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Applied and Environmental Microbiology, March 2004, p. 1487-1493, Vol. 70, No. 3
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.3.1487-1493.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Use of a Novel Fluorinated Organosulfur Compound To Isolate Bacteria Capable of Carbon-Sulfur Bond Cleavage

Jonathan D. Van Hamme,1* Phillip M. Fedorak,2 Julia M. Foght,2 Murray R. Gray,3 and Heather D. Dettman1

National Centre for Upgrading Technology, Devon, Alberta T9G 1A8,1 Department of Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9,2 Department of Chemical and Materials Engineering, University of Alberta, Edmonton, Alberta T6G 2G6, Canada3

Received 7 July 2003/ Accepted 24 November 2003


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The vacuum residue fraction of heavy crudes contributes to the viscosity of these oils. Specific microbial cleavage of CS bonds in alkylsulfide bridges that form linkages in this fraction may result in dramatic viscosity reduction. To date, no bacterial strains have been shown conclusively to cleave CS bonds within alkyl chains. Screening for microbes that can perform this activity was greatly facilitated by the use of a newly synthesized compound, bis-(3-pentafluorophenylpropyl)-sulfide (PFPS), as a novel sulfur source. The terminal pentafluorinated aromatic rings of PFPS preclude growth of aromatic ring-degrading bacteria but allow for selective enrichment of strains capable of cleaving CS bonds. A unique bacterial strain, Rhodococcus sp. strain JVH1, that used PFPS as a sole sulfur source was isolated from an oil-contaminated environment. Gas chromatography-mass spectrometry analysis revealed that JVH1 oxidized PFPS to a sulfoxide and then a sulfone prior to cleaving the CS bond to form an alcohol and, presumably, a sulfinate from which sulfur could be extracted for growth. Four known dibenzothiophene-desulfurizing strains, including Rhodococcus sp. strain IGTS8, were all unable to cleave the CS bond in PFPS but could oxidize PFPS to the sulfone via the sulfoxide. Conversely, JVH1 was unable to oxidize dibenzothiophene but was able to use a variety of alkyl sulfides, in addition to PFPS, as sole sulfur sources. Overall, PFPS is an excellent tool for isolating bacteria capable of cleaving subterminal CS bonds within alkyl chains. The type of desulfurization displayed by JVH1 differs significantly from previously described reaction results.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Microbial methods of removing sulfur from organosulfur compounds are of interest to the petroleum industry for reducing sulfur emissions and, more recently, for reducing heavy oil viscosity. As conventional crude oils are consumed throughout the world, heavier oils are being exploited which, due to their high viscosity, cannot be transported from remote field sites to refineries without adding diluents. The vacuum residue fraction of crude oils (boiling point >= 524°C [975°F]) contributes to viscosity, and recent models indicate that alkyl sulfides compose important bridges in the network of high-molecular-weight molecules in this fraction (34). Up to 40% of the sulfur in these fractions is in the form of alkyl sulfides; if these alkyl CS bonds can be selectively cleaved using a biological catalyst, reductions in molecular size and viscosity could occur.

The first requirement for developing a biological process for heavy oil viscosity reduction is obtaining a microorganism capable of alkyl CS bond cleavage without reducing the carbon value of the substrate. Precedence for this type of reaction with aromatic heterocycles can be found in the well-characterized 4S pathway that selectively removes sulfur from dibenzothiophene (DBT) (35). The dsz or sox operon (genes responsible for DBT desulfurization) (7, 38) in Rhodococcus sp. strain IGTS8 has been observed in a variety of genera that are apparently widespread in petroleum-contaminated environments (6, 8). Other genera capable of selective sulfur removal from DBT include Gordonia sp. strain CYKS1 (40), Arthrobacter sp. strain ECRD-1 (28), the thermophilic Paenibacillus sp. strain A11-2 (23, 24), and Bacillus subtilis WU-S2B (22). Strains that desulfurize alkylated DBTs (12, 13, 25, 28, 32, 33) and benzothiophene (14) have also been isolated. Recently, a metabolic pathway has been described for Rhodococcus sp. strain WU-K2R that can desulfurize naphthothiophene and benzothiophene (21).

However, there are no reports that conclusively illustrate the bacterial cleavage of alkyl CS bonds by bacteria. Van Hamme et al. (45) have shown that a variety of white-rot fungi oxidize dibenzyl sulfide to dibenzyl sulfoxide and dibenzyl sulfone prior to further degradation. However, other examples of the microbial degradation of compounds containing this type of bond (e.g., 2-chloroethyl sulfide and thiodiglycol) give no evidence for direct sulfur oxidation followed by CS bond cleavage without degradation of the alkyl or aromatic moieties (20, 29, 36, 40). For example, metabolism by Nocardioides simplex of 1-(phytanylsulfanyl)-octadecane (used as a model compound for sulfide bridges in high-molecular-weight fractions of sulfur-rich petroleum) has been described previously (18); although the sulfur was oxidized, no CS bond cleavage was observed. This bacterium oxidized the terminal alkyl groups in the model compound, which was subsequently degraded by ß-oxidations.

