Oxidation of aliphatic, branched chain, and aromatic hydrocarbons by Nocardia cyriacigeorgica isolated from oil-polluted sand samples collected in the Saudi Arabian Desert
Le Thi Nhi-Cong1, Annett Mikolasch2, Susanne Awe2, Halah Sheikhany3, Hans-Peter Klenk4 and Frieder Schauer2
1 Department of Petroleum Microbiology, Institute of Biotechnology, VAST, Hoang Quoc Viet-Cau Giay-Hanoi-Vietnam
2 Ernst-Moritz-Arndt-University Greifswald, Institute of Microbiology, Greifswald, Germany 3 Department of Soil Science, Faculity of Agriculture, Damascus, Syria 4 Department of Microbiology, DSMZ – German Collection of Microorganisms and Cell Cultures,
Braunschweig, Germany
A soil bacterium isolated from oil-polluted sand samples collected in the Saudi Arabian Desert has been determined as Nocardia cyriacigeorgica, which has a high capacity of degrading and utilizing a broad range of hydrocarbons. The metabolic pathways of three classes of hydro-carbons were elucidated by identifying metabolites in cell-free extracts analyzed by GC/MS and HPLC/UV-Vis in comparison with standard compounds. During tetradecane oxidation, tetra-decanol; tetradecanoic acid; dodecanoic acid; decanoic acid could be found as metabolites, indicating a monoterminal degradation pathway of n-alkanes. The oxidation of pristane result-ed in the presence of pristanoic acid; 2-methylglutaric acid; 4,8-dimethylnonanoic acid; and 2,6-dimethylheptanoic acid, which give rise to a possible mono- and di-terminal oxidation. In case of sec-octylbenzene, eight metabolites were detected including 5-phenylhexanoic acid; 3-phe-nylbutyric acid; 2-phenylpropionic acid; β-methylcinnamic acid; acetophenone; β-hydroxy acetophenone; 2,3-dihydroxy benzoic acid and succinic acid. From these intermediates a new degradation pathway for sec-octylbenzene was investigated. Our results indicate that N. cyria-cigeorgica has the ability to degrade aliphatic and branched chain alkanes as well as alkyl-benzene effectively and, therefore, N. cyriacigeorgica is probably a suitable bacterium for biodegradation of oil or petroleum products in contaminated soils.
Received: November 03, 2009; accepted: November 03, 2009
DOI 10.1002/jobm.200900358
Introduction*
Crude oil represents a complex mixture of hydrophobic organic compounds composed of linear and branched alkanes, alkylbenzenes, alicyclic and polyaromatic hy- Correspondence: Le Thi Nhi-Cong, Department of Petroleum Micro-biology, Institute of Biotechnology, VAST, 18-Hoang Quoc Viet-Cau Giay-Hanoi-Vietnam E-mail: [email protected] or [email protected] Phone: 0084-4-37562000 Fax: 0084-4-37564483
drocarbons as well as asphaltenes of varying structure. As a complex of naturally occurring compounds crude oil is subject to degradation by various microorganisms [1, 4]. Whereas the degradation of oil in freshwater and marine habitats or by the mesophilic microflora of soils is studied in detail only a few reports exist on microor-ganisms involved in oil degradation in very hot and dry environments. n-Alkanes are probably the least reactive class of organic compounds. Nevertheless their water insolubil-
ity, hydrophobicity and their frequent occurrence as environmental pollutants in higher amounts they are suitable carbon sources for several microorganisms. The problem of environmental contamination with crude oil is also aggravated by the branched chain al-kanes that normally accompany it. Branched alkanes are generally more recalcitrant to biodegradation than linear alkanes due to the fact that the alkyl branches either hinder the uptake of hydrocarbon into the cell or are not susceptible to the enzymes of the β−oxidation pathway [2, 8, 27, 35]. Similar to linear and branched alkanes, aromatic hydrocarbons are ubiquitous in nature and are signifi-cant environmental pollutants. Benzene and related compounds are characterized by their possession of large (negative) resonance energy. This results in a ther-modynamic stability which manifests itself in chemical properties very different from those observed for ali-phatic compounds. The appearance of electron donat-ing substituent groups onto the benzene ring increases the possibility of alternative modes of biodegradation; either ring attack or side chain attack in the case of aliphatic side chains [37]. For example, longer chain length of alkylbenzenes support growth because of β-oxidation of the side chain and several organisms may not be able to degrade the aromatic moiety [5]. Sari-aslani et al. reported on the bacterial degradation of long alkylbenzenes by initial side chain attack via ω- and β-oxidation [32]. Whereas Smith et al. [37] con-cluded from the biodegradation of alkylbenzenes in-cluding several branched side chains by Pseudomonas sp. that all of these alkylbenzenes were catabolized via ring attack, rather than side chain attack. Hitherto, though several studies have reported the isolation and characterization of various bacteria capa-ble of utilizing these substrates as a carbon and energy source, a single bacterium does not possess the capabil-ity of degrading all or even most of the organic com-pounds in a polluted soil [4]. It is generally accepted that hydrocarbon utilization is extremely common amongst the genera Corynebacterium, Mycobacterium, Rho-dococcus and Nocardia, even amongst potentially patho-genic members of this group. To explain this phenon-menon, Stephens et al. proposed the presence of a highly hydrophobic cell surface on these organisms and the possession of numerous oxygenase enzymes that might facilitate their growth and predominance in n-alkane enrichments [38]. Members of the Nocardiaceae are known for their capacity to degrade hydrocarbons. N. cyriacigeorgica (formerly named Nocardia cyriacigeorgici) is a newly rec-ognized species within the genus [44]. Still, far too little
is known about the distribution and physiology of the genus N. cyriacigeorgica and whether the species plays a role in hydrocarbons degradation is largely unknown [13, 15]. In the present work, we report on the isolation of a strain SBUG 1472 belonging to the species N. cyria-cigeorgica from oil-polluted sand samples collected in the Saudi Arabian Desert and on its capacity of biodeg-radation. We also demonstrate that the strain is able to grow on a variety of petroleum hydrocarbons ranging from n-alkanes (C6 to C16) and branched hydrocarbons (pristane) to alkylbenzenes (sec-octylbenzene). sec-Octyl-benzene was preferred to n-octylbenzene to study the oxidation mechanism at the branching point of the carbon chain which is hitherto widely unknown for bacteria. Because of its broad hydrocarbon oxidation capacity strain SBUG 1472 could be useful for applica-tion in bioremediation technologies.
