Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Swetha, V. P. Articles by Phale, P. S. Search for Related Content PubMed PubMed Citation Articles by Swetha, V. P. Articles by Phale, P. S.

 Previous Article  |  Next Article 

Journal of Bacteriology, April 2007, p. 2660-2666, Vol. 189, No. 7
0021-9193/07/$08.00+0     doi:10.1128/JB.01418-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Purification and Characterization of 1-Naphthol-2-Hydroxylase from Carbaryl-Degrading Pseudomonas Strain C4

Vandana P. Swetha, Aditya Basu, and Prashant S. Phale^*

Biotechnology Group, School of Biosciences and Bioengineering, Indian Institute of Technology, Bombay, Powai, Mumbai 400 076, India

Received 7 September 2006/ Accepted 9 January 2007

Top Abstract Introduction Materials and Methods Results Discussion References

 
Pseudomonas sp. strain C4 metabolizes carbaryl (1-naphthyl-N-methylcarbamate) as the sole source of carbon and energy via 1-naphthol, 1,2-dihydroxynaphthalene, and gentisate. 1-Naphthol-2-hydroxylase (1-NH) was purified 9.1-fold to homogeneity from Pseudomonas sp. strain C4. Gel filtration and sodium dodecyl sulfate-polyacrylamide gel electrophoresis showed that the enzyme is a homodimer with a native molecular mass of 130 kDa and a subunit molecular mass of 66 kDa. The enzyme was yellow, with absorption maxima at 274, 375, and 445 nm, indicating a flavoprotein. High-performance liquid chromatography analysis of the flavin moiety extracted from 1-NH suggested the presence of flavin adenine dinucleotide (FAD). Based on the spectral properties and the molar extinction coefficient, it was determined that the enzyme contained 1.07 mol of FAD per mol of enzyme. Although the enzyme accepts electrons from NADH, it showed maximum activity with NADPH and had a pH optimum of 8.0. The kinetic constants K_m and V_max for 1-naphthol and NADPH were determined to be 9.6 and 34.2 µM and 9.5 and 5.1 µmol min^–1 mg^–1, respectively. At a higher concentration of 1-naphthol, the enzyme showed less activity, indicating substrate inhibition. The K_i for 1-naphthol was determined to be 79.8 µM. The enzyme showed maximum activity with 1-naphthol compared to 4-chloro-1-naphthol (62%) and 5-amino-1-naphthol (54%). However, it failed to act on 2-naphthol, substituted naphthalenes, and phenol derivatives. The enzyme utilized one mole of oxygen per mole of NADPH. Thin-layer chromatographic analysis showed the conversion of 1-naphthol to 1,2-dihydroxynaphthalene under aerobic conditions, but under anaerobic conditions, the enzyme failed to hydroxylate 1-naphthol. These results suggest that 1-NH belongs to the FAD-containing external flavin mono-oxygenase group of the oxidoreductase class of proteins.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Polyaromatic compounds are highly reduced, toxic, and recalcitrant in nature due to a resonance-stabilized benzene ring. However, microorganisms have evolved or adapted the ability to utilize polycyclic aromatic compounds as a sole source of carbon and energy. This is achieved by increasing the oxidation level of the compound followed by breaking the aromaticity by incorporating molecular oxygen, with this reaction being catalyzed by the oxygenase group of the oxidoreductase class of enzymes. Oxygenases are subgrouped into ring-hydroxylating mono-oxygenases and ring-cleaving or ring-hydroxylating dioxygenases (13, 16, 23-25). Hence, these enzymes are important in the metabolism of polyaromatic compounds and responsible for releasing the locked carbon from these pollutants. One such recalcitrant compound is 1-naphthol, a high-volume industrial product widely used in the production of synthetic dyes, perfumes, and pesticides such as carbaryl (Sevin) (8, 14). It is also released by microbes into the environment as a metabolic intermediate of various polycyclic aromatic compounds, including carbaryl (6, 9, 32, 36).

