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Low-temperature biodegradation of petroleum hydrocarbons (n-alkanes, phenol, anthracene, pyrene) by four actinobacterial strains
International Biodeterioration & Biodegradation 84 (2013) 185e191
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International Biodeterioration & Biodegradation journal homepage: www.elsevier.com/locate/ibiod
Low-temperature biodegradation of petroleum hydrocarbons (n-alkanes, phenol, anthracene, pyrene) by four actinobacterial strains Rosa Margesin*, Christoph Moertelmaier, Johannes Mair Institute of Microbiology, University of Innsbruck, Technikerstrasse 25, A-6020 Innsbruck, Austria
a r t i c l e i n f o
a b s t r a c t
Article history: Received 9 January 2012 Received in revised form 4 May 2012 Accepted 6 May 2012 Available online 4 June 2012
In this study we evaluated the ability of four cold-adapted bacterial strains to degrade n-alkanes (C12 eC22), aromatic hydrocarbons (phenol) and polyaromatic hydrocarbons (anthracene, pyrene) at low temperatures. All four strains belonged to the phylum Actinobacteria and were identified as Rhodococcus erythropolis (strain BZ4), Rhodococcus cercidiphyllus (strain BZ22), Arthrobacter sulfureus (strain BZ73) and Pimelobacter simplex (strain BZ91). The strains could grow over a temperature range of 1e30 C and showed catechol-1,2-dioxyogenase activity. One of the strains, R. erythropolis BZ4, degraded all of the compounds tested. The strain utilized n-alkanes and high amounts of phenol (7.5 mM), anthracene and pyrene (50 mg l1) at 15 C. P. simplex BZ91 degraded n-alkanes as well as up to 7.5 mM phenol; phenol degradation was observed at 1e30 C. R. cercidiphyllus BZ22 fully degraded C12 (700 mg l1) at 1e20 C, while degradation of C16 and C20 was delayed and lower compared to C12 degradation. A. sulfureus BZ73 was the best phenol degrader and utilized up to 12.5 mM phenol; phenol degradation occurred over a temperature range of 1e25 C. Such strains are promising candidates for low temperature (low-energy) treatment of industrial wastewaters contaminated with hydrocarbons. Ó 2012 Elsevier Ltd. All rights reserved.
Keywords: Rhodococcus Arthrobacter Pimelobacter Cold-adapted Hydrocarbons
1. Introduction Petroleum hydrocarbons are the most widespread contaminants in the environment. Alkanes, aromatic and polyaromatic hydrocarbons are representative fractions of petroleum hydrocarbons. Alkanes can be linear (n-alkanes), cyclic (cyclo-alkanes) or branched (iso-alkanes) and can constitute up to 50% of crude oil, depending on the oil source, but they are also produced by many living organisms. Since alkanes are apolar molecules that are chemically very inert, they are characterized by low water solubility and tend to accumulate in cell membranes (Rojo, 2009). Phenol and phenolic compounds are widely distributed in nature and as environmental pollutants. They are common constituents of many industrial wastewaters such as those produced from crude oil refineries and coal gasification plants. Due to their toxicity to microorganisms, phenolic compounds may often cause the breakdown of wastewater treatment plants by inhibition of microbial growth (Ren and Frymier, 2003), even at relatively low concentrations such as 2 mM (Li and Humphrey, 1989). PAHs are frequent environmental contaminants that occur in coal, oil and tar deposits
* Corresponding author. Tel.: þ43 512 5076021; fax: þ43 512 5072929. E-mail address: [email protected] (R. Margesin). 0964-8305/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.ibiod.2012.05.004
and are produced as byproducts of fuel burning and by incomplete combustion of carbon-containing fuels. PAHs are of concern as pollutants since they are lipophilic; some compounds are mutagenic, cancerogenic and teratogenic (Samanta et al., 2002). The capacity of a broad spectrum of microorganisms to utilize hydrocarbons as the sole source of carbon and energy has been recognized very early (Zobell, 1946) and was the basis for the development of biological remediation methods. The ability to degrade hydrocarbons is widespread among soil microorganisms. They may adapt rapidly to the contamination, as demonstrated by significantly increased numbers of hydrocarbon degraders after a pollution event (Margesin and Schinner, 2001; Greer et al., 2010). The intensity of biodegradation is influenced by several factors, such as nutrients, oxygen, pH-value, composition, concentration and bioavailability of the contaminants. Temperature plays a significant role in controlling the nature and the extent of microbial hydrocarbon metabolism. Bioavailability and solubility of hydrophobic substances with low solubility, such as some aliphatic and polyaromatic hydrocarbons, are temperature-dependent. The decreased volatilization and solubility of some hydrocarbons at low temperature affects toxicity. A temperature decrease also results in a decrease in diffusion rates of organic compounds and in an increase in viscosity, which affects the degree of distribution (Whyte et al., 1998; Rojo, 2009).
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In cold and temperate climatic regions, temperatures of industrial wastewater, groundwater and soil can often decrease to temperature levels around or below 15 C due to seasonal and/or diurnal fluctuations. The activity of mesophilic degraders is severely limited at this temperature, whereas cold-adapted microorganisms have evolved a series of adaptation strategies that enable them to compensate for the negative effects of low temperatures on biochemical reactions (Feller, 2007; Margesin et al., 2008). It was the objective of this study to evaluate and to compare the capability of four cold-adapted actinobacterial strains to degrade representative fractions of petroleum hydrocarbons, i.e. alkanes, aromatic hydrocarbons (phenol) and polyaromatic hydrocarbons (anthracene, pyrene) at low temperatures. Strains with the required properties for efficient low temperature biodegradation, i.e., growth and degradation at low temperatures, are useful for the treatment of hydrocarbon-contaminated ecosystems. 2. Materials and methods 2.1. Isolation and identification of strains Bacterial strains were isolated from petroleum hydrocarbon contaminated alpine soil collected from a former industrial district in Bozen, South Tyrol, Italy. Leakage from heavy oil storage tanks were the main reason for contamination. At the time of sampling, the mean soil temperature in the sampling area was 8e10 C. The soil contained 13,300 mg hydrocarbons/kg dry soil. 40% and 60% of this contamination consisted of C10eC20 and C20eC40 hydrocarbons, respectively, which points to a high content of heavy oils. Isolation of bacterial strains was done on mineral medium agar containing diesel oil as previously described (Margesin et al., 2003b). Four bacterial strains, BZ4, BZ22, BZ73 and BZ91, were selected on the basis of their ability to grow well in the presence of petroleum hydrocarbons. Strains were routinely cultured on R2A agar and maintained as a suspension in skimmed milk (10%, w/v) at 80 C. Cell morphology was examined by phase-contrast microscopy ( 1000). The Gram-reaction was tested by Gram-staining. API strips (API 20 NE, API Coryne, API ZYM; bioMérieux) were used to determine physiological and biochemical characteristics as well as enzyme activities. Ligninolytic activity was evaluated on MM agar plates containing 0.4% (w/v) lignosulfonic acid sodium salt (Margesin et al., 2002) after 7 days at 15 C. Growth under anaerobic conditions was assessed as described (Zhang et al., 2012b). Genomic DNA of the bacterial strains was extracted using the UltraClean Microbial DNA isolation kit (Mo Bio Laboratories). The 16S rRNA genes were amplified as described earlier (Zhang et al., 2010). The 16S rRNA gene sequences were submitted for comparison and identification to the GenBank databases using the NCBI Blast N algorithm, the EMBL databases using the Fasta algorithm
