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3-Nitropropionic acid (3NPA) is a widespread nitroaliphatic toxin found in a variety of legumes and fungi. Several enzymes have been reported that can transform the compound, but none led to the mineralization of 3NPA. We report here the isolation of bacteria that grow on 3NPA and its anion, propionate-3-nitronate (P3N), as the sole source of carbon, nitrogen, and energy. Experiments with resting cells, cell extracts, and purified enzymes indicate that the pathway involves conversion of 3NPA to P3N, which upon denitration yields malonic semialdehyde, nitrate, nitrite, and traces of H2O2. Malonic semialdehyde is decarboxylated to acetyl coenzyme A. The gene that encodes the enzyme responsible for the denitration of P3N was cloned and expressed, and the enzyme was purified. Stoichiometry of the reaction indicates that the enzyme is a monooxygenase. The gene sequence is related to a large group of genes annotated as 2-nitropropane dioxygenases, but the P3N monooxygenase and closely related enzymes form a cluster within COG2070 that differs from previously characterized 2-nitropropane dioxygenases by their substrate specificities and reaction products. The results suggest that the P3N monooxygenases enable bacteria to exploit 3NPA in natural habitats as a growth substrate.
Large-scale release of synthetic nitroaromatic compounds to the biosphere followed the invention of nitrobenzene around 1830. In less than 200 years, microorganisms adapted to the presence of nitroaromatic compounds in the environment by developing catalytic pathways to exploit them as growth substrates. Such rapid development suggests that the pathways did not develop de novo but evolved from preexisting degradation pathways such as might be found in microorganisms that degrade naturally occurring compounds.
3-Nitropropionic acid (3NPA) is a widespread naturally occurring nitroaliphatic compound. It is a principal toxic component of Astragalus locoweeds and has been found in hundreds of species of legumes (20, 39) and a variety of fungi (6). The compound causes irreversible inhibition of succinate dehydrogenase, which makes it deadly to eukaryotes (1). Plants that make 3NPA also contain an enzyme, 3NPA oxidase (NPAO) (19, 20), which converts the compound to malonic semialdehyde (MSA) to protect the plant against the toxic effects of the compound (20). Given its widespread occurrence, we hypothesized that there must be bacteria in soil that degrade 3NPA and play a major role in determining the flux of the compound.
Although bacteria that degrade 3NPA have previously been sought, the focus has been on organisms that ingest 3NPA-containing plant matter. Rumen microorganisms reduce 3NPA to β-alanine (4), and in the grasshopper gut, 3NPA is bound to glycine to form inert conjugates which are then eliminated (24). The plant enzyme NPAO converts 3NPA and O2 to MSA, nitrate, nitrite, and hydrogen peroxide (19). It is similar to propionate-3-nitronate (P3N) oxidase (P3NO; EC 126.96.36.199) from Penicillium atrovenetum that converts the P3N form of the compound to MSA (36). The enzymes mentioned above are “orphan enzymes” (28), which means that the gene(s) has not been identified. None of the previously studied microorganisms can use 3NPA as a growth substrate, and the physiological roles of the enzymes have not been established.
MSA appears to be a central intermediate in the eukaryotic transformation of 3NPA and its analogs. However, the transformations involve distinctly different types of reactions and metabolites released. P3N and 3NPA release nitrate and nitrite in a 2:1 ratio when attacked by the fungal or plant oxidases. When 3NPA is reduced to β-alanine by rumen microorganisms, β-alanine is further metabolized (4), possibly by deamination to MSA (18).
We report here the isolation from soil of aerobic bacteria that grow on 3NPA as the sole source of carbon, nitrogen, and energy. The genes that encode the initial enzymes of the degradation pathway were cloned, and recombinant proteins were purified and partially characterized to allow determination of the initial steps in the catabolic pathway.
(Preliminary reports of this work have been presented previously at the 106th General Meeting of the American Society for Microbiology [32a] and the 108th General Meeting of the American Society for Microbiology [32b].)
Bacteria were isolated by selective enrichment in nitrogen-free minimal medium (BLK) (5) containing 3NPA (100 μM to 1 mM). After several transfers, the cultures were spread onto agar plates containing BLK with 3NPA (500 μM) and incubated at 30°C until colonies appeared. Cultures that grew on 3NPA plates were selected for identification and further study. 16S rRNA gene analysis was performed by MIDI Labs (Newark, DE).