Carbaryl (1-naphthyl-N-methylcarbamate) is a pesticide used in the agriculture industry. The ester bond between N-methylcarbamic acid and 1-naphthol is responsible for its toxicity. In aqueous solutions, carbaryl hydrolyzes to 1-naphthol, methylamine, and CO₂ (41). A few microbes have been reported to transform 1-naphthol by hydroxylation to either 4-hydroxy-1-tetralone (5), 3,4-dihydro-dihydroxy-1(2H)-naphthalenone (42), or 1,4-naphthoquinone (33). Pseudomonas sp. strain C4, isolated in our laboratory from soil by an enrichment culture technique, utilizes carbaryl as the sole source of carbon and energy via 1-naphthol and 1,2-dihydroxynaphthalene, as shown in Fig. 1 (36). The involvement of 1,2-dihydroxynaphthalene as a metabolic intermediate was established by conducting a series of metabolic studies and demonstrating 1-naphthol-2-hydroxylase (1-NH) and 1,2-dihydroxynaphthalene dioxygenase activities in the cell extracts of carbaryl-grown cells (36). The enzyme 1-NH, responsible for the conversion of 1-naphthol to 1,2-dihydroxynaphthalene, was found to be inducible, requires NAD(P)H plus flavin adenine dinucleotide (FAD) for its activity, and has not been purified or characterized to date.

View larger version (13K): [in this window] [in a new window]

FIG. 1. Pathway for degradation of carbaryl by Pseudomonas sp. strain C4. The enzymes involved are as follows: 1, carbaryl hydrolase; 2, 1-naphthol-2-hydroxylase; 3, 1,2-dihydroxynaphthalene dioxygenase; and 4, gentisate dioxygenase.

In the present study, we report the purification to homogeneity of 1-naphthol-2-hydroxylase from Pseudomonas sp. strain C4, its biochemical characterization, and the kinetic properties of the enzyme.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Materials. FAD, flavin mononucleotide (FMN), NADH, NADPH, 1-naphthol, 1,2-dihydroxynaphthalene, EDTA, EGTA, 1,10-phenanthroline, 2,2'-dipyridyl, Q-Sepharose (fast flow), phenyl Sepharose (fast flow), Sephacryl S-200 HR, acrylamide, and bisacrylamide were purchased from Sigma-Aldrich. Carbaryl was a gift from Bayer (India) Ltd. All other chemicals were of analytical grade and were purchased locally.

Bacterial strain and culture conditions. Pseudomonas sp. strain C4 was grown in 150 ml minimal salt medium in 500-ml baffled Erlenmeyer flasks at 30°C on a rotary shaker at 200 rpm (36). The medium was supplemented aseptically with carbaryl (0.1%) as the sole source of carbon.

Enzyme assays. 1-NH was monitored either spectrophotometrically or polarographically. The rate of disappearance of NAD(P)H at 340 nm was measured spectrophotometrically (Lambda 35; Perkin-Elmer). The reaction mixture (1 ml) contained potassium phosphate buffer (50 mM, pH 7.5), FAD (6.25 µM), substrate (50 µM), NAD(P)H (300 µM), and an appropriate amount of enzyme. The enzyme activity was calculated using the molar extinction coefficient of NAD(P)H (₃₄₀ = 6,220 M^–1·cm^–1) and expressed as µmol of NAD(P)H oxidized per min. In the polarographic assay, rates of O₂ consumption were monitored at 30°C by using an oxygraph (Hansatech, United Kingdom) fitted with a Clark's O₂ electrode. The reaction mixture (2 ml) contained potassium phosphate buffer (50 mM, pH 7.5), substrate (50 µM), FAD (6.25 µM), NAD(P)H (300 µM), and an appropriate amount of enzyme. The enzyme activity was calculated as nmol of O₂ consumed per min. The specific activity is reported as µmol min^–1·mg^–1 protein. Protein estimation was performed as described by Bradford (7), using bovine serum albumin (BSA) as the standard.

Enzyme purification. 1-Naphthol-2-hydroxylase was purified from carbaryl-grown Pseudomonas sp. strain C4 cells to apparent homogeneity by using the following steps. All steps were performed at 4°C or on ice. During purification, the enzyme activity was monitored using NADH plus FAD as the cofactor.