Relative biodegradation (%)
100
and the Ribosomal Database Project II (RDP) using its Sequence Match (Cole et al., 2005). GenBank accession numbers (National Center for Biotechnical Information, NCBI) for the 16S rRNA gene sequences of strains BZ4, BZ22, BZ73 and BZ91 are HQ588862, HQ588861, HQ588859 and HQ588857, respectively. 2.2. Growth temperature range of bacterial strains Suspensions of bacterial cells (pre-grown on R2A agar plates at 15 C) in 0.9% NaCl were used to inoculate R2A agar plates that were incubated at 1, 5, 10, 15, 20, 25, 30 and 37 C, using two replicates per strain and temperature. Growth was monitored up to an incubation time of 7e21 days. 2.3. Biodegradation of hydrocarbons The ability of the strains to degrade aerobically n-alkanes, phenol or polyaromatic hydrocarbons was determined in 100-ml Erlenmeyer flasks containing 10 ml of pH-neutral phosphatebuffered mineral medium without yeast extract (MM) (Margesin and Schinner, 1997) amended with the hydrocarbons to be tested as the sole carbon source. The medium was inoculated with a suspension of microbial cells (pre-grown in medium with the hydrocarbon to be degraded) prepared in 0.9% NaCl to give an initial OD600 of 0.05 after inoculation. Abiotic losses were monitored in sterile hydrocarbon-containing medium. All tests were performed with three replicates; the standard deviations obtained were 15%. 2.3.1. Biodegradation of n-alkanes MM was amended with a mixture of the n-alkanes C12, C16, C18, C20 and C22 (each 700 mg l1) as the sole carbon source. C18, C20 and C22 are solid at 20 C and were thus solubilized in a water bath at 55 C before addition to the medium. Recovery of n-alkanes was determined immediately after contamination of the medium (t ¼ 0). Erlenmeyer flasks with screw caps were used in order to minimize abiotic loss by volatilization. After an incubation period of 7 and 15 days at 15 C and 150 rpm, the residual concentration of the individual n-alkanes was quantified by gas chromatography after extraction with heptane (DIN EN ISO 9377-2). The best n-alkane degrader, strain BZ22, was selected to evaluate the effect of temperature on n-alkane degradation. The strain was cultured at 1, 10, 20 and 30 C with a mixture of the n-alkanes C12, C16 and C20 (each 700 mg l1) as the sole carbon source and the residual n-alkane concentration was determined after 4, 7, 11, 14 and 28 days of incubation (Fig. 2). 2.3.2. Biodegradation of phenol Fed-batch cultivation with increasing phenol concentration was used to determine the highest amount of phenol that could be degraded by the tested strains (Fig. 3). Strains were grown at 15 C 100
100
80
Rh. erythropolis BZ4
80
Rh. cercidiphyllus BZ22
80 P. simplex BZ91
60
60
60
40
40
40
20
20
20 0
0
0 C12
C16
C18
C20
C22
C12
C16
C18
C20
C22
C12
C16
C18
C20
C22
Fig. 1. Relative n-alkane degradation after 7 days (white colums) and 15 days (gray colums) at 15 C by three actinobacterial strains. The initial concentration (100%) of each of the nalkanes was 700 mg l1. Data are mean values and standard deviations of three replicates.
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80
80 C12
60
100 Residual conc. (%)
100
Residual conc. (%)
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80 60 40
1°C 10°C
20
20°C 30°C
C16
0 10
20
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C20
0
0 0
187
0
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Days
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0
Days
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20
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Days
Fig. 2. Effect of temperature on relative n-alkane degradation by Rhocococcus cercidipyhllus BZ22. The initial concentration (100%) of each of the n-alkanes was 700 mg l1; the residual concentration is shown. Data are mean values of three replicates, standard deviations were 15%.
Residual phenol (mM)
12
Rh. erythropolis BZ4
10 8 6 4 2 0
0
10
20
Residual phenol (mM)
16
30
40
50
A. sulfureus BZ73
14 12 10 8 6 4 2 0
0
Residual phenol (mM)
12
10
20
30
40
50
10 8 6 4 2 0
10
2.3.3. Biodegradation of polyaromatic hydrocarbons (PAHs) The MM contained anthracene or pyrene (20 and 50 mg l1) as the sole carbon source and was inoculated with strain BZ4. After 28 days (20 mg PAH l1) or 48 days (50 mg l1) of incubation at 15 C and 150 rpm, the residual PAH concentration was determined by gas chromatographyemass spectrometry according to DIN 38407F39 by Eurofins Umwelt Ost GmbH (Jena, Germany). 2.4. Catechol dioxygenase activity
P. simplex BZ91
0
and 150 rpm in 100-ml-Erlenmeyer flasks containing 10 ml MM amended with 1.25 mM phenol. Growth (OD600) and the residual phenol concentration were monitored at regular time intervals. After phenol disappearance, the same culture was re-contaminated with increasing phenol concentrations (2.5, 5.0, 7.5, 10, 12.5 and 15 mM phenol). With each phenol amendment, the volume was adjusted to the initial volume of 10 ml by adding fresh MM. To determine the effect of temperature on phenol degradation, the most efficient phenol degraders, strains BZ73 and BZ91, were cultured at 1, 5, 10, 15, 20, 25 and 30 C at 150 rpm in the presence of 10 mM phenol. To evaluate the effect of agitation, cultivation of strains BZ73 and BZ91 was done at 15 C without agitation, as well as on a rotary shaker at 150 rpm and 250 rpm. Growth (OD600) and the residual phenol concentration were monitored at regular intervals. Phenol concentration was determined in culture supernatants, that were filtered (0.2 mm, Minsart RC4 17821) after centrifugation. High-performance liquid chromatography (HPLC) was carried out as described by Allsop et al. (1993), using an RP-18 column (5 mm 100 mm, Lichrospher, Merck), detection at 220 nm (Shimadzu SPD-20A) and an eluent flow of 0.5 ml min1. The elution time for phenol was approx. 7.5 min. The phenol calibration curve was prepared in MM.
20
30
40
50
Days Fig. 3. Phenol degradation at 15 C in fed-batch cultures of three actinobacterial strains. Arrows indicate phenol amendments. Data are mean values of three replicates, standard deviations were 15%.
The presence of catechol dioxygenases (catechol-1,2 dioxygenase activity (C1,2D), catechol 2,3 dioxygenase activity (C2,3D)) was determined according to a modification (Margesin et al., 2005) of the methods described by Nakazawa and Nakazawa (1970) and Nozaki (1970). To determine whether catechol dioxygenase activity was constitutively and/or inductively expressed, cells grown in complex medium (R2A) and in MM with phenol as the sole carbon source were tested. One hundred ml of liquid culture containing cells grown in R2A or phenol-degrading cells were mixed with 750 ml of appropriate buffer and 50 ml of 0.2 mM catechol. After 5 min at 25 C, the reaction was stopped by cooling on ice, and the enzyme activities were determined in the supernatants by measuring the product formation from catechol at 260 nm (cis, cismuconic acid for C2,1D) or 375 nm (2-hydroxymuconic semialdehyde for C2,3D).