The challenge in isolating a bacterial strain capable of cleaving CS bonds within alkyl chains lies in finding an appropriate substrate. Commercially available sulfur-containing substrates typically have terminal alkyl or aromatic moieties that are susceptible to microbial attack. Several studies have evaluated and used fluorobenzoic acids, including pentafluorobenzoic acid, as conservative tracers in soil and ground water (2, 5, 17, 37) because of their chemical stability. Thus, we hypothesized that a short-chain alkyl sulfide that contained a stable pentafluorophenyl group at each terminus would be ideal for selecting microorganisms that cleave CS bonds.

This report outlines the synthesis and use of bis-(3-pentafluorophenylpropyl)-sulfide (PFPS) as a novel compound to select, screen, and characterize isolates capable of cleaving CS bonds within alkyl chains that may occur in the residual fractions of heavy crude oils and bitumens. An isolate (Rhodococcus sp. strain JVH1) which is distinct from the known DBT-desulfurizing strains in that it cannot use DBT as a sulfur source is described. In addition, metabolites produced by JVH1 from PFPS are shown and a metabolic pathway involving specific sulfur oxidation and CS bond cleavage is presented.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Substrates and chemicals.
The commercially unavailable compounds PFPS, bis-(3-pentafluorophenylpropyl)-disulfide (PFPSS), 3-pentafluorophenylpropan-1-ol (PFPP-OH), 3-pentafluorophenylpropanoic acid (PFPP-acid), bis-(3-pentafluorophenylpropyl)-sulfoxide (PFPSO), and bis-(3-pentafluorophenylpropyl)-sulfone (PFPSO2) were synthesized as described below.

Benzothiophene, dimethyl sulfoxide (DMSO), dioctyl sulfide, dibenzyl sulfide, dibenzyl sulfone, 1,4-dithiane, tetramethylene sulfone (also known as sulfolane and tetrahydrothiophene sulfone), DBT sulfone, pentafluorobenzoic acid, 1-phenylnaphthalene, phenyl sulfide, and thianthrene were purchased from Aldrich Chemical Company, Inc. (Milwaukee, Wis.) and were >98% pure. Dioctadecyl sulfide was obtained from TCI Chemicals (Tokyo, Japan) while dibenzyl sulfoxide and DBT were from Fluka (Buch, Switzerland). Dichloromethane (DCM), pentane, hexane, ethyl ether, and ethyl acetate were obtained from Fisher Chemicals (Fair Lawn, N.J.).

Culture sources.
Mixed cultures for selection (enriched from various hydrocarbon-contaminated samples from Canadian sites) were maintained in sulfur-free and low-salts media for 18 months in roller bottles at room temperature. The DBT-desulfurizing strains Rhodococcus erythropolis IGTS8 (ATCC 53968), Bacillus sp. strain IGTS9 (ATCC 53969), Rhodococcus sp. strain D-1 (ATCC 55309), and Rhodococcus sp. strain I-3 (ATCC 55310) were obtained from the American Type Culture Collection (Manassas, Va.).

Growth medium.
All experiments used sulfur-free glucose-acetate (SFGA) medium (adapted from MacPherson et al.) (32) containing the following ingredients (per liter of deionized water): 0.4 g of KH2PO4, 1.6 g of K2HPO4, 1.55 g of NH4Cl, 0.165 g of MgCl2 · 6H2O, 0.09 g of CaCl2 · 2H2O, 5 g of sodium acetate, and 5 g of glucose. Glucose, MgCl2 · 6H2O, CaCl2 · 2H2O, 1.0 ml of Pfenning's vitamin solution (32) and 5.0 ml of modified Wolfe's minerals (32) were added as a 0.22-µm-pore-size-filter-sterilized solution to cooled medium after autoclaving to give a final pH of 7.0. Sulfate was added as an autoclaved solution of MgSO4 · 7H2O as required. All other sulfur sources were prepared in methanol and added with a glass syringe. Dioctadecyl sulfide was dissolved in DCM and added to empty tubes. The DCM was allowed to evaporate prior to culture addition.

For isolating pure cultures, plate count agar (Difco, Becton Dickinson and Company, Sparks, Md.) was used.

Culture conditions.
SFGA medium (25 ml in 125-ml Erlenmeyer flasks) was used for culture selection, routine inoculum growth, and biomass yield experiments. Flasks were incubated at 28°C on an orbital shaker at 200 rpm. Culture screening and PFPS oxidation experiments were carried out in screw-cap test tubes (inner diameter, 16 mm; length, 150 mm) capped with Teflon-lined caps containing 5 ml of SFGA medium. Tubes were incubated at 28°C on a tube roller at 40 rpm. Plates were incubated at 28°C for 3 days.

Biomass measurements.
Optical density (OD) of growing cultures was measured at 600 nm (OD600), while dry weights were determined after washing cultures three times with distilled water and drying in a 100°C oven until a constant weight was obtained.

Extraction and analysis.
PFPS, PFPSO, PFPSO2, and PFPP-OH were quantitatively extracted from roll tubes with three 1-ml aliquots of pentane. Prior to extraction, 1-phenylnaphthalene was added as an internal standard. After each addition of pentane, tubes were vortexed at high speed for 10 s. PFPP-acid was subsequently extracted by adjusting the pH to <2.0 and extracting with 3 x 1 ml of ethyl acetate.

For routine analysis, a Hewlett-Packard 5930 gas chromatograph equipped with an HP-1 column (length, 25 m; inside diameter, 0.32 mm; film thickness, 0.17 µm) and a flame-ionization detector were used. Peak areas were measured with an HP 3390A integrator. The injector was set to 300°C, and the detector was set to 320°C. The oven program was as follows: 90°C for 5 min followed by an increase in temperature of 15°C/min to 280°C, with a final hold time of 5 min.