Materials and methods
Chemicals 2,6,10,14-Tetramethylpentadecane (pristane), purity >99% by gas chromatography (GC); pristanic acid, pu-rity >99% by GC; dimethylester pimelic acid, purity >99% by GC; methylmalonic acid, purity >99%; 2-me-thylglutaric acid, purity >98%; 3-methylglutaric acid, purity >98%; 4-methylvaleric acid, purity >99%; me-thylsuccinic acid, purity >97%; 2-phenylpropionic acid, purity >97%; 3-phenylbutyric acid, purity >98%; β-me-thylcinnamic acid, purity >99%; acetophenone, purity >99%; α-hydroxy aetophenone, purity >99%; 3-hydroxy acetophenone, purity >99%; 4-hydroxy acetophenone, purity >99%; 2,3-dihydroxy benzoic acid, purity 99%; dimethyl succinate, purity >98% were obtained from Sigma Chemical Co., Germany; sec-octylbenzene, purity >99% was obtained from Labotest OHG, Germany; tet-rahydrogeranic acid; 4,8-dimethylnonanoic acid; 2-iso-butyl-heptanedioic acid were obtained from Institute of Applied Synthetic Chemistr., Austria.
Isolation The strain was originally isolated from oil-polluted sand samples collected in the Saudi Arabian Desert by en-richment cultivation performed in 500 ml flasks con-taining 1 g sand sample with 100 ml of mineral salts medium (MSM) containing 5 g litre–1 of NH4H2PO4, 2.5 g l–1 of K2HPO4, 0.5 g l–1 of MgSO4 ⋅ 7 H2O, 0.5 g l–1 of NaCl, 0.46 g l–1 of K2SO4 [18]. This was supplemented by trace element solution [14] and 1% pristane or hexa-decane as the only source of carbon and energy. After appropriate times (3 to 7 days) of shaking at 30 °C and
180 rpm, 5 ml from the cultures were transferred to 90 ml of fresh medium and incubated under the same conditions. Identical transfers were performed five more times. Afterwards, microorganisms were obtained by plating 0.1 ml of the cultures on nutrient agar (DIFCO, Otto Nordwald, Hamburg) plates. Pure cultures were maintained on nutrient agar slants and deposited as SBUG 1472 at the strain collection of the Department of Biology of the University of Greifswald (SBUG). Among 10 different types of colonies that had grown after 3 d incubation at 30 °C, 4 were capable of using pristane as sole carbon source, only the most effective strain were used for further experiments. The strain SBUG 1472 was very effective to grow on hydrocarbons and was deposited in the strain collection SBUG and used for the following experiments.
Identification For the purpose of identification, the morphology of the strain was first analyzed under a light microscope. Polar lipids were extracted and purified by using the small-scale integrated procedure of Minnikin et al. [28]. Dried preparations were dissolved in 200 μl 2-propanol and 1–10 μl amounts were separated by HPLC without further purification and analyzed as described previ-ously [25]. Polar lipids were extracted and identified by using published procedures [29]. Fatty acid methyl-ester and mycolic acid trimethyl-silylester were prepared and analyzed as previously described [22] by using the stan-dard Microbial Identification System (MIDI Inc. Dela-ware) for automated GC analyses [34]. Numerical analy-sis of fatty acids pattern was performed according to Kämpfer and Kroppenstedt [21]. Physiological charac-terization of both strains was performed as previously described by Kämpfer et al. [20]. 16 S rRNA genes were amplified with standard PCR primer, sequenced on a Beckman Capillary sequencer and compared with the reference sequences in the ribosomal database projects [26].
Hydrocarbon biodegradation For incubation experiments, cells grown on agar plates with n-alkanes were used. The agar medium contained MSM, trace element solution and agar-agar (18 g l–1). The agar plates were supplemented with n-alkanes (C10 to C16) and pristane evaporated from a sterile filter sheet in the plate cover [24] as carbon source for 2 to 7 d at 30 °C to allow growth of cells. In case of use of n-alkanes with shorter chain lengths (C6 to C9), the inocu-lated plates were put in a larger glass with evaporating n-alkane (closed by a lid) according to method F 10 of Kreisel and Schauer [24]. Tetradecane was used as
n-alkane representive substrate to study the n-alkane degradation pathway by the isolated strain. The proce-dure was performed as the pristane degradation de-scribed below.
Pristane degradation kinetics and identification of metabolites Cultures were incubated in 500 ml Erlemeyer flasks with 50 ml MSM medium and 5 (g l–1) of pristane at 30 °C and 300 rpm. As additional controls, flasks with autoclaved cells and pristane were used. At each sam-pling period, the experiment was stopped and tetradec-ane (0.5 g l–1) was added as internal standard. Then the mixture of whole cells and supernatant was extracted three times with diethyl ether. Extracts were dried over anhydrous sodium sulfate and concentrated by rotary evaporation. The residues obtained were dissolved in hexane and analyzed by gas chromatography (GC). The cell pellet was determined with the dry weight method and the pH value was measured with the pH meter. To determinate metabolites, additional cultures with 100 ml cell suspension were used. The culture super-natant was alkalized with sodium hydroxyl (NaOH) to pH 9 and then extracted with diethyl ether. After that, the supernatant was acidified to pH 2 with hydrochlo-ric acid; extracted with diethyl ether also. Extracts were dried with anhydrous Na2SO4, filtered, and concen-trated by means of rotary evaporation. After evapora-tion of the solvent, the rests of extracts were dried un-der nitrogen, the residues were taken up in methanol and analyzed by GC and gas chromatography/ mass spec-trometry (GC/MS).The data are reported as means for three separate experiments with replicated batch cul-tures, and the standard deviation was no more than 5%.
