Functional Characterization of a Novel Member of theAmidohydrolase 2 Protein Family, 2-Hydroxy-1-Naphthoic AcidNonoxidative Decarboxylase from Burkholderia sp. Strain BC1

Piyali Pal Chowdhury, Soumik Basu, Arindam Dutta, Tapan K. Dutta

Department of Microbiology, Bose Institute, Kolkata, India

ABSTRACT

The gene encoding a nonoxidative decarboxylase capable of catalyzing the transformation of 2-hydroxy-1-naphthoic acid(2H1NA) to 2-naphthol was identified, recombinantly expressed, and purified to homogeneity. The putative gene sequence of thedecarboxylase (hndA) encodes a 316-amino-acid protein (HndA) with a predicted molecular mass of 34 kDa. HndA exhibitedhigh identity with uncharacterized amidohydrolase 2 proteins of various Burkholderia species, whereas it showed a modest 27%identity with �-resorcylate decarboxylase, a well-characterized nonoxidative decarboxylase belonging to the amidohydrolasesuperfamily. Biochemically characterized HndA demonstrated strict substrate specificity toward 2H1NA, whereas inhibitionstudies with HndA indicated the presence of zinc as the transition metal center, as confirmed by atomic absorption spectroscopy.A three-dimensional structural model of HndA, followed by docking analysis, identified the conserved metal-coordinating andsubstrate-binding residues, while their importance in catalysis was validated by site-directed mutagenesis.

IMPORTANCE

Microbial nonoxidative decarboxylases play a crucial role in the metabolism of a large array of carboxy aromatic chemicals re-leased into the environment from a variety of natural and anthropogenic sources. Among these, hydroxynaphthoic acids areusually encountered as pathway intermediates in the bacterial degradation of polycyclic aromatic hydrocarbons. The presentstudy reveals biochemical and molecular characterization of a 2-hydroxy-1-naphthoic acid nonoxidative decarboxylase involvedin an alternative metabolic pathway which can be classified as a member of the small repertoire of nonoxidative decarboxylasesbelonging to the amidohydrolase 2 family of proteins. The strict substrate specificity and sequence uniqueness make it a novelmember of the metallo-dependent hydrolase superfamily.

Decarboxylase is one of the most important classes of enzymesinvolved in a large variety of catabolic and anabolic pathways.

The majority of the decarboxylases utilize an organic cofactor or atransition metal coupled with dioxygen to activate their substratesleading to the removal of carbon dioxide (1). However, there is asmall group of transition metal-dependent decarboxylases thatcarry out decarboxylation of various aromatic acids in a nonoxi-dative manner. These nonoxidative decarboxylases act on variouslignin-derived compounds, such as 4-hydroxybenzoic acid (2),(carboxy)vanillic acid (3, 4), protocatechuic acid (5), ferulic acid(6), p-coumaric acid (7), and estrogenic phthalate (8). Likewise,oxygen-independent decarboxylases are also involved in the 2-ni-trobenzoic acid degradation pathway (9, 10), the tryptophan cat-abolic pathway (11), and the thymidine salvage pathway (12).

Nonoxidative decarboxylases, in general, can broadly be clas-sified into two major groups depending on their oxygen sensitiv-ity. Oxygen-sensitive decarboxylases, viz., 4-hydroxybenzoate de-carboxylase (2), 3,4-dihydroxybenzoate decarboxylase (5), andindole-3-carboxylate decarboxylase (13), catalyze reversible reac-tions, including both carboxylation and decarboxylation. On theother hand, oxygen-insensitive decarboxylases, such as 2,3-dihy-droxybenzoate decarboxylase (14), 5-carboxyvanillate decarbox-ylase (4), and 4,5-dihydroxyphthalate decarboxylase (8), havebeen reported to catalyze the decarboxylation reaction only. How-ever, there are a few nonoxidative oxygen-insensitive decarboxy-lases, viz., �-resorcylate decarboxylase (15), vanillate/4-hydroxy-benzoate decarboxylase (16), and salicylate decarboxylase (17),that have been documented to catalyze reversible reactions.

Hydroxynaphthoates, such as 1-hydroxy-2-naphthoic acid(1H2NA), 2-hydroxy-1-naphthoic acid (2H1NA), and 3-hy-droxy-2-naphthoic acid (3H2NA), are normally encounteredduring the bacterial degradation of polycyclic aromatic hydrocar-bons, viz., phenanthrene, anthracene, and pyrene, and are metab-olized either through ring cleavage (18–22) or by oxidative decar-boxylation (23). In addition, decarboxylation of 1H2NA to1-naphthol has been proposed, based on the identification of thelater compound during degradation of phenanthrene in a few bac-teria (24–26). Similarly, decarboxylation of phenanthrene-4,5-di-carboxylic acid to phenanthere-4-carboxylic acid has also beenreported in the pathway of pyrene degradation (19). However,there is no documented report of any nonoxidatively decarboxy-lated product produced during the metabolism of 2H1NA or

Received 19 March 2016 Accepted 1 April 2016

Accepted manuscript posted online 11 April 2016

Citation Pal Chowdhury P, Basu S, Dutta A, Dutta TK. 2016. Functionalcharacterization of a novel member of the amidohydrolase 2 protein family,2-hydroxy-1-naphthoic acid nonoxidative decarboxylase from Burkholderia sp.strain BC1. J Bacteriol 198:1755–1763. doi:10.1128/JB.00250-16.

Editor: I. B. Zhulin, University of Tennessee

Address correspondence to Tapan K. Dutta, tapan@jcbose.ac.in.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/JB.00250-16.

Copyright © 2016, American Society for Microbiology. All Rights Reserved.

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3H2NA. Despite published reports of purification and character-ization of several nonoxidative hydroxybenzoate decarboxylases(5, 14–16), there are no examples of any enzyme catalyzing non-oxidative decarboxylation of any of the hydroxynaphthoic acidisomers.

Previously, we reported a nonconventional degradation path-way of 2H1NA in Burkholderia sp. strain BC1 describing 2-naph-thol, gentisaldehyde, and gentisic acid as pathway intermediates(27). Moreover, the presence of a strictly inducible nonoxidativedecarboxylase was also observed in the cell extract of 2H1NA-grown culture catalyzing the enzymatic transformation of 2H1NAto 2-naphthol. In the present study, we describe a proteomic ap-proach-based gene cloning and functional characterization ofnonoxidative 2H1NA decarboxylase from Burkholderia sp. strainBC1. In addition, the roles of specific amino acid residues respon-sible for substrate binding and enzyme catalysis have been eluci-dated.