Metabolite derivatization and identification.
Prior to gas chromatography-mass spectrometry (GC-MS) analysis, samples were dried with anhydrous sodium sulfate. PFPS, PFPSO, PFPSO2, PFPP-OH, and PFPP-acid were all detected by GC-MS without derivatization. PFPP-OH and PFPP-acid were also derivatized in 800 µl of acetonitrile with 200 µl of N-O-bis-(trimethylsilyl)acetamide (Pierce, Rockford, Ill.) at 70°C in a water bath for 15 min prior to GC-MS. PFPP-acid was also derivatized with diazomethane in ether.

Samples were analyzed on a Hewlett-Packard 5890 series II gas chromatograph with a 5970 series mass-selective detector and a 30-m DB-5 capillary column (J&W Scientific, Folsom, Calif.). The GC temperature program used for analysis was 90°C for 5 min followed by an increase of 10°C/min to 300°C, with a final hold time of 5 min.

Fluoride and sulfate analysis.
A fluoride ion-selective electrode (Thermo Orion, Beverly, Mass.) was used for fluoride analysis after samples were mixed with an equal volume of Ionall buffer solution (Anachemia Canada Inc., Lachine, Quebec, Canada).

Sulfate was analyzed using a DX600 ion chromatography system (Dionex Corp., Sunnyvale, Calif.) equipped with an AS9-HC anion exchange column and a CD25 conductivity detector. The mobile phase was 9 mM sodium carbonate set to 1.2 ml/min, and the suppressor current was set to 50 mA.

Synthesis of PFPP-OH.
All synthetic reactions and purifications were monitored by thin-layer chromatography on silica plates (Merck, Darmstadt, Germany) (250 µm of silica gel 60 F254) with ethyl acetate in hexane for development. Spots were visualized under UV light and by oxidation with potassium permanganate. All reaction products were purified by silica gel chromatography, and purity was evaluated using 1H-nuclear magnetic resonance (NMR) analysis. All reaction solvents were distilled prior to use.

PFPP-OH was synthesized from allylpentafluorobenzene (Aldrich) by hydroboration with argon by treatment with 1 M BH3.THF (triborane) complex in tetrahydrofuran (Aldrich) followed by the addition of 30% H2O2 under alkaline conditions. The product (64% yield) was extracted at pH 2 with ethyl ether and purified to give PFPP-OH, a viscous yellow liquid (1H-NMR [300 MHz] {delta} 1.31 [1H, t, J = 3 Hz, -OH], 1.83 [2H, m, CH2], 2.79 [2H, t, J = 9 Hz, CH2], and 3.67 [2H, q, J = 6 Hz, CH2]).

Synthesis of PFPS and PFPSS.
These were synthesized by first converting PFPP-OH to 1-iodo-3-pentafluorophenyl propane. This reaction was carried out in DCM with argon at room temperature according to the method of Lange and Gottardo (27). Triphenyl phosphine (Aldrich) was dissolved followed by the addition of imidazole (Aldrich), and the ingredients were allowed to mix for 30 min prior to the addition of iodine. The reaction mixture was extracted with DCM, and filtered. The extract was concentrated and the purified iodide, a dark pink oil (1H-NMR [300 MHz] {delta} 2.1 [2H, m, CH2CH2I], 2.79 [2H, t, J = 9 Hz, CH2], and 3.17 [2H, t, J = 9 Hz, CH2]), was used to make PFPS and PFPSS with the method of Landini and Rollo (26) and using the phase transfer catalyst hexadecyltributylphosphonium bromide (Aldrich) and Na2S · 9H2O. The reaction proceeded for 2 h with argon (in water purged with argon to remove residual oxygen). The two-phase reaction mixture was extracted with ether, dried with MgSO4, and concentrated using a vacuum to yield a viscous, cloudy, orange mixed product. Purified PFPS (62% yield) (1H-NMR [300 MHz] {delta} 1.83 [2H, m, CH2CH2S], 2.53 [2H, t, J = 9 Hz, CH2], and 2.79 [2H, t, J = 9 Hz, CH2]) and PFPSS (2% yield) (1H-NMR [300 MHz] {delta} 1.96 [2H, m, CH2CH2S], 2.68 [2H, t, J = 9 Hz, CH2], and 2.79 [2H, t, 9 Hz, CH2]) were viscous, slightly yellow, transparent liquids.

Synthesis of PFPP-acid.
PFPP-acid was synthesized (using the method of Prashad et al.) (39) from PFPP-OH. In this case, PFPP-OH, acetonitrile, water, ethyl acetate, NaIO4, and RuCl3 · H2O were mixed. The reaction was monitored using thin-layer chromatography, and (following the disappearance of the PFPP-OH) the mixture was filtered to remove the white precipitate. The filtrate was adjusted to pH 2 with HCl and extracted three times with ethyl acetate. The combined extracts were dried over anhydrous sodium sulfate and concentrated. The purified PFPP-acid (45% yield) was a fluffy, white, aromatic solid (1H-NMR [300 MHz] {delta} 2.66 [2H, t, J = 7.5 Hz, CH2] and 3.02 [2H, t, J = 7.5 Hz, CH2COOH]).

