Identification of phenyldecanoic acid as a constituent of triacylglycerols and wax ester produced by Rhodococcus opacus PD630

Héctor M. Alvarez1, Heinrich Luftmann2, Roxana A. Silva1, Ana C. Cesari1, Alberto Viale3, Marc Wältermann4 and Alexander Steinbüchel4

Departamento de Bioquímica, Facultad de Ciencias Naturales, Universidad Nacional de la Patagonia San Juan Bosco, CC 1078, Km 4, 9000 Comodoro Rivadavia, Chubut, Argentina1
Institut für Organische Chemie der Westfälischen Wilhelms-Universität Münster, Corrensstraße 40, D-48149 Münster, Germany2
Departamento de Química Biológica, Facultad de Ciencias Exactas y Naturales, Universidad Nacional de Buenos Aires, Buenos Aires, Argentina3
Institut für Mikrobiologie der Westfälischen Wilhelms-Universität Münster, Corrensstraße 3, D-48149 Münster, Germany4

Author for correspondence: Héctor M. Alvarez. Tel: +54 297 4550 339. Fax: +54 297 4550 339. e-mail: halvarez{at}unpata.edu.ar


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Phenyldecane supported growth and lipid accumulation of Rhodococcus opacus PD630 during cultivation under nitrogen-limiting conditions. The results of this study suggested that the hydrocarbon phenyldecane was degraded by monoterminal oxidation, followed by ß-oxidation of the alkyl side-chain to phenylacetic acid, and by an additional degradative route for the oxidation of the latter to intermediates of the central metabolism. {alpha}-Oxidation of phenyldecanoic acid also occurred to some extent. Phenyldecanoic acid, the monoterminal oxidation product, was also utilized for the biosynthesis of a novel wax ester and novel triacylglycerols. The formation of the wax ester phenyldecylphenyldecanoate probably resulted from the condensation of phenyldecanoic acid and phenyldecanol, which were produced as metabolites during the catabolism of phenyldecane. Two types of triacylglycerol were detected in phenyldecane-grown cells of strain PD630. Triacylglycerols containing only odd- and even-numbered aliphatic fatty acids, as well as triacylglycerols in which one fatty acid was replaced by a phenyldecanoic acid residue, occurred. Other phenyl intermediates, such as phenylacetic acid, phenylpropionic acid, 4-hydroxyphenylpropionic acid, protocatechuate and homogentisic acid, were excreted into the medium during cultivation on phenyldecane. On the basis of the results obtained, pathways for the catabolism and assimilation of phenyldecane by R. opacus PD630 are discussed.

Keywords: Rhodococcus opacus PD630, phenyldecane, triacylglycerol, wax ester, phenyldecylphenyldecanoate

Abbreviations: ESI-MS, electron spray ionization mass spectrometry; TAG, triacylglycerol


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Phenylalkanes consisting of a phenyl group substituted with an alkyl residue of variable chain length are common components of crude oil and detergents (Sikkema et al., 1995 ). These compounds offer two alternative pathways for degradation: (1) oxidative attack at the aromatic ring, resulting in the formation of alkyl-catechols, and (2) oxidation of the alkyl side-chain, resulting in the formation of aromatic fatty acids. These compounds are then oxidized to dihydroxylated aromatic derivatives, which are further degraded by ring fission (Gibson & Subramanian, 1984 ). The oxidation of phenylalkanes and phenyalkanoic acids has been reported for different bacteria, such as species belonging to the genera Nocardia (Sariaslani et al., 1974 ), Rhodococcus (Warhust & Fewson, 1994 ; Alvarez et al., 1996 ) and Pseudomonas (García et al., 1999 ). According to García et al. (1999) , all bacteria able to assimilate phenylacetic acid or benzoic acid via a CoA intermediate would be able to utilize phenylalkanoic acids as sole carbon source, since phenylacetic acid is a ß-oxidation product from the biodegradation of phenylalkanes possessing an alkyl side-chain with an even number of carbon atoms.

