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. 1998 May;64(5):1884–1889. doi: 10.1128/aem.64.5.1884-1889.1998

Oxidation of Methyl-Substituted Naphthalenes: Pathways in a Versatile Sphingomonas paucimobilis Strain†

Tapan K Dutta 1,2,*, Sergey A Selifonov 3,‡, Irwin C Gunsalus 2
PMCID: PMC106246  PMID: 9572967

Abstract

Aromatic compounds with alkyl substituents are abundant in fossil fuels. These compounds become important environmental sources of soluble toxic products, developmental inhibitors, etc. principally through biological activities. To assess the effect of methyl substitution on the completeness of mineralization and accumulation of pathway products, an isolate from a phenanthrene enrichment culture, Sphingomonas paucimobilis 2322, was used. Washed cell suspensions containing cells grown on 2,6-dimethylnaphthalene in mineral medium were incubated with various mono-, di-, and trimethylnaphthalene isomers, and the products were identified and quantified by gas chromatography and mass spectrometry. The data revealed enzymes with relaxed substrate specificity that initiate metabolism either by methyl group monoxygenation or by ring dioxygenation. Congeners with a methyl group on each ring initially hydroxylate a methyl, and this is followed by conversion to a carboxyl; when there are two methyl groups on a single ring, the first reaction is aryl dioxygenation of the unsubstituted ring. Intermediates are channeled to primary ring fission via dihydrodiols to form methyl-substituted salicylates. Further evidence that there are multiple pathways comes from the fact that both phthalate and (methyl)salicylate are formed from 2-methylnaphthalene.

Alkylated aromatic hydrocarbons enter the ecosystem mostly from crude oil- and petroleum-derived materials. Many of these compounds are man-made products of large-scale industry and are used as fuels, solvents, detergent precursors, and commercial insect repellents (for example, methylnaphthalenes dispersed in aromatic solvents for mosquito control) (42, 47). Because of the abundance of methyl-substituted aromatic compounds and the formation of water-soluble products, the effects of these compounds on the biosphere may be greater than the toxic damage caused by the unsubstituted hydrocarbon counterparts (1, 10, 19, 29, 30, 37).

Removal of aromatic compounds from the biosphere is principally a microbial activity. Gibson et al. (20) first reported that unsubstituted structures undergo initial ring reductive dioxygenation to cis-dihydrodiols. With ring substituents, the positions exert a major effect on the first reaction site, the pathway, and the products. With a single methyl addition, ring dioxygenation may be retained, or methyl monoxygenation may first form the analogous benzylic alcohol. The most extensive studies have examined toluene, the simplest and most abundant methylated aromatic hydrocarbon. A recent summary of six known schemes for toluene metabolic patterns are reviewed in studies of fungal oxygenation (45). The products and genetics of both ring dioxygenation (the TOD pathway) (48) and side chain monooxygenation (the TOL pathway) (2) are well-documented. Data are also available for representative aromatized plant terpenes and homologs, including p-cymene (15), 3-ethyltoluene (28), and the xylene isomers (3, 16). The attention that has been devoted to methyl derivatives is limited (1, 5, 31, 39) compared with the attention that has been devoted to unsubstituted fused ring compounds (4, 9, 14, 27, 43). Moreover, although a very large number of strains have been isolated, few organisms have received the attention needed for even minimal biological and chemical characterization, and fewer still have been subjected to a genetic analysis to determine chromosome organization, regulation, and gene origin. The detailed data available suggest there is a limited variety of oxygenase catalytic centers in animal, plant, and microbial systems. Marked similarities among mono- and dioxygenase systems have led to the concept that there are enzyme superfamilies (36). Three oxygenase centers that contain metal (iron) prosthetic groups have been well-documented. The most abundant center, which was the first center documented, is a unique heme-thiolate monoxygenase center which now contains more that 500 related, but separate, protein sequences (36). Two other well-studied oxygenase reaction centers are the Fe2(μO)2 cluster, which was found first in methane monoxygenase and later in related nonheme di-iron enzymes (41), and disassociable ferrous, mononuclear nonheme iron coordination sites (25). The essential two-electron reducing power of both mono- and dioxygenase systems is provided primarily by flavoprotein reductases; here too, a limited number of structural patterns are found (17), and there is marked homology to the anaplerotic heterotrophic and photosynthetic flavoproteins (6, 17, 22, 23, 32).

A variety of bacterial strains and species have been used to identify products of methyl-substituted aromatic hydrocarbon metabolism. The profiles obtained are the principal sources of deduced oxygenation-oxidation patterns. Most previous studies on fused ring alkyl aromatic compounds have been concerned with the 1- and 2-methylnaphthalenes and the isomer effects on initial methyl monooxygenation (13, 31, 39, 46) compared with ring dioxygenation (7, 8, 14, 44, 46). Pseudomonas putida CSV86, for example, has been reported to convert 1- and 2-methylnaphthalenes to the respective methyl catechols by ring dioxygenation or to form naphthoic acids following methyl hydroxylation (31). In addition, the 2-methyl isomer has been reported to yield 4-hydroxymethyl catechol (31), implying that there is dual-pathway regulation. In vitro eucaryote oxygenation has also been found in hepatic microsomes (11, 26), fungi (13), and cyanobacteria (12), the role in plant cells is unclear.

