances. Contrarily, the results presented by Mosqueda et al. [19] suggested that toluene metabolism in Pseudomonas putida DOT-T1 is not involved in toluene tolerance. By studying samples taken from the different media using fluorescence microscopy, it was possible to observe cells both at the surface and inside the solvent droplets, suggesting that the uptake of substrate was achieved by direct contact between the cells and the organic solvent acting as carbon source (data not shown). Bouchez-Naı¨tali et al. [5] observed a direct interfacial uptake of n-hexadecane by four R. equi strains, which did not produce biosurfactants. The growth rates obtained were independent of the interfacial area, which was expected due to the very strong adsorption of the bacterial cells at the solvent-aqueous interface. Formation of cellular flocs was also observed due to the hydrophobicity of the strains tested. In the present study, the differences between the growth rates attained with 0.125% and 0.25% of n-dodecane, n-tetradecane and nhexadecane were between 1.5 and 7% (for n-tetradecane and n-dodecane, respectively). Since at the concentrations tested these hydrocarbons are immiscible, the interfacial area available for an initial concentration of 0.25% should double that obtained with an initial concentration of 0.125%. Thus, the growth rates obtained were almost independent of the interfacial area available, confirming that the uptake of these hydrocar-
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bons should be by direct contact between cells and substrate. In the case of the miscible methanol and ethanol, the cells were freely suspended in the aqueous phase containing the carbon source, whilst in the case of the remaining immiscible carbon sources, the cells were mainly located at the hydrocarbon–aqueous interface or even at the organic phase. Direct uptake of all carbon sources should therefore be possible, decreasing the impact of mass transfer (of the carbon source to the cell) on the substrate consumption rate. The results showed that methanol and ethanol were consumed at a higher rate than the hydrocarbons tested (Fig. 5). Since, as shown, the cells may modulate their surface hydrophobicity to improve adhesion to the organic phase, the differences observed in carbon source consumption rate were the result of particularities of the metabolism of the several compounds. This is particularly true when comparing the rates obtained for the odd- and evennumbered hydrocarbons. Metabolic engineering or directed mutagenenis could thus improve the R. erythropolis DCL14 bioremediation capabilities. Nevertheless, higher growth rates were attained with the most hydrophobic carbon sources (Fig. 2, II). A comparison between cells grown at 15 and 28 °C showed that at 15 °C the growth rates seemed more independent of the carbon source hydrophobicity, and also that there was a wider variation in the hydrophobic behaviour of the cells. The uptake of the hydrocarbon can thus be quite influenced by the temperature at which the cells proceed their metabolism, as well as by the composition of the cellular membrane. Many of the oil spills occur in ecosystems with high or moderate salinities such as the sea, estuaries, seashores, etc. Seawater contains, in average, 1.95% (w/w) chlorine and 1.08% (w/w) sodium [3]. In the present work, it was shown that strain DCL14 is able to metabolise methanol, ethanol, n-hexane, iso-octane and noctane in the presence of up to 2.5% NaCl at 15 °C. The growth rates decreased with increasing salt concentrations, but when ethanol and iso-octane were used, the growth rates observed at 2.5% NaCl were almost as high as those obtained with 1% NaCl. Apparently, at 2.5% NaCl, the influence on the cell membrane is larger and the resulting higher hydrophobicity of the cellular membrane promotes the consumption of these carbon sources. The specific contact area between the cells and the substrates should be higher at this concentration due to cell shrinkage, a well-known process of cell regulation in hypertonic media. This, allied with a higher hydrophobicity of the cellular membrane, could have improved the transport of ethanol and iso-octane from the growth media to the interior of the cell. In real oil spills in the environment, we often deal with mixtures of hydrocarbons and the presence of a single compound may influence the degradation of the others
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[13]. To assess this, degradation of Shell Helix Standard 20W-50 and Mobil super 2T, four- and two-stroke motor oil respectively, was tested at 15 and 28 °C. For some concentrations (higher than 0.5% Mobil; 0.125%, 0.25% and 2% Shell), the growth rates attained at 15 and 28 °C were quite close. Direct uptake of the compounds present in these oils was achieved by adhesion of the cells to 0.02–0.09 mm oil droplets, suggesting that strain DCL14 could be used for in situ bioremediation. Fuel oil, an even more worrying source of pollution, is composed of saturated hydrocarbons, aromatics, resins and asphaltenes. Microorganisms primarily degrade n- and iso-alkanes, followed by cycloalkanes and 1- and 3-ring aromatics. Finally, polyaromatics, asphaltenes and resins are consumed [4,24]. When R. erythropolis DCL14 cells were tested as degraders of a low sulphur content fuel oil, the first compounds to be degraded were n-hexane, cyclohexane and isooctane, followed by n-dodecane. This is in agreement with the results obtained during the assays with each of these compounds as sole carbon and energy source. This specific bacterium is therefore able to use a large number of recalcitrant compounds, both isolated and as a mixture of compounds. After 9 months of fuel oil metabolism, almost complete degradation of n-, iso- and cyclo-alkanes and aromatic present was achieved. This paper clearly shows the ability of R. erythropolis DCL14 to degrade n-alkanes, cycloalkanes, branched and monoaromatic alkanes and considerably high concentrations of alcohols at both 15 and 28 °C, even under saline conditions. Since the tested conditions are found in the Atlantic, Pacific and Indian oceans and the cells were found capable to degrade both motor and fuel oils, the large potential of strain DCL14 for bioremediation of real oil spills is also demonstrated.
