Identification and Characterization of a Tetramethylpyrazine CatabolicPathway in Rhodococcus jostii TMP1

Simonas Kutanovas,a Jonita Stankeviciute,a Gintaras Urbelis,b Daiva Tauraite,a Rasa Rutkiene,a Rolandas Meskysa

Department of Molecular Microbiology and Biotechnology, Institute of Biochemistry, Vilnius University, Vilnius, Lithuaniaa; Center for Physical Sciences and Technology,Vilnius, Lithuaniab

At present, there are no published data on catabolic pathways of N-heterocyclic compounds, in which all carbon atoms carry asubstituent. We identified the genetic locus and characterized key reactions in the aerobic degradation of tetramethylpyrazine inRhodococcus jostii strain TMP1. By comparing protein expression profiles, we identified a tetramethylpyrazine-inducible pro-tein of 40 kDa and determined its identity by tandem mass spectrometry (MS-MS) de novo sequencing. Searches against anR. jostii TMP1 genome database allowed the identification of the tetramethylpyrazine-inducible protein-coding gene. Thetetramethylpyrazine-inducible gene was located within a 13-kb genome cluster, denominated the tetramethylpyrazine degrada-tion (tpd) locus, that encoded eight proteins involved in tetramethylpyrazine catabolism. The genes from this cluster werecloned and transferred into tetramethylpyrazine-nondegrading Rhodococcus erythropolis strain SQ1. This allowed us to verifythe function of the tpd locus, to isolate intermediate metabolites, and to reconstruct the catabolic pathway of tetramethylpyr-azine. We report that the degradation of tetramethylpyrazine is a multistep process that includes initial oxidative aromatic-ringcleavage by tetramethylpyrazine oxygenase, TpdAB; subsequent hydrolysis by (Z)-N,N=-(but-2-ene-2,3-diyl)diacetamide hydro-lase, TpdC; and further intermediate metabolite reduction by aminoalcohol dehydrogenase, TpdE. Thus, the genes responsiblefor bacterial degradation of pyrazines have been identified, and intermediate metabolites of tetramethylpyrazine degradationhave been isolated for the first time.

Pyrazines, monocyclic aromatic rings with two nitrogen atomsin para position, are a class of compounds that occur almost

ubiquitously in nature. Various pyrazines can be synthesized bothchemically and biologically, including tetramethylpyrazine(TTMP), which is produced by different bacteria (1, 2) or plants(3, 4). However, there is very little information available on thebiodegradation of these N-heterocyclic compounds. While bacte-rial strains able to use various alkyl-substituted pyrazines as a solecarbon and energy source have been isolated and described (5–9),almost nothing is known about the degradation pathways of al-kylpyrazines, including tetramethylpyrazine, in which all carbonatoms carry a substituent.

Under aerobic conditions, alkylated pyrazines are metabolizedvia oxidative degradation, leading to the hydroxylation of the ringat a free position. Rhodococcus erythropolis DSM 6138 and Arthro-bacter sp. strain DSM 6137 can use 2,5-dimethylpyrazine as asource of carbon and energy. The catabolism of 2,5-dimethylpyr-azine by these microorganisms gives rise to the intermediate me-tabolite 2-hydroxy-3,6-dimethylpyrazine, which accumulates inthe medium, indicating that ring hydroxylation occurs during theinitial steps of degradation (5). However, no enzymes involved inthis bioconversion were reported in the patent that describes theaforementioned reactions (5).

The 2,5-dimethylpyrazine is also catabolized by another R.erythropolis strain, DP-45, as reported by Rappert et al. (7). TheDP-45 strain also grew on a variety of other alkylpyrazines, includ-ing 2,3-dimethylpyrazine, 2,6-dimethylpyrazine, 2-ethyl-5(6)-di-methylpyrazine, 2-ethylpyrazine, 2-methylpyrazine, and 2,3,5-trimethylpyrazine (7). As was the case with strains DSM 6138 andDSM 6137, the degradation of 2,5-dimethylpyrazine by DP-45was accompanied by the accumulation of the intermediate metab-olite 2-hydroxy-3,6-dimethylpyrazine, which then disappearedwith the release of ammonium into the medium (7). The hydrox-

ylation of 2,5-dimethylpyrazine was mediated by an inducible en-zyme, while the enzyme catalyzing the subsequent ring cleavagewas shown to be constitutively expressed (7). Based on inhibitionstudies, it was proposed that the initial hydroxylation was cata-lyzed by a flavin monooxygenase or a cytochrome P450 mono-oxygenase, while the ring cleavage required P450 monooxygenase(7). However, the identities of the enzymes remain unknown. It isknown, however, that in contrast to the degradation of pyridines,which are also metabolized via ring hydroxylation, the degrada-tion of 2,5-dimethylpyrazine does not depend on molybdenum-containing enzymes (10).

The degradation of trisubstituted pyrazines was demonstratedto follow the same metabolic pattern as disubstituted pyrazines(6). Mycobacterium sp. strain DM-11 oxidized 2,3-diethyl-5-methylpyrazine to an intermediate compound, 5,6-diethyl-2-hy-droxy-3-methylpyrazine, which was further degraded with the re-lease of ammonium into the culture medium (6). Othercompounds, including 2,3,5-trimethylpyrazine, were also used bystrain DM-11 as a sole carbon, nitrogen, and energy source (6).Exposure to 2,3-diethyl-5-methylpyrazine induced the expressionof the enzymes involved in its degradation, but these enzymeshave not been identified yet (6).

