Delineation of the Caffeine C-8 Oxidation Pathway in Pseudomonassp. Strain CBB1 via Characterization of a New Trimethyluric AcidMonooxygenase and Genes Involved in Trimethyluric AcidMetabolism

Sujit Kumar Mohanty,a Chi-Li Yu,a Shuvendu Das,b Tai Man Louie,a,b Lokesh Gakhar,c and Mani Subramaniana,b

Department of Chemical and Biochemical Engineering, University of Iowa, Iowa City, Iowa, USAa; The Center for Biocatalysis and Bioprocessing, University of Iowa,Coralville, Iowa, USAb; and Protein Crystallography Facility, University of Iowa, Iowa City, Iowa, USAc

The molecular basis of the ability of bacteria to live on caffeine via the C-8 oxidation pathway is unknown. The first step of thispathway, caffeine to trimethyluric acid (TMU), has been attributed to poorly characterized caffeine oxidases and a novel qui-none-dependent caffeine dehydrogenase. Here, we report the detailed characterization of the second enzyme, a novel NADH-dependent trimethyluric acid monooxygenase (TmuM), a flavoprotein that catalyzes the conversion of TMU to 1,3,7-trimethyl-5-hydroxyisourate (TM-HIU). This product spontaneously decomposes to racemic 3,6,8-trimethylallantoin (TMA). TmuMprefers trimethyluric acids and, to a lesser extent, dimethyluric acids as substrates, but it exhibits no activity on uric acid. Ho-mology models of TmuM against uric acid oxidase HpxO (which catalyzes uric acid to 5-hydroxyisourate) reveal a much biggerand hydrophobic cavity to accommodate the larger substrates. Genes involved in the caffeine C-8 oxidation pathway are locatedin a 25.2-kb genomic DNA fragment of CBB1, including cdhABC (coding for caffeine dehydrogenase) and tmuM (coding forTmuM). Comparison of this gene cluster to the uric acid-metabolizing gene cluster and pathway of Klebsiella pneumoniae re-vealed two major open reading frames coding for the conversion of TM-HIU to S-(�)-trimethylallantoin [S-(�)-TMA]. The firstone, designated tmuH, codes for a putative TM-HIU hydrolase, which catalyzes the conversion of TM-HIU to 3,6,8-trimethyl-2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline (TM-OHCU). The second one, designated tmuD, codes for a putative TM-OHCUdecarboxylase which catalyzes the conversion of TM-OHCU to S-(�)-TMA. Based on a combination of enzymology and gene-analysis, a new degradative pathway for caffeine has been proposed via TMU, TM-HIU, TM-OHCU to S-(�)-TMA.

Caffeine (1,3,7-trimethylxanthine), a purine alkaloid, is one ofthe most widely used psychoactive substances in the world,

with an estimated global consumption of 120,000 tons per year(13). The average consumption of caffeine in humans ranges from80 to 400 mg per person per day (14). The main mode of entry ofcaffeine and other natural xanthines into the human system andsubsequently into the environment is through coffee, tea, caffein-ated cola drinks, and chocolates. Caffeine is also a large environ-mental concern due to millions of tons of caffeine-containingagro-industrial wastes, such as husk and pulp water, which arereleased by coffee and tea manufacturers (36). The implications ofcaffeine consumption on human health and environment have re-sulted in a great deal of scientific and public interest in caffeine and itsbiodegradation (1, 2, 15; http://www.uiowa.edu/�biocat/).

Humans metabolize caffeine mainly via N-demethylation cat-alyzed by the hepatic cytochrome P450s 1A2 and 2E1 (2). Caffeinetransformation by N-demethylation has also been documented inmicroorganisms such as bacteria, fungi, and yeast (4, 6). AlthoughN-demethylase activity has been detected in these microorgan-isms as early as the 1970s (3, 41), the metabolic pathway, enzymol-ogy, and molecular genetics of N-demethylation, including ndmgenes, were elucidated only recently in Pseudomonas putida CBB5(37, 38, 43). Highly specific N1-, N3-, and N7-N-demethylasesconvert caffeine and other methylxanthines to xanthine and form-aldehyde. This organism consumes both of these compounds.

The alternate pathway of caffeine transformation via C-8 oxi-dation is much less characterized. Only two early studies (28, 29)have reported the presence of caffeine oxidases involved in C-8

oxidation. However, these enzymes and the corresponding reac-tions catalyzed by them are poorly characterized. Recently, ourgroup reported the purification and characterization of a novelcaffeine dehydrogenase (Cdh) from Pseudomonas sp. strain CBB1,a bacterium capable of growth on caffeine as the sole source ofcarbon and nitrogen. This heterotrimeric caffeine dehydrogenaseoxidized caffeine at the C-8 position to form 1,3,7-trimethyluricacid (TMU) with coenzyme Q0 as the preferred electron acceptor(42). The enzyme was specific for caffeine, much less active ontheobromine but had no activity on xanthine (42). The enzymol-ogy and genes involved in further transformation of TMU havenot been elucidated. Madyastha and Sridhar (27) reported thatTMU was further degraded to 3,6,8-trimethylallantoin (TMA) byresting cells of a mixed culture of Rhodococcus and Klebsiella.However, details of this reaction are lacking.

In the present study, we report the enzymology and molecularcharacterization of TMU conversion to 1,3,7-trimethyl-5-hy-droxyisourate (TM-HIU), catalyzed by a novel NADH-dependent

Received 9 April 2012 Accepted 9 May 2012

Published ahead of print 18 May 2012

Address correspondence to Mani Subramanian, mani-subramanian@uiowa.edu.

S.K.M. and C.-L.Y. contributed equally to this article.

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

Copyright © 2012, American Society for Microbiology. All Rights Reserved.

doi:10.1128/JB.00597-12

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trimethyluric acid monooxygenase (TmuM), a flavoprotein.TM-HIU is unstable and is readily converted nonenzymatically toracemic TMA. TmuM has no activity on uric acid. Analysis of a25.2-kb gene cluster containing genes encoding Cdh (cdhABC)and TmuM (tmuM) identified two other open reading frames(ORFs), which are proposed to be responsible for the enzymaticconversion of TM-HIU to S-(�)-TMA via 3,6,8-trimethyl-2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline (TM-OHCU). Apathway for caffeine transformation in CBB1 has been proposedthat is based on a similar analysis of the uric acid degradationpathway in Klebsiella pneumoniae (8). This is the first report ofenzymatic and genetic characterization of caffeine transformationvia C-8 oxidation.

MATERIALS AND METHODSChemicals. Caffeine, 1,3,7-trimethyluric acid (TMU), 3,7-dimethyluricacid, 1,3-dimethyluric acid, 1-methyluric acid, and uric acid were all pur-chased from Sigma-Aldrich (St. Louis, MO). Yeast nitrogen base withoutamino acids and ammonium sulfate (YNB) was obtained fromForMedium (Norfolk, United Kingdom). High-pressure liquid chroma-tography (HPLC)-grade methanol, chloroform, hexane, and ethanol wereobtained from Sigma-Aldrich.

