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Keywords:

  • adaptive evolution;
  • extradiol dioxygenase;
  • metagenome;
  • metal dependence

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Extradiol dioxygenase (EDO) catalyzes metal-dependent ring cleavage of catecholic substrates. We previously screened a metagenomic library of activated sludge used to treat industrial wastewater contaminated with phenols and cyanide to identify 43 EDO genes. Here, we have characterized the enzymes belonging to novel I.2.G, I.3.M and I.3.N subfamilies. The I.3.M and I.3.N EDOs were Fe(II) dependent and preferred bicyclic substrates, whereas the I.2.G EDOs were Mn(II) dependent, preferred monocyclic substrates and had the highest affinity for catechol reported thus far. The I.2.G EDOs were more tolerant against heat (60 °C for 1 h) and chemical inhibitors (H2O2 and NaCN) than I.3.M and I.3.N EDOs. Considering the dominance of the I.2.G EDOs over all retrieved EDOs (20 of 43 clones) and the presence of cyanide in the environment, this high affinity for substrate and structural robustness should provide survival advantages to host microorganisms. The 20 I.2.G EDOs were classified into six groups based on the amino acid sequence of the predicted ancestor, 1A1. Enzymes were chosen from each group and characterized. Two descendents, 1D2 and 5B2, each had a kcat/KM approximately twofold higher than that of 1A1 and reduced thermal stability, suggesting that descendents of 1A1 have adapted evolutionarily by a trade-off of inherent stability for increased activity.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Catechols are common intermediates in the aerobic microbial degradation of natural and xenobiotic aromatic compounds (Harayama et al., 1992; Furukawa et al., 2004). Extradiol dioxygenases (EDOs) are enzymes that play an important role in the catabolism of aromatic compounds (Zylstra & Gibson, 1989), cleaving the meta-position of the C–C bond of catechol to open up the aromatic ring. Microbial screenings revealed that EDOs are widespread in both gram-positive and -negative bacteria (Hirose et al., 1994; Eltis & Bolin, 1996; Junca et al., 2004), and further genetic analysis led to the classification of EDOs into two evolutionarily distinct families (Eltis & Bolin, 1996). Type-I is the major family, which is further divided into subfamilies depending on the amino acid sequence. Enzymes belonging to the same subfamily are defined as those that have >54% sequence identity. Among the type-I EDOs, I.2.A and I.3.A are the major subfamilies, which include catechol 2,3-dioxygenase and 2,3-dihydroxybiphenyl 1,2-dioxygenase, respectively. Despite differences in substrate specificity (i.e. monocyclic or bicyclic substrates), reactions with the substrate in both subfamilies take place at similar catalytic centres, both of which contain Fe(II) as a metal cofactor. Nine highly conserved amino acids exist in type-I EDOs (Eltis & Bolin, 1996), which include metal ligands (His153, His214 and Glu265), active site residues (His199, His246 and Tyr255) and residues indispensable for forming proper structure [Gly30, Leu172 and Pro259; residue numbers are those of catechol 2,3-dioxygenase XylE from TOL plasmid pWWO of Pseudomonas putida mt-2, Swiss-Prot P06622 (Inouye et al., 1981)]. In EDOs that prefer bicyclic substrates, the Asn–Asp sequence is conserved posterior to His246. Thus, type-I EDOs appear to have diverged from a common ancestor to adapt to environments contaminated with various aromatic compounds through accumulation of nucleotide substitutions (Eltis & Bolin, 1996). Nevertheless, how EDOs adapt to environments and the molecular basis for their evolution are still unclear.