Data on fused ring dimethyl structures are more limited; thus, the pathways are not understood as well (5, 21, 33). In early studies Barnsley (5) and other workers identified ring fission products obtained from fused ring aromatic compounds, and subsequently the process was well-documented by Eaton and Chapman (18). Barnsley (5) hypothesized that there is an initial 2-methyl hydroxylation, followed by oxidation to carboxylate and then a ring monohydroxylation to 1-hydroxy-6-methylnaphthalene-2-carboxylic acid. More recently, several workers have described dioxygenation to 1-hydroxy-2-naphthoic acid as an alternate product obtained from phenanthrene, 2-methylnaphthalene, or 2-naphthoic acid (4, 5, 34). Workers have also described methyl group oxidation of dimethylnaphthalene isomers by the naphthalene dioxygenase cloned from plasmid NAH7 (40) and by Nocardia corallina (Rhodococcus sp.) grown on aliphatic hydrocarbons (24, 38).

In this report we describe the effect of methyl-substituted aromatic isomers on the metabolic pathway(s) in washed cell suspensions of Sphingomonas paucimobilis 2322 grown on 2,6-dimethylnaphthalene (2,6-DMN) as a carbon and energy source. Our data documented that relaxed enzyme specificity and isomer substitution effects on mineralization pathways occur.

MATERIALS AND METHODS

Organism, cultures, chemicals, and products.

S. paucimobilis 2322 from the collection of Peter Chapman (Gulf Ecology Division, U.S. Environmental Protection Agency) was isolated from a creosote-contaminated soil by enrichment on phenanthrene as the carbon source. The biological systematic properties of this organism were determined and fatty acid analyses and a 16S rRNA composition analysis were performed by Midi Laboratories, Newark, Del. Cells were grown in a liquid mineral salt medium (pH 7.0) containing (per liter) 3.24 g of K2HPO4, 0.87 g of NaH2PO4, 2.0 g of NH4Cl, 123 mg of nitrilotriacetic acid, 200 mg of MgSO4 · 7H2O, 12 mg of FeSO4 · 7H2O, 3 mg of MnSO4 · H2O, 3 mg of ZnSO4 · 7H2O, and 1 mg of CoCl2 · 6H2O. Solid media contained 2% agar (Difco). Cells were grown by incubating them at 30°C on a rotary shaker (180 rpm) in 2-liter Erlenmeyer flasks containing 400 ml of mineral salt medium supplemented with 1 g of a carbon source per liter. The cells were harvested from 48-h-old cultures by centrifugation (8,000 × g, 10 min), washed twice by suspending them in a volume equal to 1/10th the growth volume, collected by centrifugation, and resuspended (final volume, 100 ml) to an optical density at 600 nm of 2.0. For fermentation studies a hydrocarbon was added to a concentration of 1 g/liter to washed cell suspensions in individual Erlenmeyer flasks, and the flasks were incubated at 30°C for up to 72 h. After incubation, cells (and undissolved substrate, when present) were removed by centrifugation, and the supernatants were acidified to pH 2 to 3 with concentrated hydrochloric acid and extracted twice with equal volumes of ethyl acetate. The extracts were dried over anhydrous sodium sulfate and evaporated under reduced pressure at 30°C, and the materials recovered were methylated with an ethereal solution of diazomethane prior to gas chromatography-mass spectrometry (GC-MS) analysis. All chemicals were of the highest purity commercially available.

Analyses.

GC-MS was performed with a Hewlett-Packard model 5890 series II gas chromatograph equipped with a model HP5971 mass-selective detector and a model HP 5965B infrared detector. Compounds were separated on a type HP-5 capillary column (25 m by 0.32 mm; film thickness, 0.25 μm) by using helium as the carrier gas at a flow rate of 23 cm/s. The column temperature was kept isothermally at 50°C for 1 min and then increased to 290°C at a rate of 5°C/min. The mass spectrometer was operated at an electron ionization energy of 70 eV, and the injector, transfer line, and analyzer temperatures were 150, 300, and 300°C, respectively. Instrumental library searches and comparison with available authentic compounds were used to identify possible metabolites. The relative amounts of metabolites were calculated from the peak areas of GC-MS total ion current chromatograms. The total ion current integrated responses (excluding the peak of residual starting material) were used as 100% internal standard values for each sample analyzed. Parallel processes without substrate were also included. Thin-layer chromatography was performed on 0.25-mm-thick silica gel (Merck Silica Gel 60 F254) plates by using methylene chloride-ethyl acetate (1:1) as the developing solvent, and the results were used for preliminary identification. Starting substrates and metabolites were detected under UV light and by exposure to iodine vapor. All of the chemicals used were of the highest purity available commercially, and identities were confirmed by GC-MS analysis.

RESULTS AND DISCUSSION

Culture growth.

