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Journal of Bacteriology, January 2007, p. 464-472, Vol. 189, No. 2
0021-9193/07/$08.00+0     doi:10.1128/JB.01310-06

Complete and Integrated Pyrene Degradation Pathway in Mycobacterium vanbaalenii PYR-1 Based on Systems Biology ^,

Seong-Jae Kim,^1, Ohgew Kweon,^1, Richard C. Jones,^2, James P. Freeman,³ Ricky D. Edmondson,² and Carl E. Cerniglia^1*

Division of Microbiology,¹ Division of Systems Toxicology,² Division of Biochemical Toxicology, National Center for Toxicological Research, U.S. Food and Drug Administration, Jefferson, Arkansas 72079³

Received 17 August 2006/ Accepted 23 October 2006

Top Abstract Introduction Materials and Methods Results Discussion References

 
Mycobacterium vanbaalenii PYR-1 was the first bacterium isolated by virtue of its ability to metabolize the high-molecular-weight polycyclic aromatic hydrocarbon (PAH) pyrene. We used metabolic, genomic, and proteomic approaches in this investigation to construct a complete and integrated pyrene degradation pathway for M. vanbaalenii PYR-1. Genome sequence analyses identified genes involved in the pyrene degradation pathway that we have proposed for this bacterium. To identify proteins involved in the degradation, we conducted a proteome analysis of cells exposed to pyrene using one-dimensional gel electrophoresis in combination with liquid chromatography-tandem mass spectrometry. Database searching performed with the M. vanbaalenii PYR-1 genome resulted in identification of 1,028 proteins with a protein false discovery rate of <1%. Based on both genomic and proteomic data, we identified 27 enzymes necessary for constructing a complete pathway for pyrene degradation. Our analyses indicate that this bacterium degrades pyrene to central intermediates through o-phthalate and the ß-ketoadipate pathway. Proteomic analysis also revealed that 18 enzymes in the pathway were upregulated more than twofold, as indicated by peptide counting when the organism was grown with pyrene; three copies of the terminal subunits of ring-hydroxylating oxygenase (NidAB2, MvanDraft_0817/0818, and PhtAaAb), dihydrodiol dehydrogenase (MvanDraft_0815), and ring cleavage dioxygenase (MvanDraft_3242) were detected only in pyrene-grown cells. The results presented here provide a comprehensive picture of pyrene metabolism in M. vanbaalenii PYR-1 and a useful framework for understanding cellular processes involved in PAH degradation.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Polycyclic aromatic hydrocarbons (PAHs) are produced by incomplete combustion of fossil fuels, waste incineration, and industrial processes and occur naturally due to forest and prairie fires (8, 62). Pyrene, a PAH containing four fused benzene rings, is commonly found as a pollutant of air, water, and soil (20). Although there is no evidence that pyrene is carcinogenic to humans, nonhuman primates, or laboratory animals, when tested on mouse skin simultaneously with benzo[a]pyrene, it enhanced the carcinogenic effects of benzo[a]pyrene (20). Pyrene is toxic to the aquatic microinvertebrate Gammarus pulex and can be transformed into quinone metabolites that are mutagenic and toxic to organisms in the environment (3, 5, 51). Microbial degradation studies have shown that it is much more difficult to remove high-molecular-weight (HMW) PAHs with four or more fused aromatic rings than it is to remove low-molecular-weight PAHs (24) since HMW PAHs are thermodynamically stable and hydrophobic and they are adsorbed to solid particles (59). Pyrene has been used as a model compound to study biodegradation of HMW PAHs since it is structurally similar to several carcinogenic PAHs (24). Although a number of bacterial isolates have been reported to grow on or mineralize pyrene, the majority of these isolates are nocardioform actinomycetes, such as members of the genera Mycobacterium (4, 12, 19, 26, 40, 48, 56, 60) and Rhodococcus (6, 61).

Mycobacterium vanbaalenii PYR-1 (29, 33) was originally isolated from oil-contaminated sediment based on its ability to degrade pyrene (18). This bacterium was the first isolate reported to mineralize pyrene by mono- and dioxygenase reactions (19). PYR-1 can also degrade or transform biphenyl, naphthalene, anthracene, fluoranthene, 1-nitropyrene, phenanthrene, benzo[a]pyrene, benz[a]anthracene, and 7,12-dimethylbenz[a]anthracene (16, 17, 27, 28, 35, 43-47). Because of its metabolic versatility, this bacterium has been thought to be a potential candidate for bioremediation of PAH-contaminated areas (41).