Advances in microbial ecology have demonstrated that a large portion of environmental microorganisms (>99%) are not easily cultured in the laboratory by standard methods (Amann et al., 1995). Because uncultured bacteria are expected to include diverse microorganisms that are distantly related to cultured ones, we are interested in the exploration of catabolic genes for aromatic compounds using a different approach from the existing microbial screening methods. Previously, we used a culture-independent approach (Healy et al., 1995; Handelsman, 2004) to screen for EDO genes in a metagenomic library of microbial DNA from activated sludge that was used for industrial wastewater treatment (Suenaga et al., 2007). By screening 96 000 fosmid clones, 91 positive clones were obtained, and 38 clones were subjected to shotgun DNA sequencing. Forty-three EDO genes were identified, and 18 genes were grouped into the established subfamilies: I.2.A (eight clones), I.2.B (four clones) and I.2.C (six clones). Cell extracts prepared from Escherichia coli expressing these clones did not have substrate specificities distinct from the homologous EDOs (Suenaga et al., 2007). Therefore, we omitted further characterization of these EDOs in this study. In addition to these common enzymes, we also identified 25 EDO genes that could not be classified into the existing subfamilies. Therefore, four new subfamilies were proposed, and the EDO genes were sorted into I.1.C (two clones), I.2.G (20 clones, including three clones carrying incomplete genes), I.3.M (two clones) and I.3.N (one clone) subfamilies. In these novel EDOs, I.2.G subfamily genes were overrepresented. Note that these 20 genes were not identical; they were separated by single-nucleotide polymorphisms (SNPs) and were classified into six groups. Using the combinations of SNPs, a possible evolutionary lineage of the I.2.G clones was reconstructed as shown in Supporting Information, Fig. S1. We propose that these genes evolved from a common ancestor (group 1) and diverged via the accumulation of various nucleotide mutations (group 2 through group 6). Furthermore, we are interested in the reason for the dominance of the I.2.G EDOs in the retrieved clones and the role of the amino acid changes observed in the I.2.G enzymes.

In this study, we selected eight EDOs: six from I.2.G and one each from the I.3.M and I.3.N subfamilies. These enzymes were produced as recombinant proteins and purified to homogeneity, and then their physical (molecular mass, subunit structure and thermal stability) and catalytic (metal dependence, substrate specificity, kinetic constants and tolerance to chemical inhibitors) properties were investigated. Based on the results, we discuss the molecular basis for the evolution of these EDOs in the environment and focus especially on the I.2.G EDOs.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Bacterial strains, plasmids and media

Escherichia coli strains BL21(DE3) and BL21(DE3) pLysS and plasmids pET-28a(+) and pET-22b(+) were purchased from Novagen (Madison, WI). Escherichia coli cells were grown in Luria–Bertani (LB) or M9 medium in the presence of appropriate antibiotics (100 μg mL−1 ampicillin, 50 μg mL−1 kanamycin and 34 μg mL−1 chloramphenicol). Tryptone and yeast extract were from Difco (Detroit, MI).

Construction of expression plasmids

EDO genes were amplified by PCR, using fosmids as the template that were purified using a FosmidMAX DNA purification kit (Epicentre Biotechnologies, Madison, WI). PCR primers are listed in Table 1. The reaction mixture contained 1 × PCR buffer, 0.2 mM of each dNTP, 0.2 μM of each primer, 100 ng of template fosmid and 1.25 U of PrimeSTAR HS DNA polymerase (Takara Bio, Shiga, Japan) in a total volume of 50 μL. The mixture was subjected to 30 rounds of thermal cycling at 98 °C for 10 s, 55 °C for 5 s and 72 °C for 60 s. The amplicon was gel-purified, digested with restriction enzymes and inserted into appropriate sites of the pET-28a(+) plasmid; this fused the I.2.G subfamily EDOs with a 6 × His tag at the N-terminus. The pET-22b(+) plasmid for 7E11 and 1F2 was used to fuse a 6 × His tag to the C-terminus.

Table 1.   PCR primers used for cloning the EDO genes
EnzymeSequence*Restriction sites
  • *

    The sites of restriction endonucleases are underlined. The start and stop codons are given in bold.

