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Originally published In Press as doi:10.1074/jbc.M401692200 on April 1, 2004

J. Biol. Chem., Vol. 279, Issue 26, 27646-27655, June 25, 2004
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Crystal Structure of 4-Chlorocatechol 1,2-Dioxygenase from the Chlorophenol-utilizing Gram-positive Rhodococcus opacus 1CP*

Marta Ferraroni{ddagger}, Inna P. Solyanikova§, Marina P. Kolomytseva§, Andrea Scozzafava{ddagger}, Ludmila Golovleva§, and Fabrizio Briganti{ddagger}

From the {ddagger}Dipartimento di Chimica, Università di Firenze, Via della Lastruccia 3, Sesto Fiorentino I-50019, Italy, and the §Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino Moscow Region 142290, Russia

Received for publication, February 16, 2004 , and in revised form, March 30, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
The crystal structure of the 4-chlorocatechol 1,2-dioxygenase from the Gram-positive bacterium Rhodococcus opacus (erythropolis) 1CP, a Fe(III) ion-containing enzyme involved in the aerobic biodegradation of chloroaromatic compounds, has been solved by multiple wavelength anomalous dispersion using the weak anomalous signal of the two catalytic irons (1 Fe/257 amino acids) and refined at a 2.5 Å resolution (Rfree 28.7%; R factor 21.4%). The analysis of the structure and its comparison with the structure of catechol 1,2-dioxygenase from Acinetobacter calcoaceticus ADP1 (Ac 1,2-CTD) highlight significant differences between these enzymes. The general topology of the present enzyme comprises two catalytic domains (one for each subunit) related by a noncrystallographic 2-fold axis and separated by a common {alpha}-helical zipper motif consisting of five N-terminal helices from each subunit; furthermore the C-terminal tail is shortened significantly with respect to the known Ac 1,2-CTD. The presence of two phospholipids binding in a hydrophobic tunnel along the dimer axis is shown here to be a common feature for this class of enzyme. The active site cavity presents several dissimilarities with respect to the known catechol-cleaving enzyme. The catalytic nonheme iron(III) ion is bound to the side chains of Tyr-134, Tyr-169, His-194, and His-196, and a cocrystallized benzoate ion, bound to the metal center, reveals details on a novel mode of binding of bidentate inhibitors and a distinctive hydrogen bond network with the surrounding ligands. Among the amino acid residues expected to interact with substrates, several are different from the corresponding analogs of Ac 1,2-CTD: Asp-52, Ala-53, Gly-76, Phe-78, and Cys-224; in addition, regions of largely conserved amino acid residues in the catalytic cleft show different shapes resulting from several substantial backbone and side chain shifts. The present structure is the first of intradiol dioxygenases that specifically catalyze the cleavage of chlorocatechols, key intermediates in the aerobic catabolism of toxic chloroaromatics.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Halogenated aliphatic and aromatic hydrocarbons are a major environmental concern because large amounts of such toxic chemicals, resistant to both chemical oxidation and biological degradation, have been released into the ecosphere in the last decades as pesticides, fire retardants, and solvents (1, 2). The substitution for hydrogen with xenobiotic halogens in readily biodegradable hydrocarbons has in fact led to substances with significantly decreased microbial decomposition and, consequently, to their environmental accumulation (3).

Specialized strains of aerobic and anaerobic bacteria have been discovered to use selected halogenated hydrocarbons as sole carbon and energy sources (1, 4, 5).

The aerobic catabolism of chloroaromatics usually occurs through two different pathways (5, 6). Compounds containing one or two chlorine atoms are usually converted to chlorocatechols and then catabolized through the modified ortho-cleavage pathway (10, 11). Aromatic compounds containing more than two chlorine atoms are converted to hydroxyquinol or chlorohydroxyquinols and then cleaved by specific intradiol dioxygenases (Scheme 1) (12-18). A few interesting exceptions have been observed (7-9).



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  		SCHEME 1.
Aerobic catabolism of monoaromatic hydrocarbons.

 
The modified ortho-cleavage pathway is a central oxidative bacterial pathway that channels chlorocatechols, derived from the degradation of chlorinated benzoic acids, phenoxyacetic acids, phenols, benzenes, and other aromatics into the energy-generating tricarboxylic acid pathway (10, 11, 19) (Scheme 1). These chlorocatechols are degraded further by the addition of molecular oxygen and the subsequent cleavage between two adjacent hydroxyl groups catalyzed by nonheme Fe(III)-dependent metalloenzymes, classified as intradiol dioxygenases (19).

Dioxygenases that cleave catechols in an intradiol fashion can be divided into two structurally different families: protocatechuate 3,4-dioxygenases (3,4-PCDs)1 and catechol 1,2-dioxygenases (1,2-CTDs). 3,4-PCDs, specific for hydroxybenzoates, are typically composed of two homologous subunits in large oligomeric complexes ({alpha}{beta}Fe3+)2-12 (22), 1,2-CTDs are dimers ({alpha}Fe3+)2 with identical or very similar subunits and generally comprise enzymes with markedly different substrate specificities: catechol, alkylated catechols, hydroxyquinols (type I enzymes) (23-27), and chlorocatechols (type II enzymes) (5, 28-41). Many of these enzymes have been characterized extensively in terms of biochemical, spectroscopic, and kinetic properties (23-41). Mechanistic and structural details have also been gathered for one 1,2-CTD (42) and several 3,4-PCDs (43-47). In particular the crystal structures of 3,4-PCDs revealed that the catalytic ferric ion is ligated by two histidine and two tyrosine residues plus a hydroxyl ion in a trigonal-bipyramidal geometry (43). These residues are conserved in all investigated intradiol dioxygenases. Further exhaustive studies by Lipscomb et al. (45, 46) described the mode of binding of substrates, substrate analogs, and inhibitors of 3,4-PCDs and allowed speculation on the subsequent interaction with dioxygen. Recently, structural information on the 1,2-CTD family was obtained, although limited to a single enzyme, the catechol 1,2-dioxygenase from Acinetobacter calcoaceticus ADP1 (Ac 1,2-CTD hereafter). The structure of this enzyme differs markedly from those of 3,4-PCDs; it is characterized by the presence of a novel structural hydrophobic helical zipper motif as a subunit linker and shows that only one subunit is used to fashion each active site cavity (42). Furthermore 1,2-CTD anaerobic adducts with catechol and 4-methylcatechol confirmed several observations seen in 3,4-PCDs such as iron tyrosinate displacement upon substrate binding (42, 45-47).