In contrast to partially substituted pyrazines, TTMP cannot be

Received 2 January 2013 Accepted 29 March 2013

Published ahead of print 5 April 2013

Address correspondence to Simonas Kutanovas, simonas.kutanovas@bchi.vu.lt.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.00011-13.

Copyright © 2013, American Society for Microbiology. All Rights Reserved.

doi:10.1128/AEM.00011-13

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degraded via initial hydroxylation to form hydroxypyrazine, sinceeach of the TTMP ring carbons carries a substituent. Müller andRappert (8) suggested that the initial step of TTMP degradationmay involve ring cleavage. They used cell extracts from a Rhodo-coccus opacus strain that can use TTMP as a single carbon, nitro-gen, and energy source and were not able to detect any interme-diates during the degradation of TTMP (8). However, with theexception of the minireview (8), no experimental data have beenpublished to support these findings. Although the TTMP-degrad-ing bacteria have been isolated, neither enzymes catalyzing TTMPbiodegradation nor the corresponding genes have been identifiedin R. opacus so far.

In this study, we report the TTMP catabolic pathway of Rho-dococcus jostii strain TMP1, a strain previously shown to useTTMP as the sole source of carbon and energy (9). The genetictpdA-tpdE locus encoding the proteins required for the initialsteps of TTMP biodegradation was identified, and the corre-sponding genes were cloned and heterologously expressed in adifferent Rhodococcus sp. strain, thus enabling it to metabolizeTTMP. The identification of the intermediate metabolites(Z)-N,N=-(but-2-ene-2,3-diyl)diacetamide (BDNA) and N-(3-oxobutan-2-yl)acetamide (OBNA) was used to characterize thereactions of aerobic TTMP degradation, which allowed us to de-scribe the catabolic pathway of TTMP metabolism in bacteria forthe first time.

MATERIALS AND METHODSBacterial strains, plasmids, primers, and standard techniques. TheTTMP-degrading bacterium R. jostii TMP1 was previously isolated from asoil sample (9). R. erythropolis strain SQ1 was chosen as the host strain forthe expression of recombinant genes in bioconversion experiments. Esch-erichia coli strain DH5� was used for cloning experiments. The TpdEprotein was overexpressed in E. coli strain BL21(DE3). The bacterialstrains, plasmids, and primers used in this study are listed in Table S1 inthe supplemental material. Standard techniques were used for DNA ma-nipulations (11).

Bacterial growth medium and conditions. Rhodococcus strains weregrown at 30°C with aeration, E. coli strains were grown at 37°C withaeration. R. jostii TMP1 was cultivated either in nutrient broth (NB) (Ox-oid) medium or in minimal medium (5 g/liter NaCl, 1 g/liter K2HPO4, 1g/liter NH4H2PO4, 0.1 g/liter MgSO4, and 0.2 g/liter yeast extract, pH 7.2)supplemented with either TTMP (0.05%) or pyridine (0.05%). For cellsuspension and bioconversion experiments, R. erythropolis SQ1 wasgrown in 1-liter flasks containing 250 ml of NB medium until the culturereached an optical density at 600 nm (OD600) of 1.6 to 2.0. Then, cells werecollected by centrifugation, washed twice, and resuspended in 10 mMpotassium phosphate, pH 7.2, to achieve 4-fold-higher cell density. E. colistrains transformed with recombinant plasmids were grown in NB me-dium supplemented with either 50 �g/ml ampicillin or 40 �g/ml kana-mycin, as required. R. erythropolis SQ1 transformed with recombinantplasmids was grown in the presence of 60 �g/ml kanamycin.

Construction of plasmids. Total DNA from R. jostii TMP1 cultivatedin NB medium was isolated as described previously (12). To create agenomic library, the DNA was digested with HindIII, and the resultingfragments were inserted into the HindIII site of pUC19. The strategiesemployed to construct the TTMP-inducible green fluorescent protein(GFP)-encoding plasmid, as well as recombinant plasmids carrying dif-ferent genes of the tetramethylpyrazine degradation locus, are describedin Table S1 in the supplemental material. R. erythropolis SQ1 and E. colistrains were transformed with plasmid DNA by electroporation.

Analysis of the protein expression profile induced by TTMP. R. jostiiTMP1 was cultivated in minimal medium supplemented with eitherTTMP or pyridine; cells were collected by centrifugation and suspended

in 50 mM potassium phosphate, pH 7.2. After the addition of silica beads(0.1-mm diameter; 0.5 g/ml), the cells were disrupted by sonication at 750W for 10 min using a VC-750 ultrasound processor (Sonics & Materials,Inc.). Cell debris was removed by centrifugation at 16,000 � g for 10 min.Proteins were separated on 14% SDS-PAGE gel and visualized by Coo-massie blue staining.