Biotransformation experiments and analytical procedures For incubation experiments, cells grown on agar plates with tetradecane and pristane were used. Biomass from agar plates was transferred to 500 ml flasks with 100 ml of sterilized MSM supplemented with trace elements and sec-octylbenzene (0.25 ml l–1) or 2-phenyl-propionic acid (0.05 ml l–1) or 3-phenylbutyric acid (0.05 ml l–1). For controls we used cells in MSM without substrate and only the substrate in MSM without cells. At intervals during the incubation, 1 ml of cell suspen-sion was removed aseptically and filtered to remove the biomass. Then 100 μl aliquots of supernatant were analyzed by high-performance liquid chromatography (HPLC) for biotransformation products. Experiments were replicated three times and the standard deviation of each product concentration was no more than 5 %.
To detect metabolites for further characterization, larger amounts of cell suspensions were used. For that purpose, ten 500 ml Erlenmeyer flasks containing 100 ml cell suspension with different substrates were incubated as mentioned before. The supernatants were extracted three times with an equal volume of ethyl acetate with pH 7 and pH 2 as the same manner de-scribed above. The residues obtained were dissolved in methanol and analyzed by HPLC and GC/MS for detec-tion and characterization of transformation products.
Chemical analysis GC analyses. A gas chromatograph (Shimadzu GC – 14A, Duisburg) with a flame-ionisation detector (FID) was used for the retention time data. Chromatographic operating conditions: DB-1 ms (J&W Scientific, U.S.A.), 0.25 mm × 30 m, ID capillary glass column coated with OV1 (methyl – silicon), 0.25 μm thick layer film, adhe-sion – preventing film: dimethylpolysiloxan 100%; temperature programmed: 5 min 40 °C, 40–60 °C, 1 °C min–1, 60–300 °C, 10 °C min–1; carrier gas flow rate (He), 2 ml min–1; detector temperature, 310 °C, injector temperature, 250 °C. For GC/MS analyses, a coupled system consisting of a GC 8000 gas chromatograph (Fisons Instruments, Mainz, Germany) equipped with a 30-m DB5-ms column (0.25-mm-by 0.33-μm film; J&W Scientific, USA) and a mass selective detector MD 800 (Fisons Instruments) operating at 70 eV or a TSQ 700 (Finnigan Corp., San Jose, Calif.) triple quadrupole mass spectrometer oper-ated in a single quadrupole mode (Q1) was used. Sepa-ration on the column was achieved by using a tempera-ture programm from 60 to 290 oC (10 oC/min). Acid extracts were derivatized by methylation with diazome-thane [12]. HPLC was performed on a Hewlett-Packard (Bad Homburg, Germany) HPLC apparatus 1050 M equipped with a quaternary pump system, a diode array detector 1040 M series I, and an HP Chemstation. The separation was achieved with a LiChroCart 125-4 RP-18 end-capped (5-μm) column (Merck, Darmstadt, Germany). Elution profile is characterized by an initial solvent composi-tion of 30% methanol-70% phosphoric acid (0.1%), reaching 100% methanol after 14 min at a flow rate of 1 ml min–1.
Results
Isolation and identification of SBUG 1472 With pristane as the sole carbon and energy source, we isolated an actinomycete strain (SBUG 1472) from soil
for its pristane-degrading. The SBUG 1472 was deter-mined by the Deutsche Sammlung von Mikroorganis-men und Zellkulturen (DSMZ) to be a novel strain be-longing to the species Nocardia cyriacigeorgica [44]. Con-form to the description of the genus Nocardia [16, 17], colonies of SBUG 1472 showed a perl white aerial mycel (RL 1013) and an ivory colored (RAL 1014) substrate mycel. Only minor differences could be observed when the strain was grown on different kinds of carbons, as well as compared to typus strain (data not shown). My-colic acids had a length of 50–58 carbon atoms, with a pattern very similar of the one known from N. otitidis-caviarum (DSM 43010), but clearly different from the C46–C54 pattern known from N. cyriacigeorgica. Different pathways and precursors in the synthesis of mycolic acids in the oil-containing habitat of SBUG 472 and the clinical environment of DSM 44484 might explain these striking differences between the two closely related strains. The pattern of unbranched, saturated and un-saturated fatty acids as well as tuberculostearic acid is diagnostic for Nocardia and close relatives, Mycobacte-rium, Rhodococcus and Gordonia. Qualitative and quanti-tative analysis of the fatty acid pattern allowed the identification of SBUG 1472 as a member of the Nocar-dia (data not shown). Based on the analysis of the com-plete 1513 nucleotides (nts) 16S rRNA sequence, strain SBUG 1472 shares over 99.9% sequence identity with the type strain of N. cyriacigeorgica (DSM 44484, AF430027; two transitions difference at positions 3 and 1494), but significantly less sequence similarity with other type strains of the genus: N. abscessus (AF218292, 98.9% sequence identity, 16 nts difference), N. pau-civorans (AF430041, 98.5% sequence identity, 22 nts difference), N. asteroids (AF430019, 98.3% sequence iden-tity, 25 nts difference), N. carneae (AF430035, 98.3% sequence identity, 26 nts difference), N. farcinica, (Z36936, 98.1% sequence identity, 28 nts difference), N. nova (AF430028, 97.9% sequence identity, 30 nts difference), N. veterana (AF430055, 97.1% sequence iden-tity, 43 nt difference), and N. abobensis, (AB126876, 97.1% sequence identity, 44 nts difference), allowing a reliable identification of SBUG 1472 as a strain belong-ing to the species N. cyriacigeorgica.