Because the substrate specificities of the chlorocatechol 1,2-dioxygenases (1,2-CCDs) (type II enzymes) characterized up until now differ considerably from those of 1,2-CTDs (type I enzymes) (28-41), we believe that structural details on chlorocatechol-cleaving dioxygenases are needed to understand further the mechanism of substrate selection.

X-ray absorption measurements on 4-chlorocatechol 1,2-dioxygenase from Rhodococcus opacus (erythropolis) 1CP (Rho 1,2-CCD hereafter) and other intradiol cleaving dioxygenases have provided the first insights into the recognition of different chemical structures by ortho-cleaving dioxygenases (48).

Diffraction quality crystals have been obtained and x-ray diffraction studies are in progress for other intradiol dioxygenases with different substrate specificities (49-54).

In this paper we report the crystal structure determination for Rho 1,2-CCD solved using a multiple wavelength anomalous dispersion (MAD) experiment and refined at 2.5 Å. This is the first dioxygenase that catalyzes the degradation of chlorocatechols for which the three-dimensional structure has been determined providing details on the conformation of the active site and providing information on which particular residues are involved in substrate selection for enzymes of the 1,2-CCD family.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Protein Preparation, Crystallization, and Data Collection—Rho 1,2-CCD was purified from R. opacus 1CP as reported previously (41). The enzyme was crystallized at 293 K using the sitting drop vapor diffusion method from a solution containing 1.6 M ammonium sulfate, 0.1 M sodium chloride, 100 mM Tris-HCl, pH 7.5, and 5-15% glycerol (53). All of the reagents were of the best purity available from Sigma.

Diffraction data extending to a maximum resolution of 2.5 Å were collected at the x-ray diffraction beamline at Elettra, Trieste. Data were collected using a MAR image plate detector and a wavelength of 1.0 Å. Crystals belong to the primitive hexagonal space group P6322 with unit cell dimensions a = b = 89.33, c = 313.39. Assuming one molecule/asymmetric unit, the solvent content is about 61% of the unit cell (VM = 3.13 Å3/Da) (21). For all the data collections, crystals of the native enzyme were cooled at 100 K adding 30% glycerol to the mother liquor solution as cryoprotectant. Because of the large unit cell dimensions an image plate detector was utilized to reach the maximal resolution of the crystal without having too many overlapping reflections.

Structure Determination and Refinement—All molecular replacement attempts, using coordinates of known intradiol dioxygenase structures as a model, failed to provide a solution for Rho 1,2-CCD.

The structure of the enzyme Rho 1,2-CCD has been solved by MAD using the weak anomalous signal of the two catalytic irons (1 Fe/257 amino acids). MAD data were collected at the BW7A beamline, EMBL, DESY, Hamburg. The data collected at three wavelengths (inflection, peak, remote) were processed and integrated with DENZO and scaled by SCALEPACK, from the HKL program suite (55). Data processing statistics are summarized in Table I.


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  		TABLE I
Summary of data collection and atomic model refinement statistics

Numbers in parentheses are for the highest resolution shell.

 
The program SOLVE (56) was used to identify the two iron sites and for phases calculation. The 3.5 Å MAD phases were improved and extended to 2.5 Å by solvent flattening and histogram mapping using the program DM from the CCP4 program suite (57). The resulting electron density map was of high quality and allowed the manual tracing of 440-514 amino acid residues using the program QUANTA (58). Refinement of the model, performed at 2.5 Å resolution, using the program Refmac 5.1.24 from the CCP4 program suite (57), resulted in R factor and Rfree values of 21.4 and 28.7%, respectively.

The electron densities corresponding to residues Met-A1, Met-B1 and to the side chains of Lys-A89, Lys-B66, Lys-B173, His-B256, and Gln-B255 were missing, and therefore the corresponding atoms and bonds were not modeled. Electron density resembling one benzoate molecule bound to the iron ions was found in both active sites. Because no benzoate-like compounds were added during the crystallization procedures a possible explanation could be that such compound is formed during cell growth. Inhibition tests revealed that benzoate is a weak competitive inhibitor for the present Rho 1,2-CCD. Electronic density corresponding to a Tris buffer molecule was also found in a surface crevice interacting with residues Thr-28, Tyr-31, Glu-32, and Met-35 (from subunit B, helix H2) and Pro-48, Asp-52, Glu-56 (subunit A, helices H3 and H4), Gln-75, Gly-76, and Pro-77 (subunit A, random coil). The final coordinates have been deposited in the Protein Data Bank (accession number 1S9A [PDB] ).

Structure Analysis—The stereochemical quality of the models was assessed using the program PROCHECK (59). The Ramachandran plot is of good quality; there are 430 nonglycine and nonproline residues; among these, 374 (87%) are in the most favored regions, 50 (11.6%) are in the additional allowed regions, 5 (1.2%) in the generously allowed regions, and 1 (His-B256) (0.2%) in disallowed regions.

The secondary structure was defined utilizing the DSSP data base and program (60).

Multiple sequence alignments were performed using the ClustalX program (61).

Global structures superimpositions were carried out by utilizing the matching algorithm implemented into the HEX 4.1 program (62). Least squares fit of the active site regions was performed using the McLachlan algorithm as implemented in the program ProFit 2.2 (www.bioinf.org.uk/software/profit/) specifying as the fitting subset the four amino acid ligands to the catalytic iron ions (63).