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3. Results 3.1. Identification of strains All four strains described in this study were representatives of the phylum Actinobacteria (Zhang et al., 2012a). They were identified as members of the species Rhodococcus erythropolis (BZ4, 99.0% identity with 16S rRNA gene sequences in GenBank databases); Rhodococcus cercidiphyllus (BZ22; 99.3% 16S rRNA gene sequence identity); Arthrobacter sulfureus (BZ73, 99.1% sequence identity) and Pimelobacter simplex (BZ91, 99.9% sequence identity). All four strains were cold-adapted, showing growth on R2A agar over a temperature range of 1e30 C (Table 1). 3.2. Enzyme activities Ligninolytic activity was noted for all four strains. Since the substrate used for this activity test, lignosulfonic acid, contains a high amount of phenolic compounds, the strains were expected to be efficient phenol degraders. Catechol dioxygenases are involved in the second step of phenol degradation and catalyze the ring cleavage of catechol (¼ohydroxyphenol). All four strains showed constitutive expression of C1,2D activity, indicating the oxidation of catechol by the ortho type of ring cleavage. None of the strains produced C2,3D, i.e. the ability to oxidize catechol by the meta type of ring cleavage. C1,2D activity of phenol-degrading strains (R. erythropolis BZ4, A. sulfureus BZ73 and P. simplex BZ91; see 3.4) was considerably higher when cells were induced with phenol. 3.3. Biodegradation of n-alkanes n-Alkanes could be recovered almost completely (95e100%) at t ¼ 0. Abiotic losses as determined in sterile controls ranged from 3 1% to 6 4% after 14 days at 15 C. Biodegradation was calculated from the difference between total hydrocarbon loss in inoculated medium and abiotic loss in sterile medium. Out of the four strains tested in this study, three strains (R. erythroplis BZ4, R. cercidiphyllus BZ22, P. simplex BZ91) were able to utilize n-alkanes as the sole source of carbon and energy. The chain length influenced n-alkane biodegradation to a significant extent. After 14 days of incubation at 15 C, C12 was completely (R. erythroplis BZ4, R. cercidiphyllus BZ22) or almost completely (93%, P. simplex BZ91) degraded. Of the initial amount of C16, 75% (R. cercidiphyllus BZ22) or 59% (R. erythroplis BZ4, P. simplex BZ91) were degraded. C18 was already degraded to a much lower extent (10e29%), and C20 and C22 were only degraded up to an amount of 18% (Fig. 1). The most efficient n-alkane degrader, R. cercidiphyllus BZ22, was further tested for the effect of temperature on biodegradation
(Fig. 2). Generally, the lag time for biodegradation increased with increasing n-alkane chain length and with decreasing temperature. Full degradation of C12 was obtained at 1, 10 and 20 C. Degradation of C16 was already delayed compared to C12 degradation, however, still 80%, 65% and 50% of the initial concentration of C16 were degraded at 20, 10 and 1 C. For C20, only a degradation of 30% of the initial concentration was obtained at 20 C and 30 C; at lower temperatures degradation was low (15% at 10 C) or absent (1 C). Despite the ability of the strain to grow in complex medium (R2A) at temperatures ranging from 1 C to 30 C, both the maximum and minimum temperature for growth were suboptimal for efficient n-alkane degradation. Degradation at 1 C, the minimum growth temperature of the strain, was only efficient for C12, delayed for C16 and absent for C20. The degradation rate of C12 at 30 C was initially comparable to that observed at 10 C, however, C12 was not fully degraded at 30 C. Similarly, C16 degradation was less efficient at 30 C than at 10 C. On the other hand, C20 degradation was comparable at 20 C and 30 C and more efficient than at 10 C. This may be explained by the fact that C20 is available in its liquid form and thus better bioavailable at temperatures above 20 C, which clearly demonstrates the influence of the aggregate state (liquid vs. solid) of the compound to be degraded on bioavailability and consequently on biodegradation. 3.4. Biodegradation of phenol The biodegradation of increasing phenol concentrations was tested by using fed-batch cultivation, which is an efficient method for the selection and acclimation of hydrocarbon-degrading microorganisms. One of the four strains, R. cercidiphyllus BZ22, was not able to utilize phenol. We observed the same pattern with all three phenol-degrading strains (R. erythropolis BZ4, A. sulfureus BZ73, P. simplex BZ91). Growth paralleled phenol biodegradation. With increasing amounts of phenol the lag-phases increased due to substrate inhibition by the toxic substrate phenol (data not shown). Changes in the residual phenol concentration during cultivation at 15 C are shown in Fig. 3. As there was no abiotic loss of phenol in sterile controls, phenol disappearance could be attributed to biodegradation. Both R. erythropolis BZ4 and P. simplex BZ91 fully degraded phenol up to a concentration of 7.5 mM within 42 and 26 days at 15 C, respectively, while A. sulfureus BZ73 even fully degraded 12.5 mM phenol at 15 C within 19 days; 15 mM phenol, however, were only partly utilized (Fig. 3). A. sulfureus BZ73 fully degraded 10 mM phenol over a temperature range of 1e25 C (Fig. 4). Phenol had completely disappeared after 2 and 3 days at 25 C and 20 C, respectively, while 5 and 9 days were needed to degrade 10 mM phenol at 15 and 10 C, respectively. Biodegradation was considerably delayed at lower temperatures, nonetheless, 10 mM phenol were fully degraded
Table 1 Properties of the strains investigated in this study. Parameter
Strain BZ4
Strain BZ22
Strain BZ73
Strain BZ91
Identification Growth temperature range (R2A agar) C1,2D (constitutive expression) C1,2D (inductive expression) C2,3D (constitutive expression) C2,3D (inductive expression) Ligninolytic activity Degradation at 15 C n-Alkanes Phenol Anthracene, pyrene
Rhodococcus erythropolis 1e30 C þ þ þ
Rhodococcus cercidiphyllum 1e30 C þ No growth with phenol No growth with phenol þ
Arthrobacter sulfureus 1e30 C þ þ þ
Pimelobacter simplex 1e30 C þ þ þ
þ þ(7.5 mM) þ(50 mg l1)
þ
þ (12.5 mM)
þ þ (7.5 mM)
C1,2D: catechol-1,2 dioxygenase. C2,3D: catechol-2,3 dioxygenase.
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A. sulfureus BZ73
1.4
189
phenol occurred after 2 days at 20 C and after 10 days at 10 C). When the strain was cultured in shaken flasks (regardless of whether 150 or 250 rpm), 5 mM phenol were fully degraded after 1 day (20 C) or 3 days (10 C).
1.2
3.5. Biodegradation of PAHs
OD600
1.0 0.8
1°C 5°C 10°C 15°C 20°C 25°C
0.6 0.4 0.2 0.0 0
10
20
30
40
50
10 Residual phenol (mM)
A. sulfureus BZ73
Only one out of the four strains, R. erythropolis BZ4, was able to utilize anthracene and pyrene as the sole source of carbon and energy in liquid culture. The 3-ring compound anthracene was almost fully degraded after 28 days at 15 C when the initial concentration was 20 mg l1. An initial concentration of 50 mg l1 resulted in a residual concentration of 13 6% after 48 days at 15 C. A similar trend was noted for degradation of the 4-ring compound pyrene of which only 11 6% (t0 ¼ 20 mg l1) and 8 2% (t0 ¼ 50 mg l1) were detected after 28 and 48 days, respectively, at 15 C. There was no abiotic loss of anthracene or pyrene in sterile controls, therefore PAH disappearance could be attributed entirely to biodegradation. 4. Discussion
8 1°C 5°C 10°C 15°C 20°C 25°C
6 4 2 0 0
10
20
30
40
50
Days Fig. 4. Effect of temperature on growth and phenol degradation by Arthrobacter sulfureus BZ73. At 30 C, growth was very low (OD600 ¼ 0.14) and phenol degradation did not occur. Data are mean values of three replicates, standard deviations were 15%.