Phenyldecane-grown cells of Rhodococcus opacus PD630, which were also able to catabolize phenylacetic acid, intracellularly accumulated oxidation products of the substrate, with phenyldecanoic acid as the major compound and also lesser amounts of phenyloctanoic, phenylhexanoic, phenylnonanoic acids and diacylglycerols (Alvarez et al., 1996 ). Members of the genus Rhodococcus are widely distributed in soil environments. These micro-organisms show a broad capacity and metabolic spectrum for the biodegradation of different kinds of pollutants, such as hydrocarbons, herbicides or other xenobiotic compounds (Finnerty, 1992 ; Warhust & Fewson, 1994 ). In addition, recent studies demonstrated that these bacteria are able to accumulate triacylglycerols (TAGs) from different carbon sources, including hydrocarbons, during cultivation under nitrogen-starvation conditions (Alvarez et al., 1996 , 1997 , 2000 ). In this context, the biodegradation and production of cellular lipids by Nocardia globerula 432 from the recalcitrant branched hydrocarbon pristane under conditions restricting growth has been reported recently (Alvarez et al., 2001 ). These results suggest that these Gram-positive bacteria may be employed to eliminate pollutants from the environment under a broad range of metabolic and environmental conditions. Several reports considered the potential of these bacteria for in situ bioremediation of contaminated environments (Yakimov et al., 1999 ; Wagner-Dobler et al., 1998 ; White et al., 1998 ).

On account of the ability of R. opacus PD630 to accumulate metabolites and lipids under unbalanced growth conditions, we investigated in this study the catabolism and assimilation of derivatives of phenyldecane in this strain.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacterial strain, media and growth conditions.
R. opacus strain PD630 (DSM 44193) was isolated and characterized in a previous study (Alvarez et al., 1996 ). Cells were grown aerobically at 25 °C in nutrient broth medium (0·8%, w/v) or in mineral salts medium, according to Schlegel et al. (1961) , with the carbon source indicated in the text. To promote accumulation of lipids, the concentration of ammonium chloride in the mineral salts medium was reduced to 0·05 g l-1. Solidified media were prepared by adding 1·8% (w/v) agar. Phenyldecane was obtained from Sigma.

Growth was monitored by noting the occurrence of visible growth (turbidity) in the culture medium, using a Klett–Summerson photometer and a filter at a wavelength of 436 nm.

Induction procedure.
Induction studies based on measuring the time-dependent release of CO2 due to the mineralization of the carbon source were carried out as described by Frederikson et al. (1991) . For this purpose, the cells were grown at 25 °C on mineral salts medium agar plates containing 0·3 ml of the respective carbon source on a filter-paper disc in the lid or containing 1% (w/v) sodium gluconate in the agar. After 48 h incubation, the cells were harvested from the plates, washed twice with sterile NaCl solution (0·85%, w/v) and subsequently resuspended to an optical density at 436 nm of 2·5 in 500 ml flasks containing 50 ml mineral salts medium plus the hydrocarbon as sole carbon source. Chloramphenicol (200 µg ml-1) was also added to prevent further protein synthesis. Controls without chloramphenicol were done under identical conditions. Each flask was equipped with a vial containing 2 ml 1 M NaOH to absorb CO2 produced by the cells; these vials were exchanged every 24 h with new vials containing fresh NaOH solution during the time course of the experiment. The flasks were tightly sealed with wrapped rubber stoppers and incubated for 24 h at 25 °C on a rotary shaker at 120 r.p.m. CO2 production was monitored by titration with 1 M HCl.

TLC of extracted aromatic compounds and lipids.
Aromatic compounds were extracted from the supernatants with methanol and were subjected to TLC on silica-gel 60F254 plates (Merck), applying chloroform/acetic acid (9:1, v/v) as the solvent system. Compounds were visualized under UV light and identified by comparison of their RF values with those of phenylacetic acid, phenylpropionic acid, homogentisic acid, 4-hydroxyphenylpropionic acid, DL-mandelic acid (Sigma) and 4-hydroxybenzoic acid (Merck), which were used as reference substances.

For identification of lipids, whole cells were extracted with a mixture of chloroform and methanol (2:1, v/v), and the extracts were separated by TLC, which was performed on silica-gel 60F254 plates (Merck) using a mixture of hexane, diethyl ether and acetic acid (80:20:1, by vol.) as the solvent system. Lipid fractions were visualized after brief exposure to iodine vapour. Palmitic acid, stearic acid, dipalmitoylglycerol, tripalmitoylglycerol and cetylpalmitate (Merck) were used as reference substances.

Qualitative determination of intermediates in supernatants.
Culture supernatants obtained by centrifugation were analysed for excreted intermediates by liquid chromatography, using HPLC apparatus (Knauer). Separation was achieved by reversed-phase chromatography on Nucleosil-100 C18 (5 µm particle size, 250 mmx4·0 mm column) with a gradient of 0·1% (v/v) formic acid (eluent A) and acetonitrile (eluent B) in a range of 20–100% (v/v) eluent B and at a flow rate of 1 ml min-1. The compounds were identified by their retention times and the corresponding spectra were obtained with a diode array detector (WellChrom Diodenarray-Detektor K-2150; Knauer) (Priefert et al., 1997 ).