Electrostatic potentials were estimated first transforming the Protein Data Bank coordinate file into a pqr file containing partial charges and radii for each atom by using the PDB2PQR web service and then solving the second order differential Poisson-Boltzmann equation, which relates the electrostatic potential in a dielectric to the charge density using the macroscopic electrostatics with atomic detail (MEAD) program package (64, 65).

3- and 4-substituted catechols were docked manually into the active site by first simulating the dissociation of Tyr-169 from the iron center. As observed previously, it was assumed that the substrates bind to the iron in a bidentate fashion and with orientations of their aromatic ring similar to those observed for catechol or 4-methyl catechol in Ac 1,2-CTD or for the benzoate ion in Rho 1,2-CCD (42). Slight rotations and/or tilts of the iron bound substrate molecules did not result in changes in the amino acid residues interacting with the substrates ring substituents.

PyMol, UCSF Chimera, and MSMS were used to produce ribbon diagrams, electrostatic potential surfaces, electron density, and other representations (66-68).


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
CONCLUSIONS
 REFERENCES
 
Overall Structure and Linker Domain—The 1,2-CCD from the Gram-positive bacterium R. opacus (erythropolis) 1CP is a homodimeric protein with overall dimensions 105 x 40 x 40 Å. The statistics for data collection, phasing, and structure refinement are summarized in Table I. The final model includes residues 2-257 for each monomer, two Fe(III) ions, two benzoate ions, two phospholipids, 284 water molecules, and one Tris buffer molecule.

The global structure resembles that of catechol 1,2-dioxygenase from A. calcoaceticus ADP1, the only known structure for this class of dioxygenases (Fig. 1) (42).



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  		FIG. 1.
Schematic representation of the overall structures of Ac 1,2-CTD (A), Rho 1,2-CCD (B), and B rotated 90° around the horizontal axis (C). The iron ions are represented as pink spheres; the phospholipid molecules are also shown.

 
The overall fold of the dimer comprises two catalytic domains made up by a number of {beta}-sheets and several random coils, related by a noncrystallographic 2-fold axis, and a linker domain composed by several {alpha}-helices coming from both monomers and located at the center of the molecule. Two phospholipids are bound inside a channel formed by the two protein monomers at the center of the linker domain (see Fig. 1C). Each subunit entirely hosts a catalytic pocket containing an iron(III) ion accessible to the substrate from the hollow side of the dimer.

The linker domain is mainly composed by two long (H1 and H2) and three short (H3-H5) {alpha}-helices supplied by each subunit: four helices from the N terminus of each monomer form a hook that is interacting with the equivalent motif from the other subunit and to the fifth helix, which elongates from its catalytic domain (Figs. 1 and 2).



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  		FIG. 2.
Schematic representation of the secondary structure of Rho 1,2-CCD assigned using the program DSSP (60).

 
ClustalX sequence alignments of Rho 1,2-CCD, with a variety of other 1,2-CCDs and the representative of 1,2-CTDs, Ac 1,2-CTD, are reported in Fig. 3. The sequence identity between Rho 1,2-CCD and Ac 1,2-CTD is about 30%. Significant differences are observable in the helical linker domain and in the C-terminal region by comparing 1,2-CCDs with Ac 1,2-CTD sequences.



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  		FIG. 3.
Amino acid sequence alignments of chlorocatechol dioxygenases and Ac 1,2-CTD (61). A, R. opacus 1CP 3-chlorocatecholspecific CCD (NCBI protein accession O67987 [GenBank] ); B, R. opacus 1CP CCD (NCBI protein accession CAD28142 [GenBank] present enzyme); C, Achromobacter xylosoxidans A8 CCD (NCBI protein accession CAD56206 [GenBank] ; D, R. eutropha NH9 CCD (NCBI protein accession BAA74530 [GenBank] ; E, Delftia acidovorans P4a CCD (NCBI protein accession AAC35836 [GenBank] ; F, Pseudomonas chlororaphis RW71 CCD (NCBI protein accession CAB52140 [GenBank] ; G, Pseudomonas sp. P51 CCD (NCBI protein accession P27098 [GenBank] ); H, Burkolderia cepacia 2a CCD (NCBI protein accession AAK81678 [GenBank] ; I, Variovorax paradoxus TV1 CCD (NCBI protein accession AAC01745 [GenBank] ; J, R. eutropha plasmid pJP4 CCD (NCBI protein accession P05403 [GenBank] ); K, P. putida plasmid pAC27 CCD (NCBI protein accession A27058 [GenBank] ); L, Ralstonia sp. JS705 CCD (NCBI protein accession CAA06968 [GenBank] ; M, Pseudomonas aeruginosa CCD (NCBI protein accession T44616 [GenBank] ); N, P. aeruginosa JB2 CCD (NCBI protein accession AAC69474 [GenBank] ; O, Pseudomonas sp. GT241-1 CCD (NCBI protein accession AAR88247 [GenBank] ; P, R. eutropha JMP134 CCD (NCBI protein accession AAC44730 [GenBank] ; Q, Burkolderia sp. NK8 CCD (NCBI protein accession BAB56009 [GenBank] ; R, Burkolderia sp. R172 CCD (NCBI protein accession AAP92136 [GenBank] ; S, Defluvibacter lusatiensis CCD (NCBI protein accession CAD60254 [GenBank] ; T, A. calcoaceticus ADP1 CTD (NCBI protein accession P07773 [GenBank] ). In the line above the alignment the strongly conserved positions are indicated in the following ways: asterisk (*), positions that have a single, fully conserved residue; colon (:), positions with fully conserved strong groups; dot (.) positions with fully conserved weaker groups, according to ClustalX (61).