MATERIALS AND METHODSBacterial strains, plasmids, and culture conditions. The strains and plas-mids used in this study are listed in Table S1 in the supplemental material.Recombinant constructs in Escherichia coli [XL1-Blue and BL21(DE3)]were routinely grown and maintained in Luria-Bertani (LB) broth (perliter) containing 10 g of Bacto tryptone, 5 g of yeast extract, and 10 g ofNaCl (pH 7.2) or on LB solid medium (1.8% [wt/vol] agar) at 37°C.Where appropriate, ampicillin (100 �g/ml), kanamycin (50 �g/ml),chloramphenicol (12.5 �g/ml), IPTG (isopropyl-�-D-thiogalactopyrano-side; 0.1 to 1 mM), or X-Gal (5-bromo-4-chloro-3-indolyl-�-D-galacto-pyranoside; 20 �g/ml) was added. For expression cloning, pET28a (No-vagen, Madison, WI) served as the expression vector.

Partial purification and gene identification of 2H1NA decarboxyl-ase. Native 2H1NA decarboxylase was purified from crude cell extract ofBC1 cells grown for 16 h at 28°C in 4 liters of mineral salt medium (MSM)(20) containing 0.5 g/liter 2H1NA. Crude cell extract was prepared asdescribed previously (27) and was then fractionated by sequential proteinprecipitation using ammonium sulfate. The 30 to 50% ammonium sulfatesaturated fraction was centrifuged at 12,000 � g for 30 min, and theresulting pellet was dissolved in buffer A (50 mM K2HPO4-KH2PO4 buf-fer [pH 7.0]) and dialyzed against buffer B [50 mM K2HPO4-KH2PO4

buffer (pH 7.0) containing 0.8 M (NH4)2SO4]. The dialyzed fraction wasthen loaded onto a column (2.5 by 10 cm), packed with phenyl-Sepharose6 Fast Flow, preequilibrated with buffer B. The column was washed with 5column volumes of buffer B, and then the adsorbed proteins were elutedin steps using 10 column volumes each of buffer A containing differentconcentrations of (NH4)2SO4 (0.8 to 0.05 M). Finally, the column waswashed with two column volumes of buffer A. All purification steps werecarried out at 4°C or on ice under aerobic conditions. Fractions exhibiting2H1NA decarboxylase activity were combined and dialyzed against bufferC (50 mM K2HPO4-KH2PO4 buffer [pH 7.0], 10% glycerol), concen-trated by ultrafiltration (Millipore, Massachusetts), and stored at �80°Cuntil further use. The purity of the protein fractions obtained after am-monium sulfate precipitation and hydrophobic interaction chromatogra-phy was evaluated by 12.5% SDS-PAGE analysis, followed by Coomassieblue staining in the presence of prestained protein molecular mass mark-ers (Puregene; Genetix, India) by standard techniques. Protein quantifi-cation was performed according to the method of Bradford (28).

For the identification of the decarboxylase, tryptic digestion of theprotein and subsequent extraction of peptides from SDS-PAGE gel ma-trices were carried out according to the methods described by Shevchenkoet al. (29), followed by matrix-assisted laser desorption ionization–time offlight (MALDI-TOF) mass spectrometry (MS) and tandem MS (MS/MS)analyses using an AutoFlex II (Bruker Daltonics, Germany) MALDI-tan-dem TOF (TOF/TOF) mass spectrometer equipped with a pulsed N2 laser(� � 337 nm, 50 Hz). The mass spectra were analyzed with Flex Analysis

Software (version 2.4; Bruker; Daltonics). From the MS/MS data, partialamino acid sequences of the peptides were determined using PEAKS stu-dio 7 (Bioinformatics Solutions, Inc., Ontario, Canada), and the peptidesequences were subjected to blastp (30) analysis for identification. Prim-ers (HNDA_F [5=-TGCTGTCGCTGACGGC-3=] and HNDA_R [5=-TTGCTGAGCAGCACGAC-3=]) were designed on the basis of conserved re-gions exhibited in multiple-sequence alignment, generated by Clustalxv1.81 (31) using amidohydrolase gene sequences of various Burkholderiaspp. (see Table S2 in the supplemental material). Using the primers, PCRwas carried out in a 50-�l reaction volume using Phusion DNA polymer-ase (Thermo Fischer) in an MJ Mini Gradient Thermal Cycler (Bio-RadLaboratories, Inc., Hercules, CA) with the following thermocycling con-ditions: 30 s at 98°C, followed by 30 cycles of 30 s at 98°C, 30 s at 55°C, and10 s at 72°C. A final extension was performed at 72°C for 7 min. Theresulting PCR product was sequenced as reported previously (27).

Enzyme assay. Nonoxidative decarboxylase activity was qualitativelydetermined by UV-visible light spectral analysis as described previously(27) while the activity was quantitatively determined based on the forma-tion of 2-naphthol, analyzed by high-pressure liquid chromatography(HPLC) using a methanol-water (50:50 vol/vol) isocratic solvent systemwith a flow rate of 1 ml/min (27). A standard curve of 2-naphthol createdby HPLC under identical analytical conditions was used for quantitativeestimation. One unit of enzyme activity is defined as the amount of en-zyme required for the production of 1 �mol of product per min. Thespecific activity is expressed as units per milligram of protein.

For measurement of carboxylase activity, a standard reaction mixturecontaining recombinant protein (100 �g), 2-naphthol (20 mM), andNaHCO3-NH4HCO3 (1.0 or 2.5 M) in a final volume of 1 ml of buffer Awas prepared, and the reaction mixture was incubated for 60 min at 35°C.To analyze the reaction product, HPLC analysis was performed as de-scribed above.

Fosmid library construction, screening, and sequence analysis ofamidohydrolase gene. A genomic library of strain BC1 was prepared in E.coli using a CopyControl HTP fosmid library production kit (Epicentre,Madison, WI) according to the manufacturer’s protocol. The resultingfosmid library was screened by PCR using HNDA_F and HNDA_R prim-ers for clones harboring 2H1NA decarboxylase gene as described above.Fosmid DNA was isolated from the PCR-positive fosmid clones using theFosmidMAX DNA purification kit (Epicentre) and digested with EcoRI,HindIII, and SacII enzymes, and the DNA fragments (1 to 8 kb) weresubcloned in pBluescript SK(�) vector. The colonies were rescreened byPCR using the same primer pair. The recombinant plasmids from thescreened colonies were individually isolated and sequenced using M13universal sequencing primers. The sequences were analyzed by BLASTanalysis (version 2.2.12; National Center for Biotechnology Information[NCBI]), and the gaps between genes were bridged by using a conven-tional primer walking method.