Analysis of neutral lipids and fatty acids.
To determine the fatty acid content of the cells and the composition of lipids, 3–5 mg lyophilized whole cells or the triacylglycerol fraction obtained from preparative TLC were subjected to methanolysis in the presence of 15% (v/v) sulfuric acid. The resulting fatty acid methyl esters were analysed by GC on a Konik HRGC3000 gas chromatograph equipped with an Innowax capillary column (30 mx0·53 mm) and a flame-ionization detector (Brandl et al., 1988 ; Alvarez et al., 1996 ). A 2 µl portion of the organic phase was analysed after split injection (split ratio 1:20), and nitrogen was used as the carrier gas at a flow rate of 50 ml min-1. The temperature of the injector and the detector was 260 °C, whereas a temperature programme was used for efficient separation of the methyl esters on the column (150 °C for 1 min, temperature increases of 5 °C min-1, 240 °C for 10 min).

Electron spray ionization mass spectrometry and tandem electron spray ionization mass spectrometry.
Electron spray ionization mass spectrometry (ESI-MS) and tandem ESI-MS/MS experiments were conducted with a Micromass type Quattro LCZ (Beverly) quadrupole mass spectrometer. In MS/MS mode, fragmentation was achieved by introducing argon in the reaction chamber in front of the second quadrupole. The lipid samples were directly transferred from TLC plates with a mixture of chloroform and methanol (5:1, v/v; flow rate 100 µl min-1) into the ESI source using a TLC plate elution probe (DGMS 2001, P31, constructed by H. Luftmann, Institut für Organische Chemie, Westfälische Wilhelms-Universität, Münster, Germany).


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Excretion of metabolites during cultivation of R. opacus on phenyldecane
The occurrence of extracellular metabolites during the cultivation of cells of R. opacus strain PD630 on phenyldecane was analysed by HPLC and TLC and the compounds were identified according to the Rt or RF values, respectively, in comparison with reference compounds. Three classes of compounds were detected: (1) phenylalkanoic acids with short-chain-length alkyl substituents, (2) monohydroxylated phenyl compounds, and (3) dihydroxylated phenyl substances. The metabolites isolated from the supernatants included the following: phenylacetic acid (Rt 11·06 and RF 0·79), phenylpropionic acid (Rt 11·30 and RF 0·79), 4-hydroxyphenylpropionic acid (Rt 5·91 and RF 0·64), protocatechuate (Rt 3·70 and RF 0·50) and homogentisic acid (2,5-dihydroxyphenylacetic acid) (Rt 2·92 and RF 0·28), in HPLC and TLC analysis, respectively. The main compounds were homogentisic acid, 4-hydroxyphenylpropionic acid and phenylacetic acid, whereas phenylpropionic acid and protocatechuate occurred as minor compounds. Other intermediates, which appeared as minor compounds after TLC analysis and which exhibited RF values of 0·19, 0·23, 0·38 and 0·44, remained unidentified. Growth experiments demonstrated that R. opacus PD630 was able to use the identified metabolites as sole carbon and energy source for growth (not shown).