n-Alkane growth range and metabolism n-Alkane degradation studies were performed by incu-bating the bacterium at 30 °C with n-alkanes (ranging in their chain length from C6 to C16). By observation, the strain has grown quickly on longer n-alkanes (from C12 to C16), after 1 day and slowly on shorter n-alkanes (C6 to C11), after 3 d (Table 1). Additionally, the cells were cultured on nutrient agar at different tempera-
(–: no growth; +: good growth; + +: very good growth; +++: excellent growth)
tures (20–50 °C) with different pH values (5.5–8.0). The optimum growth temperature of the strain is 27 °C (+/–3 grd) and the highest growth temperature was 45 °C; the pH optimum for growth was 6.3. With tetradecane as substrate, GC/MS analysis of the cell-free extracts revealed four peaks at 25.6 min, 24.9 min, 22.5 min and 18.8 min. By comparison, the retention times and the MS data with the data of com-mercial standards, we also determined the metabolites as tetradecanoic acid, 1-tetradecanol and decanoic acid, respectively (Table 2). The identity of the retention times and fragment peak distributions allow the con-clusion that tetradecanol, tetradecanoic acid, dode-canoic acid and decanoic acid were the main metabo-lites during the tetradecane degradation process.
Growth on pristane and kinetics of degradation Under aerobic conditions, the isolate was incubated with 0.01% and 0.5% of pristane as carbon and energy source. After using a concentration of 0.01% (v/v), the bacterium degraded up to 90% after 8 h; and with a concentration of 0.5% (v/v) after 3 weeks more than 84% of pristane were degraded. For kinetic studies of pristane degradation, a substrate concentration of 0.5% (v/v) was used. The typical pattern of growth (Fig. 1) showed an increase in cell concentration (biomass) associated with a strong decrease in the carbon source pristane and the pH value. Alternatively, the initial concentration of pristane decreased only 0–2% in the control flask without cells and in control flask with autoclaved cells the concentration of pristane decreased about 0–2% also. Under the used growth conditions, the lag phase was not observed and the degradation seems to start immediately.
Detection of metabolites from pristane by GC/MS During degradation experiments four major metabo-lites were observed by GC/MS analysis in the medium extracts (pH 2.0), identified as methyl ester derivatives (Table 2). The first product, which eluted at 29.7 min, had the same retention time as a sample of the com-pound pristanoic acid which is commercially available. Additionally, the mass spectrum of the product was virtually identical to that of pristanoic acid. The second product was detected at 19.2 min. In comparison with available standard, this product was confirmed as 4,8-dimethylnonanoic acid which has retention time at 19.1 min. The third methylated compound with a GC/MS retention time of 20.4 min, contained a base peak m/z 88 [(CH3)CH2COOCH3] and fragment ions at m/z 129 [(CH2)3CH(CH3)COOCH3]; 101 [–CH2CH(CH3)COOCH3]; 57 [CH3CH(CH3)CH3] and 43 [CH3CH2(CH3)]. These results were consistent with the metabolite 2,6-dimethylhep-tanoic acid methylester. On the other hand, the fourth metabolite eluted at 15.3 min had the similar retention time and mass spectrum as the commercial standard 2-methylglutaric acid (MW 174). Thus, all of these me-tabolites could be identified with the help of purchased chemical standards or from the NIST library.
Biotransformation of aromatic hydrocarbons Aromatic hydrocarbons in crude oil occur mainly as al-kylbenzenes. Therefore, we used N. cyriacigeorgica, which was precultured on tetradecane or pristane to trans-form several alkylbenzenes. In these experiments we tested the oxidation of alkylbenzenes with a branched side chain such as iso-pentylbenzene, sec-hexylbenzene or sec-octylbenzene (data are not shown in this publication). Though the isolate was shown to have capacities of bio-transformation of all these compounds, in this report, we only focused on the transformation of sec-octyl-benzene. We found that sec-octylbenzene was transform-ed by tetradecane- as well as pristane- precultivated cells, and the metabolites generated from sec-octylbenzene were characterized by using aqueous samples of the cul-ture supernatant and analyzing with HPLC. sec-Octylben-zene was transformed to a series of acidic metabolites, designated as products I, II, III and IV which had reten-tion times as 7.9, 9.1, 9.6, and 11.4 min, respectively. All of these products were identified by comparison of re-tention times and UV/Vis spectra of the HPLC elution profile as well as retention time and mass spectra of the GC/MS spectroscopy to those of veritable standards (Table 3). As a result of these comparisons, the metaboli-tes I, II, III and IV could be identified as 2-phenylpropio-nic acid (I), 3-phenylbutyric acid (II), β-methylcinnamic acid (III) and 5-phenylhexanoic acid (IV), respectively.
6 Le Thi Nhi-Cong et al. Journal of Basic Microbiology 2010, 50, 1–13
Figure 1. Pristane degradation (◊), kinetic growth (■) and pH value decreasing (▲) of N. cyriacigeorgica SBUG 1472 during the culti-vation with pristane (data are shown as the means ± SD of three independent cultures).