Acknowledgements This study was supported by a Ph.D. Grant (PRAXIS XXI/BD/21574/99) awarded to Carla da C.C.R. de Carvalho by Fundac¸a˜o para a Cieˆncia e a Tecnologia, Portugal. The authors thank J.D.R. de Carvalho for the supply of the motor oils and Helder Tavanez for supplying the fuel oil used in this study.
References [1] Alvarez, H.M. (2003) Relationship between b-oxidation and the hydrocarbon-degrading profile and actinomycetes bacteria. Int. Biodeter. Biodegr. 52, 35–42. [2] Beam, H.W. and Perry, J.J. (1974) Microbial degradation of cycloparaffinic hydrocarbons via co-metabolism and commensalism. J. Gen. Microbiol. 82, 163–169.
[3] Bearman, G. (1989) Ocean Chemistry and Deep-Sea Sediments. Pergamon Press, Sidney. [4] Blackburn, J.W. and Hafker, W.R. (1993) The impact of biochemistry, bioavailability and bioactivity on the selection of bioremediation techniques. Trends Biotechnol. 11, 328– 333. [5] Bouchez-Naı¨tali, M., Blanchet, D., Bardin, V. and Vandecasteele, J.P. (2001) Evidence for interfacial uptake in hexadecane degradation by Rhodococcus equi: the importance of cell floculation. Microbiology 147, 2537–2543. [6] de Carvalho, C.C.C.R., van Keulen, F. and da Fonseca, M.M.R. (2000) Production and recovery of limonene-1,2-diol and simultaneous resolution of a diastereomeric mixture of limonene-1,2epoxide with whole cells of Rhodococcus erythropolis DCL14. Biocatal. Biotransf. 18, 223–235. [7] de Carvalho, C.C.C.R. and da Fonseca, M.M.R. (2002) Maintenance of cell viability in the biotransformation of ( )-carveol with whole cells of Rhodococcus erythropolis. J. Mol. Catal. B: Enz. 19-20C, 389–398. [8] de Carvalho, C.C.C.R. and da Fonseca, M.M.R. (2002) Influence of reactor configuration on the production of carvone from carveol by whole cells of Rhodococcus erythropolis DCL14. J. Mol. Catal. B: Enz. 19–20C, 377–387. [9] de Carvalho, C.C.C.R., Pons, M.N. and da Fonseca, M.M.R. (2003) Principal components analysis as a tool to summarise biotransformation data: influence on cells of solvent type and phase ratio. Biocatal. Biotrans. 21, 305–314. [10] de Carvalho, C.C.C.R. and da Fonseca, M.M.R. (2003) A simple method to observe organic solvent drops with a standard optical microscope. Microsc. Res. Technol. 60, 465–466. [11] de Carvalho, C.C.C.R., Cruz, A., Pons, M.N., Pinheiro, H.M., Cabral, J.M.S., da Fonseca, M.M.R., Fernandes, P., and Ferreira, B.S. Mycobacterium sp., Rhodococcus erythropolis and Pseudomonas putida behaviour in the presence of organic solvents. Microsc. Res. Tech., in press doi: 10.1002/ jemt.20061. [12] Crawford, R.L. and Zhou, E. (1995) Effects of oxygen, nitrogen, and temperature on gasoline biodegradation in soil. Biodegradation 6, 127–140. [13] Eriksson, M., Dalhammar, G. and Borg-Karlson, A.-K. (1999) Aerobic degradation of a hydrocarbon mixture in natural uncontaminated potting soil by indigenous microorganisms at 20 and 6 °C. Appl. Microbiol. Biotechnol. 51, 532–535. [14] Fernandes, P., Ferreira, B.S. and Cabral, J.M.S. (2003) Solvent tolerance in bacteria: role of efflux pumps and cross resistance with antibiotics. Int. J. Antimicrob. Ag. 22, 211–216. [15] Heald, S.C., Branda˜o, P.F.B., Hardicre, R. and Bull, A.T. (2001) Physiology, biochemistry and taxonomy of deep-sea nitrilemetabolising Rhodococcus strains. Antonie Leeuwenhoek 80, 169–183. [16] Laane, C., Boeren, S., Vos, K. and Veeger, C. (1987) Rules for optimization of biocatalysis in organic solvents. Biotechnol. Bioeng. 30, 81–87. [17] Langdahl, B.R., Bisp, P. and Ingoorsen, K. (1996) Nitrile hydrolysis by Rhodococcus erythropolis BL1, an acetonitriletolerant strain isolated from a marine sediment. Microbiology 142, 145–154. [18] Lloyd-Jones, G. and Trudgill, P.W. (1989) The degradation of alicyclic hydrocarbons by a microbial consortium. Int. Biodeterior. 25, 197–206. [19] Mosqueda, G., Ramos-Gonzalez, M. and Ramos, J. (1999) Toluene metabolism by solvent-tolerant Pseudomonas putida DOT-T1 strain and its role in solvent impermeabilization. Gene 232, 69–76. [20] Pieper, D.H. and Reineke, W. (2000) Engineering bacteria for bioremediation. Curr. Opin. Biotechnol. 11, 262–270.
C.C.C.R. de Carvalho, M. Manuela R. da Fonseca / FEMS Microbiology Ecology 51 (2005) 389–399 [21] Rosenberg, M., Gutnick, D. and Rosenberg, E. (1980) Adherence of bacteria to hydrocarbons: a simple method for measuring cellsurface hydrophobicity. FEMS Microbiol. Lett. 9, 29–33. [22] Solano-Serena, F., Marchal, R., Lebeault, J.-M. and Vandecasteele, J.-P. (2000) Selection of microbial populations degrading recalcitrant hydrocarbons of gasoline by monitoring of cultureheadspace composition. Lett. Appl. Microbiol. 30, 19–22. [23] Sokolovska´, I., Rozenberg, R., Riez, C., Rouxhet, P.G., Agathos, S.N. and Wattiau, P. (2003) Carbon source-induced modifications in the mycolic acid content and cell wall permeability of Rhodococcus erythropolis E1. Appl. Environ. Microbiol. 69, 7019–7027.