 
Fig. 4 shows the three-dimensional structural least squares superposition of a single subunit of Rho 1,2-CCD and Ac 1,2-CTD. The first C-terminal {alpha}-helix and the following random coil region present in the conventional 1,2-CTDs are totally missing in Rho 1,2-CCD. The two long H1 and H2 helices in Rho 1,2-CCD (see Fig. 2) closely match the second and third helices of Ac 1,2-CTD, whereas the short H3 helix, which provides several residues to the active site cavity (Leu-49, Asp-52, and Ala-53), is partly shifted, and about half of the fourth helix (H4) is missing with respect to the corresponding ones from Ac 1,2-CTD. On the contrary, the secondary structure of the central section of Rho 1,2-CCD thoroughly resembles that of the 1,2-CTD family. Finally, the N-terminal region of Rho 1,2-CCD misses the seventh helix, the last long random coil, and the final {beta}-sheet present in Ac 1,2-CTD.



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  		FIG. 4.
A, least squares superimposition of a single subunit of Ac 1,2-CTD (green) and Rho 1,2-CCD (red). B, 90° horizontally rotated A.

 
The presence of two bound phospholipids seems to be a common feature of this class of enzyme (42). Their precise identity could not be determined because of the absence of the electron density of the head groups, and the length of each tail was based on the length of the electron density and the stereochemistry of known phospholipids. In the present case they were modeled utilizing a phosphatidylcholine molecule with two C14/C15 hydrophobic tails. They are placed at the interface between the two subunits, one at each end of a large hydrophobic channel made up by helices H1 and H2 from both monomers, with the head group directed outward into the solvent and the tail moieties pointing inwards toward each other (see Fig. 1, B and C). Various hypothesis have been made on the role of the hydrophobic tunnel and of the bound phospholipids, but additional studies are needed to clarify their functional role (42).

Catalytic Domain—The core of the catalytic domain is made up of a series of {beta}-sheets arranged in a {beta}-sandwich conformation and by several random coils positioned between the linker domain and the {beta}-sheets assemblies (see Figs. 1 and 4). The active site metal center is located in a random coils region flanked on one side by the {beta}-sheet assemblies of each monomer and on the other side by the {alpha}-helices of the linker domain.

The catalytic pocket of Rho 1,2-CCD is formed by several residues: Leu-49, Asp-52, and Ala-53 from helix H3; Ile-74, Gln-75, Gly-76, Pro-77, Phe-78, and Phe-79 from a single random coil; Trp-126 from sheet S3; the iron ligand Tyr-134 from a second random coil; Tyr-169 from sheet S6; Ile-171 from a third random coil; Arg-191 from sheet S7; His-194 and His-196 from sheet S8; Gln-210 from sheet S9, and Cys-224 from another random coil.

Residues Leu-49, Pro-77, Phe-78, Ile-171 (Pro-172), and the backbones of the iron ligands Tyr-134 and Tyr-169 compose the hydrophobic active site entrance.

Fig. 5 shows the electrostatic potential mapped onto the molecular surfaces of Rho 1,2-CCD (A) and Ac 1,2-CTD (B) in the surroundings of the catalytic site entrances. Contrary to what observed for 3,4-PCDs, the groups neighboring the opening of the active site for both enzymes do not provide any marked positive electrostatic potential, which is supposed to guide the substrate into the catalytic cleft for 3,4-PCDs (43). In fact in catechol-cleaving dioxygenases the substrate binding driving forces are mainly of a hydrophobic nature because the substrate catechol is mostly undissociated at physiological pH.



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  		FIG. 5.
Electrostatic potential surfaces of Rho 1,2-CCD (A) and Ac 1,2-CTD (B). The active site entrances are indicated by yellow circles. The blue color corresponds to positive and red to negative potentials.

 
A close view of the active site residues, with the corresponding Fobs - Fcalc density overlaid, is represented in Fig. 6A. The catalytic center contains a mononuclear iron(III) ion, bound to four amino acid residues Tyr-134, Tyr-169, His-194, and His-196. The His2Tyr2 coordination is typical of all intradiol ring cleaving dioxygenases (26). X-ray absorption spectroscopy data on the same 1,2-CCD indicates that the native enzyme is pentacoordinated with two spheres of atoms: either two at 1.9 Å and three at 2.1 Å, or three at 1.9 Å and two at 2.1 Å (48). In the present crystal structure, a benzoate ion is found to bind to the iron ion in a bidentate asymmetric mode in place of the metal-bound water molecule/hydroxide ion, generally found in the native enzymes, expanding the iron coordination number to six (Fig. 6A). For this reason the iron ion bond distances observed in the present crystal structure are longer than those determined by the extended x-ray absorption fine structure as well as those reported for the native Ac 1,2-CTD structure (for comparison, see Table II). His-194 exhibits the shortest bond (~2.0 Å); Tyr-134, His-196, and Tyr-169 have longer bond distances (~2.2 Å), and the benzoate molecule asymmetrically bound with O1 (~2.5 Å) and O2 (~2.1 Å) completes the iron coordination sphere (see also Table II).



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  		FIG. 6.
A, Fobs - Fcalc electron density map for the active site of Rho 1,2-CCD. The electron density is contoured at 3{sigma}. B-D, representations of the solvent-accessible surface in the active site region of Rho 1,2-CCD. The surface is color-coded on the basis of the calculated electrostatic potential. The blue color corresponds to positive and red to negative potentials. The iron ion is represented as a green sphere. The three pictures show 90° vertically rotated views of the cavity, and the active site entrance is indicated by a yellow arrow.

 


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  		TABLE II
Bond distances and angles of the catalytic iron ion coordination polyhedron for Rho 1,2-CCD

The bond distances in parentheses are those observed in Ac 1,2-CTD. Estimated bond distance errors for Rho 1,2-CCD structure are ~0.35 Å, based on Luzzati analysis (20).