I.2.G EDOs5′-TGGGAGGAACCATATGTCAAAACTCG-3′NdeI
5′-GTGCGCTCGAGGGTCACTCC-3′XhoI
7E11 EDO5′-GGAATTCCATATGGCAGCAGCGGTCAAGAGCTTG-3′NdeI
5′-AACCGGCTCGAGGCCGGCGGCCTGGGCC-3′XhoI
1F2 EDO5′-GGAATTCCATATGATCAATGCGTTGTCGTATCTCGGG-3′NdeI
5′-GGCCAAGCTTATCACGGTCGATGACCATTGCCG-3′HindIII

Metal dependence

Escherichia coli BL21(DE3), harbouring an expression plasmid, was grown at 37 °C in 3 mL of M9 medium containing kanamycin. When the OD600 nm reached 0.4, isopropyl-β-d-thiogalactopyranoside (IPTG) and Fe(NH4)2(SO4)2 or MnCl2 were added at a final concentration of 1 and 0.1 mM, respectively. After additional incubation for 24 h at 25 °C, the cells were harvested and disrupted by suspending in c. 1/20 volume of the initial cultivation volume of BugBuster Plus Benzonase nuclease (Novagen). After incubation at room temperature for 20 min, cell debris was removed by centrifugation at 20 000 g for 20 min at 4 °C. The supernatant was used to determine the metal dependence of activity at 30 °C in 170 μL of 10 mM sodium phosphate buffer (pH 7.5) containing 1 mM catechol or 1 mM 2,3-dihydroxybiphenyl.

Large-scale protein production and purification

For 7E11 and 1F2 EDOs, E. coli BL21(DE3) pLysS was used as a host. Transformants were grown at 37 °C in 100 mL of LB medium containing ampicillin and chloramphenicol until the OD600 nm reached 0.4. IPTG was added to the medium at a final concentration of 1 mM. Cells were grown for another 4 h at 25 °C and harvested by centrifugation at 4000 g for 10 min at 4 °C. For I.2.G subfamily EDOs, E. coli BL21(DE3) was used as a host. Transformants were grown at 37 °C in 100 mL of M9 medium containing kanamycin until the OD600 nm reached 0.4. IPTG and MnCl2 were then added to the medium at a final concentration of 1 and 0.1 mM, respectively. Cells were grown for another 24 h at 25 °C and harvested by centrifugation at 4000 g for 10 min at 4 °C. Cells were stored frozen at −80 °C until use.

The E. coli cell paste was thawed at room temperature, resuspended in 5 mL of 50 mM sodium phosphate buffer (pH 7.5) and disrupted by addition of c. 0.5 mL of BugBuster Plus Benzonase nuclease. After incubation at room temperature for 20 min, the cell debris was removed by centrifugation at 20 000 g for 20 min at 4 °C. The supernatant was loaded onto a Qiagen (Hilden, Germany) metal affinity chelating column (Ni-NTA Superflow, 1 mL), which was pre-equilibrated with 20 mM sodium phosphate buffer (pH 7.4) containing 0.5 M NaCl (buffer A). The column was washed with buffer A containing 25 mM imidazole, and the bound proteins were eluted with a linear gradient of imidazole from 25 to 500 mM in buffer A over 20 mL. Peak fractions contained virtually no contaminants and were combined, concentrated and buffer exchanged to 50 mM Tris-HCl (pH 7.5) using an Amicon (Bedford, MA) Ultra-15 centrifugation unit. Protein concentration was determined using a Quick Start Bradford Dye Reagent (Bio-Rad Laboratories, Hercules, CA) and bovine γ-globulin as the standard.

Molecular mass

The molecular mass of a subunit of the recombinant EDOs was determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The molecular mass of the native state EDO was estimated on a GE Healthcare (Little Chalfont, Buckinghamshire, UK) gel filtration column (Superose 6 10/300 GL, 1 × 30 cm) in 20 mM sodium phosphate buffer (pH 7.4) containing 0.5 M NaCl at a flow rate of 0.6 mL min−1. The molecular standards used were thyroglobulin (670 kDa), bovine γ-globulin (158 kDa), chicken ovalbumin (44 kDa), equine myoglobin (17 kDa) and vitamin B12 (1.35 kDa).