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[24] Sugiura, K., Ishihara, M., Shimauchi, T. and Harayama, S. (1997) Physicochemical properties and biodegradability of crude oil. Environ. Sci. Technol. 31, 45–51. [25] Whyte, L.G., Slagman, S.J., Pietrantonio, F., Bourbonnie`re, L., Koval, S.F., Lawrence, J.R., Inniss, W.E. and Greer, C.W. (1999) Physiological adaptations involved in alkane assimilation at a low temperature by Rhodococcus sp. strain Q15. Appl. Environ. Microbiol. 65, 2961–2968. [26] Wiegant, W.M. and de Bont, J.A.M. (1980) A new route for ethylene glycol metabolism in Mycobacterium E44. J. General Microb. 120, 325–331.
Degradation of hydrocarbons and alcohols at different temperatures and salinities by Rhodococcus erythropolis DCL14 Carla C.C.R. de Carvalho *, M. Manuela R. da Fonseca Centro de Engenharia Biolo´gica e Quı´mica, Instituto Superior Te´cnico, Av. Rovisco Pais, 1049-001 Lisboa, Portugal Received 24 March 2004; received in revised form 4 June 2004; accepted 30 September 2004 First published online 27 October 2004
Abstract Rhodococcus erythropolis DCL14 cells were able to metabolise C5–C16 hydrocarbons and C1–C12 alcohols as sole carbon and energy sources, both at 15 and 28 °C. Metabolic activity was also observed at 1.00%, 1.95% and 2.50% sodium chloride. Almost complete degradation of n-, iso- and cyclo-alkanes and aromatic compounds present in fuel oil was achieved after 9 months, 60% being consumed in the first three months. The results from the conditions tested here suggest that this type of bacterium could be involved in bioremediation processes in marine environments such as the Atlantic, Pacific and Indian Ocean. Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords: Bioremediation; Motor oil; Degradation; Fuel oil; Saline conditions
1. Introduction Bioremediation of sites contaminated with spillage of motor oils and fuels is needed where at the same time temperature and salinity may exclude the activity of certain microbes. Temperature has a large impact on both the degradation rates and the toxicity of the hydrocarbons present in the contaminants [12]. Tolerance mechanisms to the hydrocarbons may involve altered compositions of the cytoplasm and outer membrane of microorganisms [20], as well as efflux pumps to remove the solvent from the cells [14]. For instance, Rhodococcus sp. strain Q15 modulates its membrane fluidity in response to low temperature and hydrocarbon toxicity [25]. The prerequisite access to the hydrocarbons is affected by membrane modifications in the lipid composition altering cell surface hydrophobicity [1,25], and by *
Corresponding author. Tel.: +351 21 8417681; fax: +351 21 8419062. E-mail address: [email protected] (C.C.C.R. de Carvalho).
production of extracellular products such as polysaccharides or surfactants [22]. In this study, the strain Rhodococcus erythropolis DCL14, which was isolated from a ditch sediment sample in The Netherlands and has been shown to biodegrade terpenes in organic–aqueous systems [6,7] is used. The rough variant used does not produce extracellular polysaccharides or surfactants, and thus its interactions with the hydrocarbons and the alcohols would be most influenced by membrane properties. The cells are adapted to solvent tolerance, displaying high cell hydrophobicity as shown by migration towards the organic phase in n-dodecane–aqueous systems [10] and cell aggregation in stressful conditions [9,11]. R. erythropolis DCL14 cells also consumed some of the compounds used as organic phase in biotransformation systems [7,8]. Other hydrocarbon-degrading R. erythropolis strains with high hydrophobicity and solvent tolerance were isolated from deep sea [15] and coastal [17] sediments, showing salt tolerance. Consequently, to determine if strain DCL14 would be competent under such natural
0168-6496/$22.00 Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.femsec.2004.09.010
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conditions, one aim of this study was to assess the ability of strain DCL14 to degrade hydrocarbons and alcohols at 15 and 28 °C and in saline conditions. The ability of this strain to degrade both C5–C16 hydrocarbons and C1–C12 alcohols was compared to determine which degradation pathways are present. Two- and four-stroke engine oils and a low sulphur fuel oil were used to assess the potential of this strain to degrade contaminant mixtures that resemble motor oil spills in the environment.