Pyrene degradation pathways have been proposed for M. vanbaalenii PYR-1 (19, 30, 35, 57, 58) (Fig. 1). Based on identification of metabolites and on ¹⁸O₂ incorporation experiments, this bacterium can oxidize pyrene by two pathways. First, it can oxidize pyrene via initial dioxygenation at the C-1 and C-2 positions to form O-methylated derivatives (Fig. 1) of pyrene-1,2-diol (36), as a detoxification step. However, the predominant second pathway is dioxygenation initiated at the C-4 and C-5 positions (K-region) (19) to give cis-4,5-dihydroxy-4,5-dihydropyrene (pyrene cis-4,5-dihydrodiol). Rearomatization of the dihydrodiol and subsequent ring cleavage dioxygenation lead to the formation of 4,5-dicarboxyphenanthrene, which is further decarboxylated to 4-phenanthroate. The subsequent intermediate, cis-3,4-dihydroxyphenanthrene-4-carboxylate, is produced by a second dioxygenation reaction. Rearomatization then forms 3,4-dihydroxyphenanthrene, which is further metabolized to 1-hydroxy-2-naphthoate. The next enzymatic reactions, including intradiol ring cleavage dioxygenation, result in the production of o-phthalate (19, 35, 58). Although not yet described in members of the genus Mycobacterium, protocatechuate produced from phthalate is believed to be further transformed via the ß-ketoadipate pathway (49) to tricarboxylic acid (TCA) cycle intermediates. Since the presence of phthalate eliminates the possibility that the pathway proceeds through salicylate (37) and protocatechuate is known to be degraded through the ß-ketoadipate pathway in gram-positive bacteria (10, 23), we assume that the bacterium likely utilizes this pathway for the degradation of pyrene (Fig. 1). However, despite all the metabolic information, little is known about the biochemical and genetic mechanisms for pyrene degradation in strain PYR-1.

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FIG. 1. Complete pyrene degradation pathway in M. vanbaalenii PYR-1 based on metabolite, genome, and proteome analyses. The prefix S indicates an enzymatic step. The pyrene metabolic intermediates are as follows: P1, pyrene cis-4,5-dihydrodiol; P2, 4,5-dihydroxypyrene; P3, phenanthrene-4,5-dicarboxylate; P4, phenanthrene-4-carboxylate; P5, cis-3,4-dihydroxyphenanthrene-4-carboxylate; P6, 3,4-dihydroxyphenanthrene; P7, 2-hydroxy-2H-benzo[h]chromene-2-carboxylate; P8, 1-hydroxy-2-naphthaldehyde; P9, 1-hydroxy-2-naphthoate; P10, trans-2'-carboxybenzalpyruvate; P11, 2-carboxylbenzaldehyde; P12, phthalate; P13, phthalate 3,4-dihydrodiol; P14, 3,4-dihydroxyphthalate; P15, proto- catechuate; P16, ß-carboxy-cis,cis-muconate; P17, -carboxymuconolactone; P18, ß-ketoadipate enol-lactone; P19, ß-ketoadipate; P20, ß-ketoadipyl-CoA; P21, pyrene cis-1,2-dihydrodiol; P22, 1,2-dihydroxypyrene; P23, 1-methoxy-2-hydroxypyrene; P24, 1-hydroxy-2-methoxypyrene; P25, 1,2-dimethoxypyrene. The abbreviations for metabolic intermediates identified in M. vanbaalenii PYR-1 are italicized.

A draft genome sequence of M. vanbaalenii PYR-1 has recently been reported (http://img.jgi.doe.gov). This sequence was estimated to contain 6,012 protein-encoding sequences, and the analysis revealed the presence of many putative genes involved in catabolism of aromatic compounds. Since the initial report in 1986 on the microbial degradation and mineralization of pyrene (9), it has been anticipated that elucidation of a complete pathway for pyrene degradation should be forthcoming. Here we describe, for the first time, a complete and integrated pyrene degradation pathway, which was determined using a combination of metabolite identification and genome and proteome analyses.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Bacterial strain and culture conditions. M. vanbaalenii PYR-1 (= DSM 7251) was used in this study. Pyrene was purchased from Chem Service (Media, PA), and its purity (100%) was confirmed by gas chromatography-mass spectrometry (GC-MS). A loop of M. vanbaalenii PYR-1 cells grown on Middlebrook 7H11 agar (Remel, Lenexa, KS) was inoculated into a 250-ml flask containing 50 ml of phosphate-based minimal medium (34) supplemented with 0.5% sorbitol and incubated with shaking at 200 rpm for 4 to 5 days at 28°C. The culture was then divided and used to inoculate two 250-ml flasks, each containing 100 ml of phosphate-based minimal broth with 0.5% sorbitol. Pyrene (final concentration, 25 µM) was dissolved in dimethyl sulfoxide and added to a flask when the culture reached an optical density at 600 nm of approximately 1, as measured spectrophotometrically. The same incubation conditions were used for the control culture, except that pyrene was not added.