 
Extensive studies on inhibitors, substrates, and substrate analog adducts of 3,4-PCDs and Ac 1,2-CTD have revealed several important features of the mechanism of exogenous ligands binding to their active site: phenolate competitive inhibitors bind to iron in a monodentate way; those showing low affinity generate distorted trigonal-bipiramidal iron coordination geometries with the phenolate displacing the water/hydroxide ligand, whereas the highest affinity inhibitors bind to iron concomitantly to the solvent molecule generating six-coordinated distorted octahedral metal coordination geometries (45, 47). It has also been observed that substrates (in anaerobic conditions) or substrate analogs bind to the iron center in a asymmetric bidentate mode, resulting in distorted trigonal-bipiramidal iron geometry because of the simultaneous dissociation of the axial tyrosinate and of the water/hydroxide iron ligands, both supposed to function as proton acceptors during substrate deprotonation and binding. Therefore, in the anaerobic substrate adducts, the sixth coordination site is unoccupied, whereas in substrate analog adducts it is occupied by water or cyanide (if added to the buffer) (42, 45-47). Octahedral iron coordination is also observed in 3,4-PCDs at low pH where a sulfate ion completes the coordination sphere of the active metal center (47). In the present structure we notice a different mode of binding of the iron-chelating benzoate, which causes the dissociation of the water/hydroxide iron ligand, but it does not trigger the detachment of the tyrosinate residue thus resulting in a distorted octahedral iron coordination.

In Fig. 6, B-D, are shown different views of the solvent-exposed surface in the active site region of Rho 1,2-CCD color coded for the calculated electrostatic potential. By positioning the enzyme molecule with the iron ion at the bottom of the catalytic pocket, we observe a region, made up by the iron ligands His-194, His-196, by the backbone nitrogen of Phe-78, by the ring nitrogen of Trp-126 and, on the opposite side, by Arg-191 and Gln-210, with positive electrostatic potential surrounding the metal ion, which should support the substrate/inhibitor deprotonation and binding. The active site zone opposite to the iron ion is, in contrast, made up by the negatively charged residue Asp-52 and a negative electrostatic potential is also surrounding Glu-56, Thr-177, and Met-181. This negative potential area could have implications for the correct orientation of the catechol substrates. The intermediate active site region is made up by residues Ala-53, Ile-74, by the backbones of Gly-76 and Pro-77, and particularly by Cys-224, which has been suggested previously to interact with the chlorine substituents of the aromatic ring of the substrate molecules (42).

Fig. 7A reports the superposed active sites of Rho 1,2-CCD and native Ac 1,2-CTD obtained by minimizing onto the four amino acid iron ligand residues (Tyr-134/164, Tyr-169/200, His-194/224, and His-196/226 for Rho 1,2-CCD/Ac 1,2-CTD) (62). Several amino acid substitutions are observed in the catalytic cavity, the most evident being Asp-52/Pro-76, Ala-53/Gly-77, Phe-78/Leu-109, and Cys-224/Ala-254 (Rho 1,2-CCD/Ac 1,2-CTD), whereas the other residues are basically conserved, although significant backbone and side chain shifts are detected. As reported above, in Rho 1,2-CCD the benzoate ion is chelated asymmetrically to iron, and it is stabilized by a hydrogen-bonding network that connects the benzoate O2 atom to Arg-191 NH1 (hydrogen-bonded further to Gln-210, Asp-222, and Tyr-212) and the benzoate O1 atom to a well ordered W10 active site water molecule (hydrogen-bonded further to Gln-75 and Trp-126). The binding of benzoate also results in the absence of a second active site water molecule (W572 in Ac 1,2-CTD) caused by the conformational orientations of Arg-191 and of the hydrogen-bonded Gln-210 which turned out to be very similar to those observed when catechols bind to Ac 1,2-CTD (although benzoate is not able to trigger the dissociation of Tyr-169). Arg-191 and Gln-210, conserved across all 1,2-CTDs and 3,4-PCDs, are supposed to promote the substrate positioning and deprotonation, providing significant van der Waals interactions with the ring to prevent its rotation into two equatorial positions and/or providing an electrostatic countercharge to the buildup of electron density on a carbon atom of the ring (42, 44-47).



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  		FIG. 7.
Superposition of the active sites structure of Rho 1,2-CCD onto native Ac 1,2-CTD (orange, Protein Data Bank code 1DlM [PDB] ) (A) and onto the catechol complex of Ac 1,2-CTD (magenta, Protein Data Bank code 1DLT [PDB] ) (B).

 
The structural alignment of the active site residues of the catecholate complex of Ac 1,2-CTD and Rho 1,2-CCD is shown in Fig. 7B. The main interactions of Ac 1,2-CTD with the substrate involve residues Leu-73, Pro-76, Ile-105, Pro-108, Leu-109, Arg-221, Phe-253, and Ala-254. The catechol molecule in Ac 1,2-CD binds to the iron ion with one hydroxyl group in an axial position trans to His-226 and the other in an equatorial position trans to Tyr-164.

The benzoate observed in Rho 1,2-CCD binds to the metal center occupying the ligand position trans to His-194 which is left empty in the productive catechol complex (in Ac 1,2-CTD) for the subsequent binding of the oxygen molecule (Scheme 2; the numbering in parentheses refers to Ac 1,2-CTD). Although the mode of benzoate binding to Rho 1,2-CCD is radically different from that of catechol in Ac 1,2-CTD the aromatic ring orientation is the same (see Fig. 7B).



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  		SCHEME 2.
Modes of binding of benzoate and catechol to the active sites of 1,2-CTDs and 1,2-CCDs.

 
The catalytic mechanism of intradiol cleavage dioxygenases has been proposed to proceed via activation of the catechol substrate by iron(III) to give an iron(II) semiquinone, which reacts directly with dioxygen to give a hydroperoxide intermediate, which then undergoes Criegee rearrangement via acyl migration to give muconic anhydride (69). An alternative mechanism for migration of the electron-deficient acyl group, via a benzene-oxide-oxepin interconversion, has also been proposed (70). A comparison of Rho 1,2-CCD with 3,4-PCDs and Ac 1,2-CTD indicates that residues surrounding the postulated oxygen binding pocket are partially conserved, in particular Pro-77 (Pro-108 and Pro-15 in Ac 1,2-CTD and Pseudomonas putida 3,4-PCDs, respectively) is retained, whereas Phe-78 is replaced by Leu-109 and by Tyr-16 in Ac 1,2-CTD and P. putida 3,4-PCDs, respectively; nevertheless the side chains of such residues point toward the external part of the cavity, and the backbone portion, which is near to the proposed oxygen binding location, is mainly unaffected by such substitutions, suggesting that the oxygen molecule has similar interactions in all intradiol ring cleaving-dioxygenases (45).