MS-MS analysis. R. jostii TMP1 was cultivated in minimal mediumsupplemented with TTMP. Cells were collected, and proteins were sepa-rated on an SDS-PAGE gel. The band corresponding to the induced 40-kDa protein was excised and subjected to de novo sequencing based onmatrix-assisted laser desorption ionization–time of flight (MALDI-TOF)/TOF mass spectrometry (MS) and subsequent computational analysis atthe Proteomics Centre of the Institute of Biochemistry, Vilnius University(Vilnius, Lithuania). The sample was purified as described previously(13). Tryptic digest from the gel slice was analyzed with a 4000 QTrap (ABSciex, Framingham, MA) mass spectrometer in linear ion trap mode usinginformation-dependent acquisition (IDA) and a dynamic-exclusion pro-tocol. The acquisition method consisted of an IDA scan cycle, includingan enhanced-mass scan (EMS) as the survey scan, an enhanced-resolution(ER) scan to confirm the charge state, and six dependent enhanced-prod-uct-ion (EPI) scans (tandem mass spectrometry [MS-MS]). With thethreshold of the ion intensity at 100,000 cps, the IDA criteria were set toallow the most abundant ions in the EMS to trigger EPI scans. The surveyMS scan was set to a mass range from 400 m/z to 1,400 m/z. Dynamic ionexclusion was set to exclude precursor ions after their two occurrencesduring a 60-s interval. Peak lists were generated using Analyst software1.4.2 (AB Sciex, Framingham, MA).

Illumina sequencing, contig assembly, and inducible gene locusidentification. R. jostii TMP1 DNA was sequenced using an Illumina GA2platform (Macrogen, South Korea), and contigs were assembled usingCLC-Genomics Workbench software (CLC bio, Denmark). To identifythe TTMP-inducible gene, a search of the mass spectrometry-derived dataagainst the R. jostii TMP1 genome was performed.

Gene sequence analysis. The deduced amino acid sequences of theproteins encoded by the tpd locus were searched against the NCBI data-base using BLAST (14). Protein functions were assigned based on a se-quence similarity search against the NCBI Conserved Domain Database(15). Phylogenetic and molecular evolutionary analyses were conductedusing MEGA version 5 (16).

Qualitative and quantitative RT-PCR. R. jostii TMP1 was cultivatedin minimal medium containing glucose (0.1%), TTMP (0.05%), or pyri-dine (0.05%) as the sole carbon source until the culture reached an OD600

of 0.5. Total RNA was isolated using a ZR Soil/Fecal RNA MicroPrep kit(Zymo Research Corporation). Quantitative-PCR (qPCR) amplificationwas performed using a Rotor-Gene Q 5-plex HRM (Qiagen, Germany).qPCR was conducted in 15 �l of reaction mixture containing 1.5 �MSyto9, 7.5 �l of Verso 1-step QRT-PCR mix, 0.15 �l of Verso Enzyme,0.75 �l of RT enhancer (all from the Verso 1-step QRT-PCR Kit, ThermoScientific, USA), 200 nM each primer (see Table S1 in the supplementalmaterial), and 1 �l of the RNA tested. The qPCR was initiated with reversetranscription (RT) at 50°C for 15 min, followed by initial denaturation at95°C for 15 min, and either a subsequent 45 cycles of 95°C for 15 s, 58°Cfor 1 min, and 72°C for 10 s (for tpdA, tpdB, tpdC, tpdD, and tpdE) or asubsequent 35 cycles of 95°C for 20 s, 50°C for 1 min, and 72°C for 1 min(for 16S RNA). For qualitative evaluation, endpoint PCR products wereanalyzed using electrophoresis. For quantitative analysis, fluorescencedata were recorded after the annealing step. All experiments were carriedout in duplicate. To verify the absence of DNA in the RNA samples, theprocedure was performed without the reverse transcriptase step. Thethreshold cycle (CT) (threshold value, 0.05) values were obtained usingRotor-Gene Q Series Software 2.1.0 (build 9). Relative target RNA analysiswas performed using the 2��CT algorithm and 16S RNA as a reference fornormalization.

Enhanced GFP (EGFP) fluorescence measurement. R. jostii TMP1was transformed with either the pART3-gfp or the pART3-5=UTR-gfp

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plasmid and cultivated in 25 ml of minimal medium supplemented with0.05% TTMP, 0.05% pyridine, or 0.1% glucose for 48 h until the OD600

reached 1.0 to 1.6. Cells were collected by centrifugation, washed threetimes, and resuspended in 10 mM potassium phosphate, pH 7.2, toyield a final OD600 of 10. The suspension (200 �l/well) was transferredto a black 96-well plate (Costar), and fluorescence was measured in aFlexStationII384 fluorimeter (Molecular Devices) at an excitation wave-length (�ex) of 485 nm and an emission wavelength (�em) of 510 nm.Samples of three independent experiments were measured in triplicate.

Cell suspension experiments. R. erythropolis SQ1 transformed withpART2 recombinant plasmids carrying different genes of the tpd locuswas cultivated as described above. The cell suspension was supplementedwith 0.1 mM TTMP or 0.1 mM (Z)-N,N=-(but-2-ene-2,3-diyl)diacet-amide and incubated at 30°C for 1 hour. Bacteria were removed by cen-trifugation at 16,000 � g for 1 min, and the UV absorption spectra of eachsupernatant were recorded in a PowerWave XS plate reader (BioTek In-struments, Inc.).

Bioconversion of TTMP. R. erythropolis SQ1 transformed with eitherpART2-tpdABC or pART2-tpdAB recombinant plasmid was cultivated asdescribed above. Bioconversion reactions were carried out in a total vol-ume of 250 ml at 30°C with shaking at 180 rpm. TTMP and glucose wereadded to the reaction mixture in portions of 20 mg and 125 mg, respec-tively, while monitoring the progress of conversion by the changes in theUV absorption spectrum in the 200- to 320-nm range. The reaction wasperformed for 4 days; the total amounts of TTMP added were 240 mg forpART2-tpdABC and 140 mg for pART2-tpdAB. Accumulation of biocon-version products was monitored by thin-layer chromatography in chlo-roform-methanol (9:1) using the substrate compound as a reference.