Analytical methods. Pyrene metabolites were extracted with ethyl acetate and processed for analysis as described previously (36). GC-MS was performed with either a Finnigan TSQ 700 or 7000 triple quadrupole mass spectrometer (Thermo Finnigan, San Jose, CA) with separation of the metabolites using a J&W DB-5ms capillary column (30 m by 0.25 mm; film thickness, 0.25 µm). GC-MS analyses were performed with a capillary column temperature rate of 10°C/min and a total analysis time of 30 min or less, depending on the PAH analyzed. The retention times with the newer instrument configuration were slightly shorter due to the 10% reduction in capillary column length. Electron ionization was achieved at an electron energy of 70 eV and an ion source temperature of 150°C. Derivatization prior to a GC-electron ionization MS analysis was performed by silylation with N,O-bis(trimethylsilyl)-trifluoroacetamide with 1% trimethylchlorosilane (Regis Technologies, Morton Grove, IL). The samples were dissolved in 400 µl of acetonitrile. Two hundred microliters of a dissolved sample and 100 µl of silylation reagent were mixed in an Xpertek high-recovery 1.5-ml sample vial (P. J. Cobert, St. Louis, MO) sealed with a septum cap and were allowed to react for 1 h at 68°C. The injection volume was 0.5 µl. All mass spectrometric measurements were obtained at low resolution, and no tandem mass spectrometry methods were employed. Therefore, all fragmentation losses are reported below as assumptions based on the proposed structures and available moieties.

Protein extraction. After harvest by centrifugation, the cell pellets were washed twice with 50 mM Tris-HCl (pH 7.4) and disrupted with glass beads using a FastProtein Blue kit (Qbiogene, Carlsbad, CA). Briefly, the cells (0.2 g, wet weight) were suspended in 1 ml of BugBuster protein extraction reagent (Novagen, Madison, WI) containing 0.3% sodium dodecyl sulfate (SDS), 1 µl (25 U) Benzonase nuclease (Novagen), and 0.5 mg/ml protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO) and transferred to a FastProtein Blue tube, and this was followed by homogenization using a FastPrep instrument (Qbiogene) at a speed of 6.0 m/s for two 40-s cycles. The tubes were spun at 10,000 x g for 60 min at 4°C to remove the cell debris and lysing matrix. The protein concentration was determined using a PlusOne 2-D Quant kit (GE Healthcare Bio-Sciences, Piscataway, NJ).

One-dimensional gel electrophoresis (1-DE) proteomics. SDS-polyacrylamide gel electrophoresis was performed using the X-Cell Surelock Mini-Cell electrophoresis system (Invitrogen, Carlsbad, CA). Twenty micrograms of total protein was mixed with SDS loading buffer and reducing agent and resolved on a 4 to 12% gradient minigel (Invitrogen). Bands were visualized with the SimplyBlue Coomassie blue stain (Invitrogen), and each lane was cut into 40 equal slices using a razor blade. In-gel trypsin digestion was performed robotically (ProGest; Genomic Solutions, Ann Arbor, MI), and the resultant peptides were analyzed using an UltraPlus II LC system (Microtech, Vista, CA) coupled to an LCQ Deca XP Plus (Thermo Finnigan) ion trap tandem mass spectrometer (11, 31). Product ion data were searched against the concatenated forward and reverse M. vanbaalenii PYR-1 draft genome database using the Mascot search engine (Matrix Science, London, United Kingdom). Mascot DAT files were parsed into ProteinTrack (11), where nonredundant protein lists were generated with a protein false discovery rate (FDR) of <1% (based on the formula FDR = [number proteins matching the reverse database x 2]/[number of proteins matching the forward database + number of proteins matching the reverse database] x 100%). The criteria for inclusion for multiple peptide hits were a total protein score of 62, with any individual peptide scoring 27 (1+), 23 (2+), and 25 (3+), and a minimum peptide molecular weight of 600, and the criteria for inclusion for single peptide hits were a mascot score of 46 (2+) and 50 (3+), with singly charged single peptide hits not included. Quantitative changes for samples of M. vanbaalenii grown under different conditions were estimated by comparing the normalized total peptide counts for a given protein (where the total peptides included multiple charge states, modifications, or repeats).