Synthesis of PFPSO and PFPSO2.
For this synthesis, the method of Trost and Curran (43) was used. PFPS and methanol were cooled on ice prior to adding water containing 3-mol equivalents of Oxone (Aldrich) (potassium hydrogen persulfate; 2KHSO5 · KHSO4 · K2SO4). The reaction was allowed to proceed at room temperature for 30 min. PFPSO2 (97% yield) (1H-NMR [300 MHz] {delta} 2.13 [2H, m, CH2CH2S], 2.87 [2H, t, J = 9 Hz, CH2], and 2.97 [2H, t, J = 9 Hz, CH2]) and PFPSO (2% yield) (1H-NMR [300 MHz] {delta} 2.07 [2H, m, CH2CH2S], 2.65 [2H, m, CH2], and 2.86 [2H, t, J = 9 Hz, CH2]) were white solids.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Isolation of the PFPS-desulfurizing bacterium Rhodococcus sp. strain JVH1.
To isolate microorganisms capable of cleaving CS bonds within alkyl chains, PFPS (Table 1) was used as the sole sulfur source in enrichment cultures. Enrichment cultures derived from various petroleum-contaminated soils and sludges and maintained by passage in medium without the addition of sulfate were used as inocula. Shake flasks were prepared with SFGA and 40 µM PFPS as the only added sulfur source. Cultures were grown in subcultures (4% [vol/vol]) weekly for 5 weeks to remove solids and any associated sulfur sources. Turbid cultures were diluted and spread on agar to obtain isolated colonies. A total of 70 pure isolates, including 3 filamentous fungal isolates and 1 yeast isolate, were screened for the ability to produce biomass and remove PFPS when PFPS was provided as the sole sulfur source at a limiting concentration (40 µM). Control cultures with no added sulfur were prepared for each isolate, as some strains were able to produce biomass on the trace sulfur found in medium components. Gas chromatograph-flame-ionization detector analysis showed that 4 of the 70 isolates oxidized PFPS to PFPSO2, and 2 of the 70 were able to produce significant amounts of biomass with PFPS as the sole sulfur source.


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  		TABLE 1. Structures and mass spectrometry data for PFPS and metabolites produced by Rhodococcus sp. strain JVH1

 
One strain, JVH1, was chosen for further study on the basis of the extent of growth and PFPS removal. This organism is a gram-positive, branching rod-shaped bacterium 1 µm in width, with branches up to 5 µm in length. Strain JVH1 was sent to MIDI Labs (Newark, Del.) for full-length 16S rRNA gene sequence analysis; the strain was found to cluster with Rhodococcus, Corynebacterium, and Tsukamurella species. It had a 99% match to Rhodococcus opacus in GenBank (accession number Y11893); this isolate has been named Rhodococcus sp. strain JVH1.

Strain JVH1 cleaves CS bonds.
To find PFPS metabolites, strain JVH1 was grown in SFGA medium with PFPS (325 µM) as the sole sulfur source. Cultures were extracted with pentane at neutral pH and then ethyl acetate at pH <2 at various times during the growth and stationary phases. Concentrated extracts, and extracts derivatized with either N-O-bis-(trimethylsilyl)acetamide or diazomethane, were analyzed by GC-MS. Table 1 shows the mass spectra of PFPS and metabolites detected in culture extracts. In each case, spectra of the metabolites and derivatized metabolites were identical to those of authentic standards synthesized for this work. PFPSO and PFPSO2 were detected in pentane extracts along with PFPP-OH. Ethyl acetate extracts yielded PFPP-acid. Both PFPP-OH and PFPP-acid were derivatized to produce trimethyl silyl derivatives, while PFPP-acid was converted to a methyl ester. The spectra of these novel metabolites and the sulfur source PFPS have not been reported previously.

Samples taken daily were analyzed for fluoride ion to check for possible aromatic ring attack. At no time did fluoride concentrations exceed the detection limit of 0.2 mg/liter.

Strain JVH1 differs from known DBT-desulfurizing strains.
The detection of the sulfoxide and sulfone of PFPS indicated that strain JVH1 used a pathway similar to that described for the DBT-desulfurizing strain R. erythropolis IGTS8 and related organisms. To test the versatility of strain JVH1 regarding cleavage of CS bonds, it was incubated with various organosulfur compounds. Table 2 shows that JVH1 was able to use linear alkyl sulfides ranging from DMSO to didodecyl sulfide as sole sulfur sources. However, dioctadecyl sulfide was not used by JVH1 as a sulfur source, presumably due to low solubility and to its crystalline state in aqueous medium. In addition, dibenzyl sulfide and its sulfur oxidation products, PFPSS, 1,4-dithiane, and tetrahydrothiophene sulfone, were used as sulfur sources. In contrast, phenyl sulfide, benzothiophene, and DBT and its sulfone were not used by JVH1 to produce biomass.


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  		TABLE 2. Growth of Rhodococcus sp. strain JVH1 in the presence of various organosulfur compounds as sole sulfur sourcea

 
Because strain JVH1 could not utilize DBT as a sulfur source, further comparisons were made with four well-known DBT-desulfurizing bacteria. The dry weights of biomass produced during a 7-day incubation with various sulfur sources were determined. Figure 1 shows that JVH1 is distinct from four known DBT-desulfurizing strains in that DBT does not support its growth whereas PFPS cannot support growth of the DBT-desulfurizing strains. However, GC-MS analysis showed that strain IGTS8 was able to stoichiometrically convert PFPS to PFPSO2 via PFPSO. Conversely, JVH1 was unable to oxidize DBT to DBT-sulfoxide or DBT-sulfone or to extract sulfur from DBT-sulfone. All strains were able to grow well using DMSO and sulfate as sole sulfur sources.