Intermediate metabolite isolation. Bacteria were removed from thebioconversion reaction mixtures by centrifugation at 4,000 � g for 20min, and the supernatants were evaporated under reduced pressure. Theproduct of TpdABC was extracted from concentrated aqueous solutionwith chloroform. The product of TpdAB was isolated by sequential dis-solving in acetonitrile and chloroform. The isolated intermediate metab-olites were used for structural analysis and for the whole-cell and enzymeexperiments.

Metabolite structural analysis. The structures of the products ofTpdABC and TpdAB were determined using 1H nuclear magnetic reso-nance (NMR), 13C NMR, and MS analyses. 1H and 13C NMR spectra wererecorded on a Varian Unity Inova 300 spectrometer (300 and 75 MHz,respectively). The TpdABC product was dissolved in deuterated dimethylsulfoxide, and the TpdAB product was dissolved in CDCl3. Spectra werecalibrated with respect to the solvent signal (CDCl3, 1H � � 7.26, 13C � �77.2; DMSO-d6, 1H � � 2.50, 13C � � 39.5). High-resolution MS wasperformed on a Dual-ESI Q-TOF 6520 mass spectrometer (Agilent Tech-nologies).

TpdE expression and purification. The tpdE gene was fused with the3=-polyhistidine sequence of the pET21b() expression vector (see TableS1 in the supplemental material). E. coli BL21(DE3) was transformed withthe recombinant plasmid pET21-tpdE and cultured aerobically at 30°C in1-liter conical flasks with 200 ml of brain heart infusion (BHI) medium(Oxoid) supplemented with 50 mg/ml ampicillin. When an OD600 of 1.2was reached, 0.5 mM isopropyl--D-thiogalactopyranoside (IPTG) wasadded to induce the expression of tpdE, and the culture was incubated for4 h. Cells were collected by centrifugation; washed with 50 mM potassiumphosphate, pH 7.2; resuspended in 8 ml of the same buffer; and disruptedby sonication at 750 W for 5 min using a VC750 ultrasound processor(Sonics & Materials, Inc.). Cell debris was removed by centrifugation at16,000 � g for 10 min. Cell extracts were loaded onto a HiTrap IMAC FF5-ml nickel column (GE Healthcare), and proteins were eluted with 50mM potassium phosphate, 1 M imidazole, pH 7.2. The purity of TpdE wasconfirmed by electrophoresis on a 14% SDS-PAGE gel.

TpdE activity measurements. The NADPH-dependent ketoreductaseactivity of TpdE was determined at 30°C in a Helios gamma UV-visible(UV-Vis) spectrophotometer (Thermo Fisher Scientific) by measuring

the decrease in A340 resulting from the oxidation of NADPH (ε340 � 6,220M�1 cm�1) after the addition of the substrate OBNA. TpdE activity wasassayed in buffer containing 0.2 mM NADPH and 5 �M OBNA. One unitof activity was defined as the amount of the enzyme that catalyzed theoxidation of 1 �mol of NADPH per minute. The optimum temperaturefor TpdE activity was determined to be in the range of 15 to 50°C in 50 mMpotassium phosphate, pH 7.2. The optimal pH for TpdE activity was de-termined to be in the range of pH 6.0 to 8.5, using potassium phosphateand Tris-HCl buffers. The apparent TpdE Km values for NADPH andOBNA were determined by varying concentrations of these substratesfrom 50 to 500 �M and from 0.1 to 10 �M, respectively. Data from at leastthree independent experiments were combined. For data fitting, GraFit(Erithacus Software Ltd.) software was used.

Nucleotide sequence accession number. The R. jostii TMP1 genomefragment sequence with the TTMP degradation locus was deposited inGenBank under accession no. HF544504.

RESULTS

We previously reported that R. jostii TMP1 is capable of usingTTMP as a source of carbon and energy (9). To elucidate themetabolic pathway of TTMP, the tetramethylpyrazine degrada-tion (tpd) gene locus was identified, and intermediate metabolitesof TTMP were determined.

TTMP-inducible protein expression in R. jostii TMP1. Todetect TTMP catabolism-related enzymes, we investigatedwhether TMP1 cultivation with TTMP could cause the upregula-tion of proteins that might be involved in the TTMP metabolicpathway. SDS-PAGE analysis of cell extracts revealed severalTTMP-inducible protein bands, including a dominant 40-kDaband (Fig. 1A). The expression of inducible 40-kDa protein wasTTMP specific rather than shared with N-heterocyclic com-pounds, since TMP1 cultivation in the presence of pyridineshowed a different protein expression profile, and no upregula-tion of the 40-kDa protein was observed (Fig. 1B). These resultssuggest that the 40-kDa protein plays an important and specificrole in TTMP degradation. Therefore, to elucidate the TTMPmetabolic pathway, the 40-kDa protein was selected for furtheranalysis.

Identification of the tpd gene locus. To identify the TTMP-

FIG 1 Tetramethylpyrazine-inducible protein. (A) Cultures of R. jostii TMP1were cultivated for 18 h in NB medium supplemented with different concen-trations of tetramethylpyrazine (0.1 to 10 mM). (B) Strain TMP1 was culti-vated for 48 h in minimal medium supplemented either with 0.05% TTMP or0.05% pyridine (PYR) as a single source of carbon. SDS-PAGE gels werestained with Coomassie blue. The arrows indicate the 40-kDa protein. LanesM, molecular mass ladders (in kDa).