It was shown previously that S. paucimobilis 2322 grows well aerobically on several aromatic compounds, including 2,6-DMN, biphenyl, o-xylene, p-cymene, and acenaphthene, as well as phenanthrene (13a). To confirm that there are a number of growth substrates in the catabolic module, several compounds were tested, as shown in Fig. 1. As discussed below a number of related structures failed to support growth. With 2,6-DMN yellow metabolites were present in culture filtrates.

FIG. 1.

FIG. 1

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Growth of S. paucimobilis 2322 in minimal salt medium containing aromatic compounds as carbon and energy sources. 2,3,6-DMN, 2,3,6-trimethylnaphthalene; Phn, phenanthrene.

Metabolic pathway with 2,6-DMN.

Harvested and washed cell suspensions oxidized a variety of methylnaphthalenes whose metabolites, as detected by thin layer chromatography, were later identified; the levels of these metabolites were determined by GC-MS, as shown in Table 1. The initial methyl group oxidation, a common event with the isomers tested, led to the proposed metabolic patterns shown in Fig. 2.

TABLE 1.

GC-MS data for products formed from methylnaphthalenes by S. paucimobilis 2322 grown on 2,6-DMNa

Product (compound) Relative amt (%)b GC Rt (min) m/z of major ion peaks (%) Suggested structure
From 2,6-DMN
 1 27.5 11.34 166 (M+) (34), 135 (21), 134 (100), 106 (23), 105 (11), 78 (17), 77 (18) 5-Methylsalicylic acid methyl esterc,d
 2 3.2 16.24 172 (M+) (100), 157 (23), 143 (11), 141 (8), 129 (32), 128 (29), 115 (10) 2-Hydroxymethyl-6-methylnaphthalenee
 3 50.0 17.37 200 (M+) (71), 185 (2), 169 (100), 155 (2), 141 (53), 139 (16), 115 (27) 6-Methyl-2-naphthoic acid methyl esterd
 4 8.5 18.01 216 (M+) (100), 201 (15), 187 (10), 186 (20), 173 (29), 158 (7), 145 (13), 129 (17), 119 (17), 115 (39) 6-Hydroxymethyl-2-naphthoic acid methyl estere
 5 0.5 18.58 234 (M+) (12), 216 (29), 200 (68), 184 (27), 175 (29), 169 (100), 157 (87), 141 (60), 129 (47), 115 (52) 1,2-Dihydroxy-1-hydro-6-methyl-2-naphthoic acid methyl estere
 6 0.5 20.71 244 (M+) (70), 213 (100), 185 (47), 170 (13), 169 (17), 154 (23), 126 (39), 115 (24) Naphthalene-2,6-dicarboxylic acid dimethyl esterd
From 2,7-DMN
 7 36.1 11.34 166 (M+) (34.5), 134 (100), 106 (26), 105 (20), 78 (19), 77 (18) 4-Methylsalicylic acid methyl esterc,d
 8 1.0 16.18 172 (M+) (100), 157 (33), 143 (22), 141 (38), 129 (64), 128 (50), 115 (25) 2-Hydroxymethyl-7-methylnaphthalenee
 9 57.7 17.28 200 (M+) (75), 185 (1), 169 (100), 155 (2), 141 (60), 115 (27) 7-Methyl-2-naphthoic acid methyl estere
 10 2.2 18.54 234 (M+) (12), 216 (26), 200 (71), 184 (26), 175 (32), 169 (100), 157 (82), 141 (67), 129 (46), 115 (52) 1,2-Dihydroxy-1-hydro-7-methyl-2-naphthoic acid methyl estere
From 2,3-DMN
 11 90.9 13.38 180 (M+) (34), 165 (1), 149 (21), 148 (100), 141 (2), 120 (22), 105 (3), 91 (28) 4,5-Dimethylsalicylic acid methyl esterc,d
 12 4.2 17.30 200 (M+) (72), 185 (1), 169 (100), 155 (3), 141 (56), 138 (18), 115 (27) 3-Methyl-2-naphthoic acid methyl esterc,d
From 1,5-DMN
 13 26.0 11.47 166 (M+) (35), 135 (24), 134 (100), 106 (31), 105 (22), 78 (18), 77 (17) 6-Methylsalicylic acid methyl esterc,d
 14 44.0 17.32 200 (M+) (74), 185 (3), 169 (100), 155 (2), 141 (58), 115 (32.7) 5-Methyl-1-naphthoic acid methyl esterc
From 1,6-DMN
 15 40.0 11.30 166 (M+) (35), 134 (100), 106 (27), 105 (21), 94 (7), 78 (27), 77 (20) 5-Methylsalicylic acid methyl esterc,d
 16 40.0 11.45 166 (M+) (46), 134 (100), 106 (40), 105 (34), 94 (26), 78 (35), 77 (44) 6-Methylsalicylic acid methyl esterc,d
 17 10.8 17.29 200 (M+) (74), 185 (2), 169 (100), 155 (9), 141 (73), 128 (5), 115 (37) 5(6)-Methyl-2(1)-naphthoic acid methyl estere
From 1,8-DMN
 18 2.0 11.05 166 (M+) (41), 141 (15), 134 (100), 115 (14), 106 (65), 94 (35), 78 (40), 77 (31) 3-Methylsalicylic acid methyl esterc,d
 19 67.0 15.67 170 (M+) (75), 155 (3), 142 (52), 141 (100), 139 (21), 115 (35), 84 (10) 8-Methyl-1-naphthaldehydec
 20 3.0 16.79 172 (M+) (41), 154 (73), 153 (100), 143 (13), 141 (21), 128 (30), 115 (22) 1-Hydroxymethyl-8-methylnaphthalenee
 21 1.9 19.18 184 (M+) (80), 155 (100), 139 (11), 127 (88), 101 (6), 77 (12) 1,8-Naphthalided
From 1,2-DMN
 22 70.8 13.12 180 (M+) (38), 149 (22), 148 (100), 120 (76), 105 (8), 91 (29) 3,4-Dimethylsalicylic acid methyl esterc,d
 23 1.4 16.32 172 (M+) (95.7), 157 (45), 154 (74), 143 (74), 141 (78), 129 (100), 128 (85), 115 (57) 2(1)-Hydroxymethyl-1(2)-methylnaphthalenee
From 1,4-DMN
 24 4.2 12.74 180 (M+) (42), 165 (2), 155 (12), 149 (24), 148 (85), 120 (100), 105 (8), 91 (38) 3,6-Dimethylsalicylic acid methyl esterc,d
 25 94.0 17.37 200 (M+) (61), 185 (2), 169 (100), 141 (48), 115 (28) 4-Methyl-1-naphthoic acid methyl esterc
From 2,3,6-trimethylnaphthalene
 26 32.4 13.36 180 (M+) (33), 165 (1), 149 (21), 148 (100), 120 (23), 105 (3), 91 (29) 4,5-Dimethylsalicylic acid methyl esterc,d
 27 13.0 18.22 214 (M+) (95), 199 (8), 183 (68), 182 (70), 155 (66), 154 (100), 139 (15), 128 (26), 115 (12) 3,6-Dimethyl-2-naphthoic acid methyl estere
 28 30.0 19.07 214 (M+) (85), 199 (3), 183 (100), 169 (2.1), 155 (46), 139 (9), 128 (9), 115 (8) 6,7-Dimethyl-2-naphthoic acid methyl estere
From 2-methylnaphthalene
 29 1.6 9.47 152 (M+) (46), 121 (37), 120 (100), 93 (14), 92 (49), 65 (24) Salicylic acid methyl esterc,d
 30 2.5 11.23 166 (M+) (35), 134 (100), 106 (28), 105 (24), 78 (20), 77 (19) 4-Methylsalicylic acid methyl esterc,d
 31 0.3 13.23 194 (M+) (8), 163 (100), 149 (6), 135 (8), 115 (18), 109 (20), 77 (31) Phthalic acid dimethyl esterc,d
 32 5.8 15.77 186 (M+) (57), 155 (100), 137 (11), 127 (85), 126 (14), 115 (8) 2-Naphthoic acid methyl esterc,d
 33 7.5 16.98 202 (M+) (28), 171 (18), 170 (100), 142 (7), 115 (19), 114 (55) 1-Hydroxy-2-naphthoic acid methyl esterc,d
 34 58.0 17.31 220 (M+) (9), 202 (27), 186 (2), 170 (7), 161 (37), 155 (3), 143 (100), 131 (27), 115 (95) 1,2-Dihydroxy-1-hydro-2-naphthoic acid methyl estere
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a