Cloning, expression, and purification of recombinant proteins. Theprimers Ex_HNDA_F (5=-CCGGAATTCATGACCGACCATCACCGTATC-3=) and Ex_HNDA_R (5=-CCCAAGCTTTTGTTGTGTTGTTGCGTCAG-3=) were designed (the EcoRI and HindIII restriction endonu-clease recognition sites, respectively, are underlined) to amplify the com-plete 2H1NA decarboxylase gene (hndA) from genomic DNA of strainBC1. The amplified PCR product was digested with EcoRI and HindIIIand ligated into similarly digested pET28a expression vector to formpET28a:HndA. The resulting plasmid was transformed into E. coliBL21(DE3) and plated on LB agar plates containing kanamycin. For thepreparation of single amino acid substitution in HndA, pET28:HndA wassubjected to whole-plasmid PCR with mutagenic primers (see Table S3 inthe supplemental material) under following thermocycling conditions: 3min at 98°C, followed by 16 cycles of 30 s at 98°C, 30 s at 55°C, and 3 minat 72°C, with a final extension at 72°C for 10 min using Phusion high-fidelity DNA polymerase (Thermo Fisher Scientific). After digestion withDpnI for 2 h, the PCR product was transformed into electrocompetent E.coli Top10 and plated on LB agar plates containing kanamycin. Plasmids

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isolated from random clones were subjected to sequencing analysis toconfirm the mutation at a specified location.

For recombinant enzyme expression and purification, E. coli BL21(DE3)cells harboring pET28a:HndA or its mutant derivatives (see Table S1 inthe supplemental material) were grown in 500 ml of LB medium at 37°Cwith kanamycin to achieve an optical density at 600 nm of 0.5, followed bythe addition of 0.5 mM IPTG (final concentration), and were grownfurther at 28°C for 3 h. The cultures were harvested by centrifugation(8,000 � g) and lysed in 10 ml of lysis buffer (50 mM NaHPO4, 300 mMNaCl, 10 mM imidazole, and 10% glycerol) using a precooled Frenchpress (constant cell disruption system, One Shot model; United King-dom) at 18,000 lb/in2 for one cycle. After removal of the cell debris, thesupernatant containing the His6-tagged wild-type or mutant recombi-nant protein was purified by nickel-nitrilotriacetic acid (Ni2-NTA)–aga-rose affinity chromatography using the purification buffers (wash buffer[50 mM NaHPO4, 300 mM NaCl, 40 mM imidazole, 10% glycerol] andelution buffer [50 mM NaHPO4, 300 mM NaCl, 250 mM imidazole, 10%glycerol]) according to the manufacturer’s instructions (Qiagen). The pu-rified protein fractions were pooled, dialyzed against buffer C, and ana-lyzed by 12.5% SDS-PAGE. The dialyzed protein preparation was used inall biochemical studies.

Phylogenetic analysis. Amino acid sequences of various proteins be-longing to the amidohydrolase 1 and amidohydrolase 2 families wereretrieved from the NCBI (see Table S4 in the supplemental material) andaligned, and a phylogenetic tree was constructed using the neighbor-join-ing algorithm, as implemented in ClustalX v1.81 (31). The tree was visu-alized using the program Tree Explorer v2.12, a stand-alone version of thesame program implemented in MEGA 5 (32).

Gel filtration. Native molecular mass of the decarboxylase was esti-mated by gel filtration chromatography using a P4000 PolySep GFC col-umn (30 by 0.7 cm; Phenomenax, Torrance, CA), equilibrated with bufferA containing 200 mM NaCl. The flow rate used was 0.5 ml/min. Yeastalcohol dehydrogenase (150 kDa), conalbumin (75 kDa), ovalbumin (44kDa), carbonic anhydrase (29 kDa), and RNase A (13.7 kDa) were used asstandard proteins. Blue dextran (2,000 kDa) was used to calculate the voidvolume.

Biochemical studies. All the enzyme kinetic analyses were done at35°C and pH 7.5. For decarboxylation, the kinetic data were assessed using1.5 �g of wild-type or mutant decarboxylase against 2H1NA over theconcentration range of 0.05 to 0.5 mM. The maximum velocity (Vmax)and the Michaelis constant (Km) were determined from Lineweaver-Burkdouble-reciprocal plots using GraphPad Prism program (version 5.00 forWindows). The optimum temperature of the recombinant protein wasdetermined over the range of 10 to 70°C, and the pH profile was deter-mined at the optimal temperature determined as described above understandard conditions over the pH range of 4.0 to 9.5 using the followingbuffer systems (50 mM): citrate buffer (pH 4 to 6), sodium phosphatebuffer (pH 6 to 8), and glycine-NaOH (pH 8.0 to 9.5). The effect oftemperature on enzyme stability was determined by preincubating theenzyme at different temperatures (10 to 60°C) for 30 min and measuringthe remaining activity under standard conditions. To study the effect ofvarious metal ions and inhibitors on enzyme activity, purified 2H1NAdecarboxylase preincubated with respective metal ions or inhibitors (1 or5 mM) for 10 min at 4°C was used as an enzyme preparation. However, formetal chelators, viz., EDTA, 1,10-phenanthroline, 2,2=-bipyridyl, and

8-hydroxyquinoline-5-sulfonic acid (8-HQSA), enzyme was preincu-bated for 16 h.

For metal analysis, purified HndA (3 mg) was hydrolyzed by 65%ultrapure concentrated nitric acid (2 ml; Suprapure; Merck, Darmstadt,Germany) at 110°C for 1 h. The sample was diluted 10-fold by deionizeddouble-distilled water, and the metal content was determined by using anatomic absorption spectrometer (iCE 3000 Series; Thermo Fischer Scien-tific).

Homology modeling and docking analyses. A three-dimensional(3D) model of HndA was constructed using 2-amino-3-carboxymu-conate-6-semialdehyde decarboxylase (ACMSD) from Pseudomonas fluo-rescens (PDB accession no. 2HBV) (33) as a template employing Modeler9v7 (34). The models were checked using Prochek, Verify3D, and VADAR(35–37). The NCBI PubChem database (http://pubchem.ncbi.nlm.nih.gov/) was used to obtain coordinates of the ligand 2H1NA. Preparation ofprotein and substrate files (pdbqt files) was performed using AutoDock-Tools-1.5.6 using default parameters (38). The grid box (with the dimen-sions 50 by 50 by 50 grid points) was generated using the AutoGrid4program, keeping the metal ion coordinates (43.43 by �0.323 by 16.82) atthe center. AutoDock4 was used to perform docking using genetic algo-rithm. Docked poses were analyzed using AutoDockTools-1.5.6 to get thebest binding pose of 2H1NA with the lowest binding energy. The bindingresidues were identified and the schematic diagram of protein-ligand in-teraction was generated using LigPlot suite (version 1.4.5) (39).

Nucleotide sequence accession number. The nucleotide sequence re-ported here has been deposited in the DDBJ/EMBL/GenBank databaseunder accession number KU254672.