(i) Preparation of cell extract. Cells grown on carbaryl (3.7 liters) for 13 h were harvested by centrifugation (10,000 x g) and washed twice with buffer A (potassium phosphate [50 mM, pH 7.5], glycerol [5%], EDTA [0.5 mM]). The cells (4.5 g) were suspended in ice-cold buffer A (1:4 [wt/vol]) and sonicated with an ultrasonic processor (GE130) on ice, with 20 cycles of 15 pulses each and an output of 11 W, with a 3- to 4-min interval. The cell homogenate was centrifuged at 40,000 x g for 30 min. The clear yellow supernatant obtained was referred to as the cell extract.

(ii) Q-Sepharose anion-exchange chromatography. The cell extract was loaded onto a Q-Sepharose column (10 x 150 mm) preequilibrated with buffer A, and the matrix was washed with the same buffer to remove unbound protein. The enzyme was eluted with an increasing linear gradient (150 ml) of 0 to 1 M ammonium sulfate in buffer A. Fractions (2 ml) were collected at a flow rate of 30 ml h^–1, using a fraction collector (Redifrac 920; GE Healthcare). The enzyme was eluted in the range of 0.2 to 0.4 M ammonium sulfate. The fractions with activities of >1 U were pooled and brought to 30% ammonium sulfate saturation by adding solid ammonium sulfate. The suspension was centrifuged at 40,000 x g for 30 min. The supernatant containing the enzyme was processed further.

(iii) Phenyl Sepharose hydrophobic interaction column chromatography. The enzyme from step ii was loaded onto a phenyl Sepharose column (10 x 150 mm) preequilibrated with 30% saturated ammonium sulfate in buffer A. Unbound protein was washed with the equilibration buffer. The bound enzyme was eluted using an increasing linear gradient (100 ml) of 0 to 60% ethylene glycol in buffer A. Fractions (1.5 ml) were collected at a flow rate of 30 ml h^–1. The active fractions were pooled and concentrated using ultrafiltration (membrane cutoff, 30 kDa; Pall or Millipore Amicon).

(iv) Gel filtration chromatography. The enzyme from step iii was loaded onto a Sephacryl S-200 HR gel filtration chromatography column (10 x 950 mm; bed volume, 76 ml; void volume, 33 ml) equilibrated in buffer A. The fractions (1 ml) were collected at a flow rate of 3 ml h^–1. Active fractions were pooled and concentrated. The purified enzyme was stored at 4°C.

Determination of molecular weight. The native molecular weight of the enzyme was determined by gel filtration on a Sephacryl S-200 HR column equilibrated with buffer A. The column was calibrated with ß-amylase (200 kDa), alcohol dehydrogenase (150 kDa), BSA (66 kDa), carbonic anhydrase (29 kDa), and cytochrome c (12.4 kDa). The molecular mass of 1-NH was determined from a plot of log (molecular mass) versus V_elution/V_void (V_e/V₀). The subunit composition and molecular weight were determined by discontinuous sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) with resolving (12%) and stacking (5%) gels as described by Laemmli (21). Electrophoresis was performed at a constant current of 8 mA along with standard molecular weight markers.

Spectroscopy. The UV-visible absorption spectrum of pure 1-NH was recorded in the range of 200 to 700 nm in buffer A (Lambda 35; Perkin-Elmer). The excitation and emission spectra of 1-NH were recorded using a fluorescence spectrometer (Jasco V-750).

NH₂-terminal sequencing. 1-NH from SDS-PAGE (12%) was electroblotted onto a polyvinylidene difluoride membrane (0.45 µm; Pall) in CAPS (3-[cyclohexylamino]-1-propanesulfonic acid) buffer (10 mM, pH 11) at 200 V for 8 h, stained with Coomassie brilliant blue R250, and subjected to automated Edman degradation (Applied Biosystems 470).

Identification of the prosthetic group. The flavin cofactor was extracted from the protein by treating 1-NH (1 mg in 60 µl) in buffer A with 10 µl of 70% perchloric acid on ice for 5 min, followed by centrifugation at 22,000 x g at 4°C (34). The supernatant was then subjected to high-performance liquid chromatography (HPLC) analysis (HP 1100 series; Agilent) with an RP-C₁₈ column (250 x 4 mm), using an isocratic solvent system consisting of 40% methanol and 60% 10 mM ortho-phosphoric acid (vol/vol) in water. The eluent was identified by comparison of the retention time and UV-visible spectrum to those of authentic FAD (retention time, 3.1 min) and FMN (retention time, 3.87 min).