Analyses were performed after methylation with diazomethane. 

b

Based on the total ion current response of the mass-selective detector. 

c

Identification was based on an instrument library match. 

d

Identification was based on a match of mass spectra (fragmentation and peak intensity) and capillary GC retention times with data for authentic samples. 

e

Tentative assignment of structure. 

FIG. 2.

FIG. 2

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Proposed pathway(s) for conversion of various methylnaphthalenes by S. paucimobilis 2322 grown on 2,6-DMN. The numbers of the metabolites detected are the numbers in Table 1, and the substitution patterns of the methylnaphthalene precursors are indicated in parentheses. TCA, tricarboxylic acid.

The fact that hydroxylation of 2,6-DMN occurred at the 2-methyl position was indicated by the presence of 2-hydroxymethyl-6-methylnaphthalene and its oxidation product, 6-methyl-2-naphthoic acid; the aldehyde intermediate was not detected in the reaction mixture (5, 21, 33). In contrast to the previous report of ring monooxygenation by Barnsley (5), a potential product of dioxygenation with properties of 1,2-dihydroxy-1-hydro-6-methyl-2-naphthoic acid was observed. The data support the pathway reported for naphthalene by Eaton and Chapman (18). The ring fission product appeared to be compound 35 or compound 36 (2-hydroxy-6-methylchromene-2-carboxylate or trans-2-hydroxy-5-methylbenzylidinepyruvate) as a meta fission product of 1,2-dihydroxy-6-methylnaphthalene (Fig. 2). A GC-MS analysis of the methylated derivative gave a relative amount of 4.2% (GC Rt = 16.60 min; MS m/z [%]: 234 [M+] [3], 219 [1], 203 [6], 187 [10], 175 [100], 160 [2], 159 [5], 143 [6], 132 [10], 115 [13]). Compound 36 led to a salicylate (in this case, 5-methylsalicylate), a common intermediate. An alternate, dead end route appeared to be oxidation of compound 36 to compound 33 (2,6-naphthalene dicarboxylic acid) (Fig. 2). Further support came from a lack of growth on or product formation from 2,6-naphthalene dicarboxylic acid by washed cells grown on 2,6-DMN.