Enzyme activity

Enzyme reactions were carried out at 30 °C in 170 μL of 10 mM sodium phosphate buffer (pH 7.5) containing catechol or 2,3-dihydroxybiphenyl as a substrate. The activity was determined by monitoring the formation of reaction products (375 nm for the catechol reaction and 434 nm for the 2,3-dihydroxybiphenyl reaction) on a Molecular Devices (Sunnyvale, CA) microplate reader (VersaMax). The absorption coefficients of the ring-cleavage products were ɛ375=33 000 M−1 cm−1 for the catechol product and ɛ434=13 200 M−1 cm−1 for the 2,3-dihydroxybiphenyl product (Khan et al., 1996). One unit of enzyme activity is defined as the amount of enzyme that catalysed the formation of 1 μmol of the product per minute. Michaelis–Menten kinetics were applied to determine the kinetic constants.

Thermal inactivation

Thermal stability was determined by incubating purified enzymes at 60, 70 or 80 °C. At certain intervals, the remaining enzyme activities were measured at 30 °C as described above.

Inhibitor tolerance

Inhibitor tolerance was determined by incubating purified enzyme with 1 or 10 mM hydrogen peroxide (H2O2) or sodium cyanide (NaCN) in 10 mM sodium phosphate buffer (pH 7.5) at 4 °C. After 30 min, the remaining enzyme activities were measured at 30 °C as described above.

Homology modelling

A three-dimensional structure model of 1A1 EDO was constructed based on its homology with homoprotocatechuate 2,3-dioxygenase from Arthrobacter globiformis CM-2 [Protein Data Bank ID; 1F1R (Vetting et al., 2004)] as described previously (Suenaga et al., 2006). Sequence alignments and model construction and refinement were carried out using the homology modules of the molecular operating environment (moe) program (Chemical Computing Group, Quebec, ON, Canada).

Results and discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Metal dependence

The majority of known EDOs use Fe(II) as a cofactor, and only five EDOs have been reported that utilize metals other than Fe(II). The 3,4-dihydroxyphenylacetate 2,3-dioxygenase from Klebsiella pneumoniae uses Mg(II) (Gibello et al., 1994), and the other four enzymes use Mn(II). These are 3,4-dihydroxyphenylacetate 2,3-dioxygenase from Bacillus brevis (Que et al., 1981), 3,4-dihydroxyphenylacetate 2,3-dioxygenase from A. globiformis CM-2 (Viggiani et al., 2004), 2,3-dihydroxybiphenyl 1,2-dioxygenase from Bacillus sp. JF8 (Hatta et al., 2003) and 1,2-dihydroxynaphthalene dioxygenase from Bacillus sp. JF8 (Miyazawa et al., 2004).

First, we tested the metal dependence of the metagenomic EDOs. Recombinant E. coli was grown in M9 medium supplemented with Fe(II) or Mn(II). Activities of the crude extract were then assayed using catechol and 2,3-dihydroxybiphenyl as substrates (Table 2). Enzymes prepared in the absence of metal ions were inactive with both substrates (data not shown). Recombinant I.2.G EDOs displayed activity only when Mn(II) was added in the medium, and both catechol and 2,3-dihydroxybiphenyl served as a substrate with a slight preference for catechol. Recombinant 7E11 EDO also exhibited activity towards both catechol and 2,3-dihydroxybiphenyl, with a slight preference towards 2,3-dihydroxybiphenyl, which was Fe(II) dependent. Recombinant 1F2 EDO showed a relatively high activity with 2,3-dihydroxybiphenyl, which was also Fe(II) dependent. The activities of 1F2 EDO towards catechol and 2,3-dihydroxybiphenyl were lower than the values reported in our previous study (Suenaga et al., 2007). We believe that this was due to the different growth conditions used to prepare the cell extracts. In any case, the present results implied that I.2.G enzymes were Mn(II) dependent, while the 7E11 and 1F2 were Fe(II) dependent.