Because of the limited sensitivity of analyzing aque-ous samples by HPLC, the culture supernatant without cells were extracted at pH 9, 7 and 2, and then they were concentrated. In addition to the four upper prod-ucts, the neutral and alkaline extract analysis by HPLC and GC/MS revealed one previously undetectable com-pound, called as product V. Furthermore, another product, designated as product VI, was also identified by GC/MS analysis. In the acidic extract two others, named as VII and VIII were determined. Product V detectable in neutral and alkaline extracts pointed to a metabolite containing a ketone or an alco-hol group. The absolute identity of the UV spectrum and retention time in analysis by HPLC between prod-uct V and available standard led to the assumption that product V is acetophenone. Moreover, the mass spec-trum showed a base peak ion at m/z 120, with fragment ions at m/z 105 [C6H5 + CCH3/ + H] or 105 [C6H5 + CO], and 77 [C6H5]. By comparing mass spectra and retention time by GC/MS analysis, we confirmed the structure of acetophenone for product V. Metabolite VI was detected at a retention time (tR) of 17.2 min. According to the analytical data in the library NIST, it was suggested as α-hydroxyacetophenone. By analyzing the available standard, a tR of 16.9 min was observed. Results from comparison of the mass spectra fragments showed that metabolite VI had fragments at m/z 105 (100.0%), 91 (24.07%), 77 (56.82 %), and 51 (17.25%), meanwhile the standard had fragments at m/z 105 (100.00%), 91 (20.90%) 77 (72.55%), and 51 (23.18%). Therefore, we confirmed this metabolite as α-hydroxyacetophenone. Metabolite VII was found in very small amounts at a tR of 21.7 min by GC/MS analysis of the acid extract but was not detectable by HPLC analysis. The mass spec-trum of this compound revealed a molecular ion peak
at m/z 196 (61.63%) and contained the characteristic fragment ions m/z 167 (18.22%), 166 (51.84%), 165 (79.21%), 164 (10.81%), 163 (100.00%), 149 (28.44%), 135 (24.40%), 125 (10.27%), 123 (12.16%), 122 (37.57%), 121 (16.11%), 108 (12.53%),107 (52.80%), 92 (15.99%), 79 (26.71%), 77 (42.62%), 76 (11.85%), 65 (22.63%), 63 (14.03%), 53 (19.71%), 51 (33.86%), 50 (16.26%), and 45 (30.09%). Compound VII was confirmed as 2,3-dihyd-roxybenzoic acid by matching with the retention time and the mass spectra of the available standard which was characterized by a tR of 21.6 min and fragments ion at ions m/z 196 (62.24%), 167 (16.62 %), 166 (45.84%), 165 (80.26%), 164 (11.42%), 163 (100.00%), 149 (29.95%), 135 (27.59%), 125 (9.60%), 123 (11.26%), 122 (36.04%), 121 (17.11%), 108 (10.26%),107 (46.07%), 92 (16.19%), 79 (20.21%), 77 (40.65%), 76 (10.68%), 65 (20.64%), 63 (12.07%), 53 (14.55%), 51 (32.23%), 50 (14.81%), and 45 (30.81%). Product VIII was also found in trace amounts and was only detected by GC/MS. The tR lay at 9.2 min and fragmentation patterns of the compound were at m/z 120 (100.00%), 114 (26.28%), 87 (22.75%), 59 (59.52%), 55 (78.22%) and 45 (8.91%). By comparison with suc-cinic acid which had also a tR at 9.2 min and fragments ions at m/z 120 (100.00%), 114 (31.42%), 87 (22.46%), 59 (58.80%), 55 (80.06%) and 45 (10.37%), the data led to the identification of this intermediate as succinic acid.
Transformation kinetics of sec-octylbenzene The substrate sec-octylbenzene is insoluble in water; thereby it could not be detected in the aqueous culture supernatant by HPLC analysis. The main products were formed rapidly after 4 h incubation and represent phenylalkanoic acids with shorter branched side chain (Fig. 2). The quantity of 3-phenylbutyric acid at each measuring time was always higher than that of other occurring acids. Three other products (V, VI and VII) were not detectable in the aqueous supernatant by HPLC analysis. However, they were identified by GC/MS analysis of the extracts. It seems, therefore, these prod-ucts accumulated in only small amounts or in a very late incubation period.
Transformation of formed intermediate products 2-Phenylpropionic acid, 3-phenylbutyric acid, β-methyl-cinnamic acid and acetophenone were used as sub-strates in the biotransformation experiments of tetra-decane-precultivated cells of N. cyriacigeorgica to search for the ring cleavage products and to complete the pathway of sec-octylbenzene transformation. The in-termediates were analyzed by comparing the retention times and UV/Vis spectra of the HPLC elution profile as
8 Le Thi Nhi-Cong et al. Journal of Basic Microbiology 2010, 50, 1–13
Figure 2. Kinetics of the transformation of sec-octylbenzene (0.025%) by N. cyriacigeorgica SBUG 1472 as determined by HPLC ♦, β-methylcinnamic acid; o, 5-phenylhexanoic acid; x, 2-phenylpropionic acid; Δ, 3-phenylbutyric acid. Data are shown as the means ± SD of three independent cultures.
well as the retention times and mass spectra of the GC/MS spectroscopy with those of available standards. However, N. cyriacigeorgica could not transform 2-phe-nylpropionic acid. The 3-phenylbutyric acid was dehy-drogenized after 2 h incubation to produce β-methyl-cinnamic acid; after 72 h 34% of the substrate was transformed to β-methylcinnamic acid. A small number of acetophenone was also found in the supernatant. With β-methylcinnamic acid as a substrate, acetophe-none was obtained as the major metabolite, and 2,3-dihydroxybenzoic acid was detected in the organic ex-tract. When acetophenone was used as a substrate, in ad-dition to α-hydroxyphenone; 2,3-dihydroxybenzoic acid and succinic acid were identified by GC/MS analysis.