2,7-, 1,5-, 1,6-, and 1,8-DMNs.

The reactions of 2,7-DMN were similar to the reactions of the 2,6-DMN isomer, as indicated by the presence of 2-hydroxymethyl-7-methylnaphthalene, 7-methyl-2-naphthoic acid, 1,2-dihydroxy-1-hydro-7-methyl-2-naphthoate, and 4-methylsalicylate (Table 1 and Fig. 2); the data indicate that there was dioxygenation at the 1,2 positions, forming 6- and 7-methyl-2-naphthoates. These are characteristic fission products found in the benzoate-toluate pattern (35).

The 1,5-, 1,6-, and 1,8-DMN isomers gave analogous methylnaphthoate and methylsalicylate accumulations. When 1,6-DMN was used, GC-MS analysis revealed both 5- and 6-methylsalicylates (i.e., hydroxylation occurred at either the β-methyl group or the β-methyl group). The 1,8-DMN isomer products were “upper-pathway” products (i.e., 1-hydroxymethyl-8-methylnaphthalene, 8-methyl-1-naphthaldehyde, and 1,8-naphthalide, a spontaneously cyclized product of 8-hydroxymethyl-1-naphthoic acid obtained under acidic extraction conditions) (40).

2,3,6-Trimethylnaphthalene.

With methyl groups on both rings and a third substituent at position 3−, methyl hydroxylation occurred as the first committed step, at either carbon 2− or carbon 6−, and oxidation to the carboxylate led to compounds 27 and 28 (the 3,6-dimethyl- and 6,7-dimethyl-2-naphthoates) (Table 1). The subsequent ring dioxygenation, however, appeared to be specific for the (renumbered) 2,3-dimethyl-6-naphthoate, for only 4,5-dimethylsalicylate was present. The S. paucimobilis products obtained from 2,3,6-trimethylnaphthalene are consistent with dioxygenation of the benzoate-toluate type (35).

1,2-, 1,4-, and 2,3-DMNs.

As indicated above, placing both methyl groups on the same carbocyclic ring directed the initial reaction to dioxygenation of the unsubstituted aryl group and altered the early products. A subsequent meta ring fission should then have led to the respective dimethylsalicylates (that is, the 1,2-, 1,4-, and 2,3-DMNs and conversion to the respective 3,4-, 3,6-, and 4,5-dimethylsalicylic acids). The final products obtained from the salicylic acids converged regardless of whether one or both aryl ring substitutions occurred. An alternate oxidation, to the benzyl alcohols, yielded the apparent dead end compounds 23, 12, and 25, for neither salicylate nor phthalate derivatives were detected in the spent growth medium or after reactions with cell suspensions. The data are compatible with the presence of the phenanthrene dioxygenase, which apparently is coinduced in cells grown with 2,6-DMN.

2-Methylnaphthalene.

2-Methylnaphthalene, although not a growth substrate, was metabolized by washed cells grown on 2,6-DMN. The products indicate that initially either methyl hydroxylation or dioxygenation of an unsubstituted aryl ring occur. Thus, ring fission led to either 4-methylsalicylate or phthalate. Figure 2 shows the same dual pathways found in phenanthrene-grown cells.

Dioxygenation initiated on the unsubstituted aryl ring occurred by formation of a cis-dihydrodiol at the 1,2 carbons. As indicated in Fig. 2, the data suggest that dehydrogenation to the analogous vicinal 1,2-diphenol (a catechol) and ring fission to 4-methylsalicylate via compound 37 or 38 (2-hydroxy-7-methylchromene-2-carboxylate or trans-2-hydroxy-4-methylbenzylidinepyruvate) occur. A GC-MS analysis of the methylated product revealed a relative amount of 6% (GC Rt = 16.71 min; MS m/z [%]: 234 [M+] [4], 219 [1], 203 [6], 187 [14], 175 [100], 160 [1], 159 [5], 143 [6], 132 [12], 115 [19]).

A β-methyl hydroxylation first led to phthalate or salicylate, depending on the reaction of compound 33. The presence of 2-naphthoate and 1-hydroxy-2-naphthoate recalls the Bradley monoxygenase hypothesis (5). When 2-naphthoate (compound 32) was used as the substrate, the products indicated that both mono- and dioxygenase pathways were present, based on identification of compounds 33 and 34, which led to phthalate (compound 31) or 4-methylsalicylate (compound 30). Phthalate generation from 1-hydroxy-2-naphthoate and 2-naphthoate is a well-established pathway in phenanthrene metabolism (4, 34).

To summarize, S. paucimobilis grown on phenanthrene generates phthalate via 1-hydroxy-2-naphthoate and salicylate in trace amounts in spent growth medium and washed cell suspensions. Thus, although the phenanthrene and 2,6-DMN pathways are considered separate, the two pathways seem to be coinduced by growth on either substrate, suggesting that there is a common regulation mechanism.