Bacterial strains and plasmids. The bacterial strains and plasmidsused in the present study are listed in Table S1 in the supplemental mate-rial.

Media and growth conditions. Strain CBB1 was grown in M9 mineralsalts medium (34) with 2.5 g of caffeine liter�1 and 4 g of YNB liter�1 at30°C with shaking at 200 rpm. Cell growth was monitored by measuringthe optical density at 600 nm (OD600). Cells were harvested at the late logphase by centrifugation (13,800 � g for 10 min at 4°C) as described by Yuet al. (42). Cell pellets were washed twice with 50 mM potassium phos-phate (KPi) buffer (pH 7.5) and frozen at �80°C until required. Exceptwhere indicated, Escherichia coli strains were grown at 37°C in Luria-Bertani medium (7) or Terrific Broth medium (26). For the growth ofrecombinant E. coli strains, antibiotics were added to the following finalconcentrations: ampicillin, 100 �g ml�1; and chloramphenicol, 12.5 �gml�1. LB media were solidified with 1.5% Bacto agar (Difco Laborato-ries).

Purification of TMU-oxidizing enzyme from wild-type Pseudomo-nas sp. strain CBB1. All purification procedures were performed at 4°Cusing an automated fast protein liquid chromatography (FPLC) system(AKTA Design; Amersham Pharmacia Biotech). Chromatography col-umns and column resins were from Amersham Biosciences, Piscataway,NJ. An Amicon Ultra-15 (30K) ultrafiltration membrane (Millipore, Bed-ford, MA) was used for concentration and buffer exchange of the proteinfractions containing TMU oxidation activity.

About 8 g of frozen cells was thawed and suspended in 30 ml of 50 mMKPi buffer (pH 7.5), and DNase I was added to a final concentration of 10�g ml�1. The cell suspension was broken by passing through a chilledFrench press twice at 138 MPa. Unbroken cells and debris were removedfrom the lysate by centrifugation (20,400 � g for 20 min at 4°C). Theresulting crude cell extract (CCE) was loaded onto a 100-ml (bed volume)Q-Sepharose column (GE Healthcare) preequilibrated with 50 mM KPi

buffer (pH 7.5). After the unbound proteins were washed with the samebuffer at 1 ml min�1, bound proteins were eluted with a linear gradientfrom 0 to 0.5 M KCl in the same buffer and at the same flow rate. Fractionsexhibiting TMU oxidation activity eluted at 0.05 to 0.125 M KCl. Theactive fractions were pooled, concentrated, and exchanged into 50 mMKPi buffer (pH 7.5) by ultrafiltration.

This concentrated active fraction was loaded onto a 10-ml (bed vol-ume) Blue-Sepharose column (Capto Blue; GE Healthcare) preequili-brated with 50 mM KPi buffer (pH 7.5). After the unbound proteins werewashed from the column with 120 ml of the buffer, bound proteins wereeluted with 1 M KCl at a rate of 1 ml min�1. Fractions containing TMU

oxidation activity were pooled, concentrated, and exchanged into 50 mMKPi buffer (pH 7.5).

A 4 M ammonium sulfate solution was added to the enzyme fractionfrom Blue-Sepharose to a final concentration of 0.5 M. After 1 h on ice, theprecipitate was removed by centrifugation at 16,000 g for 20 min and thesupernatant was loaded onto a 1-ml (bed volume) Phenyl-Sepharosehigh-performance column (GE Healthcare) preequilibrated with 50 mMKPi buffer containing 0.5 M ammonium sulfate. Unbound proteins wereeluted from the column with the same buffer containing 0.5 M ammo-nium sulfate. Bound proteins were eluted with a reverse linear gradient ofammonium sulfate (0.5 to 0 M in KPi buffer) at a flow rate of 1 ml min�1.Fractions exhibiting TMU oxidation activity were pooled, concentrated,and exchanged into 50 mM KPi buffer (pH 7.5) containing 0.1 M NaCl.This preparation was loaded onto an 80-ml (bed volume) Sephacryl S-300HR column (Amersham) preequilibrated with 0.1 M NaCl in 50 mM KPi

buffer. Proteins were eluted from the column with the same buffer at aflow rate of 1 ml min�1. Fractions containing TMU oxidation activitywere combined, concentrated, and exchanged into 50 mM KPi buffer(pH 7.5).

PCR amplification of the gene catalyzing TMU oxidation fromCBB1 genome. The forward degenerate primer tmuM-degF1 was de-signed based on the lowest degeneracy of amino acids (KIGADVT) in theN-terminal sequence of the purified protein (see Table S2 and Fig. S1 inthe supplemental material). A BLASTP search of the N-terminal aminoacid sequence showed significant homology with five proteins (GenBankaccession nos. ACF60813, EAQ44903, and CAJ95547), including a FAD-binding monooxygenase (GenBank accession no. EDZ47364) and salicy-late 1-monooxygenase (GenBank accession no. AAZ62959). A CLUSTALW2 (25) alignment of these five homologous proteins identified a con-served region near the 3= end of this multiple sequence alignment (see Fig.S1 in the supplemental material). Reverse degenerate primer tmuM-degR1 was designed from this conserved region (see Table S2 and Fig. S1in the supplemental material). PCR using tmuM-degF1 and tmuM-degR1was conducted using PfuUltra DNA polymerase HF (Agilent Tech-nologies, La Jolla, CA) with fosmid DNA pMVS848 as a template. Fos-mid library construction and isolation of fosmid clone E. coliEPI300(pMVS848) are described in the supplemental material. The PCRconditions were 30 cycles of 95°C for 30 s, 62°C for 30 s, and 72°C for 1min, followed by a final extension at 72°C for 10 min. The amplified PCRproduct was cloned into vector pSC-B-amp/kan (Agilent Technologies),and three clones were chosen for DNA sequencing. The specific primerstmuM-up1, tmuM-up2, tmuM-down1, and tmuM-down2 (see Table S2in the supplemental material) designed from the DNA sequence of thisamplified PCR product were used to sequence the flanking region of thePCR product with fosmid DNA pMVS848 as a template.

Protein expression and purification. Primers tmuM-F-NdeI andtmuM-R-NotI (see Table S2 in the supplemental material) were used withthe fosmid DNA pMVS848 as the template to amplify the entire codinggene sequence of TMU-oxidizing enzyme. The reverse primer was alsodesigned by removing the wild-type stop codon so that the histidine tagcontained in the pET32a vector would be fused to the expressed protein.PCRs were carried out with PfuUltra DNA polymerase. The amplificationproduct was cloned into pSC-B-amp/kan to generate pSC-tmuM. Therestriction fragment containing the tmuM gene obtained from NdeI-NotIdigestion of pSC-tmuM was ligated into the dephosphorylated vector,pET32a (Novagen), previously digested by NdeI and NotI. DNA sequenc-ing confirmed that no mutation was introduced in the gene encoding theTMU-oxidizing enzyme in plasmid pET32a. The resulting plasmidpMVS848a (see Table S1 in the supplemental material) was electropo-rated into E. coli BL21(DE3) cells. The expression of recombinant His-tagged TMU-oxidizing enzyme in E. coli BL21(DE3) and the two-steppurification using metal affinity and ion-exchange chromatography aredescribed in the supplemental material. Protein quantitation, the deter-mination of native and subunit molecular weights, the N-terminal se-quence, the bound flavin content, the kinetic parameters, the oxygen re-

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quirement, and the stoichiometry of TMU oxidation reaction aredescribed in the supplemental material.