The residues surrounding the ring of the benzoate molecule and supposed to be involved in the correct positioning of the aromatic substrate are Leu-49, Asp-52, Ala-53, Ile-74, Gly-76, Pro-77, Phe-78, Gln-210, and Cys-224. The present enzyme is able to catalyze the ring opening of a variety of substituted catechols (41). We attempted to dock 3- or 4-chlorocatechols into the active site. In the case of the 3-chloro derivative, two different orientations are possible: if the substituent is oriented toward the internal part of the cavity it would settle into a pocket formed by residues Ile-74 and Gly-76 (both conserved in 1,2-CTDs), but if the substituent is oriented outward it would essentially interact with residue Phe-78 (substituted for Leu-109 in Ac 1,2-CTD). Nevertheless, the preferred substrates for Rho 1,2-CCD are those bearing a substituent in position 4. If the 4-substituted catecholate binds with the substituent oriented toward the inner part of the catalytic site, as observed in the 4-methylcatechol adduct of Ac 1,2-CTD, it would be hosted in a cleft modeled by Asp-52, Ala-53 (replaced with Pro-76 in Ac 1,2-CTD), Gly-76, and Cys-224 (Ala-254 in Ac 1,2-CTD); whereas if it prefers the outward orientation it would interact mainly with Leu-49 (conserved in Ac 1,2-CTD).

From the ClustalX sequence alignments of Rho 1,2-CCD, with a variety of other 1,2-CCDs (Fig. 3) it can be noticed that several regions contributing residues to the active site cavity show significant amino acid substitutions. In particular, the 1,2-CCDs from Pseudomonas putida pAC27, Ralstonia eutropha pJP4, Pseudomonas chlororaphis RW71 and the 3-chlorocatechol specific 1,2-CCD from R. opacus CP1 (same strain expressing the present enzyme) exhibit higher activities toward 3-chlorocatechol than Rho 1,2-CCD. Possible reasons for the different observed specificities could be the replacement of Phe-78, a residue expected to interact with substituents in position 3, with Tyr, and in the most internal side of the cavity Ala-53, which is exchanged with Val in all the other 1,2-CCDs described (35, 36, 41, 71). Furthermore, Cys-223 and 224, suggested to be important for interacting with chlorine substituents, are generally conserved in CCDs, as shown in Fig. 3, but in the present enzyme only Cys-224 is present, whereas residue 223 is substituted for Ser (42). It has also to be noted that in the recently crystallized 3-chlorocatechol-preferring 1,2-CCD from R. opacus CP1 both Cys-223 and Cys-224 are replaced by Val and Ala, respectively, suggesting that the selection for chlorine substituents is a complex issue (54).


   CONCLUSIONS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
REFERENCES
 
The present crystal structure evidences significant differences between Rho 1,2-CCD and 1,2-CTDs. Several secondary structure deletions are observed, but in particular a number of changed residues inside the active cleft are believed to be responsible for substrate selection. The most significant should be Asp-52, Ala-53, Gly-76, Phe-78, and Cys-224, which seem to be directly involved in interactions with the differently substituted substrates. Few amino acid differences (F78Y and/or A53V) between the present enzyme, which exhibits a marked specificity for 4-substituted substrates, and a series of other CCDs more active with 3-substituted catechols, are believed to be responsible for the observed substrate selectivity differences among 1,2-CCDs.


   FOOTNOTES
 
The atomic coordinates and structure factors (code 1S9A [PDB] ) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).

* This work was supported by Grant ICA2-CT-2000-10006 from the European Commission Research and Technological Development Programme Copernicus, Contract HPRI-CT-1999-00017 from the European Community Access to Research Infrastructure Action of the Improving Human Potential Programme to the EMBL Hamburg Outstation, Cofin 2002 funding from the Italian Ministero Università e Ricerca Scientifica, and the Gruppo Nazionale di Ricerca per la Difesa dai Rischi Chimico-Industriali ed Ecologici, Consiglio Nazionale delle Ricerche.The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Dipartimento di Chimica, Università di Firenze, Via della Lastruccia 3, I-50019 Sesto Fiorentino FI, Italy. Tel.: 39-055-457-3343; Fax: 39-055-457-3333; E-mail: fabrizio.briganti{at}unifi.it.

1 The abbreviations used are: 3,4-PCD, protocatechuate 3,4-dioxygenase; 1,2-CCD, chlorocatechol 1,2-dioxygenase; 1,2-CTD, catechol 1,2-dioxygenase; MAD, multiple wavelength anomalous dispersion. Back


   ACKNOWLEDGMENTS
 
We gratefully acknowledge the technical skills of Dr. Samuele Ciattini.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 