Discussion
In this study, we reported on a new isolated strain, named SBUG 1472 with high capabilities of degrading and transforming several types of hydrocarbons. Ac-cording to analysis of the complete 1513 nucleotides 16S rRNA sequence of strain SBUG 1472, morphology, physiology and chemotaxonomy, the bacterium was identified as a novel strain within the species Nocardia cyriacigeorgica (formerly named N. cyriacigeorgici) [44]. Though most of these strains belonging to N. cyria-cigeorgica have a record as pulmonary pathogens in studies performed in Germany, France, Japan, Thailand and the US [3, 7, 10, 19, 36, 43, 44], a set of soil bacteria recently described by Shrestha et al. (2007) is located next to the N. cyriacigeorgica cluster (sharing 99% se-
quence similarity), even closer then N. abobensis, N. far-cinica, N. nova and N. veteran, indicating the close rela-tionship of environmental as well as medically im-portant strains. Moreover, the genus Nocardia belongs to non-sporulating actinomycetes, which exhibit a high capacity to transform and degrade diverse classes of hydrocarbons, including a variety of aliphatics, aromat-ics and other xenobiotic pollutants [2, 42]. In our studies to investigate the capability of N. cyria-cigeorgica to utilize hydrocarbons such as aliphatics and aromatics, the determined metabolites suggested that tetradecane were converted via tetradecanol to tet-radecanoic acid. This pointed to a monoterminal oxida-tion pathway of n-alkanes. Di-terminal and sub-termi-nal oxidation products were not found under the given experimental conditions. The capacity of SBUG 1472 to utilize n-alkanes as energy and carbon source suggested that N. cyriacigeorgica degraded longer-chain alkanes (C12 to C16) more efficiently than the shorter-chain (C6 to C11), which is a common feature of many other alkane-utilizing microorganisms [41]. However, hitherto there are no reports on n-alkane degradation by N. cyriacigeor-gica. The isolate SBUG 1472 was not only able to use straight alkanes effectively but it also grew well on branched alkanes such as pristane. This property of utilizing pristane is a remarkable ability because many n-alkane degrading microorganisms are not capable of using branched chain hydrocarbons as carbon source. GC analysis showed that when 0.1 (g l–1) of pristane was used, 92% of it was degraded after 8 hours; and when 5 (g l–1) of pristane was used, 84% was degraded after 3 weeks. The detection of potential metabolic interme-diates of pristane catabolism from the mono- and diter-minal pathways by GC/MS during the growth of N. cyria-cigeorgica on pristane indicated that the strain oxidized pristane by both terminal and diterminal oxidation mechanisms (Fig. 3). During mono-terminal oxidation of pristane followed by β-oxidation of pristanoic acid, the product 4,8-dimethylnonanoic acid and 2,6-dime-thylheptanoic acid were accumulated. In the course of di-terminal oxidation, 2-methylpentanoic acid was formed and excreted. Pirnik et al. [30] reasoned that beside mono-terminal oxidation the di-terminal oxida-tion pathway was a significant branch of the pristane degradation pathway in Brevibacterium erythrogenes. Fur-thermore these metabolic pathways have been also reported in Nocardia globerula [2] and in some unidenti-fied species of the genera Corynebacterium [27], Mycobac-terium [31], and Rhodococcus [23]. The oxidation of sec-octylbenzene (2-phenyloctane) in N. cyriacigeorgica started from the alkyl side chain. The
Figure 3. Proposed pathways for the degradation of pristane by N.cyriacigeorgica SBUG 1472, (N) are the isolated metabolites.
detectable intermediate was 5-phenylhexanoic acid, which was further degraded by β-oxidation to 3-phenyl-butyric acid. This oxidation cascade starting at the ter-minal part of the alkyl chain of sec-octylbenzene is not surprisingly; because N. cyriacigeorgica was able to oxi-
dize straight and branched chain alkanes very quickly at terminal positions. Also, N. salmonicolor degrades 1-phenyldecane and 1-phenylnonane primarily at the terminal alkyl chain yielding phenylacetic acid and benzoic acid, respectively [32] and N. amarae oxidizes
Figure 4. Proposed pathways for the degradation of sec-octylbenzene by N. cyriacigeorgica SBUG 1472, (N) are the isolated metabolites.
Journal of Basic Microbiology 2010, 50, 1–13 Hydrocarbonoxidation by Nocardia cyriacigeorgica 11
2-phenyheptane and 4-phenylheptane at terminal posi-tions too [9]. In the latter case, no hints of a cleavage of the benzene ring could be found. Similar acids were formed with two unidentified Nocardia strains, forming from 3-phenyldodecane, 3-phenylhexanoic acid and the odd carbon chain compounds 3-phenylvaleric acid and 5-phenylheptanoic acid [6]. However, further degrad-able products were not analyzed. Alternatively, Pseudo-monas putida and Pseudomonas acidovorans oxidize 2-phe-nylbutane exclusively at the aromatic ring system lead-ing to alkylated ring splitting products [6]. Of greater interest is the catabolic pathway of the formed aromatic acids with short side chains. In our experiments, N. cyriacigeorgica formed mainly 3-phenyl-butyric acid as an intermediate. This acid was further dehydrogenated to ß-methylcinnamic acid. The branch-ing point at position 3 of this acid may hinder normal β-oxidation. N. cyriacigeorgica, however, is generally able to overcome branching points, as seen by growth on pristane (see above) and can shorten the β-methyl-cinnamic acid by splitting off a C2-unit to form aceto-phenone. Acetophenone is transformed to benzoic acid and may be simultaneously oxidized at the aromatic ring to form finally 2,3-dihydroxybenzoic acid which undergoes ring opening. This pathway for oxidation of 3-phenylbutyric acid is new and differs from degrada-tion mechanisms in other microorganisms. For in-stance, Pseudomonas sp. and Rhodococcus rhodochrous started the degradation of 3-phenylbutyric acid with a demethylation step to yield 3-phenylpropionic acid [33, 38], which is further degraded to benzoic acid. In a study on the metabolism of 1-phenylethanol and acetophenone by a Nocardia and an Arthrobacter species, Cripps et al. demonstrated that acetophenone was me-tabolized via phenylacetate, phenol and catechol [11]. However, in our case, to elucidate acetophenone trans-formation, we assumed that acetophenone was trans-formed to benzoic acid. In parallel hydroxylation of the aromatic ring occur. Because 2,3-dihydroxybenzoic acid and succinic acid were detected, it may be inferred that benzoic acid was degraded via ortho-cleavage. Transfor-mation experiments with benzoic acid were performed. All of the substrate was degraded but no metabolite was observed after 8 h incubation. This suggested that benzoic acid was used completely as energy and carbon source for the cell. The metabolite 2-phenylpropionic acid was predicted to be the product of sub-terminal oxidation However, there is no confirmation by other metabolites involved in sub-terminal pathway of sec-octylbenzene. In conclusion, our study demonstrated for the first time that a strain assigned to the species N. cyriacigeor-
gica was able to grow and degrade a wide range of n-alkanes and also multiple branched-chain alkane such as pristane. Furthermore, this strain was also able to transform aromatic hydrocarbons with branched-side chains. With the results presented here N. cyriacigeor-gica may be useful as a potential species for hydrocar-bon degradation and it may be of interest to prove whether this strain is of importance during the re-moval of specific environmental pollutants in tropical soils contaminated with alkyl-branched compounds.