after 26 days at 5 C and after 39 days at 1 C. Despite the fact that the strain could grow in complex medium at 30 C, no growth occurred at this temperature in MM with phenol as the sole carbon source. Biodegradation and growth decreased with decreasing growth temperature and were considerably delayed at 1e5 C. Biomass formation (in terms of cell density), however, was higher in the low temperature range 1e10 C than at 15e25 C (Fig. 4), which demonstrates the cold adaptation of the strain. Almost the same pattern was observed with P. simplex BZ91 (data not shown). This strain degraded 5 mM phenol over the whole growth temperature range (1e30 C). Full degradation was noticed after 1 day (30 C), 3e5 days (25e15 C), 11 days (10 C), 35 days (5 C) and 39 days (1 C). Agitation influenced growth and phenol degradation of A. sulfureus BZ73 and P. simplex BZ91 to a significant extent. Growth and biodegradation by A. sulfureus BZ73 were substantially delayed in flasks without agitation. Agitation at 250 rpm resulted in a better performance than agitation at 150 rpm. After 4 days at 20 C, 10 mM phenol was fully degraded when the strain was cultured at 250 rpm, while 51 3% of the initial concentration were measured when the strain was grown at 150 rpm and only 21 2% degradation had occurred in flasks without agitation. This pattern was also observed at a growth temperature of 10 C, where full degradation of 10 mM phenol at 250 rpm was noticed only after 19 days. Similarly, degradation of 5 mM phenol by P. simplex BZ91 was slower when cultured without agitation (full degradation of 5 mM
In this study we demonstrate the potential of four bacterial strains isolated from petroleum hydrocarbon contaminated soil to degrade n-alkanes, phenol and polyaromatic compounds at temperatures of 15 C and below. All four strains were representatives of the phylum Actinobacteria and belonged to the genera Arthrobacter (Micrococcaceae), Rhodococcus (Nocardiaceae) and Pimelobacter (Nocardiaceae). Actinobacteria are Gram-positive bacteria with a high G þ C content and are typical members of microbial communities in soils, including polar and alpine soils, where they play an important role in the degradation of organic materials. They tend to be successful in resource-limited situations and can be found in both pristine and hydrocarbon-contaminated soils (Aislabie et al., 2000; Margesin et al., 2003b; Greer et al., 2010). Representatives of the genera Arthrobacter and Rhodococcus are well-known for their ability to degrade a wide range of pollutants, including aliphatic and aromatic hydrocarbons (Ganesh and Lin, 2009; Greer et al., 2010). Representatives of the genus Pimelobacter have been isolated from contaminated soil (Qiao and Wang, 2010) and have been associated with the degradation of pyridine, quinoline and sterols (Schmidt et al., 1991; Qiao and Wang, 2010; Andryushina et al., 2011), however, there are no reports on Pimelobacter strains isolated from cold environments. Here we report the first description of a cold-adapted representative of this genus (strain P. simplex BZ91) able to degrade efficiently the n-alkanes C12 and C16 as well as high amounts (7.5 mM) of phenol at low temperatures. Alkane biodegradation is strongly influenced by alkane solubility in water, which decreases as the molecular mass increases. While short-chain alkanes (C5eC9) may be toxic to microorganisms due to their high solubility in water, long-chain alkanes (C22) are considered to be recalcitrant due to their low solubility in water (Rojo, 2009). Rhodococci have often been described as efficient degraders of a broad range of n-alkanes; the capability of alkane degradation is often accompanied by the ability to produce biosurfactants (Ganesh and Lin, 2009). Degradation of short-chain and long-chain alkanes by rhodococci at low temperatures has been previously reported (Whyte et al., 1998; Rapp and Gabriel-Jürgens, 2003; De Carvalho and da Fonseca, 2005; Lee et al., 2010). In our study, the most efficient n-alkane degraders belonged to the genus Rhodococcus (strains BZ4 and BZ22) and were able to utilize C12eC22, however, with the best efficiency toward C12eC18. Alkane degradation by strain BZ22 was clearly influenced by temperature and alkane chain length and decreased with decreasing
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temperature and increasing n-alkane chain length. Whyte et al. (1998) also observed that short-chain alkanes (C10 and C16) were mineralized to a significantly greater extent than long-chain alkanes (C28 and C32) at 0 and 5 C as a result of decreased bioavailability (and thus increased recalcitrance) of long-chain alkanes at low temperature. Takei et al. (2008) hypothesized for a Rhodococcus strain the recruitment of certain alkane hydroxylase systems for the utilization of alkanes under low temperature conditions. Rojo (2009) distinguished two groups of alkane degraders. Alkane degraders may either have a very versatile metabolism that allows them to utilize many other compounds as carbon sources in addition to alkanes (Margesin et al., 2003a; Harayama et al., 2004), while others may be highly specialized in degrading hydrocarbons (Rojo, 2009). Of the three alkane degraders described in this study, R. cercidiphyllus BZ4 (degradation of n-alkanes, phenol, anthracene and pyrene) and P. simplex BZ91 (degradation of n-alkanes and phenol) belong to the first group, while R. erythropolis BZ22 only degrades n-alkanes and thus belongs to the second group. There is comparatively little information on cold-adapted phenol degraders. Kotturi et al. (1991) demonstrated the degradation of amounts up to 10.6 mM phenol by Pseudomonas putida at 10 C. Onysko et al. (2000) reported low temperature phenol degradation by cold-adapted aquatic pseudomonads. Marine bacteria degraded 1 mM phenol (Kobayashi et al., 2012). A number of alpine yeast strains degraded phenol and phenol-related compounds at low temperatures (Bergauer et al., 2005). Coldadapted bacterial and yeast strains utilized up to 12.5 and 15 mM phenol at 10 C (Margesin et al., 2003a, 2005). The degradation of higher amounts of phenol (20e30 mM) has only been reported by mesophilic bacteria (Yap et al., 1999; Zeng et al., 2010). In our study, representatives of the genera Rhodococcus (strain BZ4) and Pimelobacter (strain BZ91) fully degraded 7.5 mM phenol at 15 C, while A. sulfureus BZ73 even fully utilized 12.5 mM phenol. We isolated previously two rhodococci strains able to degrade phenol up to 12.5 mM at 10 C, however, their performance at 1 C (Margesin et al., 2005) was lower than that observed with the strains described in this study. Arthrobacter psychrophenolicus degraded 5 mM phenol at temperatures between 1 and 25 C; the highest amount degraded was 10 mM phenol (Margesin et al., 2004a). It has been shown that phenol degradation is inhibited by extremes of pH and insufficiently dissolved oxygen (Agarry et al., 2008). In fact, we also observed accelerated phenol degradation when strains were grown under agitated conditions (in shaken flasks). Interestingly, toxicity of aromatic compounds has been shown to be significantly lower for cold-adapted bacteria at 10 C compared to 25 C, while mesophilic counterparts showed a lower susceptibility to high amounts of aromatic compounds when grown at 25 C compared to 10 C (Margesin et al., 2004b). This reflects the adaptation of coldadapted and mesophilic strains to their respective optimal growth temperature ranges. Little is known on the fate of PAHs in cold habitats. The two compounds investigated in our study, anthracene and pyrene, are known contaminants of soil and groundwater. PAHs were detected in Antarctic soil and surface marine sediment; the main sources for the main compounds (2-ring and 3-ring PAHs) were lowtemperature combustion processes (Curtosi et al., 2007). Coldtolerant isolates (Sphingomonas or Pseudomonas spp.) from oilpolluted Antarctic soils utilized naphthalene, phenanthrene and fluorene as the sole carbon and energy source (Aislabie et al., 2000) and bacterial and yeast strains from alpine habitats utilized anthracene at 10 C (Margesin et al., 2003a). A mixed enrichment culture degraded anthracene and pyrene (up to 20 mg l1) at 10 C and 25 C (Sartoros et al., 2005). In our study, one strain, R. erythreus BZ4, degraded efficiently high amounts (up to 50 mg l1) of
anthracene and pyrene at 15 C. This points to the efficiency of rhodococci to degrade a wide range of hydrocarbons. Lee et al. (2010) also described a strain belonging to the genus Rhodococcus that was able to degrade recalcitrant hydrocarbons, including pyrene. In conclusion, we clearly demonstrated in this study the potential of cold-adapted actinobacterial strains to degrade a broad range of hydrocarbons, including alkanes, aromatic and polyaromatic hydrocarbons, at low temperatures. One of the strains, R. erythropolis BZ4, degraded all of the compounds tested in this study. The capability to utilize a wide range of hydrocarbons is advantageous for the treatment of mixed pollutions. Moreover, the strains investigated in this study could grow and degrade hydrocarbons over a broad temperature range (1e25 C or 1e30 C). The application of such degraders is advantageous in cold and temperate environments that undergo (diurnal and/or seasonal) thermal fluctuations. Such strains are useful for low temperature and consequently low-energy treatment of industrial effluents contaminated with hydrocarbons. In addition, they could also be useful for the construction of biosensors for the selective, sensitive and rapid monitoring or in situ analysis of pollution (Alkasrawi et al., 1999).