RESULTSPartial purification and gene identification of 2H1NA decarbox-ylase. 2H1NA decarboxylase activity was previously reported tobe strictly inducible in the presence of 2H1NA and a differentiallyexpressing 32-kDa protein band was observed only in the cellextract of 2H1NA-grown cells compared to that of 2-naphthol-grown cells (27). For detailed characterization, 2H1NA decarbox-ylase was partially purified from crude cell extract of strain BC1grown on 2H1NA using differential protein precipitation stepsand hydrophobic interaction chromatography (Table 1). The pu-rified enzyme preparation was found to be stable during the puri-fication steps, carried out under aerobic conditions, indicating theoxygen-insensitive nature of the decarboxylase. The decarboxy-lase-active fractions from a phenyl-Sepharose column repre-sented 19-fold purification (specific activity, 3.8 U mg�1) with ayield of 16.8% and showed the presence of an 32-kDa band inSDS-PAGE, supporting our earlier observation (see Fig. S1 in thesupplemental material). To confirm its identity, the 32-kDaprotein was subjected to MALDI-TOF MS/MS analysis, where thegenerated peptide fragments showed strong sequence similarity tothe uncharacterized amidohydrolase 2 proteins of various Burk-holderia spp. in blastp analyses (see Table S5 in the supplementalmaterial). Subsequently, a 220-bp PCR product was amplified(data not shown) from the genomic DNA of strain BC1 using the

TABLE 1 Purification summary of 2H1NA decarboxylase from Burkholderia sp. BC1

Step Total protein (mg) Total activity (U)Sp acta

(U/mg) Purification (fold) Yield (%)

Cell extract 180 36.3 0.2 1 100Ammonium sulfate fractionation (30 to 50% saturation) 32 25.9 0.8 4 71.3Phenyl-Sepharose 6 (Fast Flow) chromatography 1.6 6.1 3.8 19 16.8a Sp act, specific activity.

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primers HNDA_F and HNDA_R, which on sequence analysisconfirmed the results as stated above.

Cloning and sequencing of the 2H1NA decarboxylase gene.Screening of a genomic fosmid library of strain BC1 by PCR led tothe identification of a subclone, which upon IPTG induction dis-played 2H1NA decarboxylase activity, determined in the cell-freeenzyme preparation. Complete sequence analysis of the subcloneharboring a 4.2-kb EcoRI fragment revealed the presence of anamidohydrolase gene, designated hndA for hydroxynaphthoatedecarboxylase. hndA, consisting of 951 nucleotides, encoded apolypeptide of 316 amino acids with a theoretical molecular massof 34 kDa and pI of 5.59. Moreover, the CDD (Conserved DomainDatabase) and COG (Clusters of Orthologous Groups) analyses ofHndA placed it in the amidohydrolase superfamily of the triose-phosphate isomerase (TIM)-barrel fold protein (COG2159),which includes several nonoxidative decarboxylases, including5-carboxyvanillate decarboxylase (5-CVD), 2,3-dihydroxybenzo-ate decarboxylase, and �-resorcylate decarboxylase (�-RSD) (4,14, 15). HndA showed 71 to 97% identity with the biochemicallyuncharacterized metal-dependent hydrolase proteins of severalBurkholderia species belonging to the amidohydrolase superfam-ily, listed in the NCBI database. However, among the biochemi-cally well-characterized nonoxidative decarboxylases, HndAshowed a modest identity of 27 and 24% with the �-resorcylatedecarboxylase (�-RSD) of Rhizobium sp. strain MTP-10005 (15)and ACMSD of Pseudomonas fluorescens (10), respectively. Othergenes in the 4.2-kb gene cluster include orf1, orf2, and dbpA, wherethe gene products showed 99 to 100% identity with a noncharac-terized phenol degradation protein, a LysR-type regulator, and anATP-dependent RNA helicase protein of Burkholderia multiv-orans, respectively. The genetic assembly of the 4.2-kb cluster isshown in Fig. 1A.

Phylogenetic analysis of HndA. A phylogenetic tree (Fig. 1B)constructed using multiple-sequence alignment of various pro-teins belonging to the amidohydrolase 1 and amidohydrolase 2family positioned HndA within the amidohydrolase 2 family.With the exception of 4-oxalomesaconate hydratase (OMAH)from Sphingomonas paucimobilis SYK-6 (40), the other represen-tative members belonging to this family are nonoxidative decar-boxylases, viz., isoorotate decarboxylase (IDCase) from Neuro-spora crassa (12) and 5-carboxyvanillate decarboxylase (5-CVD)from Sphingomonas paucimobilis SYK-6 (4), apart from �-RSDfrom Rhizobium sp. strain MTP-10005 (15) and ACMSD fromPseudomonas fluorescens (10). In addition, a number of uncharac-terized metal-dependent hydrolases from Burkholderia also ap-peared to belong to this enzyme family (Fig. 1B).

Despite an overall low sequence homology among the bio-chemically characterized members of amidohydrolase 2 familyproteins, the sequence alignment did display a strong residue con-servation pattern for amino acids (His10, His12, His156, andAsp269 in HndA) that are responsible for the binding of themetal cofactor, which is crucial for enzyme catalysis (Fig. 1C).Apart from the results reported above, the alignment showedanother conserved histidine residue (His204 in HndA) whichwas earlier reported to play a crucial role in enzyme catalysis forboth ACMSD and �-RSD (33, 41).

Overexpression and purification of recombinant HndA. Therecombinant 2H1NA decarboxylase was successfully overex-pressed in E. coli BL21(DE3) with a 0.5 mM IPTG concentrationand was purified by Ni2-NTA chromatography (Fig. 2A). The

purified recombinant enzyme migrated as a single band in theSDS-PAGE gel with an apparent subunit molecular mass of 38kDa, while the molecular mass of the native recombinant enzymeon gel filtration was found to be 38.1 � 0.5 kDa, suggesting themonomeric nature of the enzyme. Purified recombinant HndAcatalyzed the decarboxylation of 2H1NA to 2-naphthol, as re-vealed by both spectral and HPLC analyses (Fig. 2B and C), with aspecific activity of 9.0 U/mg of protein. Figure 2D shows the time-dependent transformation of 2H1NA to 2-naphthol by purifiedHndA over a period of 10 min.

Biochemical properties of recombinant HndA. It was ob-served that HndA did not favor carboxylation of 2-naphthol un-der the conditions tested, and thus it appears that the enzymecatalyzes an irreversible reaction (decarboxylation). Again, HndAshowed strict substrate specificity toward 2H1NA since its otherstructural isomers, 1H2NA and 3H2NA, could not be trans-formed. Also, it failed to decarboxylate mono- and dihydroxyben-zoic acids, viz., 2-hydroxybenzoic acid (salicylic acid), 3-hydroxy-benzoic acid, 4-hydroxybenzoic acid, 2,3-dihydroxybenzoic acid,2,4-dihydroxybenzoic acid, 2,5-dihydroxybenzoic acid (gentisicacid), and 2,6-dihydroxybenzoic acid (�-resorcylic acid). Simi-larly, HndA failed to transform phthalic acid, 1-naphthoic acid,and 2-naphthoic acid.