Gene prediction and annotation for pyrene metabolic pathway. To search for genes and correlate them with enzymes in the pyrene degradation pathway, the draft genome of M. vanbaalenii was screened. The draft genome data are based on the Joint Genome Institute assembly, released on 1 December 2005. The assembly contained 25 contigs (estimated genome size, 6.46 Mbp) and 6,012 putative gene products. We pooled all available gene/open reading frame (ORF) paralogs that might function in the pathway by using enzymes which have been characterized in other microbial species. We picked enzymes of interest from these organisms and BLASTP (1) searched against the M. vanbaalenii genome sequence. The similarity cutoff for match sequences was the default value (E < 1e-5), provided by the integrated microbial genome website (http://img.jgi.doe.gov/). To validate predictions of gene function, the selected ORFs were analyzed further as follows: conservation of domains or motifs or catalytically important amino acid residues, organization and location of genes with respect to other genes in the pathway, and homologies in the overall sequences. Public databases and the expression data were also correlated to improve the annotation of these genes.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Pyrene induction experiment. Before studies on differential expression of proteins involved in pyrene degradation in M. vanbaalenii PYR-1, we performed metabolism studies to confirm that the culture was highly induced. As shown in Fig. 2, pyrene was totally metabolized after 24 h of incubation. GC-MS analysis of neutral and acidic extracts, as trimethylsilyl derivatives from the pyrene-induced culture, indicated that significant levels of pyrene cis-4,5-dihydrodiol and other hydroxylated intermediates were present. Pyrene metabolic products were identified by comparison with the chromatographic retention times of authentic compounds, molecular and fragment ions in the mass spectra, and previously published values. This experiment confirmed our previous studies that described the isolation and identification of metabolites in the pyrene degradation pathway (19, 35, 57).

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FIG. 2. Pyrene degradation () and metabolite formation by M. vanbaalenii PYR-1. Metabolites were identified as pyrene cis-4,5-dihydrodiol () and dihydroxypyrene ().

Identification of pyrene catabolic enzymes. As shown in Fig. 1, we proposed a complete pyrene degradation pathway based on the key metabolic intermediates that have been identified. However, despite all this information, the pathway has yet to be found in any single organism at the molecular level. Therefore, we performed a 1-DE-based proteomic analysis (gel electrophoresis in combination with liquid chromatography-tandem mass spectrometry [GeLC/MS]) of total protein extracts from control or pyrene-induced cultures to screen and detect proteins involved in the pyrene pathway. The sorbitol culture was chosen because its catabolism was not linked to the pyrene pathway. We observed a large number of proteins with molecular masses up to 220 kDa, as estimated from molecular mass markers on the gel (Fig. 3). The proteome profiles and band intensities were similar for the control and pyrene-induced cultures, and therefore we relied on mass spectrometric analyses for quantitative information. Here, either peptide or spectral counting (39) was used as a semiquantitative method to assess relative protein abundance. The gel was cut into 40 equal slices using a grid template, in-gel digested with trypsin, and analyzed by nano-liquid chromatography-MS-MS. We used the draft PYR-1 genome sequence supplemented with previously reported genes (30, 32, 58) as a database to identify proteins by using product ion data combined with the Mascot search engine. ProteinTrack (11) was used to collate identified proteins into nonredundant lists and also to compare total peptide counts obtained from each lane. The use of a concatenated forward and reverse protein database (50) allowed the FDR to be determined, and in these experiments we employed a protein FDR of <1%. Table S1 in the supplemental material shows the identities of 903 and 827 proteins from the control and pyrene-induced samples, respectively. After protein identification data from both samples were combined, a total of 1,028 unique gene products were identified. Although peptide counting provides only a semiquantitative estimate of relative protein abundance, it can be used to indicate fold changes. Thus, we used the ratio of normalized total numbers of peptides as an approximation of the fold change. The numbers of the total peptides for all proteins identified were 7,365 and 5,991 for the control and pyrene-induced cultures, respectively, and were normalized to the pyrene-induced total. Based on this rationale, 142 proteins were determined to be at least twofold more abundant or completely absent in control cultures compared with pyrene-treated cultures. These proteins were examined first for potential candidates involved in pyrene degradation. Table 1 summarizes these 142 upregulated proteins, which were determined to include 25 proteins involved in aromatic hydrocarbon degradation.