Table 2.   Metal ion dependence and substrate preferences of metagenomic EDOs
EnzymeSubfamilyGroup*Enzyme activity (U g−1 protein)
Catechol2,3-Dihydroxybiphenyl
Fe(II)Mn(II)Fe(II)Mn(II)
  • *

    The groups are defined in the Supporting Information (Fig. S1).

1A1I.2.G13.2121.91.0107.5
1D2I.2.G22.3123.91.867.3
7B2I.2.G30.2113.60.366.7
2B9I.2.G40.195.20.288.0
5B2I.2.G52.6113.41.686.6
2C1I.2.G61.1122.90.877.7
7E11I.3.M338.32.2444.939.0
1F2I.3.N0.50.710.30.4

Subunit and ternary structures

For protein purification, 7E11 and 1F2 EDOs were produced in LB medium. There was no need to add extra Fe(II) to the medium, probably because trace Fe(II) in the powdered medium was sufficient for incorporation into the recombinant proteins. Conversely, Mn(II) was required in M9 medium for recovering I.2.G recombinant enzymes; large amounts of insoluble recombinant proteins accumulated when produced in LB medium without supplementing with Mn(II). For convenient protein purification, a His-tag was attached to the termini. For 7E11 and 1F2 EDOs, the tag was fused to the C-termini; an N-terminal tag prohibited the binding of the enzyme to the column. For I.2.G enzymes, the tag was placed at the N-terminus. Proteins were purified to homogeneity using immobilized metal affinity column chromatography.

The molecular masses of the purified recombinant proteins were determined using sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The masses were 39 kDa for I.2.G enzymes and 35 kDa for 7E11 and 1F2 enzymes; these empirically determined masses agreed with the values calculated from the deduced amino acid sequences (Table 3). Furthermore, the molecular masses of the native structure of the enzymes were determined by gel filtration column chromatography. From the elution volumes, the I.2.G EDOs were estimated to be composed of four identical subunits. Moreover, 7E11 and 1F2 were estimated to be composed of eight identical subunits, similar to the known Fe(II)-dependent 2,3-dihydroxybiphenyl 1,2-dioxygenase, which includes enzymes from Rhodococcus jostii RHA1 (Hauschild et al., 1996), Pseudomonas putida OU83 (Khan et al., 1996), Burkholderia xenovorans LB400 (Han et al., 1995) and Pseudomonas pseudoalcaligenes KF707 (Furukawa et al., 1987).

Table 3.   Molecular masses of His-tagged EDOs
EnzymeSubfamilyGroup*Amino acid replacementLength (a.a.)Molecular mass (kDa)Subunit composition
CalculatedGel filtration
  • *

    The groups are defined in the Supporting Information (Fig. S1).

  • The difference in amino acids relative to 1A1.

1A1I.2.G134839.149134Homotetramer
1D2I.2.G2A241T34839.179139Homotetramer
7B2I.2.G3D82N34839.148139Homotetramer
2B9I.2.G4K253R34839.177140Homotetramer
5B2I.2.G5I114V, V136I34839.149144Homotetramer
2C1I.2.G6I114V, V136I, L290I34839.149135Homotetramer
7E11I.3.M32135.450269Homo-octamer
1F2I.3.N31734.493307Homo-octamer

Kinetic analysis

Kinetic constants were determined using purified enzymes. When catechol was used as a substrate, the activity increased concomitantly with the concentration of the substrate, consistent with Michaelis–Menten-type kinetic profiles. The kinetic constants, KM and kcat, were determined from Lineweaver–Burk plots (Table 4). All EDOs belonging to the I.2.G subfamily showed extremely high affinities for catechol with KM values <1 μM. The known EDOs exhibiting a preference for catechol have KM values ranging from 1 to 50 μM (Cerdan et al., 1995; Okuta et al., 2003; Junca et al., 2004; Viggiani et al., 2004). Thus, I.2.G EDOs have the lowest KM values reported thus far. With regard to turnover number, kcat, known EDOs have kcat values ranging from 1 to 500 s−1 (Cerdan et al., 1995; Okuta et al., 2003; Junca et al., 2004; Viggiani et al., 2004). As for the metagenomic clones, all of them had low turnover numbers relative to known EDOs. The overall catalytic efficiency, kcat/KM, of the I.2.G EDOs exhibited equivalent values to known EDOs, but slight differences were observed among clones. The 1D2 (group 2) and 5B2 (group 5) clones showed significantly higher activities due to a lowered KM and enhanced kcat values. The 7E11 and 1F2 EDOs exhibited much lower catalytic efficiencies (approximately two orders of magnitude), probably because they preferred 2,3-dihydroxybiphenyl to catechol (Table 2).