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inducible protein, the 40-kDa band was excised from an SDS-PAGE gel and analyzed by MS-MS sequencing. The candidatepeptides were searched against the genomic Illumina database,resulting in the identification of a contig that contained the locusof the 40-kDa protein-coding gene. To verify the sequence of thetpd locus, a genomic library was constructed and a clone contain-ing a sequence of interest was identified by PCR screening usingprimers specific to the 40-kDa protein-coding gene (tpdA_F andtpdA_R [see Table S1 in the supplemental material]). Sequencingof the identified clone confirmed the nucleotide sequence of a7-kb region of the tpd locus. The remaining sequence of the tpdlocus was verified by direct sequencing of overlapping PCR prod-ucts amplified from total DNA using primers designed accordingto the Illumina contig sequence (primer pairs seq_F and seq_R[see Table S1 in the supplemental material]). Thus, a 13-kb regionthat contained the genes required for the initial steps of TTMPdegradation was sequenced. Sequence analysis of the tpd locusrevealed eight open reading frames (ORFs) that were denomi-nated tpdA, tpdB, tpdC, tpdR, tpdD, tpdE, ORF1, and ORF2(Fig. 2). The 40-kDa inducible protein was encoded by the tpdAgene. Genes tpdA, tpdB, and tpdC were arranged in tandem on thesame strand and contained very short to no intergenic spaces,suggesting that the genes are organized into the operon tpdABC

(Fig. 2). To confirm that these three genes were cotranscribed,RT-PCR analysis was performed using primers designed to am-plify the intergenic regions (AB_F and AB_R; BC_F and BC_R[see Table S1 in the supplemental material]). Endpoint PCR re-vealed that gene pairs tpdA-tpdB and tpdB-tpdC were transcribedas a contiguous transcript, confirming the tpdABC operon(Fig. 3A). The transcription of tpdABC was specifically induced byTTMP, but not by glucose or pyridine (Fig. 3A).

To verify that the expression of the tpdABC operon was regu-lated by TTMP, the fragment of the upstream region of the tpdAgene was amplified by PCR and fused to an EGFP gene from thepART3-gfp vector. Strain TMP1, transformed with the pART3-5=UTR-gfp plasmid, expressed EGFP when cultivated on TTMP,as determined by the developed bacterial fluorescence (Fig. 3B).The level of EGFP fluorescence increased 10-fold in TTMP-in-duced bacteria carrying pART3-5=UTR-gfp compared to that inglucose-induced bacteria. In contrast, no EGFP fluorescenceabove the background level was detected upon induction withpyridine. This confirmed that the upstream region of tpdA con-tained a promoter that was specifically activated upon exposure toTTMP.

To investigate whether the expression of the tpdD and tpdEgenes was also dependent on TTMP, real-time RT-PCR analysiswas performed. The results revealed that the expression of each ofthe tpdA-tpdE genes was specifically induced when R. jostii TMP1was cultivated in the presence of TTMP (Fig. 3C), suggesting thatproteins TpdD and TpdE participate in TTMP catabolism.

TTMP oxidation and subsequent hydrolysis by TpdABCproteins. Sequent analysis revealed that TpdA shows high se-quence similarity to luciferase-like monooxygenases (see Table S2and Fig. S1 in the supplemental material). In addition, TpdA con-tains conserved domains characteristic of the cl07892 superfamilyof flavin-utilizing monooxygenases and shares several commondomains with coenzyme F420-dependent flavin oxidoreductases(Table 2; see Table S3 in the supplemental material).

Flavin monooxygenases usually act as two-component enzymesystems: a larger component (an oxidase), which uses the reducedflavin nucleotides to hydroxylate substrates, and a smaller com-

FIG 2 Gene locus of R. jostii TMP1 involved in the metabolism of tetrameth-ylpyrazine. The markings at the top denote separate fragments that were se-quenced to verify the tpd locus: the clone from the HindIII genomic library(pUC19-H1) and four fragments obtained by PCR (Seq1 to Seq4). The grayarrows represent genes that are known to be involved in TTMP degradation;the white arrows represent other ORFs in the locus. *, the tpdA gene encodes a40-kDa tetramethylpyrazine-inducible protein.

FIG 3 Induction of tpd locus expression by tetramethylpyrazine in R. jostii TMP1. Strain TMP1 was cultivated in liquid minimal medium supplemented witheither 0.05% TTMP, 0.1% glucose (GLC), or 0.05% PYR as a single source of carbon. (A) RT-PCR analysis using primers designed to amplify the intergenicregions between tpdA-tpdB and tpdB-tpdC. (B) TMP1 was transformed with pART3-gfp plasmid containing the tpdA upstream region inserted as a promoter(pART-5=UTR-gfp). Bacterial EGFP fluorescence was measured in a plate reader (�ex � 485 nm; �em � 510 nm); the data are presented as averages of threeindependent experiments plus the standard deviation. RFU, relative fluorescence units. (C) Quantitative RT-PCR analysis of the transcription of tpdA genes totpdE genes. The data are presented as relative RNA amounts calculated from the threshold cycles using the threshold cycle of 16S RNA as a reference; averages ofduplicate runs plus standard deviations are presented.