Large cultures were grown in 1 liter of BLK containing twice the normal concentration of MgSO4 plus 3NPA (1 mM). When the 3NPA disappeared from the medium, it was added twice more to a final concentration of 1 mM. Some cultures were grown on 3NPA (500 μM) as the nitrogen source with succinate (7 mM) or acetate (10 mM) as the carbon source. Cultures were harvested by centrifugation, washed twice with phosphate buffer (20 mM; pH 7.0), and stored on ice until used in assays. Cells were broken by three passages through a French pressure cell followed by centrifugation at either 26,000 × g or 100,000 × g. Cell extracts were stored on ice until used.
3NPA denitrase was assayed by measurement of oxygen uptake in the presence of 3NPA or P3N. Reaction mixtures consisted of cell extracts or purified protein (1 to 500 μg protein) in phosphate (20 mM; pH 7.8) or Tris buffer (100 mM; pH 7.8). The reaction was started by addition of substrate. MSA formed from P3N was quantified spectrophotometrically by following the NADH-dependent disappearance of MSA catalyzed by β-hydroxybutyrate dehydrogenase (BDH) (36). Malonyl coenzyme A (malonyl-CoA) reductase was measured in the forward direction by following the disappearance of NADPH at 365 nm (22) and in the reverse direction by following the reduction of NADP (22). MSA decarboxylase was assayed by following the disappearance of MSA in the absence of cofactors (17). MSA oxidative decarboxylase was measured by following the CoA-dependent reduction of NADP to NADPH in the presence of MSA (29). Acetaldehyde dehydrogenase was measured by following the conversion of NAD to NADH in the presence of acetaldehyde (17).
DNA (approximately 5 mg) was extracted from cultures of Pseudomonas sp. strain JS189 with the Promega SV genomic DNA purification system (Madison, WI) according to the manufacturer's instructions. The resulting DNA was sheared by vortexing. DNA fragments approximately 30 kb in size were gel purified, ligated into the fosmid vector pCC1Fos, packaged into phage, and transfected into Escherichia coli strain EPI300-T1 according to the manufacturer's directions (CopyControl fosmid library production kit; Epicentre Biotechnologies, Madison, WI) to create a library of several thousand recombinant E. coli clones.
Clones were transferred to 96-well plates containing tryptic soy broth supplemented with NH4Cl (1 mM), chloramphenicol (20 μg/ml), and 1× concentrated fosmid induction solution (Epicentre Biotechnologies, Madison, WI) and grown for 16 h at 30°C. After growth, the library was replica plated onto BLK supplemented with NH4Cl (1 mM), chloramphenicol (20 μg/ml), 1× concentrated fosmid induction solution, P3N (200 μM), and sodium succinate (200 μM) and then incubated for 12 to 16 h at 30°C. Clones were screened for nitrite release from P3N.
The location of pnoA on the fosmid was determined by transposon mutagenesis followed by a second round of screening for loss-of-function mutants. The fosmid bearing pnoA was purified using the FosmidMax DNA purification kit (Epicentre Biotechnologies, Madison, WI) and then randomly mutated in vitro with the EZ-Tn5 <KAN-2> insertion kit (Epicentre Biotechnologies, Madison, WI). The fosmid::Ez-Tn5 <KAN-2> fusion was reintroduced into E. coli strain EPI300-T1 by electroporation. Mutants in which the transposon had inserted into the fosmid were selected by growth on LB media supplemented with chloramphenicol (20 μg/ml) and kanamycin (25 μg/ml). Kanamycin-resistant transposon mutants were screened for the ability to release nitrite from P3N. Loss-of-function mutants were sequenced using the KANREV primer (41) to determine the insertion site of the transposon. The acquired sequence was used to manually design additional primers for subsequent rounds of primer walking. Approximately 10 kb of sequence was obtained and assembled into a 2-kb contig with CAP3 (21). Open reading frames were determined by ORFFinder (http://www.ncbi.nlm.nih.gov/projects/gorf/).