Analysis of intracellular metabolites accumulated by R. opacus during growth on phenyldecane
Cells of strain PD630 accumulated neutral lipids during cultivation on phenyldecane as sole carbon source under nitrogen-limiting conditions. These lipids were separated by TLC analysis and identified by comparision of their RF values with those of reference substances and by additional chemical analysis. After 4 d cultivation of the cells on phenyldecane, four spots of lipids were detected on the TLC plates (data not shown). The spot exhibiting an RF value of 0·13 contained the diacylglycerol fraction, as revealed by comparison with the respective RF value of a dipalmitoylglycerol standard. The occurrence of diacylglycerols in phenyldecane-grown cells of strain PD630 has been reported previously (Alvarez et al., 1996 ). In addition, detailed analysis of the spots occurring in TLC in a previous study confirmed the occurrence of diacylglycerols in cells of R. opacus PD630 (Wältermann et al., 2000 ). The other compounds were identified by ESI-MS and ESI-MS/MS after separation on TLC plates. The spot exhibiting an RF value of 0·43 contained a mixture of TAGs with pseudomolecular ions between m/z [M+Na]+ 850 and 950. The mass spectrum obtained was very similar to that of TAGs accumulated by strain PD630 during cultivation on gluconate (Wältermann et al., 2000 ). These TAGs contained odd- and even-numbered aliphatic fatty acids with carbon chain lengths ranging from 13 to 19 carbon atoms, palmitic acid, margaric acid, cis{Delta}9-heptadecenoic acid and oleic acid being major components (Alvarez et al., 1996 ; Wältermann et al., 2000 ). An ESI mass spectrum of the purified TLC fraction exhibiting an RF value 0·54 revealed pseudomolecular ions in the range, for m/z [M+Na]+, 800 to 880 (Fig. 1a). The main ions represented a mixture of TAGs in which one acyl group was replaced by a phenyldecanoic acid residue. The occurrence of fatty acids with a phenyl group in the TAG of this TLC fraction was confirmed by ESI-MS/MS analysis. One ESI-MS/MS spectrum and the fragmentation pattern of the pseudomolecular ion with an m/z [M+Na]+ of 847 are presented in Fig. 1(b) as an example. The spectrum showed the occurrence of odd-numbered fatty acids among the TAGs, since the mass differences between ionic groups frequently amounted to 14. Similar results were previously reported by Wältermann et al. (2000) for TAGs accumulated by gluconate-grown cells of R. opacus PD630.



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Fig. 1. ESI-MS spectrum of TAGs isolated from cells of R. opacus strain PD630 cultivated on phenyldecane under nitrogen-starvation conditions after TLC sparation. TAGs exhibiting an RF value of 0·54 were isolated from a TLC plate and analysed. (a) Pseudomolecular ions of whole TAGs each containing one phenyldecanoic acid residue. (b) ESI-MS/MS spectrum of the main pseudomolecular ion with m/z 847 and the corresponding fragmentation pattern. All assignments correspond to [M+Na]+.

 
Finally, the compound exhibiting an RF value of 0·88 was identified as the wax ester phenyldecylphenyldecanoate. Its ESI-MS/MS spectrum and the fragmentation pattern are shown in Fig. 2. Interestingly, other wax esters, consisting, for example, of phenyldecanoic acid and decanoic acid, could not be detected.



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Fig. 2. ESI-MS/MS mass spectrum and fragmentation pattern of the aromatic wax ester phenyldecylphenyldecanoate accumulated by R. opacus strain PD630 during growth on phenyldecane. The compounds exhibiting an RF value of 0·88 were isolated from a TLC plate and analysed. All assignments correspond to [M+Na]+.

 
Regulation of phenyldecane catabolism
To reveal which enzymes are responsible for the oxidation of phenyldecane, and to evaluate whether they are constitutively expressed or induced, two-stage cultivation experiments with a different substrate in each stage, in the presence of chloramphenicol in the second stage, were done with cells of strain PD630. The amounts of CO2 formed from the carbon source in the second cultivation stage were measured. The results of these mineralization experiments are shown in Table 1. Degradation of phenyldecane was clearly an inducible process. When the cells were grown on gluconate, phenylacetic acid or phenylalanine in the first stage, they were unable to degrade phenyldecane during cultivation in the second stage. In contrast, phenyldecane itself and hexadecane induced mineralization of phenyldecane. In addition, the exposure of cells with phenyldecane, phenylacetic acid or phenylalanine induced the catabolism of phenylacetic acid. Gluconate also induced the mineralization of phenylacetic acid to some extent, probably because this last compound may be a degradation intermediate of aromatic amino acids generated in gluconate-grown cells of R. opacus PD630.


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Table 1. Mineralization of phenyldecane and phenylacetic acid by cells of R. opacus strain PD630 in the presence of chloramphenicol after previous growth of cells on various substrates

 

   DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
This study revealed the formation of hitherto undescribed wax esters and TAGs containing phenyldecanoic acid as constituents in cells of R. opacus strain PD630 during cultivation of the cells on phenyldecane.On the basis of the results of this study, three steps for the oxidation of phenyldecane by R. opacus PD630 can be distinguished. (1) The induction of phenyldecane degradation by hexadecane suggests that probably both phenylalkanes and n-alkanes are oxidized by the same enzymes. Alkane monooxygenase, alcohol dehydrogenase and aldehyde dehydrogenase probably catalyse the monoterminal oxidation of both compounds. (2) The resulting phenyldecanoic acid is then further degraded mainly by the ß-oxidation of the alkyl side-chain to phenylacetic acid. The occurrence of this catabolic route for the oxidation of phenylalkanes was previously reported for other actinomycetes (Gibson & Subramanian, 1984 ; Warhust & Fewson, 1994 ). In addition, {alpha}-oxidation must also occur to some extent, as indicated by the presence of phenylnonanoic acid and phenylpropionic acid in the cells and in the supernatants. (3) The {alpha}- and ß-oxidation products, phenylpropionic and phenylacetic acid, respectively, are probably hydroxylated prior to cleavage of the aromatic ring, which results in the formation of linear intermediates that are further converted to central intermediates. These results suggest that phenylacetic acid is, like phenylalanine, catabolized via homogentisic acid (Olivera et al., 1998 ), because the capacity to degrade phenylacetic acid was induced in phenylalanine- and also in phenyldecane-grown cells of R. opacus strain PD630. The results of this study demonstrated that strain PD630 possesses the necessary enzymes for the aromatic ring fission of phenylacetic acid; however, these enzymes were not able to attack the aromatic ring of phenyldecane. This suggested that the occurrence of an alkyl side-chain in the aromatic molecule may hinder the oxidative attack by strain PD630.

However, phenyldecanoic acid is directed not only to catabolic pathways ({alpha}- and ß-oxidation) in R. opacus PD630 but also towards anabolic routes, such as the biosynthesis of a novel wax ester and novel TAGs. In contrast, it is very interesting that (though it remains unclear why) wax esters composed of regular fatty acids and the corresponding alcohols were not synthesized when R. opacus PD630 was grown on gluconate under nitrogen starvation (Wältermann et al., 2000 ). To our knowledge, this is the first report on the formation of waxes and TAGs containing aromatic constituents in bacteria. In this context, the accumulation of unusual polyhydroxyalkanoates bearing a phenyl group by Pseudomonas species has to be mentioned (Fritzsche et al., 1990 ; García et al., 1999 ). The synthesis of such polyhydroxyalkanoates required the occurrence of a polyhydroxyalkanoate synthase with broad substrate specificity. In contrast, R. opacus PD630 is unable to store 3-hydroxyalkanoic acids as polyhydroxyalkanoates but accumulates fatty acids as neutral lipids under the culture conditions in this study (Alvarez et al., 1996 ).

Whether or not the formation of waxes and TAGs with aromatic constituents in R. opacus is physiologically relevant remains to be evaluated. As result of its broad metabolic capacity, unusual fatty acids such as phenyldecanoic acid may be generated from phenylalkanes, which may disturb the membrane fluidity if they are incorporated into phospholipids. Therefore, their incorporation into acylglycerols or wax esters may provide a defence strategy to regulate the fatty acid composition of the membrane phospholipids. Recently, Dahlqvist et al. (2000) identified an acyl-CoA-independent enzyme in plants, which is responsible for the biosynthesis of TAGs using phospholipids as acyl donor. This enzyme probably contributes only slightly to the accumulation of TAGs, but its role may be important for maintaining the functional fluidity of cellular membranes. If a similar enzyme occurs in strain PD630, it may be responsible for the biosynthesis of TAGs containing aromatic fatty acids. Assimilation of phenyldecane by strain PD630 was not complete under the cultivation conditions used in this study, since intermediates of phenyldecane oxidation were accumulated intracellularly, and others were excreted into the medium. These surplus intermediates may be further oxidized when an external carbon source becomes limiting.

In conclusion, this study provides a physiological and biochemical approach for investigating the catabolism and assimilation of phenyldecane by R. opacus strain PD630 under conditions of restricted growth. Such conditions normally predominate in natural environments. The understanding of the metabolic response of bacteria to the substances investigated in this study could be important not only for bioremediation processes but also for obtaining novel compounds, such as TAGs and wax esters containing aromatic compounds, by using biotechnological processes.


   ACKNOWLEDGEMENTS
 
H. M. Alvarez is indebted to the Deutsche Akademischer Austauschdienst for the award of a post-doctoral scholarship. This work was partially supported by a grant from the Agencia Nacional de Promoción Científica y Tecnológica, Secretaria de Ciencia y Técnica, Argentina (PICT97 no. 01-0000.01245-BID802/OC-AR). We thank Dr H. Priefert (Institut für Mikrobiologie, Westfälische Wilhelms-Universität Münster) for helpful discussions.