Table 4.   Kinetic parameters and specific activities of purified EDOs*
EnzymeSubfamilyGroupKM (μM)kcat (s−1)kcat/KM (μM−1 s−1)Relative activity to 2,3-dihydroxybiphenyl
  • *Values are expressed as average ± SD of three to five independent experiments.

  • The groups are defined in the Supporting Information (Fig. S1).

  • Activity to catechol.

  • §The specific activity of 1A1 was 0.246 U mg−1.

1A1I.2.G10.75 ± 0.150.36 ± 0.020.48 ± 0.071§
1D2I.2.G20.33 ± 0.020.38 ± 0.031.14 ± 0.601.29
7B2I.2.G30.64 ± 0.130.22 ± 0.020.34 ± 0.041.09
2B9I.2.G40.55 ± 0.170.34 ± 0.020.63 ± 0.191.58
5B2I.2.G50.46 ± 0.210.40 ± 0.030.85 ± 0.261.05
2C1I.2.G60.62 ± 0.560.32 ± 0.110.51 ± 0.191.35
7E11I.3.M45.9 ± 13.10.353 ± 0.068.0 × 10−3± 2.0 × 10−36.91
1F2I.3.N245 ± 781.00 ± 0.264.08 × 10−3± 0.22 × 10−33.01

As shown in Table 2, all EDOs used in this study exhibited activity towards 2,3-dihydroxybiphenyl. However, at substrate concentrations beyond a certain level, a reduction in the activity was observed. As for 1F2, the activity increased concomitantly with the concentration of 2,3-dihydroxybiphenyl, but began to decrease above 100 μM. As for 7E11 and I.2.G enzymes, maximal activities were obtained with 50–80 μM substrate concentrations. Substrate inhibition has also been observed in other EDOs, especially when 2,3-dihydroxybiphenyl was used (Adams et al., 1992; Heiss et al, 1995; Andújar et al., 2000). We therefore used substrate concentrations at which inhibition would be negligible. The kinetic constants for 1F2 were determined to be a KM of 25.4 μM, a kcat of 20.0 s−1 and a kcat/KM of 0.787 μM−1 s−1. The known 2,3-dihydroxybiphenyl 1,2-dioxygenases have a KM ranging from 1 to 50 μM (Hauschild et al., 1996; Khan et al., 1996; Schmid et al., 1997; Hatta et al., 2003); hence, 1F2 had a KM consistent with the reported values. The kcat/KM was comparable to those of I.2.G EDOs with catechol. As for I.2.G and 7E11 enzymes, we could not identify the appropriate conditions to determine kinetic constants. Therefore, we evaluated the reaction efficiencies from the relative activities at a fixed concentration (80 μM) of 2,3-dihydroxybiphenyl. Under this condition, 7E11 and 1F2 displayed higher activities with 2,3-dihydroxybiphenyl compared with I.2.G (Table 4), demonstrating a specificity for bicyclic substrates.

Thermostability

Thermostability was determined by measuring the remaining activities after incubating the enzymes at high temperatures (60, 70 and 80 °C). As shown in Fig. 1, 1F2 was the most thermolabile and lost activity at 60 °C within 10 min. The 7E11 enzyme was also thermolabile; incubation at 60 °C for 1 h decreased the activity to 41% of the preheated enzyme. In contrast, I.2.G enzymes were quite thermostable. Of the six I.2.G EDOs, four enzymes (1A1, 2C1, 5B2 and 7B2) retained almost full activity after heat treatment at 60 °C for 1 h. Of these, 1A1 and 7B2 were the most thermostable, retaining 25% and 17% of their activities, respectively, even after incubation at 70 °C for 1 h.

image

Figure 1.  Thermal inactivation profiles of metagenomic EDOs. Purified enzymes were incubated at 60°C (closed circle), 70°C (closed square) or 80°C (closed triangle). At various time intervals, aliquots were recovered, and the remaining activities were determined at 30°C.