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  		FIG. 1. Biomass (measured as washed cell dry weight) accumulated after 7 days of incubation in SFGA medium amended with various sulfur sources (40 µmol/liter of sulfur). ATCC 55309, Rhodococcus sp. strain D1; ATCC 55310, Rhodococcus sp. strain I-3; ATCC 53969, Bacillus sp. strain IGTS9; ATCC 53968, R. erythropolis IGTS8; JVH1, Rhodococcus sp. strain JVH1.

 
Strain JVH1 degrades PFPS in batch culture.
Growth of strain JVH1 corresponded to the disappearance of PFPS from the culture medium, as shown in Fig. 2. Growth caused an increase in pH, presumably from the consumption of acetate. After 14 days of incubation, JVH1 had removed all of the PFPS from the medium, with no accumulation of PFPSO or PFPSO2. Sulfate concentrations in the culture supernatant did not exceed that of the cell-free control (~10 µM sulfate) at any time during the incubation.



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  		FIG. 2. Time course of PFPS degradation by Rhodococcus sp. strain JVH1 grown in SFGA medium with 325 µM PFPS as the sole sulfur source. (Upper panel) Symbols: {diamondsuit}, PFPS (sterile control); {blacksquare}, PFPS; {triangleup}, PFPP-OH. (Lower panel) Symbols: {circ}, culture pH; {square}, growth (OD600). Error bars (where visible) show standard deviations (n = 3).

 
Gray et al. (15) reported that the CS bond cleavage of DBT sulfone yielded 2'-hydroxybiphenyl-2-sulfinate. In an analogous manner, we hypothesized that cleavage of the CS bond in PFPSO2 would yield PFPP-OH and a sulfinate (Fig. 3). Subsequent hydrolysis of the CS bond in the sulfinate would also yield PFPP-OH. The sulfinate was not detected by our analytical methods, but up to 18 µM of PFPP-OH accumulated in the culture medium by day 4 and then disappeared by day 6 (Fig. 2). Trace amounts of PFPP-acid were also detected. Further degradation of the acid via beta-oxidation would form pentafluorobenzoic acid, but none of this proposed metabolite was detected.



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  		FIG. 3. Proposed pathway of PFPS desulfurization by Rhodococcus sp. strain JVH1. Compounds in brackets were not identified but are potential metabolites.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
PFPS allows for the efficient selection of strains capable of cleaving CS bonds within alkyl chains. The relative ease with which isolates were obtained may indicate that this type of activity is not uncommon in the environment. However, to our knowledge this activity has not been described previously.

Based on the metabolites formed, the metabolic pathway in Fig. 3 is proposed. Here, sulfur is oxidized to produce first a sulfoxide and then a sulfone in a manner analogous to that of the 4S pathway observed with DBT-desulfurizing bacteria (35). Once activated in this manner, CS bond cleavage occurs to produce an alcohol and, presumably, a sulfinate. In the 4S pathway for DBT desulfurization, DBT-sulfone (DBTO2) is oxidized to 2-(2'-hydroxybiphenyl)-benzene sulfinate via the action of DszA, which is a DBTO2-monooxygenase (15). If a sulfinate were formed from PFPSO2, it would likely be cleaved to yield sulfite, which would then spontaneously oxidize to sulfate prior to being reduced by the microorganism for growth. Gray et al. (15) showed that the desulfinase in the 4S pathway (DszB) produces 2-hydroxybiphenyl and sulfite from 2-(2'-hydroxybiphenyl)-benzene sulfinate. Similarly, both sulfite and sulfate were detected by Greene et al. (16) in cultures of a Variovorax sp. strain growing on tetrahydrothiophene sulfone and by Bressler et al. (3) in cultures of a Pseudonocardia sp. strain with the sulfones of benzothiophene and 3- and 5-methylbenzothiophene as sole carbon and sulfur sources. In the current report, culture sulfate concentrations did not increase above those of the cell-free controls, indicating that sulfate was rapidly consumed by the growing cells.

While this would complete the sulfur extraction process, JVH1 further oxidized PFPP-OH to PFPP-acid. PFPP-acid would be metabolized to pentafluorobenzoic acid, presumably, but this metabolite has not been detected and the fate of the PFPP-acid is unknown. We have not pursued this question, because the primary goal of the study was CS bond cleavage. Defluorination and aromatic ring opening prior to sulfate release are not included in the proposed metabolic pathway, because fluoride ion was not detected in the culture medium. Even if only 1 in 10 fluoride ions per molecule of PFPS had been removed, the fluoride ion concentration would have risen to 6.2 mg/liter (30 times higher than the detection limit). Defluorination of oxidized aromatic rings has been previously reported among members of the Actinomycetales (9, 44) and Pseudomonadaceae (10, 41, 42), but (in general) fluorinated compounds are considered to be more recalcitrant than other halogenated organics (4, 11).