Enzyme assays. A typical TMU oxidation activity assay was carriedout at 30°C in 1-ml reaction mixture containing 0.5 mM NADH, 0.5 mMTMU, and appropriate amounts of crude cell extract, fractions fromFPLC, or purified protein, in 50 mM KPi buffer (pH 7.5). The reaction wasinitiated by the addition of TMU. NADH oxidation was monitored at 340nm using a UV-Visible spectrophotometer (Shimadzu UV-2450). An ex-tinction coefficient of 6,200 M�1 cm�1 for reduced minus oxidizedNADH was used in calculating the enzyme activity. One unit of activitywas defined as the amount of protein required to oxidize 1 �mol ofNADH per min under defined reaction conditions. The TMU oxidationactivity was also determined by measuring the disappearance of trimethy-luric acid from the reaction mixture by HPLC analysis. Periodically, 100�l from the reaction mixture was mixed with equal volume of acetonitrilefor quantifying trimethyluric acid.

Incorporation of 18O2 into trimethyluric acid. An 18O2 (CambridgeIsotope Laboratories, Inc., Andover, MA) incorporation experiment wasconducted in a 1-ml (total volume) reaction mixture containing, 0.5 mMTMU, 0.5 mM NADH, and 10 �l of purified recombinant TMU-oxidizingenzyme. The reaction mixture was transferred into a modified 0.5 l round-bottom flask with two side arms and a quartz cuvette fused at the bottom.The enzyme was pipetted into one side arm. After the air in the flask wasevacuated, argon was introduced. The enzyme in the side arm was tiltedinto the reaction mixture. NADH consumption was monitored at 340 nmat 30°C to confirm there is no oxidation under anaerobic conditions. 18O2

was then introduced into the reaction vessel to initiate the reaction. Theenzyme activity was monitored by the decrease of absorbance at 340 nm.After 30 min, an aliquot of the reaction mixture was mixed with an equalvolume of acetonitrile to precipitate the protein. The supernatant wasthen analyzed by an HPLC system equipped with a Shimadzu LCMS-2010EV single-stage quadruple mass analyzer. A parallel control reactionwas carried out with 16O2 (air), and the product was analyzed.

Liquid chromatography-mass spectrometry (LC-MS) analysis oftime course of product formation. The time course of TMU-oxidizingenzyme-catalyzed reaction with TMU was conducted with the recombi-nant enzyme. Aliquots sampled from the reaction mixture were mixedwith equal volumes of acetonitrile to stop the reaction. Identification andquantification of trimethyluric acid, reaction intermediates, and endproduct were conducted with a Shimadzu LC-20AT HPLC systemequipped with Shimadzu LCMS-2010EV single-stage quadrupole massanalyzer and a Hypersil BDS C18 column (4.6 by 125 mm) as describedpreviously (43). Methanol-water-acetic acid (20:80:0.5 [vol/vol/vol]) wasused as an isocratic mobile phase with a flow rate of 0.5 ml min�1. Sub-strates and products resolved by the C18 column first passed through thephotodiode array detector, during which UV-visible absorption spectrawere recorded. Then molecules were ionized by electrospray ionization(ESI) in positive-ion mode. The following compounds corresponding totheir (M�1)� ions were monitored: caffeine, m/z � 195; trimethyluricacid (TMU), m/z � 211; 1,3,7-trimethyl-5-hydroxyisourate (TM-HIU),m/z � 227; 3,6,8-trimethyl-2-oxo-4-hydroxy-4-carboxy-5-ureidoimida-zoline (TM-OHCU), m/z � 245; and 3,6,8-trimethylallantoin (TMA),m/z � 201.

Separation and identification of TMA. A 100-ml reaction, containing0.5 mM TMU, 0.5 mM NADH, and 1.5 mg of recombinant enzyme in 50mM KPi buffer (pH 7.5), was incubated at 30°C for 1 h, and the TMUdisappearance was monitored by the HPLC method described above. Atthe end, the reaction mixture was concentrated to 5 ml using a ThermoSavant AES1010 automatic environmental SpeedVac system (ThermoElectron) at room temperature. The reaction mixture was then loadedonto prep-scale Silica Gel 60 F254 plates of 1 mm (EM Science,Gibbstown, NJ) and developed with chloroform-methanol (85:15 [vol/vol]) as the solvent system. Resolved compounds were visualized by ex-posing the plates to UV light at 254 nm. TMU (Rf � 0.59) and the reactionproduct (Rf � 0.51) were subsequently extracted from a preparative-scale

TLC plate with methanol and subjected to LC-MS analysis (as describedabove). Further, nuclear magnetic resonance (NMR), high-resolutionelectro-spray ionization mass spectrometry (HRESIMS), and chiral-HPLC analyses were performed on the reaction product as describedbelow.

Analytical procedures. 1H and 13C NMR spectra of TMA were ob-tained from a Bruker AMX-600 high-field spectrometer equipped with anIBM Aspect-2000 processor (Bruker Instruments, Billerica, MA). A 2-mgportion of purified product (TMA) was dissolved in 50 �l of CDCl3 andtransferred into a 5-mm high-quality NMR tube (Goss Scientific Instru-ments, Great Baddow, Essex, United Kingdom) to acquire the spectra.Solvent CDCl3 served as an internal standard. All NMR spectra were ob-tained in chloroform-d with chemical shifts expressed in parts per million(�) and coupling constants (J) in Hz. Correlation spectroscopy (COSY),heteronuclear multiple-bond coherence (HMBC), and heteronuclearmultiple-quantum coherence (HMQC) experiments were carried out us-ing the same Bruker AMX-600 high-field spectrometer. HRESIMS wasdone using a Fisons VG-ZAB-HF reversed-geometry (BE configuration,where B is a magnetic sector and E is an electrostatic analyzer) massspectrometer (VG Analytical, Inc.) at the High Resolution Mass Spec-trometry Facility at The University of Iowa. A chiral HPLC analysis on aChiralcelpak IA column (4.6 by 250 mm; Chiral Technologies, Exton, PA)was performed using a Shimadzu LC-20AT HPLC system with an iso-cratic mobile phase (ethanol-hexane, 50:50 [vol/vol]) at a flow rate of 0.5ml min�1.

Homology modeling. A BLASTP search with TMU-oxidizing enzymeand HpxO (30) as queries against the PDB identified a flavin-containinghydroxylase involved in pyocyanin biosynthesis (PhzS) from Pseudomo-nas aeruginosa (16) as the best match, with 30% sequence identity to both.PhzS belongs to the family of flavin-dependent aromatic hydroxylases, ofwhich the best characterized is p-hydroxybenzonate hydroxylase(PHBH). Homology models for TMU-oxidizing enzyme and HpxO weregenerated by using the program Modeller 9.10 (9) and PhzS (PDB ID3C96) as the template. The homology models (ten for each protein) weresuperimposed on PhzS and PHBH (PDB ID 1PBE) using PyMOL 1.5(Schrödinger, LLC), and the FAD- and substrate-binding cavities wereidentified. The ligand-bound structure of VOIDOO (23) was used to cal-culate the solvent-accessible volumes of the active site cavities for each ofthe models and then averaged for each protein.