Identification of the reaction product. The identification of the product was achieved by carrying out bulk enzyme reactions. The reaction mixture (10 ml) contained potassium phosphate buffer (50 mM, pH 7.5), 1-naphthol (50 µM), FAD (3.75 µM), NAD(P)H (50 µM), and an appropriate amount of enzyme. The reaction mixture was incubated for 1 h at 30°C. The substrate and cofactors were added intermittently at 15, 30, and 60 min. The reaction was terminated by adjusting the pH to 2 with HCl (2 N), and the products were extracted in ethyl acetate, concentrated, and dried over anhydrous sodium sulfate. The reaction products were resolved by thin-layer chromatography (TLC) as described earlier (36) and identified by comparing R_f and UV fluorescence properties with those of authentic 1-naphthol and 1,2-dihydroxynaphthalene.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Purification of 1-naphthol-2-hydroxylase. 1-Naphthol-2-hydroxylase was purified to homogeneity from carbaryl-grown cells of strain C4 by using anion-exchange (Q-Sepharose), hydrophobic interaction (phenyl Sepharose), and gel filtration (Sephacryl S-200 HR) column chromatography. The purification steps are summarized in Table 1. The enzyme was eluted as a single peak from the gel filtration chromatography column (Fig. 2A) and was bright yellow. 1-NH was purified 9.1-fold, with a 19% yield and a specific activity of 5.3 µmol min^–1·mg^–1 protein with NADH (the specific activity with NADPH was 6.1 µmol min^–1·mg^–1). SDS-PAGE analysis of the enzyme showed a single band with a molecular mass of 66 kDa (Fig. 2B). The native molecular mass, as determined by S-200 gel filtration chromatography, was found to be 130 kDa (Fig. 2A). The pure enzyme was found to be stable in buffer A and retained 90% activity when stored at 4°C for 90 days.

View this table: [in this window] [in a new window]

TABLE 1. Purification of 1-naphthol-2-hydroxylase from Pseudomonas sp. strain C4

View larger version (31K): [in this window] [in a new window]

FIG. 2. (A) Elution profile of 1-naphthol-2-hydroxylase from Sephacryl S-200 HR gel filtration column chromatography. The hydroxylase activity is represented by crosses, and protein elution is represented by filled circles. (Inset) Plot of log (molecular mass) versus Ve/V0 for gel filtration standard molecular mass protein markers, represented by open circles (alcohol dehydrogenase [150 kDa], BSA [66 kDa], carbonic anhydrase [29 kDa], and cytochrome c [12.4 kDa]). The filled circle represents 1-NH. OD280, optical density at 280 nm. (B) SDS-PAGE analysis of 1-NH during the different stages of purification. Lane 1, cell extract; lane 2, Q-Sepharose fraction; lane 3, phenyl Sepharose fraction; lane 4, Sephacryl S-200 HR fraction; lane 5, molecular size markers (phosphorylase B [97.4 kDa], BSA [66 kDa], ovalbumin [43 kDa], carbonic anhydrase [29 kDa], soybean trypsin inhibitor [20.1 kDa], and lysozyme [14.3 kDa]).

NH₂-terminal sequencing. The NH₂-terminal amino acid sequence of the purified 1-NH from strain C4 was determined to be MLKNIFLXDEIRXVSV by automated Edman degradation. A BLAST search performed with the N-terminal sequence failed to yield any significant similarity with existing proteins in the database (NCBI).

Spectral properties of 1-NH. The purified enzyme was yellow. The UV-visible spectrum of 1-NH (0.6 mg ml^–1) gave absorption maxima at 274, 375, and 445 nm (Fig. 3A). Above 320 nm, the absorption spectrum closely resembled that of authentic FAD (Fig. 3A, inset). The ratio of absorbance at 274 nm to that at 445 nm was found to be 13.1. The addition of NADPH or sodium dithionite led to the disappearance of the absorption maximum at 445 nm (Fig. 3A, inset). Excitation of 1-NH (0.81 mg ml^–1) at 450 nm showed an emission maximum at 529 nm, and a similar emission spectrum was observed for authentic FAD (Fig. 3B). The absorbance for 1-NH (5.7 mg ml^–1) at 445 nm was 0.54. Assuming a purity of 95%, a relative molecular mass of 130 kDa, and an absorption coefficient of 11,300 M^–1 cm^–1 for the FAD moiety (43), it was estimated that 1 mol of enzyme contained 1.07 mol of FAD.