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ponent (flavin reductase), which uses NAD(P)H to reduce FAD orFMN (17). Sequence analysis predicted that a flavin reductase ofthis type may be encoded by the tpdD gene (Table 2; see Tables S2and S3 in the supplemental material).

A deduced tpdB gene product belongs to the nuclear transportfactor 2 (NTF2)-like superfamily, which contains proteins that,despite many common structural details, diverge greatly in theirfunctions, including ketosteroid isomerases and the beta subunitof the ring-hydroxylating dioxygenases (Table 2; see Fig. S2 in thesupplemental material).

The third gene of the tpdABC operon, tpdC, encodes the pro-tein that belongs to the superfamily of amidases (Table 2; see TableS3 in the supplemental material) and shares very high sequencehomology with omega-octalactam hydrolase (see Table S2 inthe supplemental material), whose function has been confirmedat the protein level (18).

To characterize Tpd proteins involved in TTMP degradationand to investigate the associated metabolic reactions, pART2 ex-pression vectors containing different genes of the tpdABC operon

and their combinations were constructed. The involvement of in-dividual genes in the metabolism of TTMP was assessed in R.erythropolis strain SQ1, which does not metabolize TTMP. Thetransformation of strain SQ1 with the pART2 vector led to effec-tive protein expression, as confirmed by EGFP fluorescence devel-oped in the cells transformed with pART2-gfp (data not shown).TTMP metabolism in resting cell suspensions was evaluated byUV absorption spectroscopy. As seen in Fig. 4, while SQ1 cellstransformed with either tpdABC or tpdAB were capable of metab-olizing TTMP, those harboring either tpdA, tpdB, or tpdC did nothave the ability to metabolize the compound. Furthermore,TTMP degradation by TpdABC leads to the formation of metab-olites different from those found for TpdAB, as was evaluated byUV absorption spectroscopy (Fig. 4) and thin-layer chromatogra-phy (data not shown).

TpdABC- and TpdAB-generated metabolites were extractedfrom the bioconversion media, and their structures were deter-mined by 1H NMR, 13C NMR, and MS analyses. The 1H NMRspectrum of the TpdABC product showed five peaks in the 1H

FIG 4 Metabolism of tetramethylpyrazine in R. erythropolis SQ1 transformed with recombinant pART2 plasmids, each carrying a different combination of genesfrom the tpd locus. Cultures of R. erythropolis SQ1 were incubated in potassium phosphate buffer supplemented with 0.1 mM tetramethylpyrazine for 1 h, andthe UV absorption spectra were recorded. The solid lines represent the initial spectrum of tetramethylpyrazine; the dotted lines indicate the final spectra of thebioconversion products.

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NMR spectrum [� 6.37 (br s, 1H, NH), 4.58 (p, J � 7.1 Hz, 1H,CH), 2.21 (s, 3H, COCH3), 1.99 (s, 3H, NHCOCH3), 1.36 (d, J �7.1 Hz, 3H, CH3)] and six peaks in the 13C NMR spectrum (�207.1, 169.7, 54.8, 26.8, 23.4, and 17.8), identifying the TpdABCproduct as OBNA. These NMR spectra were in full agreementwith previously published results (19). Meanwhile, the product ofTpdAB showed three peaks with � 8.67 (s, 1H, NH), 1.88 (s, 3H,CH3), and 1.82 (s, 3H, CH3), while the 13C NMR spectrum con-tained four peaks with � 167.7, 121.2, 23.2, and 16.4. High-resolution MS analysis confirmed the molecular formula of thecompound as C8H15N2O2 (calculated [MH] mass was171.11335, found mass was 171.1134). These NMR spectra re-vealed that TpdAB metabolized TTMP to BDNA.

Next, the intermediate metabolite BDNA was used as a sub-strate for strain SQ1 transformed with recombinant plasmids,each carrying either individual genes of the tpdABC operon ordifferent combinations of the tpdABC genes. TpdABC, TpdBC,and TpdC were able to metabolize BDNA, indicating that TpdCalone was sufficient to catalyze the conversion of BDNA (Fig. 5).The results of bioconversion experiments (summarized inTable 1) demonstrated that the catabolism of TTMP is a multistepprocess starting with TTMP oxidation by TpdAB to produceBDNA, which is then hydrolyzed by TpdC to form OBNA. BDNAhydrolysis by TpdC led to the formation of an optically activeOBNA that rotated plane-polarized light with a specific rotation at20°C and 586 nm ([a]D

20) of 102° (1.16 g in 100 ml CHCl3),indicating that the reaction is stereospecific.

OBNA reduction by TpdE. TpdE shares high homology withclassical short-chain dehydrogenases/reductases that have a Ross-mann fold NAD(P)H/NAD(P) binding (NADB) domain(Table 2; see Table S3 in the supplemental material). This domainis found in numerous dehydrogenases and other redox enzymes

that can use a wide variety of substrates, including alcohols, glu-cose, and steroids (20). OBNA, which contains a keto group,might be a substrate for short-chain dehydrogenases/reductases,since most bacterial short-chain dehydrogenases/reductases areknown to be alcohol dehydrogenases that can catalyze bidirec-tional oxidoreduction.