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Alvarez, H. M., Mayer, F., Fabritius, D. & Steinbüchel, A. (1996). Formation of intracytoplasmic lipid inclusions by Rhodococcus opacus PD630. Arch Microbiol 165, 377-386.[Medline]

Alvarez, H. M., Kalscheuer, R. & Steinbüchel, A. (1997). Accumulation of storage lipids in species of Rhodococcus and Nocardia and effect of inhibitors and polyethylene glycol. Fett/Lipid 9, 239-246.

Alvarez, H. M., Kalscheuer, R. & Steinbüchel, A. (2000). Accumulation and mobilization of storage lipids by Rhodococcus opacus PD630 and Rhodococcus ruber NCIMB 40126. Appl Microbiol Biotechnol 54, 218-223.[Medline]

Alvarez, H. M., Souto, M. F., Viale, A. & 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.[Medline]

Brandl, H., Gross, R. A., Lenz, R. W. & Fuller, R. C. (1988). Pseudomonas oleovorans as a source of poly(hydroxyalkanoates) for potential applications as biodegradable polyesters. Appl Environ Microbiol 54, 1977-1982.

Dahlqvist, A., Stähl, U., Lanman, M., Banas, A., Lee, M., Sandager, L., Ronne, H. & Stymne, S. (2000). Phosholipid:diacylglycerol acyltransferase: an enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl Acad Sci USA 12, 6487-6492.

Finnerty, W. R. (1992). The biology and genetics of the genus Rhodococcus. Annu Rev Microbiol 46, 193-218.[Medline]

Frederickson, J. K., Brockman, F. J., Workman, D. J., Li, S. W. & Stevens, T. O. (1991). Isolation and characterization of subsurface bacterium capable of growth on toluene, naphthalene, and other aromatic compounds. Appl Environ Microbiol 57, 796-803.

Fritzsche, K., Lenz, R. W. & Fuller, R. C. (1990). An unusual bacterial polyester with a phenyl pendant group. Macromol Chem 191, 1957-1965.

García, B., Olivera, E. R., Minabres, B., Fernándes-Valverde, M., Canedo, L. M., Prieto, M. A., García, J. L., Martínez, M. & Luengo, J. M. (1999). Novel biodegradable aromatic plastics from a bacterial source. J Biol Chem 274, 29228-29241.[Abstract/Free Full Text]

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Olivera, E. R., Minambres, B., García, B., Muniz, C., Morena, M. A., Ferrández, A., Díaz, E., García, J. L. & Luengo, J. M. (1998). Molecular characterization of the phenylacetic acid catabolic pathway in Pseudomonas putida U: the phenylacetyl-CoA catabolon. Proc Natl Acad Sci USA 95, 6419-6424.[Abstract/Free Full Text]

Priefert, H., Rabenhorst, J. & Steinbüchel, A. (1997). Molecular characterization of genes of Pseudomonas sp. strain HR199 involved in bioconversion of vanillin to protocatechuate. J Bacteriol 179, 2595-2607.[Abstract]

Sariaslani, F. S., Harper, D. B. & Higgins, I. J. (1974). Microbial degradation of hydrocarbons. Catabolism of 1-phenylalkanes by Nocardia salmonicolor. Biochem J 140, 31-45.[Medline]

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Sikkema, J., De Bont, J. A. M. & Poolman, B. (1995). Mechanisms of membrane toxicity of hydrocarbons. Microbiol Rev 59, 201-222.

Wagner-Dobler, I., Bennasar, A., Vancanneyt, M., Strompl, C., Brummer, I., Eichner, C., Grammel, I. & Moore, E. R. (1998). Microcosm enrichment of biphenyl-degrading microbial communities from soils and sediments. Appl Environ Microbiol 64, 3014-3022.[Abstract/Free Full Text]

Wältermann, M., Luftmann, H., Baumeister, D., Kalscheuer, R. & Steinbüchel, A. (2000). Rhodococcus opacus strain PD630 as a new source of high-value single cell oil? Isolation and characterization of triacylglycerols and other storage lipids. Microbiology 146, 1143-1149.[Abstract/Free Full Text]

Warhust, A. M. & Fewson, C. A. (1994). Biotransformation catalyzed by the genus Rhodococcus. Crit Rev Biotechnol 14, 29-73.[Medline]

White, L. G., Hawari, J., Zhou, E., Bourbonnière, L., Innis, W. E. & Greer, H. W. (1998). Biodegradation of variable-chain-length alkanes at low temperatures by a psychotrophic Rhodococcus sp. Appl Environ Microbiol 64, 2578-2584.[Abstract/Free Full Text]

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Received 28 August 2001; revised 11 December 2001; accepted 11 January 2002.



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