2. Materials and methods 2.1. Microorganism Rhodococcus erythropolis DCL14 was delivered by the Division of Industrial Microbiology of the Wageningen Agricultural University, Wageningen, The Netherlands. The strain was isolated from a sediment sample from a ditch in Reeuwijk, The Netherlands. 2.2. Chemicals The organic solvents used as carbon sources were ethanol (99.8%), butanol (>99.5%), propanol (>99.5%), n-dodecanol, cyclohexane (>99.5%) and toluene (>99.5%) from Merck; n-octane (>99%) from Merck– Schuchardt; methanol (>99.8%), pentanol (>99%), n-hexane (>99%) and iso-octane (>99.5%) from Riedel-de Hae¨n (Seelze, Germany); n-undecane (99%), ntetradecane (99%) and n-hexadecane (99%) from Sigma; cyclohexanol (99%) and n-dodecane (>99%) from Aldrich; pentane (99%) from Fluka; n-heptane (95%) from Lab-Scan and n-nonane (99%) from Acros. The four stroke motor oil tested was Shell Helix Standard 20W50. The two stoke motor oil used was Mobil Super 2T. Fuel oil was from Petrogal, SA (Portugal). 2.3. Growth Cells were grown in cylindrical 100-ml flasks closed with rubber bungs and containing 20 ml of medium (minimal salts medium, pH 7.0) [26], incubated at 15 or 28 °C and 150 rpm on a rotary shaker with an amplitude of 12.5 mm. The organic compounds used as sole carbon sources were present at an initial concentration of 0.125% or 0.25% (v/v). The extent of growth was monitored by measurement of the optical density (OD) at 600 nm and the dry weight, and, in addition, by microscopy and image analysis techniques. The initial OD at 600 nm in the assays carried out at 15 °C was 0.90, which corresponded to a dry weight of 0.30 g and an average of 311 cells per image, while in the experiments at 28 °C the initial OD was 1.24, the dry weight was 0.40 g and an average of 424 cells was observed per image. The percentages of motor oil tested were 0.13%,
0.25%, 0.50%, 1.00% and 2.00% (v/v). Assays were carried out at least in duplicate. 2.4. Carbon source consumption rate The cells were grown in 10-ml test tubes closed with rubber bungs wrapped in aluminium foil, containing 2 ml of minimal salts medium and 0.125% (v/v) of carbon source. The initial OD was 0.12, the dry weight being 0.04 g. The internal diameter of the test tubes was 12.5 mm and the depression caused on the liquid centre by the round bottom of the tubes was 2 mm. The test tubes were incubated at 28 °C and 200 rpm. During growth, at least five samples were taken at different times in order to follow the carbon source consumption. Each sample corresponded to three test tubes: one for biomass measurement and two for determination of the carbon source present. The content of these latter two tubes was extracted with 0.5 ml of ethyl acetate and, after phase separation, the ethyl acetate layer was analysed by gas chromatography (GC). Blank assays, carried out without cells, were also analysed. 2.5. GC analysis The samples were analysed by gas chromatography on a Hewlett–Packard 5890 gas chromatograph with a FID detector, connected to a HP3394 integrator. The capillary column was a SGE HT5, 25 m in length and with internal and external diameters of 0.22 and 0.33 mm, respectively. The oven temperature was 120 °C and that of the injector corresponded to 200 °C. The detector was set at 250 °C. 2.6. Fuel oil consumption Cells were grown in 100-ml flasks, closed with rubber bungs, containing 20 ml of minimal salts medium, R. erythropolis cells and 20, 40, 80, 160 or 320 ll of fuel oil adsorbed on the surface of a plastic tip. Two batches of flasks were used: on one batch, the initial OD600 was 0.78 (0.26 g dry weight); on the other, the initial OD600 was 0.39 ( 0.13 g dry weight). Fuel oil degradation was followed by extraction of the whole volume of 2 flasks with 2 ml of ethyl acetate and by extraction of 1 ml of the aqueous phase of 2 other flasks with 200 ll of ethyl acetate. The ethyl acetate layer was analysed by GC with the following oven temperature program: 50 °C in the first 4 min, followed by an increase of 10 °C min 1 up to 150 and 150 °C in the last 10 min. Several samples were taken in duplicate over a 10-month period, including blank assays without cells. 2.7. Cell hydrophobicity test Cell hydrophobicity tests were carried out according to the ‘‘microbial adhesion to hydrocarbon’’ (MATH)
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test, described by Rosenberg and co-workers [21] as follows: various volumes of n-hexadecane were added in test tubes to 1.2 ml of washed cells (collected at the beginning of the stationary phase) suspended in phosphate buffer (pH 7). After 10 min pre-incubation at 30 °C, the mixtures were agitated at full speed for 120 s on a vortex (Heidolph REAX 2000, Germany). After 15 min, the organic phase was removed and the absorbance of the aqueous phase was measured at 600 nm. Cell hydrophobicity refers to the percentage of cells that migrates towards the organic phase.
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peated injections, and are quoted for a confidence interval of 95%. Biomass concentration measurements (OD) had an associated error of ±8% based on the standard deviation and sample mean of eight repeated samples, quoted for a confidence interval of 95%. The error associated with the image analysis was ±7% based on the standard deviation and sample mean of twelve repeated images taken from the same sample, quoted for a confidence interval of 95%.
3. Results 2.8. Microscopy and image analysis 3.1. Growth on hydrocarbons at 15 and 20 °C The number of cells per image and the cell viability were measured by fluorescence microscopy, using a LIVE/DEADÒ Bac Lightä Bacterial Viability Kit from Molecular Probes. The microscope was an Olympus CX40, with an Olympus U-RFL-T burner and an UMWB mirror cube unit (excitation filter: BP450-480; barrier filter:BA515). Cell aggregates and cell positioning at the organic-aqueous interface were also observed under brightfield transmitted light. Images were captured using a COHU RGB camera. The acquisition software was Matrox Inspector 2.1. Image analysis was carried out using Visilog 5 for Windows 95 from Noesis SA, Les Ulis, France. Fluorescence microscopy allied with image analysis allowed the rapid quantification of cell growth, positioning and viability in the presence of the tested solvents. 2.9. Error analysis The error associated with the GC quantification of samples was ±6%. The errors were calculated based on the standard deviation and sample mean of seven re-
When cells of R. erythropolis DCL14 were grown on different organic solvents as the sole carbon source, growth was observed 60 h after inoculation on all substrates tested except with 0.25% toluene (data not shown). Clustering of cells was visible with the naked eye from 96 h onwards with n-dodecane and after 120 h with n-tetradecane and n-hexadecane as carbon sources (data not shown). Cell clustering was also considerable when the cells were metabolising the other carbon sources, with the exception of 0.125% toluene and iso-octane, in the presence of which no aggregation was visible. Observation of samples, taken from the different media using fluorescence microscopy, showed that the majority of R. erythropolis cells were either at the surface or inside the solvent droplets (image not shown). Growth was faster at 28 °C than at 15 °C for all the carbon sources tested, the highest growth rates being around 0.1 h 1 with n-dodecane, n-tetradecane and nhexadecane (Fig. 1). At 28 °C, toluene, n-octane and iso-octane appeared to be toxic at concentrations of
Fig. 1. Growth rates of Rhodococcus erythropolis DCL14 during the consumption of hydrocarbons as sole carbon and energy sources.