The Basic Local Alignment Search Tool (BLAST) (3) was used to perform a homology search of the amino acid sequence of PnoA against the protein database in GenBank. Several of the most similar genes (Table (Table1)1) were amplified by PCR. The reaction mixture (20 μl) contained genomic DNA (68 to 115 ng), primers (0.25 μM each), deoxynucleoside triphosphates (dNTPs; 0.4 mM each), buffer (1×), and Promega GoTaq hot-start polymerase (2 U). Amplifications (30 cycles) were carried out as follows: 95°C for 1 min, 54°C for 30 s, and 72°C for 1 min, after initial denaturation at 95°C for 10 min. The PCR products were cut with BamHI and XhoI and cloned into the pET-21a vector (Novagen, Gibbstown, NJ). The resulting constructs were transformed into E. coli Top10 (Invitrogen, Carlsbad, CA) for plasmid propagation and maintenance. The constructs were transformed into E. coli BL21 Star (DE3) (Invitrogen) for expression. Cells were grown in 30 ml of LB medium containing Overnight Express autoinduction system I (Novagen) at 37°C. When the optical density at 600 nm (OD600) reached 0.8 to 0.9, the temperature was reduced to 25°C, and the cultures were further incubated for 10 h. Cells were harvested by centrifugation, washed twice with phosphate buffer (20 mM; pH 7.5) and stored on ice until used. Trees and alignments were constructed with Geneious Pro 4.7.4.
His tag fusions were constructed using the pBAD102-D/TOPO expression system (Invitrogen). Genes of interest were cloned in frame with the N-terminal His patch thioredoxin tag and the C-terminal His tag. The host was E. coli Rosetta 2 (Novagen).
Single colonies of the clone containing pnoA were grown overnight with shaking at 30°C in LB (5 ml) supplemented with ampicillin (125 μg/ml) and chloramphenicol (20 μg/ml) (LBAC). The culture was diluted into fresh LBAC (1 liter) in a 1-liter bioreactor (Applikon, Netherlands) and grown overnight at room temperature to an OD600 of 0.6. Dissolved oxygen was maintained at 50% at a stirring rate of 500 rpm, and pH was controlled at 7.0. Arabinose (0.2% final concentration) was added to induce expression, and incubation was continued until the culture reached stationary phase (10 h). The culture was harvested by centrifugation and washed twice with phosphate buffer (20 mM; pH 7.4). The pellet was stored overnight at −80°C. The pellet was suspended in binding buffer (20 mM phosphate, 20 mM imidazole [pH 7.4]), and the cells were broken with a French press. The extract was clarified by centrifugation at 100,000 × g for 45 min at 4°C. A 5-ml HiTrap chelating HP column (GE Healthcare) was charged with Ni2+, and the protein was purified according to the manufacturer's protocol (14). The fractions were screened for nitrite release from P3N. Benchmark His-tagged protein standards used to estimate the protein size were from Invitrogen.
High-performance liquid chromatography (HPLC) was performed on an Agilent 1100 system equipped with a diode array detector and a Merck Chromolith column (100 mm by 4.6 mm). The mobile phase consisted of 95% trifluoroacetic acid (13 mM) in water (part A) and 5% trifluoroacetic acid (6.5 mM) in acetonitrile (part B) delivered at a flow rate 1 ml min−1. The autosampler and column heaters were maintained at 4 and 45°C, respectively.
Protein was measured using a Pierce bicinchoninic acid kit or a Nanodrop 1000 spectrophotometer by reading A280. Oxygen uptake was measured using a Clark-type electrode and a YSI model 3600 oxygen meter. Nitrite (40), nitrate (32), and ammonia (34) were measured colorimetrically.
Cells grown on -strength tryptic soy agar plates were suspended in phosphate buffer (20 mM; pH 7.2) to an OD600 of ~0.2. The suspension (100 μl) was spread onto agar plates so that the resulting growth would form a lawn. Blank Sensi-Discs (BBL) loaded with 3 mg of P3N (five applications of 10 μl of 500 mM P3N, then dried) were placed onto the spread plates, along with untreated control discs. The plates were incubated at 30°C for up to 10 days (33).
MSA was prepared the day of use from ethyl-3,3-diethoxypropanoate by the method of Yamada and Jakoby (42). Nitronate forms of 3NPA and other nitroalkanes were made immediately before use by addition of two equivalents of KOH (37). Ethyl-3,3-diethoxypropanoate was from Acros Organics. 3NPA, BDH (EC 188.8.131.52), 2-nitroethanol, nitropentane, nitrocyclohexane, β-alanine, and malonyl-CoA were from Sigma-Aldrich. Nitroethane, nitromethane, 1-nitropropane, and 2-nitropropane were from ChemService.