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FIG. 3. Separation of the total proteins of M. vanbaalenii PYR-1 by SDS-polyacrylamide gel electrophoresis. Lane 1, molecular mass ladder; lane 2, 20 µg proteins from noninduced control sample; lane 3, 20 µg proteins from pyrene-induced sample.

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TABLE 1. Summary of 142 proteins upregulated more than twofold identified by proteome analysis

As shown in Table 2, 27 enzymes required for the 25 steps of the complete pyrene degradation pathway were identified. The results agree well with the ORFs assigned in the genome analysis described below, suggesting that the selected ORFs are not silent but translated into proteins. These proteins include 14 proteins responsible for the degradation of pyrene to phthalate, six proteins responsible for the degradation of phthalate to protocatechuate, seven proteins responsible for the lower pathway from protocatechuate to acetyl coenzyme A (acetyl-CoA) and succinyl-CoA, and three proteins responsible for the transformation of pyrene to 1,2-dimethoxypyrene.

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TABLE 2. Proteins involved in the pyrene metabolic pathway identified in M. vanbaalenii PYR-1 grown in the presence of pyrene

Of these gene products, all proteins required for the upper pathway from pyrene to protocatechuate, except PhtAc, were among the proteins whose amount increased twofold or more. This result suggests that these enzymes are involved in pyrene degradation. The level of PhtAc, an oxygenase ferredoxin component of the ring-hydroxylating oxygenase (RHO), was not estimated to be increased. However, we included PhtAc in the degradation pathway based on functional evidence and the genetic location. Of particular note, all terminal oxygenase subunits, as well as dihydrodiol dehydrogenase and a ring cleavage dioxygenase involved in the upper pathway, were detected only in the pyrene-grown cells and not in the control, suggesting that these molecules are tightly regulated in the pyrene pathway.

Identification of pyrene catabolic genes. The genome sequence was analyzed in correlation with the pyrene metabolic pathway in M. vanbaalenii PYR-1 (Fig. 4 and Table 2). Using the genome sequence and previously published genetic data for PYR-1, genes/ORFs were assigned to each enzymatic step required for the pathway, which confirmed the previous metabolite studies. As shown in Table 2 and Fig. 4, a search of the PYR-1 genome assembly revealed 27 genes/ORFs organized in four contigs, which encode all the steps for the degradation of pyrene to acetyl-CoA and succinyl-CoA or to 1,2-dimethoxypyrene (Fig. 1). The data in Table 2 are data for genes/ORFs whose expression was confirmed by a proteomic analysis by GeLC/MS as described above.

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FIG. 4. Organization of the gene clusters involved in the catabolism of pyrene in M. vanbaalenii PYR-1. Genes/ORFs are represented by arrows. The gene products identified as proteins involved in pyrene degradation are represented by shaded arrows with numbers that correspond to enzymatic steps in the pathway shown in Fig. 1. The numbers above the arrows indicate the ORF locus tag numbers (Table 2) from the PYR-1 draft genome (http://img.jgi.doe.gov). Genes identified in previous studies are in italics. The numbers of contigs are also indicated. Vertical lines indicate that genes are not adjacent in the genome. The gene sizes are not proportional to the sizes of the arrows.

The assignment of genes/ORFs was straightforward when direct experimental results were available from M. vanbaalenii PYR-1. For example, nidAB (enzymatic step S1), nidA3B3 (S1/22), phtAaAb (S13), phtB (S14), and MvanDraft_0992 (S24/25) can be assigned to their functional steps in the pathway as shown in Fig. 2 and 4 and Table 2. The involvement of these enzymes in the pathway was confirmed previously; nidAB, nidA3B3, and phtAaAb encoding terminal oxygenase subunits and phtAcAd encoding ferredoxin/reductase components of the RHO enzyme were expressed in Escherichia coli, and their substrate specificities were determined (30, 36, 57). The enzymatic functions of phtB and MvanDraft_0992, encoding cis-3,4-dihydro-3,4-dihydroxyphthalate dehydrogenase and catechol O-methyltransferase, respectively, were also determined (30, 36, 57).