Nucleotide sequence accession numbers. The nucleotide sequencesof cdhA, cdhB, cdhC, tmuM, tmuH, and tmuD deposited at the GenBanknucleotide sequence database were assigned accession numbers ADH15879,ADH15880, ADH15881, JQ743481, JQ743482 and JQ743483, respectively.

RESULTSPurification of TMU-oxidizing enzyme from cell extracts ofstrain CBB1. The consumption of trimethyluric acid in crude cellextract was dependent on the presence of NADH (data notshown). While 0.2 mM TMU was completely utilized in 15 min bycrude cell extract (200 �g/ml) in the presence of NADH, onlymarginal activity was observed without the cofactor. Activity wasnot observed with boiled cell extract. TMU-oxidizing enzyme waspurified 55-fold, but with a low recovery of 0.2%. In the absence ofTMU, no NADH oxidation was observed with the purified en-zyme. The specific activity of the purified enzyme was 4128 mUper mg (Table 1, Fig. S2). The molecular mass of TMU-oxidizingenzyme as estimated by SDS-PAGE gel electrophoresis was �43kDa (see Fig. S2 in the supplemental material), and its N-terminalsequence was determined to be SRPLRVTIIGAGIGGLSAAVALRKIGADVT.

Cloning, expression, and purification of TMU-oxidizing en-zyme from CBB1. PCR using the primers tmuM-degF1 andtmuM-degR1 successfully amplified a single, 900-bp PCR productfrom the fosmid DNA pMVS848 (see Table S1 in the supplemen-

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tal material). A BLASTX search using the DNA sequence of this900-bp product showed that this amplified partial gene has signif-icant homology to the FAD-binding monooxygenase family ofproteins. Further sequencing of the flanking region of this partialgene revealed a complete ORF with 1,191 nucleotides (nt). Thegene was designated tmuM, and it encoded a 396-amino-acid pro-tein with a theoretical Mr of 42,619 and a pI of 6.12. N-terminalsequence derived from the tmuM gene matched the experimen-tally determined N terminus of the purified TMU-oxidizing en-zyme from CBB1. The amplified gene was cloned into the pET32aplasmid with a His6 tag at the C-terminal end. The recombinantprotein was overexpressed in a soluble form and purified to elec-trophoretic homogeneity using immobilized-metal affinity chroma-tography and Q-Sepharose chromatography (see Fig. S2 in thesupplemental material). The specific activity of the recombinantTMU-oxidizing enzyme was 9,700 mU per mg, which is twice thatof the purified enzyme from CBB1. Given the higher activity of therecombinant enzyme, this preparation was used for further char-acterization.

Biochemical characterization of TMU-oxidizing enzyme.The recombinant TMU-oxidizing enzyme was similar to the en-zyme purified from CBB1 in terms of the native (�45-kDa) andsubunit (�43-kDa) molecular mass, and UV-visible absorptionspectrum (see Fig. S2 in the supplemental material). Both the na-tive and the recombinant enzymes were monomeric with UV-visible absorption spectrum maxima at 450, 380, and 280 nm,which is characteristic of a bound flavin cofactor. HPLC analysisof the cofactor released from the recombinant enzyme upon heatdenaturation was identified as FAD with TmuM:FAD stoichiom-etry (1:1; data not shown). The apparent Km for TMU and the kcat

of the recombinant enzyme in 50 mM KPi (pH 7.5) at 30°C were10.2 � 2.2 �M and 448.9 � 21.7 min�1, respectively (Table 2).

Characterization of the enzyme as TmuM. When the enzymereaction was carried out in a completely anaerobic environment,

no TMU disappearance or NADH oxidation was observed. Uponthe introduction of 18O2 into the reaction chamber, activity wasimmediately observed and was monitored spectrophotometri-cally. The end product of the reaction was characterized by ESI-LC/MS analysis as TMA. TMA produced in the presence of atmo-spheric O2 had an m/z of 201, which agreed with the (M�1)� ionof the compound (Fig. 1b). TMA produced in the presence of 18O2

had an m/z of 203, indicating the incorporation of one atom ofisotopic oxygen (Fig. 1a). Further characterization of TMA is de-scribed in detail in the product characterization and product iden-tification sections below. Monitoring the oxygen consumption ina Clark-type oxygen electrode and simultaneously quantifyingTMU disappearance by HPLC established the stoichiometry of thereaction. After 5 min of reaction, 190.5 nmol of O2 was consumed,while 185 nmol of TMU was degraded. NADH to TMU stoichi-ometry was also established as 1:1 in a separate experiment bymonitoring both NADH oxidation and TMU disappearance.Based on these results, the TMU-oxidizing enzyme was designatedas TMU monooxygenase (TmuM).

Substrate specificity of TmuM. The apparent kinetic parame-ters of the recombinant enzyme with various tri-, di-, andmonomethyluric acids and uric acid are shown in Table 2. Initialactivity assays with wild-type TmuM and recombinant TmuM-His6 suggested that the enzyme was active on TMU and othermethyluric acids, but not on uric acid (data not shown). The cat-alytic activity kcat (turnover number) of TmuM-His6 for TMU wasfound to be 2.5, 3.8, and 15.4 times higher than that of 1,3- and3,7-dimethyluric acid and 1-methyluric acid, respectively (Table2). These results suggested that TMU is the preferred substrate forTmuM, followed by 1,3-dimethyluric acid, 3,7-dimethyluric acidand 1-methyluric acid. In contrast, the catalytic efficiency kcat/Km

for 3,7-dimethyluric acid was two times higher than that of TMUdue to an order of magnitude lower Km. 1-Methyluric acid was theleast preferred substrate, although its Km was only an order ofmagnitude lower than that of TMU (Table 2).

Detailed characterization of the product of TmuM-catalyzedreaction. The time course of TMU oxidation and product forma-tion catalyzed by recombinant TmuM-His6 was monitored byLC-MS (Fig. 1d). Initial HPLC analysis showed that TMU wascompletely consumed while NADH was converted to NAD� (datanot shown). Since no product peak could be detected when UVabsorption was used as the detection method during HPLC anal-ysis, the intermediates and product formation were subsequentlymonitored by ESI-MS (Fig. 1d). The ESI-LC/MS analysis datawere monitored in the positive-ion mode at m/z � 211, 227, 245,or 201, based on the expected products formed from TMU trans-formation, which were hypothesized to be TM-HIU (molecularweight [MW], 226), TM-OHCU (MW, 244), or TMA (MW, 200),

TABLE 1 Purification of trimethyluric acid oxidizing enzyme from Pseudomonas sp. strain CBB1

Purification step Amt of protein (mg)

Activitya

Yield (%) Purification (fold)Specific (mU/mg) Total (mU)

Crude cell extract 903 74 66,806 100 1Q-Sepharose 83 407 33,703 50 5Blue-Sepharose 20.3 1,228 24,916 37 16Phenyl-Sepharose HP 0.97 3,577 3,478 5 48Sephacryl-S300 0.03 4,128 116.4 0.17 55a One unit of activity was defined as the amount of protein required to oxidize 1 �mol of NADH per min under the reaction conditions described in Materials and Methods.