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0.25%, the growth rate decreasing more than 50% in comparison with that observed at concentrations of 0.125%. At 15 °C, cell growth appeared to be generally higher with 0.25% than with 0.125%, probably due to a lower solubility of the tested solvents in the aqueous phase. Overall, R. erythropolis DCL14 exhibited a nearly 10fold lower growth rate on hydrocarbons with an odd number of carbons than in those with an even number (Fig. 1), and its lag phase was longer on toluene than on the other carbon sources tested. According to the MATH test, the cells hydrophobicity was higher than 80% when growing on cyclohexane, n-hexane, n-octane and iso-octane, at both 15 and 28 °C (Fig. 2). Cells grown on toluene showed values of 54.9% and 92.1% at 15 and 28 °C, respectively. In general, the cell membrane hydrophobicity increased with increasing temperature, except for cells grown on n-hexadecane, which showed a hydrophobicity of 92.1% and 73.7% at 15 and 28 °C, respectively. Fig. 2(I) shows the cell hydrophobicity as a function of the log P of the carbon source, which is a measure of the hydrophobicity and toxicity of the solvent. According to Laane et al. [16], a log P of 4 is the minimum value for an organic solvent to be considered biocompatible. Observation of the results indicates that the data can be grouped in three clusters, corresponding to cells grown on carbon sources with an even (A and B) or odd (C) number of carbon atoms, and with a log P value lower (a) or higher (b) than 6. Cells grown on odd-numbered carbon sources showed lower hydrophobicity than cells grown on carbon sources with an even number of carbons. Considering the same type of carbon source (same cluster in Fig. 2), the cell surface hydrophobicity of strain DCL14 increased with increasing log P values, i.e., with increasing hydrophobicity of the solvent and with an increasing number of carbon atoms on the hydrocarbon chain. At 28 °C, solvents with an even
number of carbon atoms and with log P values higher than 6 allowed higher growth rates than those with log P values lower than 6, but with the latter, cell hydrophobicity was higher (Fig. 2, II). Cells grown on carbon sources with an odd number of carbons grew at almost the same rate, but cell hydrophobicity varied between 49.1 and 62.6%. 3.2. Growth on alcohols Considering growth of R. erythropolis DCL14 cells on several alcohols, the results from the growth experiments indicated that the cells were able to grow on C1–C12 alcohols as sole carbon and energy source at both 28 °C (Fig. 3) and 15 °C, although at less than half the rate when growing at 15 °C as compared to 28 °C (data not shown). No growth was observed on either of the n-octanol concentrations tested. At 15 °C, the growth rates attained with 0.125% of methanol and ethanol were 0.05 and 0.04 h 1, respectively. At 28 °C and with an initial substrate concentration of 0.125%, the growth rate decreased with increasing number of carbon atoms in the carbon source (Fig. 3). Also, with increasing number of carbon atoms, the growth rates also showed less variance up to concentrations of 1% of each carbon source. The cells were able to grow at concentration up to 1% cyclohexanol, 2% butanol and pentanol, 5% dodecanol, 15% methanol and 20% ethanol. These concentrations were achieved without any strategy to adapt the cells to high concentrations. The growth rates for 15% methanol and ethanol were 0.01 and 0.02 h 1, respectively, and for 20% ethanol it was 0.01 h 1. It was possible to achieve a linear relation between: (i) the highest concentration of the tested alcohols at which the cells were still able to grow completely (comprising lag, exponential and stationary phases) within 30 h and (ii) the difference in the growth rate observed between the highest and the lowest substrate concentration
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showing a correlation of 0.99 (Fig. 4). Higher differences in growth rates were achieved obviously for the highest concentrations. These results show that there is a relation between the maximum growth rate (which will determine the differences shown in Fig. 4) and the highest substrate concentration at which the cells can still grow. R. erythropolis DCL14 cells were found to be rather hydrophilic when grown on alcohols: only 10% of the cells grown on methanol, around 40% of those grown on ethanol and nearly 60% of cells grown on n-dodecanol migrated towards the n-hexadecane phase during the MATH test. The other alcohols tested as carbon sources produced cells with intermediate values, cell surface hydrophobicity increasing with the number of carbon atoms on the carbon source. Furthermore, no
clusters were observed during growth of cells on these carbon sources, at both 15 and 28 °C, except for the highest concentrations of methanol and ethanol used. The specific carbon source consumption rate was higher for methanol and ethanol than for the hydrocarbons tested (Fig. 5). Ethanol was consumed 30% faster than methanol and its consumption rate was double the average consumption rate of the hydrocarbons with an even number of carbons. 3.3. Growth under saline conditions at 15 °C Since the natural sites at which bioremediation has to take place may present saline conditions to the microorganisms, growth of R. erythropolis DCL14 cells was tested in the presence of different salt concentrations.