To demonstrate that TpdE is indeed a short-chain dehydroge-nase/reductase, as predicted by the sequence analysis, His6-taggedTpdE protein was overproduced in E. coli and purified by affinitychromatography with a yield of 2,600 U per liter of culture. Apurified protein migrated as an �30-kDa band, which agreed withthe expected size of the recombinant His6-TpdE protein (27.9kDa) (see Fig. S3 in the supplemental material). The specific ke-

FIG 5 Metabolism of (Z)-N,N=-(but-2-ene-2,3-diyl)diacetamide in R. erythropolis SQ1 transformed with recombinant pART2 plasmids, each carrying adifferent combination of genes from the tpd locus. Cultures of R. erythropolis SQ1 were incubated in potassium phosphate buffer supplemented with 0.1 mM(Z)-N,N=-(but-2-ene-2,3-diyl)diacetamide for 1 h, and the UV absorption spectra were recorded. The solid lines represent the initial spectrum of (Z)-N,N=-(but-2-ene-2,3-diyl)diacetamide; the dotted lines represent the final spectra of the bioconversion products.

TABLE 1 Products of resting-cell reactions of R. erythropolis SQ1 cellstransformed with tpd locus genesa

Plasmid Expressed gene(s)

Product on substrateb:

TTMP BDNA

pART2 ND NDpART2-tpdA tpdA ND NApART2-tpdB tpdB ND NApART2-tpdC tpdC ND OBNApART2-tpdAB tpdA, tpdB BDNA NApART2-tpdBC tpdB, tpdC ND OBNApART2-tpdABC tpdA, tpdB, tpdC OBNA OBNAa The cultures of transformed bacteria were incubated in the presence of either 0.1 mMTTMP or 0.1 mM BDNA. Substrate degradation was observed by UV absorptionspectroscopy. The reaction products BDNA and OBNA were determined by either 1HNMR, 13C NMR, and MS analysis (TTMP conversion) or by UV spectrum comparisonand thin-layer chromatography (BDNA conversion).b ND, bioconversion not detected; NA, bioconversion not analyzed.

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toreductase activity of TpdE, with OBNA as a substrate andNADPH as an electron donor, was 26 U/mg. The optimum pH ofTpdE was 7.2, and the optimum temperature was 35°C. The ap-parent Km for NADPH calculated from steady-state analysis was379 � 73 �M in the presence of 6 mM OBNA, and the apparentKm for OBNA was 5.2 � 1.5 �M in the presence of 500 �MNADPH, with kcats of 7.7 s�1 and 2.5 s�1, respectively. The en-zyme has an absolute requirement for NADPH, yielding no de-tectable OBNA reduction in the presence of NADH.

Hypothetical proteins encoded by ORF1 and ORF2, whichwere found upstream of tpdE, may contribute to further degrada-tion of TTMP. ORF1 encodes the putative large subunit of N,N-dimethylformamidase, while a hypothetical protein encoded byORF2 is homologous to peptidases (Table 2; see Tables S2 and S3in the supplemental material).

DISCUSSION

To determine in detail the degradation pathway of TTMP in R.jostii TMP1, we isolated and characterized the genes involved inTTMP metabolism. Based on the identity of the TTMP-inducibleprotein, we identified the tetramethylpyrazine degradation locusand described the enzymes encoded in this locus.

TTMP degradation was demonstrated to be a multistep pro-

cess, involving oxidative aromatic ring cleavage by the TpdA-TpdB complex and subsequent hydrolysis catalyzed by TpdC,followed by keto group reduction by TpdE (Fig. 6). Flavinmonooxygenases, such as TpdA, are capable of oxidizing varioussubstrates, including some N-heterocyclic compounds (17, 21).To oxidize TTMP, TpdA requires TpdB in a manner similar tothat observed in bacterial luciferases, which act as a complex of �and subunits. However, while luciferase � and subunits arehomologous to each other (17), the TpdB protein is considerablysmaller than TpdA and shares no sequence homology withluciferase-like monooxygenases.

TpdB shares sequence homology with proteins belonging tothe families of ketosteroid isomerases and the beta-subunit ofthe ring-hydroxylating dioxygenases (15). Since TTMP oxida-tion might be facilitated by the functions of the enzymes ofboth of these families, the function of TpdB cannot be inferredfrom sequence similarity alone. Ketosteroid isomerases cata-lyze double-bond migration reactions of steroid substrates(22); therefore, such a function of TpdB might facilitate TTMPconversion through the destabilization of the aromatic ring.The enzyme PhzA/B, a homologue of TpdB, has been shown toparticipate in the condensation of two amino ketone molecules(23). This condensation is accompanied by PhzA/B-mediated

TABLE 2 Functional annotations of deduced Tpd proteins

ProteinSize (aminoacids/kDa) Putative function

Superfamily designation

SuperfamilyAccessionno. E value Conserved domaina

TpdA 387/42.9 Flavin-utilizing monooxygenase Flavin-utilizingmonooxygenases

cl07892 5.3e�16 Not described

TpdB 136/15.2 Unknown NTF2-like cl09109 7.4e�09 Not describedTpdC 484/51.4 Amidase Amidase cl11426 2.3e�63 Not describedTpdR 787/86.2 LuxR family transcriptional regulator LuxR_C-like cl17315 1.4e�14 Sequence-specific DNA binding domains

AAAb cl17189 1.2e�4 Conserved nucleotide phosphate-binding motif

TpdD 179/19.3 Flavin reductase Flavin reductases cl00801 5.7e�26 FMN-binding domain found inNAD(P)H-flavin oxidoreductases

TpdE 260/26.9 Short-chain dehydrogenase/reductase NADB Rossmann cl09931 2.4e�73 Rossmann-fold NADB domainORF1 747/81.2 Large subunit of

N,N-dimethylformamidaseORF2 123/13.6 Peptidase GAT_1 cl00020 8.9e�16 Not describeda The conserved domains of putative proteins encoded by the tpd locus were analyzed against the Conserved Domain Database at the NCBI website.b AAA, ATPases associated with a wide variety of cellular activities.