Acknowledgement
We thank P. Bednarski (Institute of Pharmacy, Univer-sity of Greifswald) for help in preparing the manu-script.
References
[1] Abed, R.M., Safi, N.M., Koster, J., de Beer, D., El-Nahhal, Y., Rullkotter, J. and Garcia-Pichel F., 2002. Microbial di-versity of a heavily polluted microbial mat and its com-munity changes following degradation of petroleum com-pounds. Appl. Environ. Microbiol., 68, 1674–83.
[2] Alvarez, H.M., Souto, M.F., Viale, A. and Pucci, O.H., 2001. Biosynthesis of fatty acids and triacylglycerols by 2,6,10,14-tetramethyl pentadecane-grown cells of Nocardia globerula 432. FEMS Microbiol. Lett., 200, 195–200.
[3] Akcaglar, S., Yilmaz, E., Heper, Y., Alver, O., Akalin, H., Ener, B., Tore, O., Ersoy, C. and Imamoglu, S., 2008. No-cardia cyriacigeorgica: pulmonary infection in a patient with Basedow-Graves disease and a short review of report-ed cases. Int. J. Infect. Dis., 12, 335–8.
[4] Atlas, R.M., 1975. Effects of temperature and crude oil composition on petroleum biodegradation. Appl. Micro-biol., 30, 396–403.
[5] Awe, S., Mikolasch, A., Hammer, E. and Schauer, F., 2008. Degradation of phenylalkanes and characterization of aromatic intermediates acting as growth inhibiting sub-stances in hydrocarbon utilizing yeast Candida maltosa. Int Biodeterio. and Biodegrad. DOI: 10.1016:1-7.
[6] Baggi, G., Catelani, D., Galli, E. and Treccani, V., 1972. The microbial degradation of phenylalkanes. 2-Phenylbutane, 3-phenylpentane, 3-phenyldodecane and 4-phenylhepta-ne. Biochem. J., 126, 1091–1097.
[7] Beaman. B.L. and Tam, S., 2008. An unusual murine be-havior following infection with log-phase Nocardia aster-oides type 6 strain GUH-2 (Nocardia cyriacigeorgica GUH-2). Microbes Infect., 10, 840–843.
[8] Berekaa, M.M. and Steinbüchel, A., 2000. Microbial deg-radation of the multiply branched alkane 2,6,10,15,19, 23-hexamethyltetracosane (Squalane) by Mycobacterium fortui-tum and Mycobacterium ratisbonense. Appl. Environ. Micro-biol., 66, 4462–4467.
[9] Bhatia, M. and Singh, D.H., 1996. Biodegradation of com-mercial linear alkyl benzenes by Nocardia amarae. J. Bio-sci., 21, 487–496.
[10] Conville, P.S. and Witebsky, F.G., 2007. Organisms de-signated as Nocardia asteroides drug pattern type VI are members of the species Nocardia cyriacigeorgica. J. Clin. Mi-crobiol., 45, 2257–2259.
[11] Cripps, R.E., Trudgill, P.W. and Whateley, J.G., 1978. The metabolism of 1-phenylethanol and acetophenone by No-cardia T5 and an Arthrobacter species. Eur. J. Biochem., 86, 175–186.
[12] De Boe, T.D. and Backer, H.J., 1956. Diazonethane. Org. Synth., 36, 14–16.
[13] Elsayed, S., Kealey, A., Coffin, C.S., Read, R., Megran, D. and Zhang, K., 2006. Nocardia cyriacigeorgica septicemia. J. Clin. Microbiol., 44, 280–282.
[14] Fritsche, W., 1968. Der Einfluβ der Kohlenstoffquelle auf das Wachstum, den Proteingehalt und das Enzymmuster von Candida guilliermondii. Z. Allg. Mikrobiol., 8, 91–99.
[15] Fux, C., Bodmer, T., Ziswiler, H.R. and Leib, S.L., 2003. Nocardia cyriacigeogici: first report of invasive human in-faction. Dtsch. Med. Wochenschr., 128, 1038–1041.
[16] Goodfellow, M., Lechevalier, M.P., 1987. Genus Nocardia Trevisian 1898. In: Bergey's Manual of Systematic Bacteri-ology. Williams & Wilkinson, Baltimore, 2, 1459–1465.
[17] Gordon, R.E. and Mihm, J.M., 1962. Identification of No-cardia caviae (Erikson) nov. comb. Ann. NY. Acad. Sci., 98, 628–636.
[18] Hundt, K., Wagner, M., Becher, D., Hammer, E. and Schauer, F., 1998 Effect of selected environmental factors on degradation and mineralization of biaryl compounds by the bacterium Ralstonia pickettii in soil and compost. Chemosphere, 36, 2321–2335.