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Highest concentration, % (v/v) Fig. 4. Differences in the growth rates observed between an initial alcohol concentration of 0.125% and the maximum concentration at which the R. erythropolis DCL14 cells grew as function of this maximum concentration.
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Carbon source consumption (g/h.g d.w. )
0.7 0.6 0.5 0.4 0.3 0.2 0.1
Hexadecane
Tetradecane
Carbon source
Dodecane
Undecane
Nonane
Iso-octane
Octane
Toluene
Heptane
Hexane
Cyclohexane
Pentane
Ethanol
Methanol
0
Fig. 5. Carbon source consumption rates attained during growth of R. erythropolis DCL14 cells on alcohols and hydrocarbons as single carbon and energy source.
The lag phase of the cells increased with increasing concentrations of salt: around 50, 100 and 200 h (150 h for ethanol) at an initial NaCl concentration of 1.00%, 1.95% and 2.50%, respectively. At 1.00% NaCl, the highest growth rates (around 0.04 h 1) were observed when the cells were using ethanol, n-hexane and iso-octane as sole carbon sources, whereas the growth rate was almost half for cells growing on methanol and n-octane (Fig. 6). For an initial NaCl concentration of 1.95%, the growth rates attained with all the carbon sources tested only ranged between 0.017 h 1 for cells grown on methanol and 0.026 h 1 for cells grown on iso-octane. Growth rates decreased to nearly 0.014 h 1 at a salt concentration of 2.5% when the cells used methanol, nhexane and n-octane. However, a growth rate almost
as high as that attained when the salt concentration was 1% was observed with cells metabolising ethanol and iso-octane. In these two carbon sources, the cell hydrophobicity also decreased from around 40% at a salt concentration of 2% to less than 10% at a NaCl concentration of 2.5% (data not shown). Fluorescence microscopy and image analysis showed that the cells in contact with the highest salt concentration had decreased their size to about 30% as compared to cells in medium without additional NaCl. 3.4. Degradation of two- and four-stroke motor oils When the cells were incubated at 28 °C with two- and four-stroke motor oils, the optical density of the cultures
0.05 NaCl 1% NaCl 1.95%
0.04 Growth rate (h -1)
NaCl 2.5% 0.03
0.02
0.01
0 Methanol
Ethanol
n-Hexane
Iso-octane
n-Octane
Carbon source Fig. 6. Growth rates of R. erythropolis cells, carrying out the metabolism of alcohols and hydrocarbons as sole carbon and energy sources, observed in salted media at 28 °C.
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doubled during the first 20 h at initial oil fractions lower than 0.5% Shell Helix standard 20W-50 and for all Mobil Super 2T concentrations tested. With increasing oil concentrations, however, the growth rate decreased (Fig. 7). Significant cell aggregation was observed after 15 h with Shell Helix oil fractions higher than 0.5% and after 30 h with lower oil fractions. In the presence of Mobil Super 2T, cell clustering was observed after 40 h of cell growth. Thus quantification of growth by measurement of the optical density became impossible, and the results were calculated using the data obtained via fluorescence microscopy and image analysis and also by dry weight measurements. At 15 °C, the growth rate increased with increasing concentrations of Mobil Super 2T up to a fraction of 0.5% and remained almost constant with higher oil fractions at a value of nearly 0.04 h 1 (Fig. 7). When the cells were using Shell Helix, a sharp linear decrease of
the growth rate with increasing oil fractions was observed. At lower oil fractions, cell growth started at a higher rate when the cells were using Shell Helix but at higher fractions, cell growth was faster when cells were using Mobil 2T. Very small clusters were visible after 50 h in all growth experiments with both oils. The higher the oil fraction, the larger the cell clusters were. Images taken with a microscope showed cells gathered around small oil droplets of both oils (data not shown). The droplet diameter ranged between 0.02 and 0.09 mm. The MATH hydrophobicity test showed that, for both oils, cells were less hydrophobic when grown on 0.5% oil fraction (data not shown). The most hydrophobic cells were those grown on either the lowest (0.125%) or the highest (2%) initial oil fraction. Furthermore, cells grown on Mobil Super 2T were more hydrophobic than those consuming Shell Helix, at identical initial fractions.
0.09 15˚ C, Shell Helix
0.08
15˚ C, Mobil Super 2T
0.07 Growth rate (h -1)
28˚ C, Shell Helix 0.06
28˚ C, Mobil Super 2T
0.05 0.04 0.03 0.02 0.01 0 0
0.5
1
1.5
2
2.5
Concentration (%, v/v) Fig. 7. Growth rates attained with R. erythropolis DCL14 cells growing on motor oils at 15 and 28 °C.
100
Oil remaining (%)
80
60
40
20
0 0
2
4
395
6
8
10
Time (month) Fig. 8. Percentage of fuel oil remaining at different times during its biodegradation by R. erythropolis cells.
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In the presence of 2.5% NaCl, the growth curves followed similar trends to those shown in Fig. 7, but the growth rates were half of those achieved in media without salt (data not shown). 3.5. Degradation of fuel oil Fuel oil is composed of saturated hydrocarbons, aromatics, resins and asphaltenes. GC analysis only allowed the detection of alkanes and aromatics in the fuel oil used. The asphaltenes and resins were probably retained at the injector. The first compounds to be degraded were n-hexane, cyclohexane and iso-octane, followed by ndodecane. Almost complete degradation of the n-, isoand cyclo-alkanes, as well as aromatics present in fuel oil, was achieved by R. erythropolis DCL14 after 9 months of biodegradation (Fig. 8). After 3 months, more than 60% of the hydrocarbons had already been consumed. The percentage of oil remaining decreased with time according to a second order polynomial curve with a correlation of 0.99.