TABLE 2 Kinetic parameters and substrate specificities of purifiedTmuM-His6

Substrate

Avg � SDa

kcat (min�1) Km (�M)kcat/Km

(min�1 �M�1)

Trimethyluric acid 448.9 � 21.7 10.2 � 2.2 44.1 � 2.11,3-Dimethyluric acid 185.0 � 16.4 126.5 � 29.3 1.5 � 0.13,7-Dimethyluric acid 118.2 � 8.4 1.3 � 0.6 89.4 � 6.31-Methyluric acid 29.1 � 2.4 1.2 � 0.5 24.5 � 2.0Uric acid NA NAa Averages and standard deviations were derived from three independent assays. Initialreaction rates were determined by monitoring substrate-dependent NADH oxidation at340 nm. NA, no activity.

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respectively. Corresponding nonmethylated metabolites have beenreported for the uric acid pathway (8, 31, 32). Selected ion monitor-ing (SIM) revealed the presence of substrate TMU, metabolite II, andmetabolite IV with (M�1)� ions at m/z � 211, m/z � 227 (Fig. 1c),and m/z � 201 (Fig. 1b), respectively. The observed molecularmasses of 226 (with m/z � 227) for metabolite II and 200 (withm/z � 201) for metabolite IV matched to that of TM-HIU andTMA, respectively. Metabolite II at m/z � 227 could be detectedinitially but disappeared after 10 min (Fig. 1d). In contrast, TMAat m/z � 201 continued to accumulate until the end of the reaction(Fig. 1d). Based on these results, and the observation of 5-hy-droxyuric acid in the uric acid degradation pathway (22), metab-olite II (TM-HIU) was proposed as the true product of TMUmonooxygenase reaction. Metabolite II (TM-HIU) was unstableunder the reaction condition and spontaneously converted toTMA (possibly via metabolite III), as is the case in the uric acid

degradation pathway (8, 31, 32). Metabolite III (TM-OHCU)could not be detected.

Identification of metabolite IV as racemic TMA. The struc-ture of metabolite IV (TMA), purified from the TmuM reactionby using preparative-scale thin-layer chromatography, was estab-lished by spectroscopic analyses. HRESIMS showed that metabo-lite IV M� was m/z � 200.0907, indicating an empirical formula ofC7H12N4O3 (calculated value, 200.0909). Thus, metabolite IVcontained two hydrogen atoms more and one carbon atom lessthan TMU. The 1H NMR spectrum of metabolite IV contained allsignals of TMU, in addition to two extra signals (Table 3). Theseincluded signals for a new secondary amine proton at 5.77 ppm forH-8 (singlet, broad) and a doublet at 5.19 ppm (J � 7.77 Hz) forH-5 coupled to the H-1 signal of the secondary amine group. The13C NMR spectra of metabolite IV showed one less signal thanTMU but contained all three methyl group signals at 40.7 ppm

FIG 1 Mass spectra of metabolite IV (TMA) from 18O2 (a) and 16O2 (b) incorporation. (c) Metabolite II (proposed TM-HIU) formed from TMU byrecombinant TmuM-His6. (d) Time course of selected ion monitoring (SIM)/LC-MS analysis of TMU oxidation and product formation by TmuM-His6. The ionintensities of metabolite I and II are associated with the primary y axis. The ion intensity of metabolite IV is associated with secondary y axis. Symbols: �, TMU(I);o, TM-HIU(II); Œ, TMA(IV).

TABLE 3 1H and 13C NMR data of metabolite IV

H Chemical shift (ppm) Peak type J coupling (Hz) C Chemical shift (ppm)

H-1 (1H) 6.51 Doublet 7.77 C-2 (CO) 171.5H-5 (1H) 5.19 Doublet 7.77 C-4 (CO) 156.1H-8 (1H) 5.77 Singlet, broad C-5 (CH) 65.9H-9 (3H) 2.98 Singlet C-7 (CO) 158.3H-10 (3H) 2.88 Singlet C-9 (CH3) 40.7H-11 (3H) 2.69 Doublet 4.6 C-10 (CH3) 26.8

C-11 (CH3) 24.9

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(C-9), 26.8 ppm (C-10), and 24.9 ppm (C-11) (Table 3). Fromthese data it is clear that loss of the carbon atom in metabolite IVwas not due to N-demethylation but occurs via ring rupture. The13C NMR spectrum of metabolite IV also showed two carbonylgroup signals at 171.5 ppm (C-2) and 158.3 ppm (C-7) and onealtered carbonyl group signal at 156.1 ppm (C-4), which is fartherdownfield than the original TMU signal. In addition, the spectrumof metabolite IV contained signal for a new secondary carbonatom at 65.93 ppm (C-5). Signal assignments and connectivitieswere confirmed using COSY, HMBC, and HMQC spectral edit-ing. COSY analysis correlated H-8 with H-11 and correlated H-1with H-5. HMBC showed 3-bond correlations between H-11 andC-7, i.e., H-9 between C-2 and C-4, H-10 between C-5 and C-7,and H-5 between C-2 and C-10. Based on all of these data, metab-olite IV was characterized as TMA. It should be pointed out thatthere is no commercial sample of TMA available.

The specific rotation ([]25D) of metabolite IV was close to zero(�0.003°, concentration of 0.2 g/100 ml, CHCl3). This indicates thatthe stereochemistry of the C-5 atom (in metabolite IV) does not cor-respond to one specific enantiomer but rather is a racemic mixture oftwo enantiomers. The racemic nature of the product was furtherconfirmed by chiral stationary-phase HPLC with a ChiralcelpakIA column. Pure metabolite IV, which eluted as a single peak atretention time 8.14 min with reversed-phase C18 column, resolved

into two separate peaks of equal peak area with retention times of10.56 and 12.35 min, respectively (see Fig. S3a in the supplementalmaterial). The ESI-LC-MS analysis confirmed the molecular massof both peaks as 200 (see Fig. S3b in the supplemental material).These results further suggest that metabolite IV (TMA) isolatedfrom the TmuM reaction is a racemic mixture of two enantiomersof TMA.