Genes/ORFs were also confidently assigned when enzymes in the pathway were functionally characterized in other microorganisms. For example, PdoA2B2 was selected from Mycobacterium sp. strain 6PY1 to search for the enzyme responsible for the conversion of 4-phenanthroate to cis-3,4-dihydroxyphenanthrene-4-carboxylate in the pathway (enzymatic step S5). The PdoA2B2 enzyme was shown to have the same dioxygenase activity in strain 6PY1 (38). A BLAST search of the PdoA2 sequence of 6PY1 with the genome sequence detected a total of 18 paralogs encoding dioxygenases which exhibit levels of sequence similarity starting at 99%. The product of the second highest ORF exhibits 63% sequence similarity with PdoA2. We assigned the first ORF to step S5. In this way, ORFs for steps S4/15 (decarboxylase), S10 (1-hydroxy-2-naphthoate dioxygenase), S11 (trans-2'-carboxybenzalpyruvate hydratase-aldolase), S12 (2-carboxybenzaldehyde dehydrogenase), S13 (phthalate 3,4-dioxygenase), S16 (protocatechuate 3,4-dioxygenase), S18/19 (-carboxymuconolactone decarboxylase/ß-ketoadipate enol-lactone hydrolase), and S20 (ß-ketoadipate succinyl CoA transferase) were successfully annotated.

Many genes involved in aromatic degradation were found to be clustered together. In this case, studies have shown that the function of a gene can often be deduced from the gene organization or arrangement (14, 15, 52, 55). We analyzed the genomic contexts to search missing ORFs for which functional evidence is lacking. For example, in the assignment of dihydrodiol dehydrogenase, which is the enzyme for steps S2, S6, and S23 (Fig. 1), PhdE from Nocardioides sp. strain KP7 was used to identify the corresponding homologs in the M. vanbaalenii PYR-1 genome. Although phdE, encoding a dihydrodiol dehydrogenase, was not functionally characterized, the gene was adjacent to other genes in the same locus for phenanthrene degradation in Nocardioides sp. strain KP7 (55). We found two possible paralogs similar to PhdE and identified the ORF, MvanDraft_0815 (76% sequence similarity), as an ORF encoding a potential enzyme for these steps. Inspection of the location of this ORF revealed that it is located along with other genes encoding enzymes for steps S4 (decarboxylase) and S5 (phenanthrene RHO, subunit). The second ORF candidate, MvanDraft_0715, belongs to a chromosomal cluster whose genes are predicted to be required for biphenyl degradation. Likewise, the gene homologs for ring cleavage dioxygenase (S3/7), hydratase-aldolase (S8), aldehyde dehydrogenase (S9), ß-carboxy-cis,cis-muconate cycloisomerase (S17), and ß-ketoadipyl-CoA thiolase (S21) were assigned in this way. The selected ORFs for these enzymes were also detected downstream or upstream of other ORFs in the pyrene pathway, as shown by the genetic organization (Fig. 4).

Top Abstract Introduction Materials and Methods Results Discussion References

 
The ability of M. vanbaalenii PYR-1 to degrade pyrene has been extensively studied, and the metabolic intermediates have been identified. Based on careful stoichiometric and flux balance analyses, plausible metabolic pathways have been proposed (19, 35, 57, 58). However, to our knowledge, a complete pyrene metabolic pathway from pyrene to the central TCA cycle intermediates has not been described previously. In this investigation, we identified all the proteins involved in pyrene degradation by proteomic approaches, which has never been reported for any organism, including strain PYR-1. We also identified all the genes involved in the pyrene pathway using genome sequence analysis, and the results matched the proteome results well. The genomic and proteomic analyses in the current work showed that pyrene is degraded through the ß-ketoadipate pathway in M. vanbaalenii PYR-1 at the molecular level.

Previously, parts of 12 of 27 genes responsible for pyrene degradation were found, and it was speculated that these genes are involved in phenanthrene degradation based on simple sequence comparisons (58). However, most of the genes in the previous work were annotated without observation of protein expression or functional evidence. The recent draft genome sequence provided a new framework for elucidation of the genomic and enzymatic basis of pyrene metabolism in M. vanbaalenii PYR-1. An integrative systemic approach using metabolic, genomic, and proteomic data could then be conducted to construct the pyrene degradation pathway in strain PYR-1.