A comment on the growth substrates of strain 2322 is in order. In general, the 2-substituted dimethylnaphthalene isomers are suitable growth substrates, as are the 2,3,6-trimethyl structure and phenanthrene. Naphthalene and mono-substituted 2-naphthalenes, however, do not support growth. Similarly, in strains of this type, neither 1-substituted dimethylnaphthalenes nor 1- or 2-naphthoate supports growth. The persistence of these compounds in nature requires more detailed consideration, along with the effects of other microbic strains (31), but this was beyond the scope of this work.

ACKNOWLEDGMENTS

This work was supported in part by a National Research Council Resident Research Associateship Award (to T.K.D.) and by cooperative agreement CR823946-01-0 with the U.S. Environmental Protection Agency (to S.A.S.).

We thank Maureen Downey (Avanti Corporation) for carrying out GC-MS analyses. We also thank Peter J. Chapman for supplying strain 2322, for making reference chemicals available, and for helpful discussions. We are grateful to Peter J. Chapman for reviewing the manuscript and making insightful suggestions.

Footnotes

†

Contribution 1024 from the NHEERL, Gulf Ecology Division, U.S. Environmental Protection Agency, Gulf Breeze, Fla.

REFERENCES

  • 1.Alexander R, Kagi R I, Rowland S J, Sheppard P N, Chirila T V. The effect of thermal maturity on distribution of dimethylnaphthalenes and trimethylnaphthalenes in some ancient sediments and petroleums. Geochim Cosmochim Acta. 1985;49:385–395. [Google Scholar]
  • 2.Assinder S J, Williams P A. The TOL plasmids: determinants of the catabolism of toluene and xylenes. Adv Microb Physiol. 1990;31:1–69. doi: 10.1016/s0065-2911(08)60119-8. [DOI] [PubMed] [Google Scholar]
  • 3.Baggi G, Barbieri P, Galli E, Tollari S. Isolation of a Pseudomonas stutzeri strain that degrades o-xylene. Appl Environ Microbiol. 1987;53:2129–2132. doi: 10.1128/aem.53.9.2129-2132.1987. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Barnsley E A. Phthalate pathway of phenanthrene metabolism: formation of 2′-carboxybenzalpyruvate. J Bacteriol. 1983;154:113–117. doi: 10.1128/jb.154.1.113-117.1983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Barnsley E A. Metabolism of 2,6-dimethylnaphthalene by flavobacteria. Appl Environ Microbiol. 1988;54:428–433. doi: 10.1128/aem.54.2.428-433.1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Batie C J, Ballou D P, Correll C J. Phthalate di-oxygenase reductase and related flavin-iron-sulfur containing electron transferases. In: Muller F, editor. Chemistry and biochemistry of flavoenzymes. Boca Raton, Fla: CRC Press; 1991. pp. 543–556. [Google Scholar]
  • 7.Bestetti G, Di Gennaro P, Galli E, Leoni B, Pelizzoni F, Sello G, Bianchi D. Bioconversion of substituted naphthalenes to the corresponding salicylic acids. Appl Microbiol Biotechnol. 1994;40:791–793. [Google Scholar]
  • 8.Bestetti G, Bianchi D, Bosetti A, Di Gennaro P, Galli E, Leoni B, Pelizzoni F, Sello G. Bioconversion of substituted naphthalenes to the corresponding 1,2-dihydro-1,2-dihydroxy derivatives. Determination of the regio- and stereochemistry of the oxidation reactions. Appl Microbiol Biotechnol. 1995;44:306–313. [Google Scholar]
  • 9.Boldrin B, Tiehm A, Fritzsche C. Degradation of phenanthrene, fluorene, fluoranthene, and pyrene by a Mycobacterium sp. Appl Environ Microbiol. 1993;59:1927–1930. doi: 10.1128/aem.59.6.1927-1930.1993. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Boylan D B, Tripp B W. Determination of hydrocarbons in sea water extracts of crude oil and crude oil fractions. Nature (London) 1971;230:44–47. doi: 10.1038/230044a0. [DOI] [PubMed] [Google Scholar]
  • 11.Breger R K, Franklin R B, Lech J J. Metabolism of 2-methylnaphthalene to isomeric dihydrodiols by hepatic microsomes of rat and rainbow trout. Drug Metab Dispos. 1981;9:88–93. [PubMed] [Google Scholar]
  • 12.Cerniglia C E, Freeman J P, Athaus J R, Van Baalen C. Metabolism and toxicity of 1- and 2-methylnaphthalene and their derivatives in cyanobacteria. Arch Microbiol. 1983;136:177–183. [Google Scholar]
  • 13.Cerniglia C E, Lambert K J, Miller D W, Freeman J P. Transformation of 1- and 2-methylnaphthalene by Cunninghamella elegans. Appl Environ Microbiol. 1984;47:111–118. doi: 10.1128/aem.47.1.111-118.1984. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13a.Chapman, P. Personal communication.