Identification of cdhA, cdhB, cdhC, tmuM, tmuH, and tmuDgenes in CBB1. Previously, caffeine dehydrogenase (Cdh) puri-fied from CBB1 was proposed to have an � ( [90-kDa], [32-kDa], and � [20-kDa])-subunit structure (42). PCR based onthe primers (see Table S2 in the supplemental material) designedfrom the N-terminal protein sequence of the large (90-kDa) andmedium (40-kDa) subunits of caffeine dehydrogenase resulted inamplification of a 2,392-nt DNA fragment from CBB1 genome(see Table S2 in the supplemental material). This amplified frag-ment contained a 2,376-nt ORF, which encoded an 87-kDa prod-uct. Its deduced protein sequence was identical to the N-terminalprotein sequence of Cdh large subunit. Therefore, this 2,376-ntORF was designated cdhA (GenBank accession no. ADH15879).Using sequences derived from cdhA as probes to screen a genomiclibrary of CBB1, a clone, E. coli EPI300(pMVS848), containingcdhA was found. DNA sequencing of the flanking regions of thisclone revealed two ORFs of 888 and 528 nt immediately down-

FIG 2 Physical map of genes for caffeine transformation in a 25.2-kb gene cluster in Pseudomonas sp. strain CBB1. Genes are denoted by arrows, and the extentsand directions of transcription are indicated. The two genetic modules are denoted by the substrate upon which the gene products act: caffeine, black; andtrimethyluric acid, dark gray. The sequence similarity and function assignment of the ORF-encoded proteins are indicated in the table.

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stream of cdhA (Fig. 2). While the first ORF encoded a 32.5-kDaproduct with its deduced protein sequence matching the N-termi-nal protein sequence of Cdh -subunit, the second ORF encodedan 18.4-kDa product, which matched the molecular weight of Cdhsmall (�) subunit (42). Based on these results, the 888-nt ORF wasdesignated cdhB (GenBank accession no. ADH15880), and the528-nt ORF was designated cdhC (GenBank accession no.ADH15881). A BLASTX search of full-length cdhA, cdhB, andcdhC against the nonredundant protein sequence (nr) databaseidentified the molybdopterin-binding subunit of a putative xan-thine dehydrogenase (GenBank accession no. YP_003395893),the medium subunit of a bifunctional hydratase/alcohol dehydro-genase (GenBank accession no. ADV16272) involved in anaerobictransformation of cyclohexanol, and the small subunit of a puta-tive aldehyde oxidase (GenBank accession no. YP_002521823) asbest hits, with 49, 39, and 59% identities, respectively. Searchagainst the Swiss-Prot database identified the large (GenBankaccession no. P19919), medium (GenBank accession no. P19914),and small (GenBank accession no. P19915) subunits of carbonmonoxide dehydrogenase as the best hit with 32, 31, and 47%identities, respectively (Fig. 2). Xanthine dehydrogenase, hydra-tase/alcohol dehydrogenase, aldehyde oxidase, and carbon mon-oxide dehydrogenase have an �- subunit structure like that ofCdh and belong to the molybdopterin-binding oxidoreductasefamily containing molybdopterins, FAD, and [2Fe-2S] clusters ascofactors (5, 19). The location of the tmuM gene encoding theenzyme TmuM was also identified and amplified from the fosmidDNA pMVS848 (see Table S1 in the supplemental material). A

BLASTX search of tmuM against the nr database identified a FAD-binding monooxygenase (GenBank accession no. YP_003741647)from Erwinia billingiae Eb661 (Fig. 2) and HpxO (GenBank acces-sion no. ACF60813) from Klebsiella pneumonia as the best hits,with sequence identities of 38 and 37%, respectively.

Identification of the genes for the first two reactions of the C-8oxidation pathway for caffeine metabolism (Fig. 3) in fosmidDNA pMVS848 prompted us to sequence the flanking regions ofcdhA, cdhC, and tmuM genes. A total of 25.2 kb of the DNA se-quence was obtained from the fosmid DNA pMVS848. Compu-tational analysis of this 25.2-kb genomic DNA sequence identified21 complete ORFs, including the cdhA, cdhB, cdhC, and tmuMgenes and two incomplete ORFs (Fig. 2). A BLASTX search oftmuH, the first complete ORF in the gene cluster (Fig. 2), againstthe nr database identified a putative hydroxyisourate (HIU) hy-drolase from Rhizobium leguminosarum bv. Trifolii WSM1325(GenBank accession no. YP_00297642) as the best match with50% identity. When the same search was performed against theSwiss-Prot database, putative HIU hydrolase 2 from Sinorhizo-bium meliloti 1021 (GenBank accession no. Q92UG5) was identi-fied as the best match with 44% identity. Similarly, a BLASTXsearch of tmuD, the second complete ORF (Fig. 2) in the genecluster, against the nr database identified putative OHCU decar-boxylase from Starkeya novella DSM 506 (GenBank accession no.YP_003695277) with 39% sequence identity. The best hit from theSwiss-Prot database was putative OHCU decarboxylase fromHaloferax volcanii DS2 (GenBank accession no. D4GPU8) with37% identity (Fig. 2). Thus, on the basis of these sequence simi-

FIG 3 Proposed caffeine transformation pathway in Pseudomonas sp. strain CBB1. Caffeine dehydrogenase (Cdh) oxidizes caffeine to TMU (I), which is oxidizedto TM-HIU (II) by trimethyluric acid monooxygenase (TmuM). (a) Spontaneous conversion of TM-HIU to racemic TMA through the TM-OHCU (III)intermediate. (b) Proposed pathway for the enzymatic conversion of TM-HIU to S-(�)-TMA. The gene representation is in italics. Metabolites: I, 1,3,7-trimethyluric acid (TMU); II, 1,3,7-trimethyl-5-hydroxyisourate (TM-HIU); III, 3,6,8-trimethyl-2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline (TM-OHCU); IV, 3,6,8-trimethylallantoin (TMA).

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larities, tmuH encoding a 114-amino-acid product and tmuD en-coding a 291-amino-acid product were designated putative 1,3,7-trimethyl-5-hydroxyisourate hydrolase (TM-HIU hydrolase) andputative 3,6,8-trimethyl-2-oxo-4-hydroxy-4-carcoxy-5-ureido-imidazoline decarboxylase (TM-OHCU decarboxylase), respec-tively.

DISCUSSION

In this report, we have proposed a new pathway for the degrada-tion of caffeine via C-8 oxidation in a caffeine-degrading bacte-rium, Pseudomonas sp. strain CBB1 (Fig. 3). The first reaction ofthe proposed pathway is the hydrolytic oxidation of caffeine toTMU, catalyzed by a novel caffeine dehydrogenase (Cdh) (42).Previously, Yu et al. had established by sequence homology thatcdhA, the large ( [90-kDa]) subunit of Cdh, contained themolybdopterin cofactor (42). In the present study, total sequenceanalysis of cdhB and cdhC revealed that cdhB had significant ho-mology with FAD binding medium subunits of similar enzymessuch as bifunctional hydratase/alcohol dehydrogenase and carbonmonoxide dehydrogenase. In addition, cdhC was homologous tothe corresponding iron-sulfur containing small subunits of all ofthe heterotrimeric enzymes. This suggests that the (40-kDa)-subunit of Cdh contains FAD cofactor and that the �-subunitcontains the iron-sulfur cluster. Cdh is similar to the knownNAD�-dependent xanthine dehydrogenases in terms of catalysis,subunit structure (�) and cofactor content. However, this en-zyme is quinone dependent, does not use NAD(P)�, and has noactivity with xanthine (42). This is the first report of completesequence of Cdh.