View larger version (17K): [in this window] [in a new window]

FIG. 3. Spectral properties of 1-naphthol-2-hydroxylase from Pseudomonas sp. strain C4. (A) Absorption spectra of native 1-NH (0.6 mg ml–1) in buffer A (solid line), trichloroacetic acid-precipitated protein dissolved in buffer A (dashed line), and trichloroacetic acid supernatant (dotted line). (Inset) Magnified visible spectrum of oxidized 1-NH (0.6 mg ml–1 [solid line]), which disappeared when the enzyme was treated with NADPH (8 mM, dotted line) under anaerobic conditions or with a few milligrams of sodium dithionite (dashed-dotted-dashed line) or authentic FAD (100 µM [dashed line]). The lag between the addition of NADPH or sodium dithionite and recording of spectra was 15 s, and no absorbance was detected beyond 500 nm. (B) Fluorescence emission spectra of native 1-NH (0.81 mg ml–1 [solid line]) and authentic FAD (100 nM [dotted line]) excited at 445 nm in buffer A.

Cofactor requirement and apoenzyme preparation. The cofactor requirements for 1-NH are summarized in Table 2. The enzyme exhibited 45% more activity with NADPH plus FAD than with NADH plus FAD (Table 2). In the presence of NADPH and 1-naphthol, 1-NH activity increased 17% upon the addition of external FAD but decreased slightly (1%) upon the addition of external FMN. HPLC analysis of the extracted flavin moiety from 1-NH showed a retention time of 3.09 min, which closely corresponded to the retention time of authentic FAD (3.1 min). These results indicate the presence of FAD as the prosthetic group in 1-NH. In the absence of substrate, the enzyme did not oxidize NAD(P)H or consume oxygen, suggesting the absence of NAD(P)H oxidase activity (Table 2). Furthermore, in the presence of either NADPH or 1-naphthol, the enzyme did not produce hydrogen peroxide.

View this table: [in this window] [in a new window]

TABLE 2. Cofactor requirements for 1-naphthol-2-hydroxylase

Preparation of the apoenzyme (FAD-free protein) was attempted using various dialysis protocols. Purified 1-NH (2 mg ml^–1) was dialyzed against buffer A (four changes of 250 ml each [every 12 h]) containing KBr, urea, or ammonium sulfate at pH 6 or 7.5 for 36 to 48 h at 4°C. The enzyme dialyzed against KBr or ammonium sulfate at pH 6.0 led to an 65% or 30% loss of the activity, respectively, while dialysis against urea (2 and 4 M) led to an 90% loss in the activity. External addition of FAD or FMN to the assay mixture or dialysis against buffer A containing FAD failed to reconstitute the activity. Dialysis against ammonium sulfate at pH 7.5 retained 92% activity, with absorption at 274, 375, and 445 nm, comparable to that of the nondialyzed enzyme. Activity (100%) could be recovered after external addition of FAD. The enzyme (2 mg) treated with trichloroacetic acid (10%) lost its activity, with protein precipitation. The supernatant was pale yellow, with characteristic absorption maxima at 375 and 445 nm (Fig. 3A), while the precipitated protein in buffer A was colorless and showed no characteristic FAD absorbance spectrum (Fig. 3A).

pH optimum and effects of metal ions and metal chelators. The pH optimum of the 1-NH reaction was investigated using sodium citrate buffer (50 mM, pH 4.0 to 6.0), KH₂PO₄/K₂HPO₄ buffer (50 mM, pH 6.0 to 8.0), and Tris-HCl (50 mM, pH 7.5 to 9.0). The enzyme exhibited activity at pH 7 to 9, with maximal activity detected at pH 8.0. The activity in potassium phosphate buffer was twofold higher than that observed in Tris-HCl buffer.