Acknowledgments This research work was supported by a grant from the “Autonome Provinz Bozen, Südtirol”.
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International Biodeterioration & Biodegradation journal homepage: www.elsevier.com/locate/ibiod
Low-temperature biodegradation of petroleum hydrocarbons (n-alkanes, phenol, anthracene, pyrene) by four actinobacterial strains Rosa Margesin*, Christoph Moertelmaier, Johannes Mair Institute of Microbiology, University of Innsbruck, Technikerstrasse 25, A-6020 Innsbruck, Austria
a r t i c l e i n f o
a b s t r a c t
Article history: Received 9 January 2012 Received in revised form 4 May 2012 Accepted 6 May 2012 Available online 4 June 2012
In this study we evaluated the ability of four cold-adapted bacterial strains to degrade n-alkanes (C12 eC22), aromatic hydrocarbons (phenol) and polyaromatic hydrocarbons (anthracene, pyrene) at low temperatures. All four strains belonged to the phylum Actinobacteria and were identified as Rhodococcus erythropolis (strain BZ4), Rhodococcus cercidiphyllus (strain BZ22), Arthrobacter sulfureus (strain BZ73) and Pimelobacter simplex (strain BZ91). The strains could grow over a temperature range of 1e30 C and showed catechol-1,2-dioxyogenase activity. One of the strains, R. erythropolis BZ4, degraded all of the compounds tested. The strain utilized n-alkanes and high amounts of phenol (7.5 mM), anthracene and pyrene (50 mg l1) at 15 C. P. simplex BZ91 degraded n-alkanes as well as up to 7.5 mM phenol; phenol degradation was observed at 1e30 C. R. cercidiphyllus BZ22 fully degraded C12 (700 mg l1) at 1e20 C, while degradation of C16 and C20 was delayed and lower compared to C12 degradation. A. sulfureus BZ73 was the best phenol degrader and utilized up to 12.5 mM phenol; phenol degradation occurred over a temperature range of 1e25 C. Such strains are promising candidates for low temperature (low-energy) treatment of industrial wastewaters contaminated with hydrocarbons. Ó 2012 Elsevier Ltd. All rights reserved.
Keywords: Rhodococcus Arthrobacter Pimelobacter Cold-adapted Hydrocarbons
1. Introduction Petroleum hydrocarbons are the most widespread contaminants in the environment. Alkanes, aromatic and polyaromatic hydrocarbons are representative fractions of petroleum hydrocarbons. Alkanes can be linear (n-alkanes), cyclic (cyclo-alkanes) or branched (iso-alkanes) and can constitute up to 50% of crude oil, depending on the oil source, but they are also produced by many living organisms. Since alkanes are apolar molecules that are chemically very inert, they are characterized by low water solubility and tend to accumulate in cell membranes (Rojo, 2009). Phenol and phenolic compounds are widely distributed in nature and as environmental pollutants. They are common constituents of many industrial wastewaters such as those produced from crude oil refineries and coal gasification plants. Due to their toxicity to microorganisms, phenolic compounds may often cause the breakdown of wastewater treatment plants by inhibition of microbial growth (Ren and Frymier, 2003), even at relatively low concentrations such as 2 mM (Li and Humphrey, 1989). PAHs are frequent environmental contaminants that occur in coal, oil and tar deposits
* Corresponding author. Tel.: þ43 512 5076021; fax: þ43 512 5072929. E-mail address: [email protected] (R. Margesin). 0964-8305/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.ibiod.2012.05.004
and are produced as byproducts of fuel burning and by incomplete combustion of carbon-containing fuels. PAHs are of concern as pollutants since they are lipophilic; some compounds are mutagenic, cancerogenic and teratogenic (Samanta et al., 2002). The capacity of a broad spectrum of microorganisms to utilize hydrocarbons as the sole source of carbon and energy has been recognized very early (Zobell, 1946) and was the basis for the development of biological remediation methods. The ability to degrade hydrocarbons is widespread among soil microorganisms. They may adapt rapidly to the contamination, as demonstrated by significantly increased numbers of hydrocarbon degraders after a pollution event (Margesin and Schinner, 2001; Greer et al., 2010). The intensity of biodegradation is influenced by several factors, such as nutrients, oxygen, pH-value, composition, concentration and bioavailability of the contaminants. Temperature plays a significant role in controlling the nature and the extent of microbial hydrocarbon metabolism. Bioavailability and solubility of hydrophobic substances with low solubility, such as some aliphatic and polyaromatic hydrocarbons, are temperature-dependent. The decreased volatilization and solubility of some hydrocarbons at low temperature affects toxicity. A temperature decrease also results in a decrease in diffusion rates of organic compounds and in an increase in viscosity, which affects the degree of distribution (Whyte et al., 1998; Rojo, 2009).
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In cold and temperate climatic regions, temperatures of industrial wastewater, groundwater and soil can often decrease to temperature levels around or below 15 C due to seasonal and/or diurnal fluctuations. The activity of mesophilic degraders is severely limited at this temperature, whereas cold-adapted microorganisms have evolved a series of adaptation strategies that enable them to compensate for the negative effects of low temperatures on biochemical reactions (Feller, 2007; Margesin et al., 2008). It was the objective of this study to evaluate and to compare the capability of four cold-adapted actinobacterial strains to degrade representative fractions of petroleum hydrocarbons, i.e. alkanes, aromatic hydrocarbons (phenol) and polyaromatic hydrocarbons (anthracene, pyrene) at low temperatures. Strains with the required properties for efficient low temperature biodegradation, i.e., growth and degradation at low temperatures, are useful for the treatment of hydrocarbon-contaminated ecosystems. 2. Materials and methods 2.1. Isolation and identification of strains Bacterial strains were isolated from petroleum hydrocarbon contaminated alpine soil collected from a former industrial district in Bozen, South Tyrol, Italy. Leakage from heavy oil storage tanks were the main reason for contamination. At the time of sampling, the mean soil temperature in the sampling area was 8e10 C. The soil contained 13,300 mg hydrocarbons/kg dry soil. 40% and 60% of this contamination consisted of C10eC20 and C20eC40 hydrocarbons, respectively, which points to a high content of heavy oils. Isolation of bacterial strains was done on mineral medium agar containing diesel oil as previously described (Margesin et al., 2003b). Four bacterial strains, BZ4, BZ22, BZ73 and BZ91, were selected on the basis of their ability to grow well in the presence of petroleum hydrocarbons. Strains were routinely cultured on R2A agar and maintained as a suspension in skimmed milk (10%, w/v) at 80 C. Cell morphology was examined by phase-contrast microscopy ( 1000). The Gram-reaction was tested by Gram-staining. API strips (API 20 NE, API Coryne, API ZYM; bioMérieux) were used to determine physiological and biochemical characteristics as well as enzyme activities. Ligninolytic activity was evaluated on MM agar plates containing 0.4% (w/v) lignosulfonic acid sodium salt (Margesin et al., 2002) after 7 days at 15 C. Growth under anaerobic conditions was assessed as described (Zhang et al., 2012b). Genomic DNA of the bacterial strains was extracted using the UltraClean Microbial DNA isolation kit (Mo Bio Laboratories). The 16S rRNA genes were amplified as described earlier (Zhang et al., 2010). The 16S rRNA gene sequences were submitted for comparison and identification to the GenBank databases using the NCBI Blast N algorithm, the EMBL databases using the Fasta algorithm