  1. Ghosal, D., You, I.-S., Chatterjee, D. K., and Chakrabarty, A. M. (1985) Science 228, 135-142[Abstract/Free Full Text]
  2. Alexander, M. (1981) Science 211, 132-138[Abstract/Free Full Text]
  3. Haggblom, M. M., and Valo R. (1995) in Microbial Transformation and Degradation of Toxic Organic Chemicals (Young, L. Y., and Cerniglia, C., eds) pp. 389-439, John Wiley & Sons, Inc.
  4. Frantz, B., and Chakrabarty, A. M. (1986) in The Bacteria (Gunsalus, I, C., Sokatch, J. R., and Ornston, L. N., eds) Vol. 10, Academic Press, New York
  5. Reineke, W., and Knackmuss, H.-J. (1988) Annu. Rev. Microbiol. 42, 263-287[CrossRef][Medline] [Order article via Infotrieve]
  6. Chaudhry, G. O., and Chapalamadugu, S. (1991) Microbiol. Rev. 55, 59-79[Abstract/Free Full Text]
  7. van der Meer, J. R., van Neerven, A. R. W., de Vries, E. J., de Vos, W. M., and Zehnder, A. J. B. (1991) J. Bacteriol. 173, 6-15[Abstract/Free Full Text]
  8. Potrawfke, T., Armengaud, J., and Wittich, R.-M. (2001) J. Bacteriol. 183, 997-1011[Abstract/Free Full Text]
  9. Kaschabek, S. R., Kasberg, T., Muller, D., Mars, A. E., Janssen, D. B., and Reineke, W. (1998) J. Bacteriol. 180, 296-302[Abstract/Free Full Text]
  10. Tiedje, J. M., Duxbury, J. M., Alexander, M., and Dawson, J. E. (1969) J. Agric. Food Chem. 17, 1021-1026[CrossRef]
  11. Duxbury, J. M., Tiedje, J. M., Alexander, M., and Dawson, J. E. (1970) J. Agric. Food Chem. 18, 199-201[CrossRef][Medline] [Order article via Infotrieve]
  12. Apajalahti, J. H. A., and Salkinoja-Salonen, M. S. (1987) J. Bacteriol. 169, 5125-5130[Abstract/Free Full Text]
  13. Joshi, D. K., and Gold, M. H. (1993) Appl. Environ. Microbiol. 59, 1779-1785[Abstract/Free Full Text]
  14. Kozyreva, L. P., Shurukhin, Y. U., Finkel'shtein, Z. I., Baskunov, B. P., and Golovleva, L. A. (1993) Mikrobiologiya 62, 78-85
  15. Li, D.-Y., Eberspächer, J., Wagner, B., Kuntzer, J., and Lingens, F. (1991) Appl. Environ. 57, 1920-1928
  16. Sangodkar, M., Chapman, P. J., and Chakrabarty, A. M. (1988) Gene (Amst.) 71, 267-277[CrossRef][Medline] [Order article via Infotrieve]
  17. Sze, I. S., and Dagley, S. (1984) J. Bacteriol. 159, 353-359[Abstract/Free Full Text]
  18. Travkin, V. M., Jadan, A. P., Briganti, F., Scozzafava, A., and Golovleva, L. A. (1997) FEBS Lett. 407, 69-72[CrossRef][Medline] [Order article via Infotrieve]
  19. Bollag, J.-M., Briggs, G. G., Dawson, J. E., and Alexander, M. (1968) J. Agric. Food Chem. 16, 829-833[CrossRef]
  20. Luzzati, V. (1952) Acta Crystallogr. 5, 802-810[CrossRef]
  21. Mattews, B. W. (1968) J. Mol. Biol. 33, 491-497[Medline] [Order article via Infotrieve]
  22. Que, L., Jr., and Ho, R. Y. N. (1996) Chem. Rev. 96, 2607-2624[CrossRef][Medline] [Order article via Infotrieve]
  23. Hammer, A., Stolz, A., and Knackmuss, H. (1996) Arch. Microbiol. 166, 92-100[CrossRef][Medline] [Order article via Infotrieve]
  24. Nakai, C., Horiike, K., Kuramitsu, S., Kagamiyama, H., and Nozaki, M. (1990) J. Biol. Chem. 265, 660-665[Abstract/Free Full Text]
  25. Maltseva, O. V., Solyanikova, I. P., and Golovleva, L. A. (1991) Biochemistry (Mosc.) 56, 1548-1555
  26. Briganti, F., Pessione, E., Giunta, C., and Scozzafava, A. (1997) FEBS Lett. 416, 61-64[CrossRef][Medline] [Order article via Infotrieve]
  27. Briganti, F., Pessione, E., Giunta, C., Mazzoli, R., and Scozzafava, A. (2000) J. Protein Chem. 19, 709-716[Medline] [Order article via Infotrieve]
  28. Moisseeva, O. V., Linko, E. V., Baskunov, B. P., and Golovleva, L. A. (1999) Microbiology 68, 400-405
  29. Moisseeva, O. V., Belova, O. V., Solyanikova, I. P., Schlomann, M., and Golovleva, L. A. (2001) Biochemistry (Mosc.) 66, 548-555[CrossRef][Medline] [Order article via Infotrieve]
  30. Solyanikova, I. P., Maltseva, O. V., Vollmer, M. D., Golovleva, L. A., and Schlömann, M. (1995) J. Bacteriol. 177, 2821-2826[Abstract/Free Full Text]
  31. Gorlatov, S. N., Maltseva, O. V., Shevchenko, V. I., and Golovleva, L. A. (1989) Microbiology 58, 647-651
  32. Dorn, E., and Knackmuss, H.-J. (1978) Biochem. J. 174, 73-84[Medline] [Order article via Infotrieve]
  33. Dorn, E., and Knackmuss, H.-J. (1978) Biochem. J. 174, 85-94[Medline] [Order article via Infotrieve]
  34. Ngai, K.-L., and Ornston, L. N. (1988) J. Bacteriol. 170, 2412-2413[Abstract/Free Full Text]
  35. Pieper, D., Reineke, W., Engesser, K.-H., and Knackmuss, H.-J. (1988) Arch. Microbiol. 150, 95-102[CrossRef]
  36. Broderick, J. B., and O'Halloran, T. V. (1991) Biochemistry 30, 7349-7358[CrossRef][Medline] [Order article via Infotrieve]
  37. Solyanikova, I. P., Maltseva, O. V., and Golovleva, L. A. (1992) Biochemistry (Mosc.) 57, 1310-1316
  38. Hinteregger, C., Loidl, M., and Streichsbier, F. (1992) FEMS Microbiol. Lett. 97, 261-266[CrossRef]