4. Discussion Nowadays, all kinds of motors are widely present in our environment and accidental oil spills are frequent. The bioremediation of sites contaminated with hydrocarbons, resulting from e.g. spillage of motor oils, diesel or jet fuels, may have to be carried out in the natural environment and thus at temperatures considerably below the optimum and/or under saline conditions. Microorganisms that present the ability to degrade these contaminants under different environmental conditions can represent a useful tool for the remediation of contaminated soil or for dedicated effluent treatment. Bacteria and fungi initiate the aerobic metabolism of alkanes using mono-oxygenases, which by a radical mechanism introduce a hydroxyl group in the alkane molecule through a partial reduction of O2. The resulting alkanol is further oxidised and metabolised in the b-oxidation pathway. In actinomycetes (including Rhodococcus, Mycobacterium and Nocardia), the b-oxidation pathway can be both a catabolic pathway and a source of fatty acids for the production of a large variety of lipids needed for survival and growth under adverse conditions [1]. Comparing the results obtained in this study for the degradation of the alkane with those of the corresponding alcohol, it is observed that on pentanol R. erythropolis DCL14 cells grew at a rate 5-fold higher than on n-pentane, whilst on cyclohexanol and dodecanol the DCL14 cells grew at a third of the rate obtained with cyclohexane and dodecane, respectively. No growth was observed on n-octanol, even at an initial concentration of 0.125%. These results indicate that during pentane metabolism in the tested strain, the forma-
tion of pentanol from pentane must be the limiting step. During the consumption of other hydrocarbons, the degradation of the corresponding alcohol, which should be more toxic according to the log P scale, is the limiting step since rather low concentrations of these compounds may damage the cells. However, at 28 °C, the cells were able to grow up to concentrations of 1% cyclohexanol, 2% butanol and pentanol, 5% dodecanol, 15% methanol and 20% ethanol. Growth on these high concentrations was achieved without any strategy to adapt the cells to increasing concentrations of either carbon source. Studies carried out by Solano-Serena et al. [22] showed that growth on cyclohexane and iso-octane may be difficult: most of the populations able to grow on solid medium using cyclohexane as substrate could not grow in liquid medium on this substrate; the only strain that was able to grow on iso-octane solely was isolated from a gasoline-polluted sample. Degradation of cyclohexane was achieved by co-metabolism [2] and by a mixed microbial population, which by several unknown interactions, was able to perform an accordant attack on the substrate [18]. R. erythropolis DCL14 was able to degrade these two recalcitrant compounds and to grow in medium with only these substrates as carbon and energy sources, showing the great potential of this strain for bioremediation. Furthermore, toluene, another toxic compound, was degraded at an initial concentration of 0.125%, although the lag phase was longer than with the other carbon sources. In a previous study, de Carvalho et al. [11] observed that when R. erythropolis DCL14 cells were incubated with toluene, non-viable cells were twice as large as the viable ones, indicating that toluene should increase the fluidity of the cellular membrane. After incubation with 2.5% toluene, the colonies growing on agar plates were yellow, i.e., the cells that survived produced phenotypically different colonies. Sokolovska´ et al. [23] found a clear correlation between the carbon source and the mycolic acid profiles of R. erythropolis E1: there was a lack of odd-numbered carbon chains in cells growing on linear alkanes with an even number of carbon atoms; mycolic acids were present with both even and odd-numbered carbon chains in cells growing on compounds with an odd number of carbon atoms, branched alkanes or mixtures of these compounds. These results, and the available literature, led the authors to conclude that there are two different pathways for the synthesis of mycolic acids in this strain: one by addition of acetyl coenzyme A and the other from addition of propionyl coenzyme A. According to Alvarez [1], during cultivation of the strain R. erythropolis 17 on n-alkanes, the cells produced fatty acids with a chain length associated with the chain length of the growth substrate, as well as with other fatty acids derived from the b-oxidation pathway. The strain used
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by this author [1] showed a more efficient mechanism for the production of the intermediate propionyl-CoA, precursor of fatty acids containing an odd-number of carbon atoms. However, in our case, the results indicate that R. erythropolis DCL14 cells have a less efficient mechanism for producing the precursor propionylCoA than other strains of the Rhodococcus genus, as the growth rates obtained with carbon sources with an odd number of carbon atoms were 10-fold lower than those obtained with an even number of carbons. The absence of a clear and linear relation between the hydrophobicity of the carbon source used and the observed cell hydrophobicity, suggests that the latter is the result of modifications in the membrane composition and not of direct incorporation of solvent molecules at the cellular membrane. Since a rough variant was chosen, i.e., the strain did not produce extracellular polymeric substances, modifications at the cell surface hydrophobicity by changes of the lipid composition of the membrane were probably the only available mechanism for protection from the toxic effects of the solvents. The maximum growth rate attained with a certain carbon source tested was related to the highest substrate concentration at which R. erythropolis DCL14 cells could still grow. Thus, the results clearly indicate that there is a relation between the DCL14 ability to degrade a certain compound and the cellÕs resistance to that compound, which may be crucial for the degradation of toxic substances. Contrarily, the results presented by Mosqueda et al. [19] suggested that toluene metabolism in Pseudomonas putida DOT-T1 is not involved in toluene tolerance. By studying samples taken from the different media using fluorescence microscopy, it was possible to