For 2H1NA decarboxylase, the values of Km and Vmax weredetermined to be 0.17 mM and 0.02 �mol/min, respectively. Thekcat/Km value for 2H1NA was 47.05 mM�1 s�1. The optimum pHand temperature of the protein were found to be 7.5 and 35°C,respectively (see Fig. S2A and B in the supplemental material). Theenzyme was found to be stable up to 45°C and retained 58% ofinitial activity when incubated at 50°C for 30 min. However, theenzyme completely lost its activity when incubated above 60°C(see Fig. S2C in the supplemental material).

The effect of various metal ions as well as inhibitors on enzymeactivity is shown in Table S6 in the supplemental material. Nosignificant change in enzyme activity was observed with majorityof the metal ions, inhibitors, and metal ion chelators, individuallyincubated for 10 min. However, activity of the enzyme was inhib-ited by AgNO3 and HgCl2, as suggested for �-RSD (15, 42). Nev-ertheless, a modest inhibition in HndA activity was observed whenthe enzyme was incubated for 16 h individually with metal chela-tors, including 8-hydroxy-quinoline-5-sulfonic acid, a zinc metal-specific inhibitor. This observation suggests the possible presenceof a deeply embedded metal ion, inadequately accessible by metalchelators. Diethylpyrocarbonate, a histidine residue modifier, alsoshowed a reasonable decrease in enzymatic activity, suggesting thepresence of active-site histidine residues in HndA, as describedearlier (27). To identify the metal center in the active site of HndA,atomic absorption spectroscopy analysis was performed that re-vealed the presence of zinc at 0.95 � 0.1 mol per mol of protein.This result corroborated well with results determined for othernonoxidative decarboxylases belonging to the amidohydrolase su-perfamily that possess zinc as the transition metal center (33, 41).

Structural modeling and functional analysis of HndA mu-tants. A 3D structural model of HndA showed the presence of a(�/�)8 barrel fold with eight parallel � strands flanked by eight �helices on the outer face. The structural proximity of the con-served His10, His12, His156, and Asp269 residues in the 3D modelof HndA was fully compatible with their putative role in forminga metal ion binding motif (Fig. 3A). In order to confirm the func-tional roles of these amino acid residues, site-directed mutants

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HndAH10Q, HndAH156Q, and HndAD269V were constructed andexpressed as soluble proteins (see Fig. S3 in the supplemental ma-terial), and their specific activities against 2H1NA were deter-mined. HndAH10Q and HndAD269V showed no activity against2H1NA, whereas HndAH156Q retained only 7.23% (0.65 U/mg)of the wild-type decarboxylase activity. To analyze the role ofthe conserved histidine residue His204, the mutant proteinHndAH204Q was studied and showed a complete loss of enzymatic

activity, suggesting a critical role of this residue in HndA-medi-ated catalysis.

In order to determine other important amino acid residuesresponsible for substrate binding, docking analysis was performedusing 2H1NA as a ligand. Docking analysis revealed the role of twoimportant amino acid residues, Arg33 and Tyr272, which werefound to interact with the carboxyl functional group of 2H1NA byhydrogen bonding (Fig. 3B). In addition to the carboxyl group,

FIG 1 (A) Gene organization of 4.2-kb EcoRI fragment showing gene designations. orf1, partial phenol degradation protein; hndA, 2H1NA decarboxylase; orf2,LysR-type-like regulator; dbpA, ATP-dependent RNA helicase. (B) Phylogenetic tree based on protein sequences from amidohydrolase 1 and amidohydrolase 2family of proteins. Numbers at the nodes indicate the levels of bootstrap support based on neighbor-joining analysis of 100 resampled data sets. Bootstrap valuesbelow 50% are not shown. The scale bar represents 0.1 substitutions per nucleotide position. GenBank or PDB accession numbers are indicated withinparentheses. Amidohydrolase 3 from Burkholderia sp. strain lig30 (KDB07616) was used as an outgroup. (C) Multiple-sequence alignment of protein sequencesof the representative members of amidohydrolase 2 protein family. Metal coordinating residues are shaded.

FIG 2 (A) SDS-PAGE analysis of overexpressed recombinant HndA protein. Lane 1, crude extract of E. coli BL21(DE3) carrying empty pET28a vector; lane 2,crude extracts of induced E. coli BL21(DE3) carrying pET28a:HndA; lane 3, purified recombinant HndA protein; lane M, molecular mass marker (Puregene). (B)Spectral changes during transformation of 2HINA by purified recombinant HndA protein. The sample and reference cuvettes contained 50 mM potassiumphosphate buffer (pH 7.0) in a 1-ml volume. The sample cuvette also contained 220 nmol of 2H1NA. Spectra were recorded every 1 min after the addition of 10�g of protein to both cuvettes. The up and down arrows indicate increasing and decreasing absorbances, respectively. (C) HPLC chromatogram showingtransformation of 2H1NA to 2-naphthol by purified HndA in a reaction mixture (final volume, 1 ml) containing 0.5 mM 2H1NA and 5 �g of protein in bufferA incubated for 10 min at 35°C. UV-visible light spectra of 2H1NA and 2-naphthol are shown in insets. (D) Time-dependent transformation of 2H1NA to2-naphthol by purified HndA. The concentrations of 2H1NA (Œ) and 2-naphthol (�) were determined by HPLC from the reaction mixtures (as described inpanel C) during enzymatic transformation over 10 min.

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Tyr272 was also found to interact with the hydroxyl group of2H1NA (Fig. 3B). To further assess their role in substrate binding,HndAR33L and HndAY272F mutant proteins were generated (seeFig. S3 in the supplemental material), and subsequently theiractivities were tested against 2H1NA. The mutant enzymes,HndAR33L and HndAY272F, showed specific activities of 4.1 and 2.8U/mg of protein, respectively. Kinetic parameters determined forthese mutants showed negligible change in Km for HndAH156Q butshowed a clear increase in Km for mutants HndAR33L andHndAY272F, suggesting their role as substrate-binding residues.Also, the kcat/Km values for the mutants decreased by nearly 6- to15-fold with respect to wild-type protein, indicating that all ofthese residues are essential for 2H1NA decarboxylation reaction(Table 2).

DISCUSSION

Decarboxylation is ubiquitous in nature and is of fundamentalbiological importance. Among the different classes of decarboxy-lases, nonoxidative decarboxylases have received specific atten-tion primarily because they catalyze transition metal-dependentdecarboxylation without using molecular oxygen as a cosubstrate(1). Microorganisms expressing these enzymes not only play asignificant role in biodegradation and/or bioremediation of soil,water, and sediment contaminated with lignin-related com-pounds and benzene derivatives of industrial origin but also act asbiocatalysts in industrial biotransformation reactions (4, 15, 17).

Nonoxidative decarboxylases belonging to structurally distinctprotein families that catalyze either reversible or irreversible reac-tions differ in their oxygen sensitivities (42). In aromatic acid me-tabolism, the presence of a variety of hydroxybenzoic acid non-oxidative decarboxylases has been detected, and some have beenpurified and characterized (2, 5, 14, 15, 42). To the best of ourknowledge, the oxygen-insensitive 2H1NA nonoxidative decar-boxylase described in the present study is the first bacterial enzymebelonging to the amidohydrolase superfamily that catalyzes anirreversible decarboxylation of a hydroxynaphthoic acid.