Initially, we started annotating genes for the enzymatic steps of pyrene degradation using our previous gene and enzyme expression studies (7, 30, 32, 57, 58). However, since direct genetic and enzymatic information about M. vanbaalenii PYR-1 was still unavailable for several steps of the pathway, we screened genome sequences to identify pyrene degradation enzymes. We collected many candidate genes/ORFs that might have functions in pyrene metabolism. Since genes for enzymes functionally identical to enzymes for steps in the pathway have been experimentally identified in PYR-1, as well as in other microorganisms, including Mycobacterium sp. strain 6PY1 (pyrene degradation pathway) (38), Nocardioides sp. strain KP7 (phenanthrene degradation pathway) (21, 22, 55), and Terrabacter sp. strain DBF63 (phthalate degradation pathway) (14), we were able to search for putative genes for the pathway enzymes. The enzymes of interest from these bacteria were then subjected to BLAST searches against the M. vanbaalenii genome sequence. The ORFs recovered were analyzed further to deduce their most probable functions.

The draft genome analysis of M. vanbaalenii PYR-1 predicted the existence of a pyrene catabolic pathway. However, genomic information reflects only a static state, and analysis of specific protein expression in the enzymatic pathway in response to pyrene exposure is a prerequisite to elucidating the mechanism of degradation. The genome sequence revealed that M. vanbaalenii PYR-1 possesses multiple ORF homologs with similar functions, which are distributed throughout several chromosomal gene clusters. For example, at least 22 and 7 ORF homologs encoding terminal oxygenase subunits of the RHO enzyme and ring cleavage dioxygenase, respectively, were found in the genome. Since it is not considered likely that all of these ORFs are either responsive to or involved in pyrene degradation, functional assignment based on simple sequence comparisons might result in misleading interpretations of the enzymes involved in the pathway. The detailed proteomics used in this study of M. vanbaalenii PYR-1 complemented the predictive information obtained by genome analysis and enabled us to unambiguously identify the pyrene metabolic pathway utilized by the bacterium when it is grown on pyrene.

A total cell extract of M. vanbaalenii PYR-1 grown on the medium containing pyrene was used for this proteomic study. In previous two-dimensional (2-DE) studies of M. vanbaalenii PYR-1, we detected proteins responding to several PAHs (30, 31), some of which were involved in pyrene or fluoranthene degradation (30, 32). However, we could not determine all the proteins involved in PAH degradation, mainly because sequence data for the bacterium were limited at the time. In addition, hundreds of protein spots displayed on the 2-DE gel also made accurate detection of induced proteins difficult. Although 2-DE in combination with spot intensity is widely used for resolution of complex protein mixtures and is often a powerful tool for identifying significantly expressed marker proteins, identification of proteins that are poorly soluble or not abundant or that have a low molecular mass is limited (13). Therefore, with the availability of the genome sequence, we used a 1-DE-based proteomic approach (GeLC/MS) to screen and detect the proteins induced by pyrene.

Our proteome analyses identified a total of 1,028 protein species, including 142 proteins that showed upregulation upon exposure of the bacterium to pyrene. These proteins, which were estimated to be induced more than twofold, were initially screened as candidates involved in pyrene degradation. A striking result of this work was the observation that 16 of 17 enzymes required for the conversion of pyrene to protocatechuate were detected among the 142 upregulated proteins. Induction of proteins involved in pyrene metabolism has been observed in M. vanbaalenii PYR-1(18, 19, 30-32). In contrast, it was not clear whether the synthesis of the proteins in the ß-ketoadipate pathway responsible for protocatechuate degradation was induced.

The proteomic data along with the genome sequence led to identification of potential enzymes involved in the 21 enzymatic steps required for degradation of pyrene to TCA cycle intermediates in M. vanbaalenii PYR-1. The results are consistent with the proposed pyrene pathway, as well as the enzymatic functional studies. In the pathway (Fig. 1), enzymatic steps S1, S4, S13, and S22 were identified as steps that are catalyzed by RHO, a multicomponent enzyme often consisting of four polypeptides, terminal oxygenase subunits and ß and two electron transfer components, ferredoxin and ferredoxin reductase (42). The two electron transfer components have often been found to be essential for the enzyme activity. For these four steps, we identified NidAB2, MvanDraft_0816/0817, PhtAaAb, and NidA3B3, respectively, as terminal oxygenases. We also identified PhtAc and PhtAd, a ferredoxin and ferredoxin reductase, respectively, as electron transfer components for these four steps. Since phtAc is expected to be in the same transcription unit as phtAd, whose expression was increased 5.3-fold, it is curious that phtAc was not found to be upregulated, as phtAd was in the proteome results. However, we included the gene in the pathway because the protein was functionally confirmed and phtAcAd was the only gene set encoding electron transfer components of the RHO enzyme that was detected in the genome. This kind of numerical imbalance between oxygenase and electron transfer components has been reported previously; multiple gene copies for terminal oxygenase were found with only one set of genes for the electron transfer components, reductase and ferredoxin (52, 54). It was also shown previously that the one copy of the electron transfer component gene is shared by the multiple terminal oxygenases for full enzyme activity (25, 52). In a previous experiment, a biotransformation study with E. coli revealed that PhtAcAd could function with PhtAaAb and convert phthalate into its dihydrodiol intermediate (57). Since NidAB and another terminal oxygenase, NidA3B3 (32), also transformed PAHs into their dihydrodiols in cooperation with PhtAcAd (data not shown), it is assumed that the single-copy gene product, PhtAcAd, can function with these terminal oxygenases in M. vanbaalenii PYR-1.