Download figure to PowerPoint

Effects of chemical inhibitors

Table 5 shows the tolerance of EDOs against the chemical inhibitors H2O2 and NaCN. The oxidizing reagent H2O2 (10 mM) led to concentration-dependent inactivation, and reduced the activities to between 21% and 26% that of the initial activity for I.2.G EDOs; H2O2 decreased the activity of 7E11 to 13% of the initial value, and the activity of 1F2 to 16% of the preincubated enzyme. Differences in tolerance between I.2.G EDOs and the other enzymes were more significant at 1 mM concentrations. The metal-binding reagent NaCN selectively inhibited 7E11 and 1F2 by 65–68% at 10 mM. No loss of activity was observed for I.2.G enzymes with NaCN under identical conditions. From these results and the heat inactivation experiments, I.2.G EDOs were structurally more robust than 7E11 and 1F2 enzymes.

Table 5.   Effects of inhibitors on the activity of purified EDO*
EnzymeSubfamilyGroupRemaining activity (mM)
H2O2NaCN
110110
  • *

    Values are the average of three independent experiments.

  • The groups are defined in the Supporting Information (Fig. S1).

1A1I.2.G181 ± 621 ± 198 ± 198 ± 2
1D2I.2.G279 ± 526 ± 299 ± 1108 ± 11
7B2I.2.G370 ± 821 ± 495 ± 2113 ± 9
2B9I.2.G484 ± 626 ± 388 ± 8103 ± 12
5B2I.2.G559 ± 224 ± 397 ± 9112 ± 12
2C1I.2.G663 ± 1021 ± 497 ± 6113 ± 9
7E11I.3.M17 ± 413 ± 187 ± 431 ± 4
1F2I.3.N26 ± 116 ± 691 ± 1035 ± 6

Basis for the dominance of the I.2.G EDOs in the retrieved EDOs

As described previously, I.2.G EDOs were over-represented among the retrieved EDOs; 20 of 43 clones belonged to this novel subfamily. The I.2.G EDOs are characterized by their unique metal dependence. Unlike many other EDOs that are Fe(II) dependent, I.2.G EDOs preferred Mn(II). We believe that this is the primary reason for the dominance of the I.2.G EDOs. Coke-plant wastewater contains phenolic compounds and other toxic compounds, including NaCN (Chao et al., 2006; Chang et al., 2008). Although the actual concentration of NaCN in wastewater is unclear, coke-oven wastewater usually contains 0.5–3 mM NaCN (Chao et al., 2006; Chang et al., 2008). Under such conditions, the use of Mn(II) instead of Fe(II) should favour enzyme resiliency to NaCN stress.

The structural robustness of Mn(II)-dependent enzymes is supported by many studies. The Fe(II)-dependent EDO from Pseudomonas ovalis is rapidly inactivated by H2O2 and NaCN, but this is not the case with Mn(II)-dependent enzymes (Que et al., 1981; Whiting et al., 1996), and high thermostability of an Mn(II)-dependent EDO was reported by Hatta et al. (2003). Other Mn(II)-dependent enzymes also demonstrate a capacity to withstand stress. Mn(II)-dependent superoxide dismutase was shown to be more tolerant to NaCN and H2O2 than Fe(II)-dependent enzymes (Asada et al., 1975; Lumsden et al., 1976), and the thermostability of xylose isomerase from Bacillus licheniformis is most strikingly enhanced in the presence of Mn(II) compared with other metal ions such as Mg2+ and Co2+ (Vieille & Zeikus, 2001).