The novelty of the desulfurization activity exhibited by strain JVH1 is best illustrated by comparison to known DBT-desulfurizing bacteria unable to utilize alkyl sulfides other than DMSO as sulfur sources (19). Interestingly, Arensdorf et al. (1) reported that a Dsz-negative mutant of strain IGTS8 can transform DMSO, indicating that the DBT-desulfurization pathway is not required for this activity. In the results seen with this study, the detection of PFPSO in trace amounts (and the near stoichiometric recovery of PFPSO2 from the four DBT-desulfurizing cultures) supports the hypothesis that these strains are unable to cleave CS bonds in alkyl chains. Assuming that these strains employ the dsz operon for DBT desulfurization (6, 7, 31), it would appear that DszC, a sulfoxide-sulfone monooxygenase (30) that catalyzes the conversion of DBT to DBT-sulfoxide to DBT-sulfone (7, 38), has sufficiently broad substrate specificity to catalyze the first two steps of PFPS oxidation. This observation agrees with the work by Lei and Tu (30), who found that purified DszC from IGTS8 is able to oxidize benzyl sulfide and benzyl sulfoxide to benzyl sulfone.

In contrast, it would appear that DszA is not active against PFPSO2. These results are in agreement with recent work by Arensdorf et al. (1), who used a chemostat to isolate mutants of strain IGTS8 that could utilize octyl sulfide, 5-methylbenzothiophene, and benzothiophene as sulfur sources. In their work, specific single-nucleotide changes in DszA were responsible for expanding the normally narrow substrate range of this enzyme. Once the enzymes involved in desulfurization by strain JVH1 are isolated, comparative studies can be undertaken.

In conclusion, the novel organosulfur compound PFPS was effective in selecting microorganisms capable of cleaving CS bonds in alkyl chains. This activity will be valuable in desulfurization and viscosity reduction studies (provided that preferential use of more-bioavailable sulfur compounds can be controlled). To this end, reaction mechanisms involved in PFPS oxidation and cleavage are being investigated, and current work is focused on locating the novel desulfurization genes and enzymes in strain JVH1.


   ACKNOWLEDGMENTS
 
Partial funding for NCUT has been provided by the Canadian Program for Energy Research and Development (PERD), the Alberta Research Council, and the Alberta Energy Research Institute. Additional support for this study came from the University of Alberta, AEC Oil and Gas, Imperial Oil Resources, and PanCanadian Resources.