The thrust of this work is on the second reaction, hydroxyla-tion of TMU to 1,3,7-trimethyl-5-hydroxyisourate (TM-HIU;metabolite II) catalyzed by a novel NADH-dependent trimethyl-uric acid monooxygenase (TmuM), a flavoprotein. A time courseof this reaction, analyzed by ESI-MS in the positive-ion mode,revealed transient accumulation of unstable TM-HIU and contin-uous accumulation of TMA (Fig. 1d). Therefore, the true oxida-tion product of TmuM catalyzed reaction was identified as TM-HIU (Fig. 3). One atom of molecular oxygen incorporated intoTMU, forming TM-HIU, was detected in TMA (Fig. 1a). Thesecond oxygen was incorporated into water. TM-HIU is unstableand spontaneously decomposes to racemic TMA during thecourse of the reaction (Fig. 1d and 3a). The racemic TMA wascharacterized by chiral-HPLC (see Fig. S3a in the supplementalmaterial), NMR (Table 3), 18O2 incorporation (Fig. 1a), and op-tical rotation (data not shown). Transient accumulation of thehypothesized second unstable metabolite in the C-8 caffeine oxi-dation pathway (Fig. 3b; metabolite III, TM-OHCU with a m/z of245) could not be detected during the course of the reaction. Thismay be due to not enough positive or negative ions generatedunder the reaction condition or due to rapid decarboxylation ofTM-OHCU. This supposition is similar to the corresponding uricacid degradation pathway reaction (32), where the oxidation ofuric acid by urate oxidase results in the transient accumulation of5-hydroxyisourate (HIU) (30). HIU formed from uric acid by thetraditional urate oxidase (not a monooxygenase) from soybeanroot nodules was also shown to be unstable, with a half-life of lessthan 30 min at neutral pH (20–22). This unstable compoundspontaneously hydrolyzes to 2-oxo-4-hydroxy-4-carboxy-5-ure-idoimidazoline (OHCU), which gets further decarboxylatedspontaneously to racemic allantoin, the stable end product of

urate oxidase reaction (32). The instability of TM-HIU can beattributed to its structural similarity to HIU. An 18O2 incorpora-tion experiment with TmuM was conducted as an endpoint reac-tion (Fig. 1a and b) due to sampling difficulties from the enrichedatmosphere. Thus, isotopic oxygen-labeled 18O-TM-HIU couldnot be detected at the end of this experiment. Nevertheless, theretention of isotopic oxygen in 18O-TMA with a m/z of 203 (Fig.1a) conclusively established TmuM as a monooxygenase.

The gene encoding TmuM (tmuM) is 1,191 nt in length. Se-quence analysis of tmuM revealed that this enzyme belongs to thefamily of FAD containing aromatic-ring hydroxylases, which in-cludes FAD-dependent urate oxidase (HpxO; GenBank accessionno. ACF60813) from Klebsiella pneumoniae (30), a flavin-depen-dent hydroxylase (PhzS; GenBank accession no. AAG07605)from Pseudomonas aeruginosa (16), salicylate 1-monooxygenase(GenBank accession no. AAZ62959), and other uncharacterized/putative FAD-binding monooxygenases (GenBank accession nos.EAQ44903, CAJ95547, and EDZ47364). A multiple sequencealignment of these homologous proteins revealed that the primarysequence of TmuM contains conserved residues that include anFAD and NADH-binding domains (see Fig. S1 in the supplemen-tal material). Phylogenetic analysis of these seven homologousflavoprotein monooxygenases (data not shown) revealed TmuMand HpxO clustered together in a clade, distinct from the otherfive flavin monooxygenases. TmuM and HpxO appear divergingfrom a common ancestor.

There are two types of enzymes that catalyze reactions similarto TmuM. The first one is the most prevalent urate oxidase (EC1.7.3.3), which does not require any cofactor (20). The coproductof this reaction, according to the enzyme nomenclature, is hydro-gen peroxide. The second type of enzyme is NADH-dependent,FAD-containing urate oxidase. This enzyme follows a catalyticmechanism similar to flavin containing aromatic-ring hydroxy-lases (18). HpxO from K. pneumoniae (30) and TmuM are exam-ples of the second type. TmuM is closely related to HpxO in termsof sequence similarity, phylogeny, cofactor content, NADH de-pendence and, likely, the catalytic mechanism. However, the en-zymatic conversion of urate to HIU catalyzed by HpxO, althoughshown to be oxygen dependent (30), has not been characterized asa monooxygenase. It has been mis-assigned as urate oxidase. Hy-drogen peroxide is not one of the products of this reaction. Basedon the similarity of HpxO catalyzed reaction to TmuM, we feelthat HpxO should be designated as uric acid monooxygenase.Both TmuM and HpxO belong to a unique class of (methyl) uratemonooxygenases. However, TmuM is distinct from HpxO; it hasno activity on uric acid. TmuM has the highest catalytic activity fortrimethyluric acid compared to di- and monomethyluric acids(Table 2). A comparison of the active-site cavities of TmuM andHpxO based on homology models (Fig. 4I) reveals a much biggercavity for TmuM (244 � 45 Å3) compared to the cavity for HpxO(121 � 10 Å3). Also, the GRAVY value based on the Kyte andDoolittle hydropathy scale (24) for the residues lining the putativeactive site reveals a much more hydrophobic cavity for TmuM(0.673) (Fig. 4IIA) compared to HpxO (�0.218) (Fig. 4IIB).These results are consistent with the enzyme data (Table 2) andprovide an initial explanation of the specificity of TmuM to thelarger TMU and 3,7-dimethyluric acid.

The metabolites downstream of TMU were characterized bygene cluster analysis of the 25.2-kb genomic DNA segment ofCBB1, which harbored all of the genes of the pathway (Fig. 2 and

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3). We undertook this method due to (i) the lack of availability ofthe metabolites TM-HIU, TM-OCHU, and racemic and S-(�)-TMA proposed in this pathway (Fig. 3) and (ii) the lack of pub-lished synthetic methods for these compounds. Homology andgene cluster-based analysis is a valid tool for pathway analysis andhas also been used extensively in recent years for elucidation ofuric acid degradation pathway in various microbial strains, in-cluding Bacillus subtilis (35), Klebsiella pneumoniae (8, 17), andKlebsiella oxytoca (31). Analysis of the hpx gene cluster in K. pneu-moniae (8) identified seven genes involved in oxidation of hypo-xanthine to allantoin via uric acid. Of particular relevance to thepresent work are three genes, hpxO, hpxT, and hpxQ, encodingurate oxidase (HpxO) (30), HIU hydrolase, and OHCU decarbox-ylase, respectively. To our knowledge, this is the first report on thethree-step enzymatic conversion of uric acid to S-(�)-allantoin inbacteria based on gene cluster analysis. This led to the functionalexpression of hpxO, hpxT, and hpxQ in E. coli and the crystalstructure elucidation of KpHIUH (hpxT) (11) and KpOHCU de-carboxylase (hpxQ) (10). Even though the substrates of KpHIUHand KpOHCU are unstable, the catalytic activities of these en-zymes were confirmed by the generation of substrates in situ. Ra-

mazzina et al. (32) identified HIU hydrolase (MuraH) and OHCUdecarboxylase (MuraD) from mouse by phylogenetic comparisonof genomes and sequence analysis. This led to the cloning of thesegenes in E. coli and the establishment, by 13C NMR, of the conver-sion of HIU to S-(�)-allantoin via OHCU (32).