The I.2.G EDOs were further characterized by their high affinities with the substrate (Table 4). The KM values of I.2.G EDOs with catechol (<1 μM) were the lowest values reported so far. Microorganisms carrying the I.2.G EDO gene are thought to effectively utilize the aromatic compounds to advantage in activated sludge-treated coke-plant wastewater.

Adaptive evolution in I.2.G EDOs

Next, we assessed the capacity of I.2.G EDOs to evolve based on their individual enzymatic properties. The catalytic properties of EDO were reported to be influenced by single amino acid changes (Junca et al., 2004). Pseudomonad catechol 2,3-dioxygenase carrying a Tyr at amino acid position 218 spreads both at highly and slightly contaminated sites with benzene and toluene, which is consistent with a low turnover number and a high affinity for catechol. However, a variant carrying His at position 218 can be retrieved only from a site highly contaminated with the same compounds, which is consistent with a high turnover number and a low affinity for catechol. Collectively, this suggests that the enzymes were positively selected in the environment in response to pollutant conditions.

As for the metagenomic I.2.G EDOs, the catalytic activities varied from group to group due to apparently minor amino acid substitutions (Table 4). 1A1 and 7B2 had the highest thermostabilities, while 1D2 and 5B2 had significantly higher catalytic efficiencies compared with 1A1 and 7B2, which was accompanied by a reduction in stability (Table 4 and Fig. 1). Hence, an apparent trade-off exists between activity and stability. This could also be verified by the moderate character of both the catalytic activity and the thermostability in 2C1. Recently, Bloom et al. (2006) reported that cytochrome P450 BM3 mutants with higher stabilities had more chances to acquire new or improved functions via random mutagenesis. Based on this result, they concluded that protein stability promotes adaptive protein evolution. Similarly, in I.2.G EDO genes, the most thermostable ancestral 1A1 may have evolved towards more active 1D2 and 5B2 through 2C1 by sacrificing thermostability in the wastewater treatment plant operated at natural ambient temperatures. Note that I.2.G clones that acquired higher activities (groups 2 and 5) were more frequently discovered in the retrieved I.2.G clones (eight group 2 enzymes and three group 5 enzymes), which likely reflects the allele frequencies in the environment.

Finally, we attempted to unveil the mechanisms of activation or stability based on the three-dimensional structure. The three-dimensional structure of I.2.G EDO (1A1) was modelled using the homoprotocatechuate 2,3-dioxygenase from A. globiformis (MndD) as a template, which shared 30% amino acid sequence identity (Vetting et al., 2004) (Fig. 2). Based on the modelled structure, all the mutations (Asp82, Val136, Ala241, Lys253 and Leu290) were located on the surface of the protein far from the catalytic centre. Therefore, at present, we could not clarify the structural basis for enzyme evolution. However, as reported by Junca et al. (2004), Tyr/His218, which is located away from the active site, modulated the activity of catechol 2,3-dioxygenase to fit the surrounding environment. Similarly, the activity of the I.2.G subfamily enzymes appears to have been modulated by favourable mutations during evolution that allowed adaptation to their environment.

image

Figure 2.  Homology model of 1A1 EDO. The amino acids shown in red are presumed to be in the active site. Amino acid substitutions relative to 1A1 [i.e. A241T (identified in 1D2), D82N (2B9), K253R (2C1), I114V, V136Ie (5B2 and 7B2) and Leu290Ile (7B2)] are shown in green, except for Leu290Ile, which is invisible because the residue is buried in the molecule.

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Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

We thank Dr Masatoshi Goto (Kyushu University) for help with protein homology modelling. We also thank Dr Yoichi Kamagata (AIST) and Dr Yuji Nagata (Tohoku University) for critically reviewing the manuscript.

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  3. Introduction
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  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Table S1. Evolutionary lineage of EDOs in the I.2.G subfamily. The clone names, the groups and the frequencies of the retrieved EDO clones are indicated in the boxes. Amino acid substitutions are indicated by the arrows.

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FEM_719_sm_figS1.tiff41KSupporting info item

Please note: Wiley Blackwell is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.