FIG 6 Proposed tetramethylpyrazine catabolic pathway in R. jostii TMP1. 1, tetramethylpyrazine; 2, BDNA; 3, OBNA; 4, N-(3-hydroxybutan-2-yl)acetamide;5, 5-3-amino-2-butanol. The dashed arrows indicate hypothetical reactions. TpdAB, TTMP oxygenase; TpdD, flavin reductase; TpdC, BDNA hydrolase; TpdE,aminoalcohol dehydrogenase.

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rearrangement of double bonds, yielding a phenazine precur-sor that is structurally similar to tetraalkylpyrazine (23). Theanalogous rearrangement of double bonds by TpdB may con-tribute to the destabilization of TTMP, facilitating its oxidationby TpdA.

Even though the product of TpdAB-catalyzed TTMP oxidationresembles that of dioxygenases (24), similar products may also beformed via the cleavage of the aromatic ring by flavin monooxy-genases, such as 2-methyl-3-hydroxypyridine-5-carboxylic acidoxygenase or 5-pyridoxic acid oxygenase (21). These monooxy-genases open the aromatic ring of the substrate and incorporateonly one oxygen atom from O2, whereas another atom comesfrom an H2O molecule. The exact mechanism of aromatic ringoxidation catalyzed by the TTMP oxygenase TpdAB requires fur-ther investigation, such as oxygen tracer experiments.

Flavin monooxygenases homologous to TpdA act as part of atwo-component enzyme system that also includes flavin reduc-tase, such as that encoded by tpdD. Therefore, the oxidation ofTTMP should require both TpdA and TpdD acting together.However, in R. erythropolis SQ1, tpdD was not required to oxidizeTTMP (Fig. 4). This is not surprising, since Rhodococcus strainsencode many flavin reductases (as seen from published genomes[25, 26]) that may have substituted tpdD.

Oxidized TTMP is further metabolized via a hydrolysis reac-tion catalyzed by TpdC (Fig. 6). 1H NMR spectra (in CDCl3) (datanot shown) revealed that BDNA may exist in different tautomericforms. Since BDNA hydrolysis yields optically active OBNA,rather than a racemate, we believe that the imine form of BDNA,rather than amine, is a true substrate for TpdC. Nevertheless, fur-ther analysis is required to elucidate the precise mechanism ofTpdC-mediated cleavage of OBNA.

Subsequent OBNA reduction is catalyzed by the amino alcoholdehydrogenase TpdE (Fig. 6), which uses NADPH as an electrondonor and presumably converts OBNA into amino alcohol. Thedetailed biochemical characterization of TpdE is currently inprogress.

Further steps of TTMP metabolism may involve the enzymesencoded by ORF1 and ORF2. Both ORF1 and ORF2 may recog-nize the amidic bond in the TpdE product and may hydrolyze it toproduce 3-amino-2-butanol (Fig. 6).

In R. jostii TMP1, TTMP degradation is an inducible processinvolving the induction of TpdA expression (Fig. 1 and 3). A pu-tative regulator of the tpd locus is encoded by tpdR. The analysis ofthe deduced amino acid sequence of TpdR revealed that the pro-tein contains a number of domains that are characteristic of tran-scription regulators belonging to the LuxR family. However, toascertain the ability of TpdR to regulate the tpd locus, furtherresearch is required.

In summary, here, we identified for the first time the geneticlocus responsible for bacterial degradation of pyrazines, namely,tetramethylpyrazine. Our data revealed that, in accordance withprevious reports on TTMP metabolism by Rhodococcus sp. strains(8), the initial step of TTMP degradation in R. jostii TMP1 is ringoxidation and cleavage. However, in contrast to the data pub-lished by Müller and Rappert (8) suggesting that TTMP is oxi-dized by a cytochrome P450-type enzyme, the TTMP oxidasecharacterized here is a putative flavoenzyme.

While identification of the TTMP catabolism pathway willundoubtedly advance the field of bacterial degradation of pyr-azines, one must bear in mind that the reactions described here

may be specific only to TTMP and similar pyrazines that carrya substituent at each carbon of the ring. Usually, the degrada-tion of semisubstituted pyrazines starts with hydroxylation ofthe pyrazine ring at a free ring position (2, 8) and thereforeemploys different enzymes than those identified in this study.

Identification of the genes responsible for bacterial degra-dation of TTMP and isolation of its intermediate metabolitesallowed us for the first time to describe the catabolic pathway ofalkylpyrazine metabolism in bacteria, providing fundamentalknowledge about the biodegradation of these N-heterocycliccompounds and revealing various enzymes that may be em-ployed in the future for selective and specific bioconversionreactions.

ACKNOWLEDGMENTS

This research was funded by a grant (no. MIP-046/2011) from the Re-search Council of Lithuania.

We thank Urte Neniskyte and Laura Kaliniene for helping to preparethe manuscript. We are grateful to Marija Ger for performing peptidesequence analysis and Maksim Bratchikov for carrying out real-time RT-PCR analysis.

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