  39. Bhat, M. A., Ishida, T., Horiike, K., Vaidyanathan, C. S., and Nozaki, M. (1993) Arch. Biochem. Biophys. 300, 738-746[CrossRef][Medline] [Order article via Infotrieve]
  40. Miguez, C. B., Greer, C. W., and Ingram, J. M. (1993) Can. J. Microbiol. 39, 1-5
  41. Maltseva, O. V., Solyanikova, I. P., and Golovleva, L. A. (1994) Eur. J. Biochem. 226, 1053-1061[Medline] [Order article via Infotrieve]
  42. Vetting, M. W, and Ohlendorf, D. H. (2000) Struct. Fold Des. 39, 429-440
  43. Ohlendorf, D. H., Lipscomb, J. D., and Weber, P. C. (1988) Nature 336, 403-405[CrossRef][Medline] [Order article via Infotrieve]
  44. Orville, A. M., Elango, N., Lipscomb, J. D., and Ohlendorf, D. H. (1997) Biochemistry 36, 10039-10051[CrossRef][Medline] [Order article via Infotrieve]
  45. Orville, A. M., Lipscomb, J. D., and Ohlendorf, D. H. (1997) Biochemistry 36, 10052-10066[CrossRef][Medline] [Order article via Infotrieve]
  46. Elgren, T. E., Orville, A. M., Kelly, K. A., Lipscomb, J. D., Ohlendorf, D. H., and Que, L., Jr. (1997) Biochemistry 36, 11504-11513[CrossRef][Medline] [Order article via Infotrieve]
  47. Vetting, M. W., D'Argenio, D. A., Ornston, L. N., and Ohlendorf, D. H. (2000) Biochemistry 39, 7943-7955[CrossRef][Medline] [Order article via Infotrieve]
  48. Briganti, F., Mangani, S., Pedocchi, L., Scozzafava, A., Golovleva, L. A., Jadan, A. P., and Solyanikova, I. P. (1998) FEBS Lett. 433, 58-62[CrossRef][Medline] [Order article via Infotrieve]
  49. Benvenuti, M., Briganti, F., Scozzafava, A., Golovleva, L., Travkin, V. M., and Mangani, S. (1999) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 901-903[CrossRef][Medline] [Order article via Infotrieve]
  50. Earhart, C. A., Radhakrishnan, R., Orville, A. M., Lipscomb, J. D., and Ohlendorf, D. H. (1994) J. Mol. Biol. 236, 374-376[CrossRef][Medline] [Order article via Infotrieve]
  51. Earhart, C. A., Hall, M. D., Michaud-Soret, I., Que, L., Jr., and Ohlendorf, D. H. (1994) J. Mol. Biol. 236, 377-378[CrossRef][Medline] [Order article via Infotrieve]
  52. Ludwig, M. L., Weber, L. D., and Ballou, D. P. (1984) J. Biol. Chem. 259, 14840-14842[Abstract/Free Full Text]
  53. Ferraroni, M., Ruiz Tarifa, M. Y., Briganti, F., Scozzafava, A., Mangani, S., Solyanikova, I. P., Kolomytseva, M. P., and Golovleva, L. A. (2002) Acta Crystallogr. Sect. D Biol. Crystallogr. 58, 1074-1076[CrossRef][Medline] [Order article via Infotrieve]
  54. Ferraroni, M., Ruiz Tarifa, M. Y., Scozzafava, A., Solyanikova, I. P., Kolomytseva, M. P., Golovleva, L., and Briganti, F. (2003) Acta Crystallogr. Sect. D Biol. Crystallogr. 59, 188-190[CrossRef][Medline] [Order article via Infotrieve]
  55. Terwillinger, T. C., and Berendzen, J. (1999) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 849-861[CrossRef][Medline] [Order article via Infotrieve]
  56. Otwinowski, Z., and Minor, W. (1997) Methods Enzymol. 276, 307-326
  57. Collaborative Computational Project 4 (1994) Acta Crystallogr. Sect. D Biol. Crystallogr. 50, 760-763[CrossRef][Medline] [Order article via Infotrieve]
  58. Accelrys Inc. (1997) QUANTA Software, Accelrys Inc., San Diego, CA
  59. Laskowski. R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993) J. Appl. Crystallogr. 26, 283-291[CrossRef]
  60. Kabsch, W., and Sander, C. (1983) Biopolymers 22, 2577-2637[CrossRef][Medline] [Order article via Infotrieve]
  61. Thompson, J. D., Gibson, T. J., Plewniak, F., Jeanmougin, F., and Higgins, D. G. (1997) Nucleic Acids Res. 24, 4876-4882
  62. Ritchie, D. W., and Kemp, G. J. L. (1999) J. Comp. Chem. 20, 383-395[CrossRef]
  63. McLachlan, A. D. (1982) Acta Crystallogr. Sect. A 38, 871-873[CrossRef]
  64. Nielsen, J. E., Andersen, K. V., Honig, B., Hooft, R. W., Klebe, G., Vriend, G., and Wade, R. C. (1999) Protein Eng. 12, 657-662[Abstract/Free Full Text]
  65. Bashford, D. (1997) in Scientific Computing in Object-oriented Parallel Environments (Ishikawa, Y., Oldehoeft, R. R., Reynders, J. V. W., and Tholburn, M., eds) pp. 233-240, Springer-Verlag, Berlin
  66. DeLano, W. L. (2002) The PyMOL Molecular Graphics System, DeLano Scientific, San Carlos, CA
  67. Huang, C. C., Couch, G. S., Pettersen, E. F., and Ferrin, T. E. (1996) Pacific Symp. Biocomputing 1, 724
  68. Sanner, M. F., Olson, A. J., and Spehner, J. C. (1996) Biopolymers 38, 305-320[CrossRef][Medline] [Order article via Infotrieve]
  69. Jang, H. G., Cox, D. D., and Que, L., Jr. (1991) J. Am. Chem. Soc. 113, 9200-9204[CrossRef]
  70. Eley, K. L., Crowley, P. J., and Bugg, T. D. H. (2001) J. Org. Chem. 66, 2091-2097[Medline] [Order article via Infotrieve]
  71. Potrawfke, T., Timmis, K. N., and Wittich, R.-M. (1998) Appl. Environ. Microbiol. 64, 3798-3806[Abstract/Free Full Text]

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