A similar gene homology and cluster-based analysis of the25.2-kb genomic DNA segment of CBB1 revealed the presence oftmuH and tmuD upstream of tmuM (Fig. 2). The gene tmuH had50 and 44% identities, respectively, to putative HIU hydrolasesfrom Rhizobium leguminosarum bv. Trifolii and Sinorhizobiummeliloti. Gene tmuD had 39 and 37% identity, respectively, withputative OHCU decarboxylases from Starkeya novella andHaloferax volcanii (Fig. 2). Comparison of the organization oftmuH and tmuD (Fig. 2) to the corresponding (characterized)genes hpxO, hpxT, and hpxQ of the uric acid pathway in the hpxgene cluster from K. pneumoniae (8) and the 22.872-kb purineutilization gene cluster from K. oxytoca (31) reveals several inter-esting features. Although hpxO from hpx gene cluster of Klebsiellapneumoniae (8) had 38% identity to tmuM, the hpxT and hpxQgenes from the same cluster had 29 and 19% sequence identities totmuH and tmuD, respectively. A BLASTP analysis of tmuH and

FIG 4 (I) Homology models for TmuM and HpxO. The best homology models for TmuM (purple) and HpxO (green) are superimposed with the modelingtemplate PhzS (PDB ID 3C96, wheat). Blue and red spheres indicate the N- and C-terminal regions of the protein that were either observed in the X-ray structureor could be modeled reliably. FAD (orange sticks) is modeled into place based on PHBH (1PBE) with the isoalloxazine ring in the “in” position, as would beexpected in the presence of the ligand. TMU (purple sticks) is modeled in the ligand-binding pocket based on the ligand position in PHBH (1PBE). (II) Active-sitecavities for TmuM and HpxO homology models. Residues lining the modeled active site for TmuM (A) and HpxO (B) are shown as sticks colored by hydropathyvalues based on the Kyte-Doolittle scale (24), with the most hydrophobic in red and the least hydrophobic in blue. TmuM has a bigger, more hydrophobicactive-site cavity (trimethyluric acid modeled is in purple) than HpxO (uric acid modeled is in green).

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hpxT revealed that both of the proteins belong to transthyretin-like superfamily (11). The organization of tmuM, tmuH, andtmuD in the caffeine gene cluster of CBB1 (Fig. 2) is similar to thatof the corresponding hpxO, hpxT, and hpxQ genes in the hpx geneclusters in Klebsiella in terms of orientation, the direction of tran-scription, and gene location. The genes tmuH and tmuD are nextto each other, with the same direction of transcription and sepa-rated from the oppositely transcribing gene tmuM by a single1,188-nt ORF (Fig. 2). This is the same as the organization ofcorresponding the hpxT and hpxQ genes and their separationfrom the oppositely transcribing hpxO by hpxP (8, 31). Further, inthe K. oxytoca gene cluster, hpxB, located �8 kb downstream ofhpxO, codes for an allantoinase (31). In the caffeine gene cluster(Fig. 2), orf1 with homology to allantoinases, along with othergenes (orf2 and orf3, Fig. 2), are 14 kb downstream of tmuM. It istempting to propose that these genes are involved in furtherdownstream metabolism of S-(�)-TMA, but this needs furthersubstantiation. The larger separation of orf1 from tmuM can beattributed to the recruitment of 3.765-kb cdhABC genes, codingfor heterotrimetic caffeine dehydrogenase (Cdh) (Fig. 2). Basedon this, tmuH is proposed to catalyze the N1-C6 hydrolysis andring opening reaction of TM-HIU to TM-OHCU (Fig. 3), andtmuD is proposed to catalyze the stereoselective decarboxylationof TM-OHCU to S-(�)-TMA (Fig. 3). Another strong piece ofevidence for the formation of S-(�)-TMA is that only S-(�)-allantoin seems to be formed in cells (39) capable of utilizing uricacid. These organisms rely on the enzymatic conversion of HIU toS-(�)-allantoin via OHCU (32). Our contention is that the puta-tive TM-HIU hydrolase and a putative TM-OHCU decarboxylasein CBB1 are specific for methyl substituents in the respective sub-strates (Fig. 3). This is based on the preference of both Cdh andTmuM to trimethylated substrates with no activity on xanthineand uric acid. Hence, in the proposed conversion of TMU toS-(�)-TMA, all enzymes are designated with preference for(tri)methylated substrates (Fig. 3).

Further metabolism of S-(�)-TMA in CBB1 is under investi-gation. Organisms capable of utilizing uric acid as a source ofcarbon and nitrogen possess allantoinase and allantoin recemase(17, 33). Recently, Guzmán et al. (17) reported the presence ofanother gene cluster in K. pneumoniae, which harbors genes re-sponsible for further transformation of allantoin. In particular,the two genes hpxA and hpxB, which encode allantoin recemaseand allantoinase, respectively, were identified. This led to the crys-tal structure determination of heterologously expressed hpxAgene product KpHpxA (12). Thus, it is well established that (R)-allantoin is converted by allantoin recemase to (S)-enantiomer(12), which then gets further transformed to allantoate by allan-toinase (33, 39, 40). Very likely, in CBB1, (R)-TMA is converted to(S)-TMA and then to trimethylallantoate. This compound couldpotentially be transformed to glyoxylic acid, dimethylurea, andmonomethylurea (27), as in the uric acid pathway (40). Our initialanalysis of DNA sequence downstream of Cdh genes has identifiedorf1 with homology to allantoinase. However, we are unable tolocate a trimethylallantoin racemase. Any clarity on the biochem-istry of formation and metabolism of S-(�)-TMA (Fig. 3) willcome from functional expression of tmuH and tmuD, along withidentification of the genes responsible for its transformation.

In summary, a new pathway has been proposed for transfor-mation of caffeine via C-8 oxidation to S-(�)-TMA. The first twosteps, i.e., conversion of caffeine to TMU by Cdh and TMU to

TM-HIU by TmuM, are based on detailed enzymology and geneanalysis. Subsequent transformation of TM-HIU to S-(�)-TMAis based on gene cluster analysis and comparison to uric acidtransformation, which is based on gene cluster analysis as well. Toour knowledge, this is the first report of biochemical and geneticanalysis of caffeine transformation via C-8 oxidation.

ACKNOWLEDGMENTS

This research was supported by the University of Iowa Research Funds.We thank Amnon Kohen and his graduate student Eric Koehn in the

Department of Chemistry, University of Iowa, for providing assistance in18O-incorporation experiment. We also thank Santhana Velupillai of theNMR Central Research Facility and High Resolution Mass SpectrometryFacility of the University of Iowa for assistance with instrumentation.

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