Pseudomonas aeruginosa PAO1 was a gift from Ronald Olsen. Angela Sessitsch and Jerzy Nowak generously provided Burkholderia phytofirmans PsJN.
The sequences for msaD and pnoA have been deposited in GenBank under accession number GU014557. The 16S rRNA gene sequences for JS189 and JS190 have been deposited under accession numbers GU354208 and GU354209, respectively.
Bacteria were isolated from soil and water samples collected from several locations around Atlanta, GA. Isolates that grew and accumulated nitrite and/or nitrate in the culture medium with 3NPA as the sole source of carbon, nitrogen, and energy were presumed to be 3NPA-degrading bacteria. A total of 48 isolates were represented by six different colony morphologies. Two isolates from garden soil were selected for further study. Analysis of partial 16S rRNA gene sequences identified the two strains as Cupriavidus sp., designated JS190, and Pseudomonas sp., designated JS189. When strain JS190 was grown on 3NPA as the sole source of carbon, nitrogen, and energy, 70% of the initial nitrogen accumulated as nitrate, and 2% accumulated as nitrite (Fig. (Fig.1).1). Ammonia was not detected. Results were similar for JS189. Cultures grew on 3NPA at concentrations up to 2 mM, the highest concentration tested. The molar growth yield of JS190 was 24.5 g (dry weight) per mol of 3NPA.
Like many other nitroaliphatic compounds, 3NPA exists in solution in two ionic forms (23), the neutral (acid) form and an anionic nitronate form. At neutral pH, only 1% of the compound is in the nitronate form (2); therefore, in cultures, the predominant form of the compound is likely to be the acid form. Resting cells of 3NPA-grown JS189 and JS190 were 2 to 10 times more active with the nitronate form than with the acid form (Table (Table2),2), as measured by stimulation of oxygen uptake. Rates of oxygen uptake with 3NPA were similar when cells were grown on acetate or succinate (data not shown) with 3NPA as the nitrogen source. Succinate (data not shown)- and acetate (Table (Table2)-grown2)-grown cultures exhibited constitutive activity against P3N but not against 3NPA. No activity was detected with any other substrate tested. Cell extracts were specific for the nitronate (Table (Table3)3) and required no additional cofactors for oxidation of P3N. Dialysis of cell extracts did not affect the denitration reaction. The stoichiometry of the reaction was 0.87 ± 0.14 mol of O2 consumed per mol of P3N.
Fosmid libraries constructed from JS189 and JS190 were screened for the ability to release nitrite from 3NPA. One clone from Pseudomonas sp. strain JS189 was selected for further study. Nitrite release was localized to a 10-kb BamHI-HindIII fragment of a pUC19 subclone. Several colonies that had lost the ability to release nitrite from P3N after transposon mutagenesis were identified. The gene disrupted by Tn5 was designated pnoA.
The purified overexpressed protein encoded by pnoA catalyzed the denitration of P3N with a stoichiometry of 0.95 ± 0.05 mol of O2 consumed per mol of P3N. The substrate range was identical to that of extracts from wild-type cells. The theoretical molecular size of the His-tagged protein was predicted to be 52.5 kDa (Fig. (Fig.2).2). The absorbance spectrum showed maxima at 275 and 446 nm with a shoulder at 368 nm, consistent with that of a flavoprotein (38). The purified protein was stabilized by the addition of bovine serum albumin (BSA; 0.1 to 1.0 mg/ml) to enzyme assays. Rates of reactions in Tris buffer were 5 to 53% of the rates of reactions in phosphate buffer, except when BSA was present in the reaction mixture (data not shown). Activity of the purified protein in phosphate buffer was two to four times that of extracts from wild-type cells (Table (Table3)3) over a range from 5 μM to 10 mM. The Km for P3N was estimated to be 168 μM. When the pnoA gene was amplified by PCR and recloned with the His tag but without the His patch thioredoxin tag, the specific activity improved 6-fold, and the Km was estimated to be 30 μM. Cell extracts from JS189 and JS190 were active over similar concentration ranges and showed the same sensitivity to Tris buffer. Both the purified protein and the wild-type cell extracts gradually lost the ability to catalyze further denitration after multiple (three to four) additions of P3N. The agreement between wild type and the purified protein in all aspects examined indicates that pnoA is responsible for the denitration of the P3N by wild-type cells.