The amidohydrolase superfamily is comprised of functionallydiverse enzymes that catalyze the cleavage of the C–N, C–C, C–O,C–Cl, C–S, or O–P bond of structurally distinct organic com-pounds (43–45). Generally, the members of the amidohydrolasesuperfamily share a signature for mono- or binuclear metal centerembedded within the TIM-like barrel fold in the catalytic domain

FIG 3 (A) Schematic representation of the structural model of HndA showing the enzyme active site. The inset shows the metal coordinating residues His10,His12, His156, and Asp269. (B) Surface topology of HndA showing the binding of 2H1NA within the catalytic pocket via electrostatic interaction with active-siteresidues Tyr272 and Arg33 based on docking analysis.

TABLE 2 Kinetic constants of wild-type and mutant 2H1NAdecarboxylases

EnzymeKm

(mM)Vmax

(�mol min�1)kcat

(s�1)kcat/Km

(mM�1 s�1)

HnD 0.17 0.02 7.99 47.27HnDH10Q NDa ND ND NDHnDH156Q 0.17 0.001 0.56 3.31HnDD269V ND ND ND NDHnDH204Q ND ND ND NDHnDR33L 0.78 0.01 6.33 8.06HnDY272F 0.31 0.007 3.02 9.71a ND, not determined (product could not be detected).

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(43, 44). Within the amidohydrolase superfamily, members of theamidohydrolase 1 family catalyze hydrolytic reactions, while theamidohydrolase 2 family proteins are primarily involved in non-hydrolytic C-C bond cleavage (44). The gene encoding the2H1NA decarboxylase (hndA) displayed similarities with themembers of the amidohydrolase 2 family. In this family, ACMSDfrom Pseudomonas fluorescens is the first characterized memberinvolved in 2-nitrobenzoic acid degradation pathway (10). Othermembers belonging to this family include 5CVD, �-RSD, sali-cylic acid decarboxylase, OMAH, and Idcase (4, 15, 17, 40, 46).Since none of the enzymes belonging to the amidohydrolase 2family possess hydrolase activity, Liu and Zhang (43) had pro-posed to rename this family the ACMSD-related protein fam-ily. Based on sequence similarity and phylogenetic analysis(Fig. 1), we propose HndA to be a new member of the ACMSD-related protein family.

Multiple-sequence alignment of ACMSD-related protein fam-ily members, including HndA, revealed a strict conservation pat-tern for key amino acid residues which act as important metal-binding protein ligands (43). For HndA, His10 and His12constitute the conserved “HXH” metal-binding motif, whereasH156 and D269 are the other two endogenous metal-binding li-gands (Fig. 3A). Enzyme inhibition by histidine residue-specificinhibitor diethylpyrocarbonate (DEPC) and site-directed mu-tagenesis studies revealed that the conserved histidine residuesconstitute the active-site protein ligands (Table 2). Interestingly,substitution of another conserved histidine residue (His204), notdirectly involved in metal binding, leads to the complete inactiva-tion of the protein. This result is similar to that observed inACMSD, where the corresponding residue (His228) was sug-gested to play the role of an acid-base catalyst involved in depro-tonation of the metal-bound water, facilitating the decarboxyl-ation of ACMS (47). On the other hand, docking analysis revealedthat carboxyl and hydroxyl groups of 2H1NA are hydrogenbonded with Arg33/Tyr272 and Tyr272 of HndA, respectively.Interestingly, the importance of the arginine residue in substratebinding via carboxylate group has been suggested in ACMSD (33,48). Similarly, the role of the active-site tyrosine residue in thebinding of the hydroxyl group of lactic acid was studied in flavo-cytochrome b2 or L-lactate dehydrogenase, and it was reported toplay a role in converting lactic acid to pyruvic acid (49). The sig-nificance of Arg33 and Tyr 272 for efficient substrate binding inHndA was also confirmed by mutational analysis (Table 2).

Being a member of metallo-dependent hydrolase superfamily,HndA did not show any enhancement in activity when a set ofmetal ions were individually supplemented externally. Again,common divalent metal chelators had only mild inhibitory effectson this enzyme (see Table S6 in the supplemental material) evenafter prolonged incubation, suggesting a probable deeply buriedmetal center within the protein molecule. Modest inhibition by azinc metal-specific inhibitor, 8-hydroxy-quinoloine-5-sulfonicacid (see Table S6 in the supplemental material), that suggestedthe possible presence of a zinc metal center within the enzyme wasverified by atomic absorption spectroscopy. Additional biophysi-cal investigations on HndA will provide further insights on thecatalytic mechanism and structure-function relationships of thisunique transition metal-dependent, oxygen-independent 2H1NAdecarboxylase.

ACKNOWLEDGMENTS

We acknowledge Gautam Basu for editing the manuscript.Financial support for this study was provided by the Bose Institute,

Kolkata, India. P.P.C. was supported by fellowships from the Council ofScientific and Industrial Research, Government of India; S.B. and A.D.were supported by fellowships from the Bose Institute.

REFERENCES1. Li T, Huo L, Pulley C, Liu A. 2012. Decarboxylation mechanisms in

biological system. Bioorg Chem 43:2–14. http://dx.doi.org/10.1016/j.bioorg.2012.03.001.

2. Huang J, He Z, Wiegel J. 1999. Cloning, characterization, and expressionof a novel gene encoding a reversible 4-hydroxybenzoate decarboxylasefrom Clostridium hydroxybenzoicum. J Bacteriol 181:5119 –5122.

3. Chow KT, Pop MK, Davies J. 1999. Characterization of a vanillic acidnon-oxidative decarboxylation gene cluster from Streptomyces sp. D7. Mi-crobiology 145:2393–2403. http://dx.doi.org/10.1099/00221287-145-9-2393.

4. Peng X, Masai E, Kitayama H, Harada K, Katayama Y, Fukuda M. 2002.Characterization of the 5-carboxyvanillate decarboxylase gene and its rolein lignin-related biphenyl catabolism in Sphingomonas paucimobilisSYK-6. Appl Environ Microbiol 68:4407– 4415. http://dx.doi.org/10.1128/AEM.68.9.4407-4415.2002.

5. He Z, Wiegel J. 1996. Purification and characterization of an oxygen-sensitive, reversible 3,4-dihydroxybenzoate decarboxylase from Clostrid-ium hydroxybenzoicum. J Bacteriol 178:3539 –3543.

6. Gu W, Li X, Huang J, Duan Y, Meng Z, Zhang KQ, Yang J. 2011.Cloning, sequencing, and overexpression in Escherichia coli of the Entero-bacter sp. Px6-4 gene for ferulic acid decarboxylase. Appl Microbiol Bio-technol 89:1797–1805. http://dx.doi.org/10.1007/s00253-010-2978-4.