Sulfate or chloride salts of the metal ions Zn²⁺, Mg²⁺, Mn²⁺, and Fe²⁺ at 0.1 mM did not show any effect on the enzyme activity. Cu²⁺ showed a marginal decrease (35%) in the activity at 0.1 mM. NaCl at 50 and 100 mM showed 50 and 67% inhibition, respectively. The metal chelators (2.5 mM) EDTA and EGTA did not inhibit the enzyme activity, but 1,10-phenanthroline and 2,2'-dipyridyl inhibited activity by 60%.

Substrate specificity. The activity of 1-NH was monitored spectrophotometrically as well as polarographically by using various mono- and diaromatic compounds (Table 3). Compared to the activity with 1-naphthol (100%), 1-NH exhibited activity with 4-chloro-1-naphthol (62%) and 5-amino-1-naphthol (54%). However, the enzyme failed to show any significant activity on carbaryl, naphthalene, substituted naphthalenes, phenol, or its derivatives (Table 3).

View this table: [in this window] [in a new window]

TABLE 3. Activity of 1-naphthol-2-hydroxylase on various substrates

Kinetic constants. The initial reaction velocities with various concentrations of either 1-naphthol, NADH, or NADPH were determined spectrophotometrically. A representative substrate saturation plot for 1-naphthol, using NADPH as an electron donor, is depicted in Fig. 4. Increasing the concentration of 1-naphthol showed a linear increase in the activity up to 20 µM. Further increases in the 1-naphthol concentration up to 100 and 200 µM showed 25 and 40% decreases in the activity, indicating substrate inhibition. Similar patterns of inhibition were observed with 4-chloro-1-naphthol and 5-amino-1-naphthol. The kinetic constants for 1-NH are summarized in Table 4. 1-NH showed a K_m of 9.6 µM for 1-naphthol in the presence of NADPH, compared to 10.05 µM in the presence of NADH. The inhibition constant K_i for 1-naphthol was determined to be 79.8 and 57.6 µM in the presence of NADPH and NADH, respectively. The enzyme showed K_m values of 34.2 and 118 µM for NADPH and NADH, respectively (Table 4), indicating that NADPH was the preferred coenzyme. 1-NH showed the highest k_cat and k_cat/K_m values for 1-naphthol plus NADPH compared to other combinations.

View larger version (15K): [in this window] [in a new window]

FIG. 4. Reaction velocity (v) versus [S] plot for 1-naphthol-2-hydroxylase. The reactions were performed with enzyme (5 µg), NADPH (200 µM), and FAD (6.25 µM), with various concentrations of 1-naphthol from 0.5 µM to 200 µM. The graph was fitted using the model for substrate inhibition (noncompetitive) with the following equation: v={Vmax/[1+(Km/S)+(S/Ki)]}

View this table: [in this window] [in a new window]

TABLE 4. Kinetic properties of 1-naphthol-2-hydroxylasea

Stoichiometry and reaction product identification. The stoichiometry of NAD(P)H to oxygen consumption was monitored by varying the substrate (1-naphthol) concentration. The enzyme was found to consume 1 mole of oxygen per mole of NAD(P)H. To identify the reaction product, the enzyme reaction was scaled up and performed under aerobic and anaerobic conditions, using Thunberg tubes and appropriate controls as described earlier (36). The reaction product was isolated and identified by TLC. Under aerobic conditions, two major spots, with R_f values of 0.10 (black quench when exposed to UV light) and 0.70 (brown-black quench when exposed to UV light), were observed on TLC, and these were identical to standard 1-naphthol (R_f, 0.71; brown-black quench) and 1,2-dihydroxynaphthalene (R_f, 0.11; black quench). Under anaerobic conditions, a spot corresponding to the substrate, 1-naphthol (R_f, 0.70; brown-black quench), was detected, indicating that 1-NH failed to transform 1-naphthol. These results suggest that 1-NH requires molecular oxygen to catalyze the hydroxylation of 1-naphthol to 1,2-dihydroxynaphthalene and can thus be classified as an oxygenase.