Thanks to John Vederas and Andrew Sutherland (Department of Chemistry, University of Alberta) for donating time and assistance with chemical syntheses. Thanks to Sara Ebert and to Kathlyn Kirkwood for useful discussions and to Eddie Wong for help with some of the culture work.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biological Sciences, University College of the Cariboo, 900 McGill Rd., P.O. Box 3010, Kamloops, B.C. V2C 5N3, Canada. Phone: (250) 377-6064. Fax: (250) 828-5450. E-mail: jvanhamme{at}cariboo.bc.ca. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Arensdorf, J. J., A. K. Loomis, P. M. DiGrazia, D. J. Monticello, and P. T. Pienkos. 2002. Chemostat approach for the directed evolution of biodesulfurization gain-of-function mutants. Appl. Environ. Microbiol. 68:691-698.[Abstract/Free Full Text]
  2. Becker, M. W., and A. M. Shipiro. 2000. Tracer transport in fractured crystalline rock: evidence of nondiffusive breakthrough tailing. Water Resour. Res. 36:1677-1686.
  3. Bressler, D. C., B. K. Leskiw, and P. M. Fedorak. 1999. Biodegradation of benzothiophene sulfones by a filamentous bacterium. Can. J. Microbiol. 45:360-368.[CrossRef][Medline]
  4. Carvalho, M. F., C. C. T. Alves, M. I. M. Ferreira, P. De Marco, and P. M. L. Castro. 2002. Isolation and initial characterization of a bacterial consortium able to mineralize fluorobenzene. Appl. Environ. Microbiol. 68:102-105.[Abstract/Free Full Text]
  5. Dahan, O., R. Nativ, E. A. Adar, B. Berkowitz, and Z. Ronen. 1999. Field observation of flow in fracture intersecting unsaturated chalk. Water Resour. Res. 35:3315-3326.
  6. Denis-Larose, C., D. Labbé, H. Bergeron, A. M. Jones, C. W. Greer, J. Al-Hawari, M. J. Grossman, B. M. Sankey, and P. C. K. Lau. 1997. Conservation of plasmid-encoded dibenzothiophene desulfurization genes in several rhodococci. Appl. Environ. Microbiol. 63:2915-2919.[Abstract]
  7. Denome, S. A., C. Oldfield, L. J. Nash, and K. D. Young. 1994. Characterization of the desulfurization genes from Rhodococcus sp. strain IGTS8. J. Bacteriol. 176:6707-6716.[Abstract/Free Full Text]
  8. Duarte, G. F., A. S. Rosado, L. Seldin, W. de Araujo, and J. D. van Elsas. 2001. Analysis of bacterial community structure in sulfurous-oil-containing soils and detection of species carrying dibenzothiophene desulfurization (dsz) genes. Appl. Environ. Microbiol. 67:1052-1062.[Abstract/Free Full Text]
  9. Engesser, K. H., M. A. Rubio, and D. W. Ribbons. 1988. Bacterial metabolism of side chain fluorinated aromatics: cometabolism of 4-trifluoromethyl(TFM)-benzoate by 4-isopropylbenzoate grown Pseudomonas putida JT strains. Arch. Microbiol. 149:198-206.[CrossRef][Medline]
  10. Engesser, K. H., R. B. Cain, and H. J. Knackmuss. 1988. Bacterial metabolism of side chain fluorinated aromatics: cometabolism of 3-trifluoromethyl(TFM)-benzoate by Pseudomonas putida (arvilla) mt-2 and Rhodococcus rubropertinctus N657. Arch. Microbiol. 149:188-197.[CrossRef][Medline]
  11. Fetznar, S., and F. Lingens. 1994. Bacterial dehalogenases: biochemistry, genetics, and biotechnological applications. Microbiol. Rev. 58:641-685.[Abstract/Free Full Text]
  12. Folsom, B. R., D. R. Schieche, P. M. DiGrazia, J. Werner, and S. Palmer. 1999. Microbial desulfurization of alkylated dibenzothiophenes from a hydrodesulfurized middle distillate by Rhodococcus erythropolis I-19. Appl. Environ. Microbiol. 65:4967-4972.[Abstract/Free Full Text]
  13. Furuya, T., K. Kirimura, K. Kino, and S. Usami. 2001. Thermophilic biodesulfurization of dibenzothiophene and its derivatives by Mycobacterium phlei WU-F1. FEMS Microbiol. Lett. 204:129-133.[CrossRef][Medline]
  14. Gilbert, S. C., J. Morton, S. Buchanan, C. Oldfield, and A. McRoberts. 1998. Isolation of a unique benzothiophene-desulphurizing bacterium, Gordona sp. strain 213E (NCIMB 40816), and characterization of the desulphurisation pathway. Microbiology 144:2545-2553.[Abstract]
  15. Gray, K. A., O. S. Pogrebinsky, G. T. Mrachko, L. Xi, D. J. Monticello, and C. H. Squires. 1996. Molecular mechanisms of biocatalytic desulfurization of fossil fuels. Nat. Biotechnol. 14:1705-1709.[CrossRef][Medline]
  16. Greene, E. A., P. H. Beatty, and P. M. Fedorak. 2000. Sulfolane degradation by mixed cultures and a bacterial isolate identified as a Variovorax sp. Arch. Microbiol. 174:111-119.[CrossRef][Medline]
  17. Jaynes, D. B. 1994. Evaluation of fluorobenzoate tracers in surface soils. Ground Water 32:532-538.
  18. Jenisch-Anton, A., P. Adam, W. Michaelis, J. Connan, D. Herrmann, M. Rohmer, and P. Albrecht. 2000. Molecular evidence for biodegradation of geomacromolecules. Geochim. Cosmochim. Acta 64:3525-3537.[CrossRef]
  19. Kayser, K. J., B. A. Bielaga-Jones, K. Jackowski, O. Odusan, and J. J. Kilbane. 1993. Utilization of organosulphur compounds by axenic and mixed cultures of Rhodococcus rhodochrous IGTS8. J. Gen. Microbiol. 139:3123-3129.
  20. Kilbane, J. J., and K. Jackowski. 1996. Biocatalytic detoxification of 2-chloroethyl ethyl sulfide. J. Chem. Tech. Biotechnol. 65:370-374.[CrossRef]
  21. Kirimura, K., T. Furuya, R. Sato, Y. Ishii, K. Kino, and S. Usami. 2002. Biodesulfurization of naphthothiophene and benzothiophenes through selective cleavage of carbon-sulfur bonds by Rhodococcus sp. strain WU-K2R. Appl. Environ. Microbiol. 68:3867-3872.[Abstract/Free Full Text]