7. Cavin JF, Barthelmebs L, Divies C. 1997. Molecular characterization ofan inducible p-coumaric acid decarboxylase from Lactobacillus planta-rum: gene cloning, transcriptional analysis, overexpression in Escherichiacoli, purification, and characterization. Appl Environ Microbiol 63:1939 –1944.

8. Pujar BG, Ribbons DW. 1985. Phthalate metabolism in Pseudomonasfluorescens PHK: purification and properties of 4,5-dihydroxyphthalatedecarboxylase. Appl Environ Microbiol 49:374 –376.

9. Muraki T, Taki M, Hasegawa Y, Iwaki H, Lau PCK. 2003. Prokaryotichomologs of the eukaryotic 3-hydroxyanthranilate 3,4-dioxygenase and2-amino-3-carboxymuconate-6-semialdehyde decarboxylase in the 2-ni-trobenzoate degradation pathway of Pseudomonas fluorescens strain KU-7.Appl Environ Microbiol 69:1564 –1572. http://dx.doi.org/10.1128/AEM.69.3.1564-1572.2003.

10. Li T, Iwaki H, Fu R, Hasegawa Y, Zhang H, Liu A. 2006. �-Amino-�-carboxymuconic-ε-semialdehyde decarboxylase (ACMSD) is a new mem-ber of the amidohydrolase superfamily. Biochemistry 45:6628 – 6634.http://dx.doi.org/10.1021/bi060108c.

11. Colabroy KL, Begley TP. 2005. Tryptophan catabolism: identificationand characterization of a new degradative pathway. J Bacteriol 187:7866 –7869. http://dx.doi.org/10.1128/JB.187.22.7866-7869.2005.

12. Smiley JA, Kundracik M, Landfried DA, Barnes VR, Sr, Axhemi AA.2005. Genes of the thymidine salvage pathway: thymine-7-hydroxylasefrom a Rhodotorula glutinis cDNA library and isoorotate decarboxylasefrom Neurospora crassa. Biochim Biophys Acta 1723:256 –264. http://dx.doi.org/10.1016/j.bbagen.2005.02.001.

13. Yoshida T, Fujita K, Nagasawa T. 2002. Novel reversible indole-3-carboxylate decarboxylases catalyzing non-oxidative decarboxylation.Biosci Biotechnol Biochem 66:2388 –2394. http://dx.doi.org/10.1271/bbb.66.2388.

14. Santha R, Savithri HS, Rao NA, Vaidyanathan CS. 1995. 2,3-Dihydroxybenzoic acid decarboxylase from Aspergillus niger: a novel de-carboxylase. Eur J Biochem 230:104 –110. http://dx.doi.org/10.1111/j.1432-1033.1995.0104i.x.

15. Yoshida M, Fukuhara N, Oikawa T. 2004. Thermophilic, reversible�-resorcylate decarboxylase from Rhizobium sp. strain MTP-10005: puri-fication, molecular characterization, and expression. J Bacteriol 186:6855– 6863. http://dx.doi.org/10.1128/JB.186.20.6855-6863.2004.

16. Lupa B, Lyon D, Shaw LN, Sieprawska-Lupa M, Wiegel J. 2008. Prop-erties of the reversible nonoxidative vanillate/4-hydroxybenzoate decar-boxylase from Bacillus subtilis. Can J Microbiol 54:75– 81. http://dx.doi.org/10.1139/W07-113.

Pal Chowdhury et al.

1762 jb.asm.org June 2016 Volume 198 Number 12Journal of Bacteriology

on July 1, 2018 by guesthttp://jb.asm

.org/D

ownloaded from

17. Kirimura K, Gunji H, Wakayama R, Hattori T, Ishii Y. 2010. EnzymaticKolbe-Schmitt reaction to from salicylic acid from phenol: enzymaticcharacterization and gene identification of a novel enzyme, Trichosporonmoniliforme salicylic acid decarboxylase. Biochem Biophys Res Commun394:279 –284. http://dx.doi.org/10.1016/j.bbrc.2010.02.154.

18. Iwabuchi T, Harayama S. 1998. Biochemical and molecular character-ization of 1-hydroxy-2-naphthoate dioxygenase from Nocardioides sp.KP7. J Biol Chem 273:8332– 8336. http://dx.doi.org/10.1074/jbc.273.14.8332.

19. Vila J, Lopez Z, Sabate J, Minguillon C, Solanas AM, Grifoll M. 2001.Identification of a novel metabolite in the degradation of pyrene by My-cobacterium sp. strain AP1: actions of the isolate on two- and three-ringpolycyclic aromatic hydrocarbons. Appl Environ Microbiol 67:5497–5505.

20. Mallick S, Chatterjee S, Dutta TK. 2007. A novel degradation pathway inthe assimilation of phenanthrene by Staphylococcus sp. strain PN/Y viameta-cleavage of 2-hydroxy-1-naphthoic acid: formation of trans-2,3-dioxo-5-(2=-hydroxyphenyl)-pent-4-enoic acid. Microbiology 153:2104 –2115. http://dx.doi.org/10.1099/mic.0.2006/004218-0.

21. Kanaly RA, Harayama S. 2000. Biodegradation of high-molecular-weightpolycyclic aromatic hydrocarbons by bacteria. J Bacteriol 182:2059 –2067.http://dx.doi.org/10.1128/JB.182.8.2059-2067.2000.

22. Kanaly RA, Harayama S. 2010. Advances in the field of high-molecular-weight polycyclic aromatic hydrocarbon biodegradation by bacteria. Mi-crob Biotechnol 3:136 –164. http://dx.doi.org/10.1111/j.1751-7915.2009.00130.x.

23. Pinyakong O, Habe H, Supaka N, Pinpanichkarn P, Juntongjin K,Yoshida T, Furihata K, Nojiri H, Yamane H, Omori T. 2000. Identifi-cation of novel metabolites in the degradation of phenanthrene by Sphin-gomonas sp. strain P2. FEMS Microbiol Lett 191:115–121. http://dx.doi.org/10.1111/j.1574-6968.2000.tb09327.x.

24. Tao XQ, Lu GN, Dang Z, Yang C, Yi XY. 2007. A phenanthrene-degrading strain Sphingomonas sp. GY2B isolated from contaminatedsoils. Process Biochem 42:401– 408.

25. Prabhu Y, Phale PS. 2003. Biodegradation of phenanthrene by Pseu-domonas sp. strain PP2: novel metabolic pathway, role of biosurfactantand cell surface hydrophobicity in hydrocarbon assimilation. Appl Mi-crobiol Biotechnol 61:342–351. http://dx.doi.org/10.1007/s00253-002-1218-y.