Top Abstract Introduction Materials and Methods Results Discussion References

 
1-Naphthol-2-hydroxylase is the second enzyme of the carbaryl metabolic pathway in Pseudomonas strain C4. The enzyme hydroxylates 1-naphthol at the ortho position to yield 1,2-dihydroxynaphthalene, which is further metabolized to Kreb's cycle intermediates through a series of enzymatic reactions. The enzyme is inducible and showed maximum activity when cells were grown on carbaryl (36). The purification and characterization of the enzyme responsible for ortho hydroxylation of 1-naphthol to 1,2-dihydroxynaphthalene have not been reported. Here we describe the purification and characterization of 1-NH from Pseudomonas sp. strain C4.

1-Naphthol-2-hydroxylase was purified to homogeneity using conventional column chromatographic techniques. The enzyme is a homodimer with a native molecular mass of 130 kDa and a subunit molecular mass of 66 kDa. 1-NH showed optimum activity at pH 8.0. Besides 274 nm, the pure enzyme showed absorption peaks at 375 and 445 nm, similar to other reported flavin hydroxylases. Aromatic flavin hydroxylases have been identified as either single- or two-component enzymes (3, 26-28, 38). These enzymes are known to initiate the biodegradation of several aromatic contaminants in the environment (17, 25, 27). Enzymes such as orcinol hydroxylase, 2,4-dichlorophenol hydroxylase, hydroquinone hydroxylase, and 2-hydroxybiphenyl-3-monooxygenase are single-component, either monomeric, homodimeric, or homotetrameric flavin monooxygenases and accept electrons from external electron donors, including NADPH and NADH (4, 12, 30, 35). Phenol hydroxylase and p-hydroxyphenylacetate-3-hydroxylase are two-component enzymes consisting of the catalytic oxygenase component, responsible for hydroxylation of the substrate, and the reductase component, responsible for electron transfer from NAD(P)H to the oxygenase component (1, 2, 20, 31). The biochemical properties of 1-NH provide compelling evidence that the enzyme is a single-component external flavin hydroxylase.

Spectral analysis indicates that 1-NH contains 1 mole of FAD per mole of protein. Preparation of apoenzyme by dialyzing the enzyme against urea, KBr, or acid ammonium sulfate (pH 6) resulted in the complete or partial loss of the enzyme's activity, with a loss in the characteristic flavin absorption spectra. External addition of FAD or FMN failed to reactivate the enzyme. This could be due to partial denaturation or subtle changes in the protein structure leading to loss of the flavin moiety and the enzyme activity. Upon dialysis against ammonium sulfate in buffer A at pH 7.5, 1-NH retained 90% activity, and external addition of FAD led to 100% recovery. The enzyme exhibited significantly higher activity with FAD than with FMN. These observations were further supported by the identification of FAD by HPLC after treating 1-NH with perchloric acid. The gel filtration profile, dialysis experiments, and identification of FAD as the prosthetic group indicate that the flavin moiety is tightly but not covalently bound to the protein and that its removal leads to a loss of enzyme activity. Some aromatic hydroxylases accept electrons exclusively from NADPH, while others also utilize NADH for the reduction of flavin (37). Flavin hydroxylases such as orcinol hydroxylase, 2,4-dichlorophenol hydroxylase, and 3-hydroxyphenylacetate 6-hydroxylase have been reported to accept electrons from NADPH as well as from NADH (4, 30, 39). 1-NH could accept electrons from both NADPH and NADH, but the enzyme exhibited significantly higher activity with NADPH (145%). The K_m for NADPH was determined to be 34.2 µM, compared to 118 µM for NADH, suggesting that the enzyme has a high affinity for NADPH. The addition of various chloride and sulfate salts of Fe, Zn, Mg, and Mn or of metal chelators, such as EDTA and EGTA, did not show any effect on the 1-NH activity, suggesting that the external addition of metal ions is not required. Inhibition with 1,10-phenanthroline and 2,2'-dipyridyl (60% at 2.5 mM) could be due to a nonspecific hydrophobic interaction between the chelator and 1-NH.