  22. Kirimura, K., T. Furuya, Y. Nishii, Y. Ishii, K. Kino, and S. Usami. 2001. Biodesulfurization of dibenzothiophene and its derivatives through the selective cleavage of carbon-sulfur bonds by a moderately thermophilic bacterium Bacillus subtilis WU-S2B. J. Biosci. Bioeng. 91:262-266.
  23. Konishi, J., T. Onaka, Y. Ishii, K. Okumura, and M. Suzuki. 2000. Demonstration of the carbon-sulfur bond targeted desulfurization of benzothiophenes by thermophilic Paenibacillus sp. strain A11-2 capable of desulfurizing dibenzothiophene. FEMS Microbiol. Lett. 187:151-154.[CrossRef][Medline]
  24. Konishi, J., Y. Ishii, T. Onaka, K. Okumura, and M. Suzuki. 1997. Thermophilic carbon-sulfur bond-targeted biodesulfurization. Appl. Environ. Microbiol. 63:3164-3169.[Abstract]
  25. Kropp, K. G., J. T. Anderson, and P. M. Fedorak. 1997. Biotransformation of three dimethyldibenzothiophenes by pure and mixed bacterial cultures. Environ. Sci. Technol. 31:1547-1554.[CrossRef]
  26. Landini, D., and F. Rollo. 1974. A convenient synthesis of primary and secondary dialkyl sulfides in the presence of phase-transfer catalysts. Synthesis 8:565-566.[CrossRef]
  27. Lange, G. C., and C. Gottardo. 1980. Facile conversions of primary and secondary alcohols to alkyl iodides. Synth. Comm. 20:1473-1479.
  28. Lee, M. K., J. D. Senius, and M. J. Grossman. 1995. Sulfur-specific microbial desulfurization of sterically hindered analogs of dibenzothiophene. Appl. Environ. Microbiol. 61:4362-4366.[Abstract]
  29. Lee, T.-S., S.-H. Chan, W. A. Weigand, and W. E. Bentley. 2001. A metabolic model for thiodiglycol degradation: capacity constraint leads to byproduct accumulation. Bioprocess Biosyst. Eng. 24:33-38.
  30. Lei, B., and S.-C. Tu. 1996. Gene overexpression, purification, and identification of a desulfurization enzyme from Rhodococcus sp. strain IGTS8 as a sulfide/sulfoxide monooxygenase. J. Bacteriol. 178:5699-5705.[Abstract/Free Full Text]
  31. Li, M. Z., C. H. Squires, D. J. Monticello, and J. D. Childs. 1996. Genetic analysis of the dsz promotor and associated regulatory regions of Rhodococcus erythropolis IGTS8. J. Bacteriol. 178:6409-6418.[Abstract/Free Full Text]
  32. MacPherson, T., C. W. Greer, E. Zhou, A. M. Jones, G. Wisse, P. C. K. Lau, B. Sankey, M. J. Grossman, and J. Hawari. 1998. Application of SPME/GC-MS to characterize metabolites in the biodesulfurization of organosulfur model compounds in bitumen. Environ. Sci. Technol. 32:421-426.[CrossRef]
  33. Matsui, T., T. Onaka, Y. Tanaka, T. Tezuka, M. Suzuki, and R. Kurane. 2000. Alkylated benzothiophene desulfurization by Rhodococcus sp. strain T09. Biosci. Biotechnol. Biochem. 64:596-599.[CrossRef][Medline]
  34. Murgich, J., J. A. Abanero, and O. P. Strausz. 1999. Molecular recognition in aggregates formed by asphaltene and resin molecules from the Athabasca oil sand. Energy Fuels 13:278-286.[CrossRef]
  35. Oldfield, C., N. T. Wood, S. C. Gilbert, F. D. Murray, and F. R. Faure. 1998. Desulphurization of benzothiophene and dibenzothiophene by actinomycete organisms belonging to the genus Rhodococcus, and related taxa. Antonie Leeuwenhoek 74:119-132.
  36. Omori, T., Y. Saiki, K. Kasuga, and T. Kodama. 1995. Desulfurization of alkyl and aromatic sulfides and sulfonates by dibenzothiophene-desulfurizing Rhodococcus sp. strain SY1. Biosci. Biotechnol. Biochem. 59:1195-1198.
  37. Pearson, R. J., W. P. Inskeep, J. M. Wraith, S. D. Comfort, and H. M. Gaber. 1996. Observed and simulated solute transport under varying water regimes: I. Bromide and pentafluorobenzoic acid. J. Environ. Qual. 25:646-653.[Abstract/Free Full Text]
  38. Piddington, C. S., B. R. Kovacevich, and J. Rambosek. 1995. Sequence and molecular characterization of a DNA region encoding the dibenzothiophene desulfurization operon of Rhodococcus sp. strain IGTS8. Appl. Environ. Microbiol. 61:468-475.[Abstract]
  39. Prashad, M., Y. Lu, H.-Y. Kim, B. Hu, O. Repic, and T. J. Blacklock. 1999. An improved and practical Sharpless oxidation of primary alcohols to the carboxylic acids. Synth. Comm. 29:2937-2942.
  40. Rhee, S.-K., J. H. Chang, Y. K. Chang, and H. N. Chang. 1998. Desulfurization of dibenzothiophene and diesel oils by a newly isolated Gordona strain, CYKS1. Appl. Environ. Microbiol. 64:2327-2331.[Abstract/Free Full Text]
  41. Schlömann, M., E. Schmidt, and H.-J. Knackmuss. 1990. Different types of dienelactone hydrolase in 4-fluorobenzoate-utilizing bacteria. J. Bacteriol. 172:5112-5118.[Abstract/Free Full Text]
  42. Schlömann, M., P. Fischer, E. Schmidt, and H.-J. Knackmuss. 1990. Enzymatic formation, stability, and spontaneous reactions of 4-fluoromuconolactone, a metabolite of the bacterial degradation of 4-fluorobenzoate. J. Bacteriol. 172:5119-5129.[Abstract/Free Full Text]
  43. Trost, B. M., and D. P. Curran. 1981. Chemoselective oxidation of sulfides to sulfones with potassium hydrogen persulfate. Tetrahedron Lett. 22:1287-1290.[CrossRef]
  44. Uotila, J. S., V. H. Kitunen, T. Saastamoinen, T. Coote, M. M. Häggblom, and M. S. Salkinoja-Salonen. 1992. Characterization of aromatic dehalogenases of Mycobacterium fortuitum CG-2. J. Bacteriol. 174:5669-5675.[Abstract/Free Full Text]
  45. Van Hamme, J. D., E. T. Wong, H. Dettman, M. R. Gray, and M. A. Pickard. 2003. Dibenzyl sulfide metabolism by white-rot fungi. Appl. Environ. Microbiol. 69:1320-1324.[Abstract/Free Full Text]

Applied and Environmental Microbiology, March 2004, p. 1487-1493, Vol. 70, No. 3
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.3.1487-1493.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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