26. Samanta SK, Chakraborti AK, Jain RK. 1999. Degradation of phenan-threne by different bacteria: evidence for novel transformation sequencesinvolving the formation of 1-naphthol. Appl Microbiol Biotechnol 53:98 –107. http://dx.doi.org/10.1007/s002530051621.

27. Chowdhury PP, Sarkar J, Basu S, Dutta TK. 2014. Metabolism of2-hydroxy-1-naphthoic acid and naphthalene via gentisic acid by dis-tinctly different sets of enzymes in Burkholderia sp. strain BC1. Microbi-ology 160:892–902. http://dx.doi.org/10.1099/mic.0.077495-0.

28. Bradford MM. 1976. A rapid and sensitive method for the quantitation ofprotein utilizing the principle of protein-dye binding. Anal Biochem 72:248 –254. http://dx.doi.org/10.1016/0003-2697(76)90527-3.

29. Shevchenko A, Tomas H, Havlis J, Olsen JV, Mann M. 2006. In-geldigestion for mass spectrometric characterization of proteins and pro-teomes. Nat Protoc 1:2856 –2860.

30. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. 1990. Basic localalignment search tool. J Mol Biol 215:403– 410. http://dx.doi.org/10.1016/S0022-2836(05)80360-2.

31. Thompson JD, Gilson TJ, Plewniak F, Jeanmougin F, Higgins, DG.1997. The ClustalX windows interface: flexible strategies for multiple se-quence alignment aided by quality analysis tools. Nucleic Acids Res 24:4876 – 4882.

32. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, Kumar S. 2011.MEGA 5: molecular evolutionary genetics analyses using maximum like-lihood, evolutionary distance, and maximum parsimony methods. MolBiol Evol 28:2731–2739. http://dx.doi.org/10.1093/molbev/msr121.

33. Martynowski D, Eyobo Y, Li T, Kun Yang K, Liu A, Zhang H. 2006.Crystal structure of �-amino-�-carboxymuconate-ε-semialdehyde de-

carboxylase: insight into the active site and catalytic mechanism of a noveldecarboxylation reaction. Biochemistry 45:10412–10421. http://dx.doi.org/10.1021/bi060903q.

34. Sali A, Blundell TL. 1993. Comparative protein modelling by satisfactionof spatial restraints. J Mol Biol 234:779 – 815. http://dx.doi.org/10.1006/jmbi.1993.1626.

35. Luthy R, Bowie JU, Eisenberg D. 1992. Assessment of protein modelswith three-dimensional profiles. Nature 356:83– 85. http://dx.doi.org/10.1038/356083a0.

36. Laskowski RA, MacArthur MW, Moss DS, Thornton JM. 1993.Procheck: a program to check the stereochemical quality of proteinstructures. J Appl Crystallogr 26:283–291. http://dx.doi.org/10.1107/S0021889892009944.

37. Willard L, Ranjan A, Zhang H, Monzavi H, Boyko RF, Sykes BD,Wishart DS. 2003. VADAR: a web server for quantitative evaluation ofprotein structure quality. Nucleic Acids Res 31:3316 –3319. http://dx.doi.org/10.1093/nar/gkg565.

38. Morris GM, Huey R, Lindstrom W, Sanner MF, Belew RK, GoodsellDS, Olson AJ. 2009. Autodock4 and AutoDockTools4: automated dock-ing with selective receptor flexibility. J Comput Chem 30:2785–2791. http://dx.doi.org/10.1002/jcc.21256.

39. Laskowski RA, Swindells MB. 2011. LigPlot: multiple ligand-proteininteraction diagrams for drug discovery. J Chem Infect Model 51:2778 –2786. http://dx.doi.org/10.1021/ci200227u.

40. Hara H, Masai E, Katayama Y, Fukuda M. 2000. The 4-oxalomesaconatehydratase gene, involved in the protocatechuate 4,5-cleavage pathway, isessential to vanillate and syringate degradation in Sphingomonas paucimo-bilis SYK-6. J Bacteriol 182:6950 – 6957. http://dx.doi.org/10.1128/JB.182.24.6950-6957.2000.

41. Goto M, Hayashi H, Miyahara I, Hirotsu K, Yoshida M, Oikawa T.2006. Crystal structures of nonoxidative zinc-dependent 2,6-dihydroxybenzoate (�-resorcylate) decarboxylase from Rhizobium sp.strain MTP-10005. J Biol Chem 281:34365–34373. http://dx.doi.org/10.1074/jbc.M607270200.

42. Ishii Y, Narimatsu Y, Iwasaki Y, Arai N, Kino K, Kirimura K. 2004.Reversible and nonoxidative �-resorcylic acid decarboxylase: character-ization and gene cloning of a novel enzyme catalyzing carboxylation ofresorcinol, 1,3-dihydroxybenzene, from Rhizobium radiobacter. BiochemBiophys Res Commun 324:611– 620. http://dx.doi.org/10.1016/j.bbrc.2004.09.091.

43. Liu A, Zhang H. 2006. Transition metal-catalyzed nonoxidative decar-boxylation reactions. Biochemistry 45:10407–10411. http://dx.doi.org/10.1021/bi061031v.

44. Seibert CM, Raushal FM. 2005. Structural and catalytic diversity withinthe amidohydrolase superfamily. Biochemistry 44:6384 – 6391.

45. Kim GJ, Lee DE, Kim HS. 2000. Functional expression and character-ization of the two cyclic amidohydrolase enzymes, allantoinase and a novelphenyl hydantoinase, from Escherichia coli. J Bacteriol 182:7021–7028.http://dx.doi.org/10.1128/JB.182.24.7021-7028.2000.

46. Xu S, Li W, Zhu J, Wang R, Li Z, Xu GL, Ding J. 2013. Crystal structuresof isoorotate decarboxylases reveal a novel catalytic mechanism of 5-car-boxyl-uracil decarboxylation and shed light on the search for DNA decar-boxylase. Cell Res 23:1296 –1309. http://dx.doi.org/10.1038/cr.2013.107.

47. Huo L, Fielding AJ, Chen Y, Li T, Iwaki H, Hosler JP, Chen L, HasigawaY, Que L, Jr, Liu A. 2012. Evidence for a dual role of an active site histidinein �-amino-�-carboxymuconate-ε-semialdehyde decarboxylase. Bio-chemistry 51:5811–5821. http://dx.doi.org/10.1021/bi300635b.

48. Huo L, Davis I, Chen L, Liu A. 2013. The power of two: arginine 51 andarginine 239* from a neighboring subunit are essential for catalysis in�-amino-�-carboxymuconate-ε-semialdehyde decarboxylase. J BiolChem 288:30862–30871. http://dx.doi.org/10.1074/jbc.M113.496869.

49. Gondry M, Dubois J, Terrier M, Lederer F. 2001. The catalytic role oftyrosine 254 in flavocytochrome b2 (L-lactate dehydrogenase from baker’syeast): comparison between the Y254F and Y254L mutant proteins. Eur JBiochem 268:4918 – 4927.

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