The products of the denitration catalyzed by purified PnoA are consistent with those of P3NO from Penicillium atrovenetum (36) and those of NPAO from Hippocrepis comosa (19), where the enzymes oxidize P3N and 3NPA to MSA with the loss of the nitro group (Table (Table4).4). BDH catalyzes the hydrogenation of MSA to 3-hydroxypropanoate (36). Both authentic MSA and the product of PnoA-catalyzed denitration of P3N resulted in the oxidation of NADH to NAD+ in assays with BDH; no activity was detected when any of the reaction components was omitted. Addition of catalase to reaction mixtures after P3N was completely consumed released an additional 2 to 4% of the original O2 consumed. The stoichiometry of the reaction catalyzed by PnoA with P3N was 178 ± 0.4 P3N + 215 ± 10 O2 → 145 ± 70 NO3− + 45 ± 5 NO2− + 140 ± 31 MSA + 16 ± 7 H2O2. Based upon the similarities of the reaction stoichiometries and the ability of BDH to catalyze NADH oxidation in mixtures with authentic MSA and the product of the PnoA-catalyzed reaction, we concluded that the product was MSA.
MSA was transformed by extracts of cells grown on 3NPA but not by extracts of cells grown on acetate (Table (Table5).5). Assays for various enzymes with the ability to transform MSA (Fig. (Fig.3)3) showed that MSA oxidative decarboxylase is highly upregulated in 3NPA-grown cells but that MSA decarboxylase, acetaldehyde dehydrogenase, and malonyl-CoA reductase are not.
MSA decarboxylase was examined closely because a gene, msaD, similar to the genes that encode MSA decarboxylase (35) was found upstream of pnoA on a 0.5-kb fragment (Fig. (Fig.4A).4A). The gene was cloned with a His tag and a His patch thioredoxin tag. The purified protein rapidly converted MSA to acetaldehyde with no additional cofactors. However, cell extracts from JS189 and JS190 grown on 3NPA had only trace levels of such activity. In addition, acetaldehyde decarboxylase activity was not detected in any cell extracts. The results indicate that MSA oxidative decarboxylase, but not the MSA decarboxylase, is responsible for the metabolism of MSA during 3NPA degradation. The location of the gene that encodes MSA oxidative decarboxylase is currently under investigation.
The proteins most similar to PnoA all belong to COG2070, the 2-nitropropane dioxygenase-like proteins. We cloned and expressed several genes with the highest identity to pnoA (Table (Table1).1). Induced cells and cell extracts catalyzed oxygen uptake and release of nitrite from P3N, but not 3NPA, 2-nitropropane, or propyl-2-nitronate (Table (Table3).3). The wild-type cells all released nitrite from P3N.
JS189, JS190, Burkholderia phytofirmans PsJN, P. aeruginosa PAO1, Cupriavidus necator JMP134, Pseudomonas putida F1, P. putida KT2440, and E. coli BL21 were tested by auxanography for the ability to grow on 3NPA. The proteobacteria were chosen because a BLAST search included their putative 2-nitropropane dioxygenases in the 50 closest matches to PnoA. E. coli BL21(DE3) was included as a control strain lacking a member of COG2070 and because it was the cloning host for the pnoA homologs. All strains except E. coli used 3NPA as the sole nitrogen source. JS189, JS190, and PsJN used 3NPA as the sole source of nitrogen and carbon. When succinate and ammonia were present, there were no detectable effects on the lawns of bacteria, except for strain PsJN, with which there was inhibition close to the source of 3NPA and a heavier growth ring slightly farther away. PsJN also was slightly inhibited by 3NPA in a lawn spread on -strength tryptic soy agar. The results indicate clearly that the genes annotated as 2-nitropropane dioxygenases in the tested strains actually encode P3N monooxygenases.