Relative biodegradation (%)
100
and the Ribosomal Database Project II (RDP) using its Sequence Match (Cole et al., 2005). GenBank accession numbers (National Center for Biotechnical Information, NCBI) for the 16S rRNA gene sequences of strains BZ4, BZ22, BZ73 and BZ91 are HQ588862, HQ588861, HQ588859 and HQ588857, respectively. 2.2. Growth temperature range of bacterial strains Suspensions of bacterial cells (pre-grown on R2A agar plates at 15 C) in 0.9% NaCl were used to inoculate R2A agar plates that were incubated at 1, 5, 10, 15, 20, 25, 30 and 37 C, using two replicates per strain and temperature. Growth was monitored up to an incubation time of 7e21 days. 2.3. Biodegradation of hydrocarbons The ability of the strains to degrade aerobically n-alkanes, phenol or polyaromatic hydrocarbons was determined in 100-ml Erlenmeyer flasks containing 10 ml of pH-neutral phosphatebuffered mineral medium without yeast extract (MM) (Margesin and Schinner, 1997) amended with the hydrocarbons to be tested as the sole carbon source. The medium was inoculated with a suspension of microbial cells (pre-grown in medium with the hydrocarbon to be degraded) prepared in 0.9% NaCl to give an initial OD600 of 0.05 after inoculation. Abiotic losses were monitored in sterile hydrocarbon-containing medium. All tests were performed with three replicates; the standard deviations obtained were 15%. 2.3.1. Biodegradation of n-alkanes MM was amended with a mixture of the n-alkanes C12, C16, C18, C20 and C22 (each 700 mg l1) as the sole carbon source. C18, C20 and C22 are solid at 20 C and were thus solubilized in a water bath at 55 C before addition to the medium. Recovery of n-alkanes was determined immediately after contamination of the medium (t ¼ 0). Erlenmeyer flasks with screw caps were used in order to minimize abiotic loss by volatilization. After an incubation period of 7 and 15 days at 15 C and 150 rpm, the residual concentration of the individual n-alkanes was quantified by gas chromatography after extraction with heptane (DIN EN ISO 9377-2). The best n-alkane degrader, strain BZ22, was selected to evaluate the effect of temperature on n-alkane degradation. The strain was cultured at 1, 10, 20 and 30 C with a mixture of the n-alkanes C12, C16 and C20 (each 700 mg l1) as the sole carbon source and the residual n-alkane concentration was determined after 4, 7, 11, 14 and 28 days of incubation (Fig. 2). 2.3.2. Biodegradation of phenol Fed-batch cultivation with increasing phenol concentration was used to determine the highest amount of phenol that could be degraded by the tested strains (Fig. 3). Strains were grown at 15 C 100
100
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Fig. 1. Relative n-alkane degradation after 7 days (white colums) and 15 days (gray colums) at 15 C by three actinobacterial strains. The initial concentration (100%) of each of the nalkanes was 700 mg l1. Data are mean values and standard deviations of three replicates.
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80 C12
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Fig. 2. Effect of temperature on relative n-alkane degradation by Rhocococcus cercidipyhllus BZ22. The initial concentration (100%) of each of the n-alkanes was 700 mg l1; the residual concentration is shown. Data are mean values of three replicates, standard deviations were 15%.
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2.3.3. Biodegradation of polyaromatic hydrocarbons (PAHs) The MM contained anthracene or pyrene (20 and 50 mg l1) as the sole carbon source and was inoculated with strain BZ4. After 28 days (20 mg PAH l1) or 48 days (50 mg l1) of incubation at 15 C and 150 rpm, the residual PAH concentration was determined by gas chromatographyemass spectrometry according to DIN 38407F39 by Eurofins Umwelt Ost GmbH (Jena, Germany). 2.4. Catechol dioxygenase activity
P. simplex BZ91
0
and 150 rpm in 100-ml-Erlenmeyer flasks containing 10 ml MM amended with 1.25 mM phenol. Growth (OD600) and the residual phenol concentration were monitored at regular time intervals. After phenol disappearance, the same culture was re-contaminated with increasing phenol concentrations (2.5, 5.0, 7.5, 10, 12.5 and 15 mM phenol). With each phenol amendment, the volume was adjusted to the initial volume of 10 ml by adding fresh MM. To determine the effect of temperature on phenol degradation, the most efficient phenol degraders, strains BZ73 and BZ91, were cultured at 1, 5, 10, 15, 20, 25 and 30 C at 150 rpm in the presence of 10 mM phenol. To evaluate the effect of agitation, cultivation of strains BZ73 and BZ91 was done at 15 C without agitation, as well as on a rotary shaker at 150 rpm and 250 rpm. Growth (OD600) and the residual phenol concentration were monitored at regular intervals. Phenol concentration was determined in culture supernatants, that were filtered (0.2 mm, Minsart RC4 17821) after centrifugation. High-performance liquid chromatography (HPLC) was carried out as described by Allsop et al. (1993), using an RP-18 column (5 mm 100 mm, Lichrospher, Merck), detection at 220 nm (Shimadzu SPD-20A) and an eluent flow of 0.5 ml min1. The elution time for phenol was approx. 7.5 min. The phenol calibration curve was prepared in MM.
20
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Days Fig. 3. Phenol degradation at 15 C in fed-batch cultures of three actinobacterial strains. Arrows indicate phenol amendments. Data are mean values of three replicates, standard deviations were 15%.
The presence of catechol dioxygenases (catechol-1,2 dioxygenase activity (C1,2D), catechol 2,3 dioxygenase activity (C2,3D)) was determined according to a modification (Margesin et al., 2005) of the methods described by Nakazawa and Nakazawa (1970) and Nozaki (1970). To determine whether catechol dioxygenase activity was constitutively and/or inductively expressed, cells grown in complex medium (R2A) and in MM with phenol as the sole carbon source were tested. One hundred ml of liquid culture containing cells grown in R2A or phenol-degrading cells were mixed with 750 ml of appropriate buffer and 50 ml of 0.2 mM catechol. After 5 min at 25 C, the reaction was stopped by cooling on ice, and the enzyme activities were determined in the supernatants by measuring the product formation from catechol at 260 nm (cis, cismuconic acid for C2,1D) or 375 nm (2-hydroxymuconic semialdehyde for C2,3D).