We also screened the genome sequence to see whether enzymes involved in the alternative routes of pyrene degradation are present since metabolic diversity has been reported in many other bacteria. However, no ORFs for enzymes such as 1-hydroxy-2-naphthoate hydroxylase, which is necessary for 1-hydroxy-2-naphthoate to be degraded through salicylate (37), were detected. In Pseudomonas spp., 1-hydroxy-2-naphthoate is hydroxylated to form 1,2-dihydroxynaphthalene, which is further mineralized into central metabolites via salicylate and catechol (37). ORFs for phthalate 4,5-dioxygenase and protocatechuate 4,5-dioxygenase, which are found in the optional routes for oxidation of phthalate (2) and the meta cleavage of protocatechuate (53), respectively, also were not detected in the genome sequence. These findings are in agreement with the pyrene pathway predicted from the metabolic information. Additionally, we checked the proteins identified in Table S1 to determine whether other proteins that might function in the pathway were present or induced. For example, as mentioned above, a total of 22 paralogs for a gene encoding terminal oxygenase of the RHO enzyme were found in the genome, and seven oxygenase copies were identified in the gels; of these, four enzymes (NidAB2, the MvanDraft_0817/0818 product, PhtAaAb, and the ORF25/26 product) were determined to be induced. We list three (NidAB2, the MvanDraft_0817/0818 product, and PhtAaAb) of these four induced proteins in Table 2 as enzymes involved in the pyrene pathway. The physiological role of the fourth induced protein, the ORF25/26 product, in pyrene degradation was unclear. The other three enzymes appeared to be constitutively expressed. Two of these enzymes, the MvanDraft_0806/0807 and MvanDraft_0811/0812 products, were not identified on the basis of sequence homology to be involved in pyrene degradation. The last enzyme, MvanDraft_0800/0801 (NidA3B3), was previously identified as a terminal oxygenase of the RHO enzyme, which showed enzyme activity for several PAHs, including fluoranthene (32). The 142 induced proteins also include proteins with different functions (Table 2). Elucidation of the physiological roles of these proteins requires further study.

In conclusion, this study is believed to be the first study to elucidate a complete pathway for the conversion of pyrene to TCA cycle intermediates. The metabolic pathway was analyzed in combination with genomic and proteomic approaches, which provided the first comprehensive picture of pyrene degradation in M. vanbaalenii PYR-1. The pathway that we describe in this work is unlike what one might predict simply based on the analysis of the genome. This study also provides a strong basis for understanding of the metabolic versatility for degradation of all other PAHs in this bacterium.

 
We thank John B. Sutherland, Mark Hart, and Ashraf A. Khan for critical reviews of the manuscript. We express our special thanks to Charles D. Miller and Ronald C. Sims at Utah State University for their efforts in organizing the genome project.

This work was supported by an appointment to the Postgraduate Research Program at the National Center for Toxicological Research administered by the Oak Ridge Institute for Science and Education through an interagency agreement between the U.S. Department of Energy and the U.S. Food and Drug Administration.

 
* Corresponding author. Mailing address: Division of Microbiology, NCTR/US FDA, 3900 NCTR Road, Jefferson, AR 72079. Phone: (870) 543-7341. Fax: (870) 543-7307. E-mail: Carl.Cerniglia{at}fda.hhs.gov.

Published ahead of print on 3 November 2006.

Supplemental material for this article may be found at http://jb.asm.org/.

S.-J.K., O.K., and R.C.J. contributed equally to this work.

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Journal of Bacteriology, January 2007, p. 464-472, Vol. 189, No. 2
0021-9193/07/$08.00+0     doi:10.1128/JB.01310-06

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