A variety of enzymes that transform 3NPA and P3N (Table (Table4)4) have been purified from eukaryotes. All are flavoproteins with either FMN or FAD as a tightly bound flavin cofactor. The subunit sizes fall within a narrow range, but the number of subunits of the functional enzyme varies. Nitronate monooxygenase (NMO; EC 184.108.40.206, formerly called 2-nitropropane dioxygenase, EC 220.127.116.11) and nitroalkane oxidase (NAO; EC 18.104.22.168) have broad substrate ranges but greatly prefer nitroalkanes to nitropropionic acid (15, 25-27). P3NO and NPAO have much narrower substrate ranges and are much more active with nitropropionic acid than with nitroalkanes (19, 36). All the enzymes that preferentially attack 3NPA release twice as much nitrate as nitrite from 3NPA, whereas reactions catalyzed by NMO and NAO release no nitrate. The striking similarity of the substrates and products of the PnoA-catalyzed reaction to those of the P3NO-catalyzed reaction from Penicillium atrovenetum (36) suggests that PnoA is a P3NO. However, the lack of stoichiometric release of H2O2 indicates that the enzyme is a monooxygenase (13, 30), and we have tentatively designated it P3N monooxygenase. Full characterization of P3N monooxygenase is currently under way to rigorously establish the reaction mechanism and thus the proper EC designation.
Several lines of evidence indicate that the P3N monooxygenase does not catalyze the first step in the pathway. The fact that acetate-grown cells transform P3N but not 3NPA whereas 3NPA-grown cells transform both forms suggests that the P3N monooxygenase is constitutive. Neither cell extracts nor purified PnoA transformed 3NPA. The results suggest that the initial step in 3NPA degradation is transformation of 3NPA to the nitronate by an unidentified enzyme that is upregulated by growth on 3NPA. It is possible that the failure of acetate-grown cells to transform 3NPA indicates a transport problem rather than lack of induction of an enzyme. Even if that were so, the specificity of cell extracts and purified PnoA for P3N still would suggest a requirement for an enzyme in the 3NPA degradation pathway to catalyze the initial conversion to the nitronate. NMO (Table (Table4)4) from Neurospora crassa (10) is the only enzyme that readily transforms both nitronate and neutral forms of nitroalkanes, although nitronates are the preferred substrates. When the neutral nitroalkane is the substrate, the initial step after enzyme-substrate complex formation is removal of a proton to convert the substrate to a nitronate (11). Thus, N. crassa NMO has the function of both tautomerase and dioxygenase.
The lack of ammonia accumulation during growth of bacteria on 3NPA indicates that 3NPA is not reduced to β-alanine as in cattle and sheep rumen (4). Purified PnoA catalyzed transformation of P3N with the release of nitrate and nitrite in a 2:1 ratio, whereas growing cells released only 2% of the nitrogen as nitrite. The observations suggest that JS189 and JS190 incorporate nitrite as the nitrogen source, which accounts for the missing nitrogen in culture fluids.
MSA is an important intermediate in multiple anabolic and catabolic pathways, and a number of enzymes that transform MSA have been described (Fig. (Fig.3).3). The lack of MSA decarboxylase activity was due to a lack of expression of msaD, which is reminiscent of the myo-inositol degradation pathway in Lactobacillus casei (43). The myo-inositol operon carries genes for MSA oxidative decarboxylase (iolA) and MSA decarboxylase (iolK), but only iolA is expressed.
Based on the above results, we propose that the pathway for degradation of 3NPA (Fig. (Fig.4B)4B) is initiated by an inducible, but as yet unidentified, enzyme that converts 3NPA to P3N. The key step in the pathway is the denitration of P3N by the action of a constitutive P3N monooxygenase, encoded by the pnoA gene. An inducible MSA oxidative decarboxylase then converts MSA to acetyl-CoA, which enters central metabolic pathways. Additional sequencing will be required to locate the genes that encode the MSA oxidative decarboxylase and the hypothesized initial enzyme. The facile isolation of bacteria that grow on 3NPA suggests a highly evolved and widespread degradation pathway that may have evolved to exploit plant or fungal production of 3NPA.
The function of all other enzymes that transform 3NPA and nitroalkanes has been ascribed to detoxification or protection, but their physiological roles have not been established. P3NO and 3NPAO have been found only in organisms that also produce 3NPA. The presence of 3NPA in plants has been attributed to antiherbivory strategies, while the presence of 3NPA in fungi remains to be explained. In contrast to the other enzymes, PnoA in Cupriavidus sp. JS190 and Pseudomonas sp. JS189 clearly serves as a means to exploit 3NPA as a growth substrate. The P3N monooxygenase described here is the only member of the group whose physiological role has been established and the first P3N monooxygenase for which a gene sequence has been reported.