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3. Results 3.1. Identification of strains All four strains described in this study were representatives of the phylum Actinobacteria (Zhang et al., 2012a). They were identified as members of the species Rhodococcus erythropolis (BZ4, 99.0% identity with 16S rRNA gene sequences in GenBank databases); Rhodococcus cercidiphyllus (BZ22; 99.3% 16S rRNA gene sequence identity); Arthrobacter sulfureus (BZ73, 99.1% sequence identity) and Pimelobacter simplex (BZ91, 99.9% sequence identity). All four strains were cold-adapted, showing growth on R2A agar over a temperature range of 1e30 C (Table 1). 3.2. Enzyme activities Ligninolytic activity was noted for all four strains. Since the substrate used for this activity test, lignosulfonic acid, contains a high amount of phenolic compounds, the strains were expected to be efficient phenol degraders. Catechol dioxygenases are involved in the second step of phenol degradation and catalyze the ring cleavage of catechol (¼ohydroxyphenol). All four strains showed constitutive expression of C1,2D activity, indicating the oxidation of catechol by the ortho type of ring cleavage. None of the strains produced C2,3D, i.e. the ability to oxidize catechol by the meta type of ring cleavage. C1,2D activity of phenol-degrading strains (R. erythropolis BZ4, A. sulfureus BZ73 and P. simplex BZ91; see 3.4) was considerably higher when cells were induced with phenol. 3.3. Biodegradation of n-alkanes n-Alkanes could be recovered almost completely (95e100%) at t ¼ 0. Abiotic losses as determined in sterile controls ranged from 3 1% to 6 4% after 14 days at 15 C. Biodegradation was calculated from the difference between total hydrocarbon loss in inoculated medium and abiotic loss in sterile medium. Out of the four strains tested in this study, three strains (R. erythroplis BZ4, R. cercidiphyllus BZ22, P. simplex BZ91) were able to utilize n-alkanes as the sole source of carbon and energy. The chain length influenced n-alkane biodegradation to a significant extent. After 14 days of incubation at 15 C, C12 was completely (R. erythroplis BZ4, R. cercidiphyllus BZ22) or almost completely (93%, P. simplex BZ91) degraded. Of the initial amount of C16, 75% (R. cercidiphyllus BZ22) or 59% (R. erythroplis BZ4, P. simplex BZ91) were degraded. C18 was already degraded to a much lower extent (10e29%), and C20 and C22 were only degraded up to an amount of 18% (Fig. 1). The most efficient n-alkane degrader, R. cercidiphyllus BZ22, was further tested for the effect of temperature on biodegradation
(Fig. 2). Generally, the lag time for biodegradation increased with increasing n-alkane chain length and with decreasing temperature. Full degradation of C12 was obtained at 1, 10 and 20 C. Degradation of C16 was already delayed compared to C12 degradation, however, still 80%, 65% and 50% of the initial concentration of C16 were degraded at 20, 10 and 1 C. For C20, only a degradation of 30% of the initial concentration was obtained at 20 C and 30 C; at lower temperatures degradation was low (15% at 10 C) or absent (1 C). Despite the ability of the strain to grow in complex medium (R2A) at temperatures ranging from 1 C to 30 C, both the maximum and minimum temperature for growth were suboptimal for efficient n-alkane degradation. Degradation at 1 C, the minimum growth temperature of the strain, was only efficient for C12, delayed for C16 and absent for C20. The degradation rate of C12 at 30 C was initially comparable to that observed at 10 C, however, C12 was not fully degraded at 30 C. Similarly, C16 degradation was less efficient at 30 C than at 10 C. On the other hand, C20 degradation was comparable at 20 C and 30 C and more efficient than at 10 C. This may be explained by the fact that C20 is available in its liquid form and thus better bioavailable at temperatures above 20 C, which clearly demonstrates the influence of the aggregate state (liquid vs. solid) of the compound to be degraded on bioavailability and consequently on biodegradation. 3.4. Biodegradation of phenol The biodegradation of increasing phenol concentrations was tested by using fed-batch cultivation, which is an efficient method for the selection and acclimation of hydrocarbon-degrading microorganisms. One of the four strains, R. cercidiphyllus BZ22, was not able to utilize phenol. We observed the same pattern with all three phenol-degrading strains (R. erythropolis BZ4, A. sulfureus BZ73, P. simplex BZ91). Growth paralleled phenol biodegradation. With increasing amounts of phenol the lag-phases increased due to substrate inhibition by the toxic substrate phenol (data not shown). Changes in the residual phenol concentration during cultivation at 15 C are shown in Fig. 3. As there was no abiotic loss of phenol in sterile controls, phenol disappearance could be attributed to biodegradation. Both R. erythropolis BZ4 and P. simplex BZ91 fully degraded phenol up to a concentration of 7.5 mM within 42 and 26 days at 15 C, respectively, while A. sulfureus BZ73 even fully degraded 12.5 mM phenol at 15 C within 19 days; 15 mM phenol, however, were only partly utilized (Fig. 3). A. sulfureus BZ73 fully degraded 10 mM phenol over a temperature range of 1e25 C (Fig. 4). Phenol had completely disappeared after 2 and 3 days at 25 C and 20 C, respectively, while 5 and 9 days were needed to degrade 10 mM phenol at 15 and 10 C, respectively. Biodegradation was considerably delayed at lower temperatures, nonetheless, 10 mM phenol were fully degraded
Table 1 Properties of the strains investigated in this study. Parameter
Strain BZ4
Strain BZ22
Strain BZ73
Strain BZ91
Identification Growth temperature range (R2A agar) C1,2D (constitutive expression) C1,2D (inductive expression) C2,3D (constitutive expression) C2,3D (inductive expression) Ligninolytic activity Degradation at 15 C n-Alkanes Phenol Anthracene, pyrene
Rhodococcus erythropolis 1e30 C þ þ þ
Rhodococcus cercidiphyllum 1e30 C þ No growth with phenol No growth with phenol þ
Arthrobacter sulfureus 1e30 C þ þ þ
Pimelobacter simplex 1e30 C þ þ þ
þ þ(7.5 mM) þ(50 mg l1)
þ
þ (12.5 mM)
þ þ (7.5 mM)
C1,2D: catechol-1,2 dioxygenase. C2,3D: catechol-2,3 dioxygenase.
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A. sulfureus BZ73
1.4
189
phenol occurred after 2 days at 20 C and after 10 days at 10 C). When the strain was cultured in shaken flasks (regardless of whether 150 or 250 rpm), 5 mM phenol were fully degraded after 1 day (20 C) or 3 days (10 C).
1.2
3.5. Biodegradation of PAHs
OD600
1.0 0.8
1°C 5°C 10°C 15°C 20°C 25°C
0.6 0.4 0.2 0.0 0
10
20
30
40
50
10 Residual phenol (mM)
A. sulfureus BZ73
Only one out of the four strains, R. erythropolis BZ4, was able to utilize anthracene and pyrene as the sole source of carbon and energy in liquid culture. The 3-ring compound anthracene was almost fully degraded after 28 days at 15 C when the initial concentration was 20 mg l1. An initial concentration of 50 mg l1 resulted in a residual concentration of 13 6% after 48 days at 15 C. A similar trend was noted for degradation of the 4-ring compound pyrene of which only 11 6% (t0 ¼ 20 mg l1) and 8 2% (t0 ¼ 50 mg l1) were detected after 28 and 48 days, respectively, at 15 C. There was no abiotic loss of anthracene or pyrene in sterile controls, therefore PAH disappearance could be attributed entirely to biodegradation. 4. Discussion
8 1°C 5°C 10°C 15°C 20°C 25°C
6 4 2 0 0
10
20
30
40
50
Days Fig. 4. Effect of temperature on growth and phenol degradation by Arthrobacter sulfureus BZ73. At 30 C, growth was very low (OD600 ¼ 0.14) and phenol degradation did not occur. Data are mean values of three replicates, standard deviations were 15%.