  • 14.Dean-Raymond D, Bartha R. Biodegradation of some polynuclear aromatic petroleum components by marine bacteria. Dev Ind Microbiol. 1975;16:97–110. [Google Scholar]
  • 15.Defrank J J, Ribbons D W. p-Cymene pathway in Pseudomonas putida: initial reactions. J Bacteriol. 1977;129:1356–1364. doi: 10.1128/jb.129.3.1356-1364.1977. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Duggleby C J, Williams P A. Purification and some properties of the 2-hydroxy-6-oxohepta-2,4-dienoate hydrolase (2-hydroxymuconic semialdehyde hydrolase) encoded by the TOL plasmid pWW0 from Pseudomonas putida mt2. J Gen Microbiol. 1986;132:717–726. [Google Scholar]
  • 17.Dutta T K, Gunsalus I C. Reductase gene and protein structures: p-cymene methyl hydroxylase. Biochem Biophys Res Commun. 1997;233:502–506. doi: 10.1006/bbrc.1997.6493. [DOI] [PubMed] [Google Scholar]
  • 18.Eaton R W, Chapman P J. Bacterial metabolism of naphthalene: construction and use of recombinant bacteria to study ring cleavage of 1,2-dihydroxynaphthalene and subsequent reactions. J Bacteriol. 1992;174:7542–7554. doi: 10.1128/jb.174.23.7542-7554.1992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Fedorak P M, Westlake D W S. Microbial degradation of aromatics and saturates in Prudhoe Bay crude oil as determined by glass capillary chromatography. Can J Microbiol. 1981;27:432–443. doi: 10.1139/m81-066. [DOI] [PubMed] [Google Scholar]
  • 20.Gibson D T, Hensley M, Yoshioka H, Mabry T J. Formation of (+)-cis-2,3-dihydroxy-1-methylcyclohexa-4,6-diene from toluene by Pseudomonas putida. Biochemistry. 1970;9:1626–1630. doi: 10.1021/bi00809a023. [DOI] [PubMed] [Google Scholar]
  • 21.Griffoll M, Selifonov S A, Gatlin C V, Chapman P J. Actions of a versatile fluorene-degrading bacterial isolate on polycyclic aromatic compounds. Appl Environ Microbiol. 1995;61:3711–3723. doi: 10.1128/aem.61.10.3711-3723.1995. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Gunsalus I C, Francesconi S C, Dutta T K. Abstracts of the UIB-GBF-CSIC-TUB Symposium on Biodegradation of Organic Pollutants. 1996. Gene-enzyme superfamily effects on biosphere chemistry; pp. 15–16. [Google Scholar]
  • 23.Harayama S, Kok M. Functional and evolutionary relationships among diverse oxygenases. Annu Rev Microbiol. 1992;46:565–601. doi: 10.1146/annurev.mi.46.100192.003025. [DOI] [PubMed] [Google Scholar]
  • 24.Jamison V W, Raymond R L, Hudson J O. Microbial hydrocarbon co-oxidation. III. Isolation and characterization of an α,α′-dimethyl-cis,cis-muconic acid-producing strain of Nocardia corallina. Appl Microbiol. 1969;17:853–856. doi: 10.1128/am.17.6.853-856.1969. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Jiang H, Parales R E, Lynch N A, Gibson D T. Site-directed mutagenesis of conserved amino acids in the alpha subunit of toluene dioxygenase: potential mononuclear nonheme iron coordination sites. J Bacteriol. 1996;178:3133–3139. doi: 10.1128/jb.178.11.3133-3139.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Kaubisch N, Daly J W, Jerina D M. Arene oxides as intermediates in the oxidative metabolism of aromatic compounds. Isomerization of methyl-substituted arene oxides. Biochemistry. 1972;11:3080–3088. doi: 10.1021/bi00766a022. [DOI] [PubMed] [Google Scholar]
  • 27.Kiyohara H, Torigoe S, Kaida N, Asaki T, Iida T, Hayashi H, Takizawa N. Cloning and characterization of a chromosomal gene cluster, pah, that encodes the upper pathway for phenanthrene and naphthalene utilization by Pseudomonas putida OUS82. J Bacteriol. 1994;176:2439–2443. doi: 10.1128/jb.176.8.2439-2443.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Kunz D A, Chapman P J. Catabolism of pseudocumene and 3-ethyltoluene by Pseudomonas putida (arvilla) mt2: evidence for new function of the TOL (pWW0) plasmid. J Bacteriol. 1981;146:179–191. doi: 10.1128/jb.146.1.179-191.1981. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Lee C C, Craig W K, Smith P J. Water soluble hydrocarbons from crude oil. Bull Environ Contam Toxicol. 1974;12:212–216. doi: 10.1007/BF01684964. [DOI] [PubMed] [Google Scholar]
  • 30.Lee Y-Z, Leighton E A, Peakall D B, Norstorm R J, O’Brien P J, Payne J F, Rahimtula A D. Effects of ingestion of Hibernia and Prudhoe Bay crude oils on hepatic and renal mixed function oxidase in nesting herring gulls (Larus argentutus) Environ Res. 1985;36:248–255. doi: 10.1016/0013-9351(85)90022-2. [DOI] [PubMed] [Google Scholar]