Gene sequences that encode dioxygenases related to 2-nitropropane dioxygenase constitute COG2070, which comprises 53 proteins distributed among 30 genomes (http://www.ncbi.nlm.nih.gov/COG/grace/wiew.cgi?COG2070), with over 2,100 nucleotide sequences in GenBank (Fig. (Fig.5).5). Many organisms contain multiple genes annotated as encoding 2-nitropropane dioxygenase. Despite the apparent widespread distribution of the COG, prior to this study, only proteins from Williopsis saturnus var. mrakii (formerly Hansenula mrakii) (27, 31), Neurospora crassa (11, 15), Pseudomonas aeruginosa (16), and Streptomyces achromogenes (9, 44) had been purified and characterized to various degrees. Although called 2-nitropropane dioxygenase, no evidence established 2-nitropropane as the physiological substrate of any of the enzymes included in COG2070. The recent reclassification of 2-nitropropane dioxygenase as nitronate monooxygenase (13) was based on the recharacterization of purified proteins from the ascomycetes.
The P3N monooxygenase is biochemically distinct from the well-characterized nitronate monooxygenases and falls within COG2070, but with only ~20 to 25% amino acid identity to the biochemically characterized enzymes. Many of the current annotations are wrong (12), and the situation with 2-nitropropane dioxygenase seems to be another example of annotation based only on modest sequence similarity without functional information. Our preliminary investigation of putative nitronate monooxygenases from B. phytofirmans PsJN and P. aeruginosa PAO1 confirms that the three closely related genes encode P3N monooxygenase rather than 2-nitropropane dioxygenase, based on the specificity of the enzymes for P3N and the lack of activity for 2-nitropropane. Ha et al. crystallized a 2-nitropropane dioxygenase from P. aeruginosa PAO1 and analyzed and aligned the amino acid sequence with several closely related enzymes (16) but did not determine the physiological substrate of the enzyme. Six motifs were described, and 10 highly conserved residues that interact with FMN were identified. When the P3N monooxygenases described here were added to the alignment, only three of the highly conserved residues interacting with FMN along with the His152 identified as the catalytic base were conserved across both 2-nitropropane dioxygenase and P3N monooxygenase (Fig. (Fig.6).6). Motifs II and IV, the latter being described as the most highly conserved motif by Ha et al., are disrupted in the P3N monooxygenases. Taken together, the evidence suggests that P3N monooxygenases form a separate cluster within COG2070 and that many of the proteins annotated as “2-nitropropane dioxygenase” are in fact P3N monooxygenases (Fig. (Fig.5).5). The argument is supported by the fact that 3NPA is much more likely to be widespread in natural ecosystems than are 2-nitropropane and other nitroalkanes.
The facile isolation of bacteria that grow on 3NPA suggests a highly evolved and widespread degradation pathway that may have evolved to exploit plant or fungal production of 3NPA. The ability of the pnoA-containing strains to grow on 3NPA as a nitrogen source but not always as a carbon source suggests the lack of a complete degradation pathway or an inability to regulate such a pathway in some strains. It is possible that, in some bacteria, the presence of P3N monooxygenase might be a detoxification mechanism similar to the proposed function in fungi and plants. It is also a plausible mechanism of scavenging nitrogen. There may be a continuum of enzymes with functions from protection to growth represented in the diversity of genes in COG2070. The 3NPA-degrading bacteria are the only organisms reported to grow well on 3NPA as a sole source of carbon, nitrogen, and energy. An Alcaligenes sp. was reported to produce nitrate and nitrite from 3NPA; however, after 28 days, only a fraction of the initial 3NPA was consumed, and minimal growth was observed (7). The failure of E. coli to either grow on or be inhibited by 3NPA suggests alternate means of detoxification of 3NPA or insensitivity to its effects. We are currently more fully characterizing the biochemistry of the P3N monooxygenase and ecological roles of bacteria that degrade 3NPA.
This work was supported by the Defense Threat Reduction Agency and Army Research Office grant W911NF-07-1-0077.
We thank Giovanni Gadda and Kevin Francis for helpful discussions and for reviewing the manuscript.
Published ahead of print on 9 April 2010.