[19] Kageyama, A., Hoshino, Y., Yazawa, K., Poonwan, N., Takeshita, N., Maki, S. and Mikami, Y., 2005. Nocardia cyri-acigeorgica is a significant pathogen responsible for nocar-diosis in Japan and Thailand. Mycopathologia, 160, 15–19.
[20] Kämpfer, P., Dott, W. and Kroppenstedt, R.M., 1990. Nu-merical classification and identification of some nocardio-form bacteria. J. Gen. Microbbiol., 36, 309–331.
[21] Kämpfer, P. and Kroppenstedt, R.M., 1996. Numerical analysis of fatty acid patterns of coryneform bacteria and related taxa. Can. J. Microbiol., 42, 989–1005.
[23] Koma D., Kubota, K., Fujii, Y., Hasumi, F., Chung, S.Y. and Kubo, M., 2003. Degradation of car engine base oil by Rho-dococcus sp. NDKK48 and Gordonia sp. NDKY76A, 67, 1590–1593.
[24] Kreisel, H. and Schauer, F., 1987. Methoden des mykologi-schen Laboratoriums. Gustav Fischer Verlag, Stuttgart, Germany, 103–104.
[25] Kroppenstedt, R.M., 1985. Fatty acid and menaquinone analysis of actinomycetes and related organisms. In:
Chemical Methods in Bacterial Systematics (M. Goodfel-low and D.E. Minnikin, eds.). London: Academic Press. SAB Technical Series, 20, 173–199.
[27] McKenna, E.J. and Kallio, R.E., 1971. Microbial metabo-lism of the isoprenoid alkane pristane. Proc. Natl. Acad. Sci. USA., 68, 1552–1154.
[28] Minnikin, D.E., O’Donnell, A.G., Goodfellow, M., Alder-son, A., Athalye, M., Schaal, K.P. and Parlett, J.H., 1984. An integrated procedure for the extraction of isoprenoid quinones and polar lipids. J. Microbiol. Methods, 2, 233–241.
[29] Minnikin, D.E., Patel, V., Alshamaony, L. and Goodfellow, M., 1977. Polar lipid composition in the classification of Nocardia and related bacteria. Int. J. Syst. Bacteriol., 27, 104–117.
[30] Pirnik, M.P., Atlas, R.M. and Bartha, R., 1974. Hydrocar-bon metabolism by Brevibacterium erythrogenes: normal and branched alkanes. J. Bacteriol., 119, 868–878.
[31] Sakai, Y., Takahashi, H., Wakasa, Y., Kotani, T., Yurimoto, H., Miyachi, N., Van Veldhoven, P. P. and Kato N., 2004. Role of alpha-methylacyl coenzyme A racemase in the degradation of methyl-branched alkanes by Mycobacterium sp. strain P101. J. Bacteriol., 186, 7214–7220.
[32] Sariaslani, F.S., Harper, D.B. and Higgins, I.J., 1974. Mic-robial degradation of hydrocarbons. Catabolism of 1-phenylalkanes by Nocardia salmonicolor. Biochem. J., 140, 31–45.
[33] Sariaslani, F.S., Sudmeier, J.L. and Focht, D.D., 1982. De-gradation of 3-phenylbutyric acid by Pseudomonas sp. J. Bacteriol., 152, 411–421.
[34] Sasser, M., 1990. Identification of bacteria by gas chroma-tography of cellular fatty acids. USFCC Newsletter, 20, 1–6.
[35] Sayavedra-Soto, L.A., Chang, W.N., Lin, T.K., Ho, C.L. and Liu, H.S., 2006. Alkane utilization by Rhodococcus strain NTU-1 alone and in its natural association with Bacillus fu-siformis L-1 and Ochrobactrum sp. Biotechnol. Prog., 22, 1368–1373.
[36] Schlaberg, R., Huard, R.C., Della-Latta, P., 2008. Nocardia cyriacigeorgica, an emerging pathogen in the United States. J. Clin. Microbiol., 46, 265–273.
[37] Shrestha, P.M., Noll, M. and Liesack, W., 2007. Phyloge-netic identity, growth-response time and rRNA operon copy number of soil bacteria indicate different stages of community succession. Environ. Microbiol., 9, 2464–2474.
[38] Simoni, S., Klinke, S., Zipper, C., Angst, W. and Kohler H.P., 1996. Enantioselective metabolism of chiral 3-phe-nylbutyric acid, an intermediate of linear alkylbenzene degradation, by Rhodococcus rhodochrous PB1. Appl. Envi-ron. Microbiol., 62, 749–755.
[39] Smith, M.R., 1990. The biodegradation of aromatic hydro-carbons by bacteria. Biodegrad., 1, 191–206.
[40] Stephens, G.M. and Dalton, H., 1987. Is toxin production by coryneform bacteria linked to their ability to utilize hydrocarbons? Tibtech., 5, 5–7.
[41] van Beilen, J.B. and Witholt, B., 1994. Genetics of alkane oxidation by Pseudomonas oleovorans, 3–4, 161–74.
[42] Webley, D.M., Duff, R.B. and Farmer, V.C., 1956. Evidence for β-oxidation in the metabolism of saturated aliphatic hydrocarbons by soil species of Nocardia. Nature, 178, 1467–1468.
[43] Witebsky, F.G., Conville, P.S., Wallace, R.J. jr, Brown-Elliott, B.A., Huard, R.C., Schlaberg, R. and Della-Latta, P., 2008. Nocardia cyriacigeorgica - an established rather than an emerging pathogen. J. Clin. Microbiol., 46, 2469.
[44] Yassin, A.F., Rainey, F.A. and Steiner, U., 2001. Nocardia cyriacigeorgici sp. nov. Int. J. Syst. Evol. Microbiol., 51, 1419–1423.