after 26 days at 5 C and after 39 days at 1 C. Despite the fact that the strain could grow in complex medium at 30 C, no growth occurred at this temperature in MM with phenol as the sole carbon source. Biodegradation and growth decreased with decreasing growth temperature and were considerably delayed at 1e5 C. Biomass formation (in terms of cell density), however, was higher in the low temperature range 1e10 C than at 15e25 C (Fig. 4), which demonstrates the cold adaptation of the strain. Almost the same pattern was observed with P. simplex BZ91 (data not shown). This strain degraded 5 mM phenol over the whole growth temperature range (1e30 C). Full degradation was noticed after 1 day (30 C), 3e5 days (25e15 C), 11 days (10 C), 35 days (5 C) and 39 days (1 C). Agitation influenced growth and phenol degradation of A. sulfureus BZ73 and P. simplex BZ91 to a significant extent. Growth and biodegradation by A. sulfureus BZ73 were substantially delayed in flasks without agitation. Agitation at 250 rpm resulted in a better performance than agitation at 150 rpm. After 4 days at 20 C, 10 mM phenol was fully degraded when the strain was cultured at 250 rpm, while 51 3% of the initial concentration were measured when the strain was grown at 150 rpm and only 21 2% degradation had occurred in flasks without agitation. This pattern was also observed at a growth temperature of 10 C, where full degradation of 10 mM phenol at 250 rpm was noticed only after 19 days. Similarly, degradation of 5 mM phenol by P. simplex BZ91 was slower when cultured without agitation (full degradation of 5 mM
In this study we demonstrate the potential of four bacterial strains isolated from petroleum hydrocarbon contaminated soil to degrade n-alkanes, phenol and polyaromatic compounds at temperatures of 15 C and below. All four strains were representatives of the phylum Actinobacteria and belonged to the genera Arthrobacter (Micrococcaceae), Rhodococcus (Nocardiaceae) and Pimelobacter (Nocardiaceae). Actinobacteria are Gram-positive bacteria with a high G þ C content and are typical members of microbial communities in soils, including polar and alpine soils, where they play an important role in the degradation of organic materials. They tend to be successful in resource-limited situations and can be found in both pristine and hydrocarbon-contaminated soils (Aislabie et al., 2000; Margesin et al., 2003b; Greer et al., 2010). Representatives of the genera Arthrobacter and Rhodococcus are well-known for their ability to degrade a wide range of pollutants, including aliphatic and aromatic hydrocarbons (Ganesh and Lin, 2009; Greer et al., 2010). Representatives of the genus Pimelobacter have been isolated from contaminated soil (Qiao and Wang, 2010) and have been associated with the degradation of pyridine, quinoline and sterols (Schmidt et al., 1991; Qiao and Wang, 2010; Andryushina et al., 2011), however, there are no reports on Pimelobacter strains isolated from cold environments. Here we report the first description of a cold-adapted representative of this genus (strain P. simplex BZ91) able to degrade efficiently the n-alkanes C12 and C16 as well as high amounts (7.5 mM) of phenol at low temperatures. Alkane biodegradation is strongly influenced by alkane solubility in water, which decreases as the molecular mass increases. While short-chain alkanes (C5eC9) may be toxic to microorganisms due to their high solubility in water, long-chain alkanes (C22) are considered to be recalcitrant due to their low solubility in water (Rojo, 2009). Rhodococci have often been described as efficient degraders of a broad range of n-alkanes; the capability of alkane degradation is often accompanied by the ability to produce biosurfactants (Ganesh and Lin, 2009). Degradation of short-chain and long-chain alkanes by rhodococci at low temperatures has been previously reported (Whyte et al., 1998; Rapp and Gabriel-Jürgens, 2003; De Carvalho and da Fonseca, 2005; Lee et al., 2010). In our study, the most efficient n-alkane degraders belonged to the genus Rhodococcus (strains BZ4 and BZ22) and were able to utilize C12eC22, however, with the best efficiency toward C12eC18. Alkane degradation by strain BZ22 was clearly influenced by temperature and alkane chain length and decreased with decreasing
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temperature and increasing n-alkane chain length. Whyte et al. (1998) also observed that short-chain alkanes (C10 and C16) were mineralized to a significantly greater extent than long-chain alkanes (C28 and C32) at 0 and 5 C as a result of decreased bioavailability (and thus increased recalcitrance) of long-chain alkanes at low temperature. Takei et al. (2008) hypothesized for a Rhodococcus strain the recruitment of certain alkane hydroxylase systems for the utilization of alkanes under low temperature conditions. Rojo (2009) distinguished two groups of alkane degraders. Alkane degraders may either have a very versatile metabolism that allows them to utilize many other compounds as carbon sources in addition to alkanes (Margesin et al., 2003a; Harayama et al., 2004), while others may be highly specialized in degrading hydrocarbons (Rojo, 2009). Of the three alkane degraders described in this study, R. cercidiphyllus BZ4 (degradation of n-alkanes, phenol, anthracene and pyrene) and P. simplex BZ91 (degradation of n-alkanes and phenol) belong to the first group, while R. erythropolis BZ22 only degrades n-alkanes and thus belongs to the second group. There is comparatively little information on cold-adapted phenol degraders. Kotturi et al. (1991) demonstrated the degradation of amounts up to 10.6 mM phenol by Pseudomonas putida at 10 C. Onysko et al. (2000) reported low temperature phenol degradation by cold-adapted aquatic pseudomonads. Marine bacteria degraded 1 mM phenol (Kobayashi et al., 2012). A number of alpine yeast strains degraded phenol and phenol-related compounds at low temperatures (Bergauer et al., 2005). Coldadapted bacterial and yeast strains utilized up to 12.5 and 15 mM phenol at 10 C (Margesin et al., 2003a, 2005). The degradation of higher amounts of phenol (20e30 mM) has only been reported by mesophilic bacteria (Yap et al., 1999; Zeng et al., 2010). In our study, representatives of the genera Rhodococcus (strain BZ4) and Pimelobacter (strain BZ91) fully degraded 7.5 mM phenol at 15 C, while A. sulfureus BZ73 even fully utilized 12.5 mM phenol. We isolated previously two rhodococci strains able to degrade phenol up to 12.5 mM at 10 C, however, their performance at 1 C (Margesin et al., 2005) was lower than that observed with the strains described in this study. Arthrobacter psychrophenolicus degraded 5 mM phenol at temperatures between 1 and 25 C; the highest amount degraded was 10 mM phenol (Margesin et al., 2004a). It has been shown that phenol degradation is inhibited by extremes of pH and insufficiently dissolved oxygen (Agarry et al., 2008). In fact, we also observed accelerated phenol degradation when strains were grown under agitated conditions (in shaken flasks). Interestingly, toxicity of aromatic compounds has been shown to be significantly lower for cold-adapted bacteria at 10 C compared to 25 C, while mesophilic counterparts showed a lower susceptibility to high amounts of aromatic compounds when grown at 25 C compared to 10 C (Margesin et al., 2004b). This reflects the adaptation of coldadapted and mesophilic strains to their respective optimal growth temperature ranges. Little is known on the fate of PAHs in cold habitats. The two compounds investigated in our study, anthracene and pyrene, are known contaminants of soil and groundwater. PAHs were detected in Antarctic soil and surface marine sediment; the main sources for the main compounds (2-ring and 3-ring PAHs) were lowtemperature combustion processes (Curtosi et al., 2007). Coldtolerant isolates (Sphingomonas or Pseudomonas spp.) from oilpolluted Antarctic soils utilized naphthalene, phenanthrene and fluorene as the sole carbon and energy source (Aislabie et al., 2000) and bacterial and yeast strains from alpine habitats utilized anthracene at 10 C (Margesin et al., 2003a). A mixed enrichment culture degraded anthracene and pyrene (up to 20 mg l1) at 10 C and 25 C (Sartoros et al., 2005). In our study, one strain, R. erythreus BZ4, degraded efficiently high amounts (up to 50 mg l1) of
anthracene and pyrene at 15 C. This points to the efficiency of rhodococci to degrade a wide range of hydrocarbons. Lee et al. (2010) also described a strain belonging to the genus Rhodococcus that was able to degrade recalcitrant hydrocarbons, including pyrene. In conclusion, we clearly demonstrated in this study the potential of cold-adapted actinobacterial strains to degrade a broad range of hydrocarbons, including alkanes, aromatic and polyaromatic hydrocarbons, at low temperatures. One of the strains, R. erythropolis BZ4, degraded all of the compounds tested in this study. The capability to utilize a wide range of hydrocarbons is advantageous for the treatment of mixed pollutions. Moreover, the strains investigated in this study could grow and degrade hydrocarbons over a broad temperature range (1e25 C or 1e30 C). The application of such degraders is advantageous in cold and temperate environments that undergo (diurnal and/or seasonal) thermal fluctuations. Such strains are useful for low temperature and consequently low-energy treatment of industrial effluents contaminated with hydrocarbons. In addition, they could also be useful for the construction of biosensors for the selective, sensitive and rapid monitoring or in situ analysis of pollution (Alkasrawi et al., 1999).
Acknowledgments This research work was supported by a grant from the “Autonome Provinz Bozen, Südtirol”.
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