  • 31.Mahajan M C, Phale P S, Vaidyanathan C S. Evidence for the involvement of multiple pathways in the biodegradation of 1- and 2-methylnaphthalene by Pseudomonas putida CSV86. Arch Microbiol. 1994;161:425–433. doi: 10.1007/BF00288954. [DOI] [PubMed] [Google Scholar]
  • 32.Mason J R, Cammack R. The electron transport proteins of hydroxylating bacterial dioxygenases. Annu Rev Microbiol. 1992;46:277–305. doi: 10.1146/annurev.mi.46.100192.001425. [DOI] [PubMed] [Google Scholar]
  • 33.Miyachi N, Tanaka T, Suzuki T, Hotta Y, Omori T. Microbial oxidation of dimethylnaphthalene isomers. Appl Environ Microbiol. 1993;59:1504–1506. doi: 10.1128/aem.59.5.1504-1506.1993. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Morawski B, Eaton R W, Rossiter J T, Guoping S, Griengl H, Ribbons D W. 2-Naphthoate catabolic pathway in Burkholderia strain JT 1500. J Bacteriol. 1997;179:115–121. doi: 10.1128/jb.179.1.115-121.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Neidle E L, Hartnett C, Ornston L N, Bairoch A, Rekik M, Harayama S. Nucleotide sequences of Acinetobacter calcoaceticus benABC genes for benzoate 1,2-dioxygenase reveal evolutionary relationships among multicomponent oxygenases. J Bacteriol. 1991;173:5385–5395. doi: 10.1128/jb.173.17.5385-5395.1991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Nelson D R, Koymans L, Kamataki T, Stegeman J J, Feyereisen R, Waxman D J, Waterman M R, Gotoh O, Coon M J, Estabrook R W, Gunsalus I C, Nebert D W. P450 superfamily: update on new sequences, gene mapping, accession numbers and nomenclature. Pharmacogenetics. 1995;6:1–42. doi: 10.1097/00008571-199602000-00002. [DOI] [PubMed] [Google Scholar]
  • 37.Radke M. Organic geochemistry of aromatic hydrocarbons. In: Brooks J, Welte D, editors. Advances in petroleum geochemistry. Vol. 2. London, United Kingdom: Academic Press; 1987. pp. 141–202. [Google Scholar]
  • 38.Raymond R L, Jamison V W, Hudson J O. Microbial hydrocarbon co-oxidation. I. Oxidation of mono- and dicyclic hydrocarbons by soil isolates of the genus Nocardia. Appl Microbiol. 1967;15:857–865. doi: 10.1128/am.15.4.857-865.1967. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Rogoff M H, Wender I. Methylnaphthalene oxidations by pseudomonads. J Bacteriol. 1959;77:783–788. doi: 10.1128/jb.77.6.783-788.1959. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Selifonov S A, Griffoll M, Eaton R W, Chapman P J. Oxidation of naphthenoaromatic and methyl-substituted aromatic compounds by naphthalene 1,2-dioxygenase. Appl Environ Microbiol. 1996;62:507–514. doi: 10.1128/aem.62.2.507-514.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Shu L, Nesheim J C, Kauffmann K, Munck E, Lipscomb J D, Que L., Jr An Fe2IVO2 diamond core structure for the key intermediate Q of methane monooxygenase. Science. 1997;275:515–518. doi: 10.1126/science.275.5299.515. [DOI] [PubMed] [Google Scholar]
  • 42.Sparling J, Chittim B, Clegg B S, Safe S, Crocker J F S. The tissue distribution and clearance of Aerotex 3470, an aromatic hydrocarbon solvent. Chemosphere. 1978;7:607–614. [Google Scholar]
  • 43.Stringfellow W T, Aitken M D. Competitive metabolism of naphthalene, methylnaphthalenes, and fluorene by phenanthrene-degrading pseudomonads. Appl Environ Microbiol. 1995;61:357–362. doi: 10.1128/aem.61.1.357-362.1995. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Treccani V, Flecchi A. Ossidazione microbica della 2-metilnaftalina. Ann Microbiol. 1958;8:36–44. [Google Scholar]
  • 45.Weber F J, Hage K C, de Bont J A M. Growth of the fungus Cladosporium sphaerospermum with toluene as the sole carbon and energy source. Appl Environ Microbiol. 1995;61:3562–3566. doi: 10.1128/aem.61.10.3562-3566.1995. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Williams P A, Catterall F A, Murray K. Metabolism of naphthalene, 2-methylnaphthalene, salicylate and benzoate by Pseudomonas PG: regulation of tangential pathways. J Bacteriol. 1975;124:679–685. doi: 10.1128/jb.124.2.679-685.1975. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Wirtz R A, Turrentine J D, Jr, Fox R C. Area repellents for mosquitoes (Diptera: Culicidae): identification of the active ingredients in a petroleum oil fraction. J Med Entomol. 1981;18:126–128. doi: 10.1093/jmedent/18.2.126. [DOI] [PubMed] [Google Scholar]
  • 48.Zylstra G J, Gibson D T. Toluene degradation by Pseudomonas putida F1: nucleotide sequence of the todC1C2BADE genes and their expression in E. coli. J Biol Chem. 1989;264:14940–14946. [PubMed] [Google Scholar]

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