observe cells both at the surface and inside the solvent droplets, suggesting that the uptake of substrate was achieved by direct contact between the cells and the organic solvent acting as carbon source (data not shown). Bouchez-Naı¨tali et al. [5] observed a direct interfacial uptake of n-hexadecane by four R. equi strains, which did not produce biosurfactants. The growth rates obtained were independent of the interfacial area, which was expected due to the very strong adsorption of the bacterial cells at the solvent-aqueous interface. Formation of cellular flocs was also observed due to the hydrophobicity of the strains tested. In the present study, the differences between the growth rates attained with 0.125% and 0.25% of n-dodecane, n-tetradecane and nhexadecane were between 1.5 and 7% (for n-tetradecane and n-dodecane, respectively). Since at the concentrations tested these hydrocarbons are immiscible, the interfacial area available for an initial concentration of 0.25% should double that obtained with an initial concentration of 0.125%. Thus, the growth rates obtained were almost independent of the interfacial area available, confirming that the uptake of these hydrocar-
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bons should be by direct contact between cells and substrate. In the case of the miscible methanol and ethanol, the cells were freely suspended in the aqueous phase containing the carbon source, whilst in the case of the remaining immiscible carbon sources, the cells were mainly located at the hydrocarbon–aqueous interface or even at the organic phase. Direct uptake of all carbon sources should therefore be possible, decreasing the impact of mass transfer (of the carbon source to the cell) on the substrate consumption rate. The results showed that methanol and ethanol were consumed at a higher rate than the hydrocarbons tested (Fig. 5). Since, as shown, the cells may modulate their surface hydrophobicity to improve adhesion to the organic phase, the differences observed in carbon source consumption rate were the result of particularities of the metabolism of the several compounds. This is particularly true when comparing the rates obtained for the odd- and evennumbered hydrocarbons. Metabolic engineering or directed mutagenenis could thus improve the R. erythropolis DCL14 bioremediation capabilities. Nevertheless, higher growth rates were attained with the most hydrophobic carbon sources (Fig. 2, II). A comparison between cells grown at 15 and 28 °C showed that at 15 °C the growth rates seemed more independent of the carbon source hydrophobicity, and also that there was a wider variation in the hydrophobic behaviour of the cells. The uptake of the hydrocarbon can thus be quite influenced by the temperature at which the cells proceed their metabolism, as well as by the composition of the cellular membrane. Many of the oil spills occur in ecosystems with high or moderate salinities such as the sea, estuaries, seashores, etc. Seawater contains, in average, 1.95% (w/w) chlorine and 1.08% (w/w) sodium [3]. In the present work, it was shown that strain DCL14 is able to metabolise methanol, ethanol, n-hexane, iso-octane and noctane in the presence of up to 2.5% NaCl at 15 °C. The growth rates decreased with increasing salt concentrations, but when ethanol and iso-octane were used, the growth rates observed at 2.5% NaCl were almost as high as those obtained with 1% NaCl. Apparently, at 2.5% NaCl, the influence on the cell membrane is larger and the resulting higher hydrophobicity of the cellular membrane promotes the consumption of these carbon sources. The specific contact area between the cells and the substrates should be higher at this concentration due to cell shrinkage, a well-known process of cell regulation in hypertonic media. This, allied with a higher hydrophobicity of the cellular membrane, could have improved the transport of ethanol and iso-octane from the growth media to the interior of the cell. In real oil spills in the environment, we often deal with mixtures of hydrocarbons and the presence of a single compound may influence the degradation of the others
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[13]. To assess this, degradation of Shell Helix Standard 20W-50 and Mobil super 2T, four- and two-stroke motor oil respectively, was tested at 15 and 28 °C. For some concentrations (higher than 0.5% Mobil; 0.125%, 0.25% and 2% Shell), the growth rates attained at 15 and 28 °C were quite close. Direct uptake of the compounds present in these oils was achieved by adhesion of the cells to 0.02–0.09 mm oil droplets, suggesting that strain DCL14 could be used for in situ bioremediation. Fuel oil, an even more worrying source of pollution, is composed of saturated hydrocarbons, aromatics, resins and asphaltenes. Microorganisms primarily degrade n- and iso-alkanes, followed by cycloalkanes and 1- and 3-ring aromatics. Finally, polyaromatics, asphaltenes and resins are consumed [4,24]. When R. erythropolis DCL14 cells were tested as degraders of a low sulphur content fuel oil, the first compounds to be degraded were n-hexane, cyclohexane and isooctane, followed by n-dodecane. This is in agreement with the results obtained during the assays with each of these compounds as sole carbon and energy source. This specific bacterium is therefore able to use a large number of recalcitrant compounds, both isolated and as a mixture of compounds. After 9 months of fuel oil metabolism, almost complete degradation of n-, iso- and cyclo-alkanes and aromatic present was achieved. This paper clearly shows the ability of R. erythropolis DCL14 to degrade n-alkanes, cycloalkanes, branched and monoaromatic alkanes and considerably high concentrations of alcohols at both 15 and 28 °C, even under saline conditions. Since the tested conditions are found in the Atlantic, Pacific and Indian oceans and the cells were found capable to degrade both motor and fuel oils, the large potential of strain DCL14 for bioremediation of real oil spills is also demonstrated.
Acknowledgements This study was supported by a Ph.D. Grant (PRAXIS XXI/BD/21574/99) awarded to Carla da C.C.R. de Carvalho by Fundac¸a˜o para a Cieˆncia e a Tecnologia, Portugal. The authors thank J.D.R. de Carvalho for the supply of the motor oils and Helder Tavanez for supplying the fuel oil used in this study.
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