Flavin monooxygenases have been reported to accept different substrates and substrate analogs, thus exhibiting broad substrate specificity (4, 12, 22, 35, 44). 1-NH showed activity with 1-naphthol and partial activity with 4-chloro-1-naphthol and 5-amino-1-naphthol. It failed to show activity with phenol, 3-hydroxybenzoate, and 4-hydroxybenzoate, suggesting that 1-NH is a new enzyme and is different from previously reported phenol or hydroxybenzoate hydroxylases. The inability of 1-NH to accept various other hydroxylated aromatic substrates suggests that the enzyme exhibits a limited substrate range. This property has previously been reported for other flavin hydroxylases (15, 18, 39). 1-NH showed a K_m of 9.6 µM for 1-naphthol. The affinity for 1-naphthol (K_m) remained unchanged irrespective of the electron donor used. However, the k_cat value was higher in the presence of NADPH (20.58 s^–1). An increase in the 1-naphthol concentration above 20 µM led to a decrease in the activity, indicating the substrate inhibition of the enzyme. Similar inhibition patterns were observed with 4-chloro-1-naphthol and 5-amino-1-naphthol. It has been reported that flavin hydroxylases are subject to substrate or substrate analog inhibition, a commonly observed phenomenon (10, 11, 19, 29, 40). 1-NH showed a K_i of 79.8 µM in the presence of NADPH, compared to 57.6 µM in the presence of NADH. A comparison of kinetic constants for 1-naphthol plus NADPH and 1-naphthol plus NADH clearly indicated that NADPH is a preferred coenzyme over NADH and that the enzyme is comparatively less sensitive to higher concentrations of substrate. The detection of 1,2-dihydroxynaphthalene by TLC under aerobic conditions and the consumption of 1 mol of molecular oxygen per mole of NAD(P)H suggest that the enzyme belongs to the oxygenase group of the oxidoreductase class of proteins.

In conclusion, 1-naphthol-2-hydroxylase, responsible for the conversion of 1-naphthol to 1,2-dihydroxynaphthalene, was purified to homogeneity from carbaryl-degrading Pseudomonas sp. strain C4. The enzyme is a homodimer, contains FAD as a prosthetic group, prefers NADPH as a cofactor, acts on 1-naphthol, and consumes 1 mole of O₂ per mole of NADPH. These results suggest that 1-NH belongs to the FAD-containing external flavin monooxygenase group of the oxidoreductase class of proteins. The results presented in this paper are a good basis for further characterization of the enzyme with respect to the events taking place during the catalytic cycle at the active site and the structural features determining the substrate specificity of the enzyme.

 
We thank N. Appaji Rao, Biochemistry Department, IISc, Bangalore, India, for constructive discussions.

This work was supported by the Department of Biotechnology, Government of India. A senior research fellowship to A.B. from IIT-B is gratefully acknowledged.

 
* Corresponding author. Mailing address: Biotechnology Group, School of Biosciences and Bioengineering, Indian Institute of Technology, Bombay, Powai, Mumbai 400 076, India. Phone: 91-22-2576 7836. Fax: 91-22-2572 3480. E-mail: pphale{at}iitb.ac.in.

Published ahead of print on 19 January 2007.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Arunachalam, U., and V. Massey. 1994. Studies on the oxidative half-reaction of p-hydroxyphenylacetate 3-hydroxylase. J. Biol. Chem. 269:11795-11801.[Abstract/Free Full Text]
2
2. Arunachalam, U., V. Massey, and S. M. Miller. 1994. Mechanism of p-hydroxyphenylacetate-3-hydroxylase. A two-protein enzyme. J. Biol. Chem. 269:150-155.[Abstract/Free Full Text]
3
3. Ballou, D. P., B. Entsch, and L. J. Cole. 2005. Dynamics involved in catalysis by single-component and two-component flavin-dependent aromatic hydroxylases. Biochem. Biophys. Res. Commun. 338:590-598.[CrossRef][Medline]
4
4. Beadle, C. A., and A. R. Smith. 1982. The purification and properties of 2,4-dichlorophenol hydroxylase from a strain of Acinetobacter species. Eur. J. Biochem. 123:323-332.[Medline]
5
5. Bollag, J. M., E. J. Czaplicki, and R. D. Minard. 1975. Bacterial metabolism of 1-naphthol. J. Agric. Food Chem. 23:85-90.[CrossRef][Medline]
6
6. Boyd, C., M. J. Larkin, K. A. Reid, N. D. Sharma, and K. Wilson. 1997. Metabolism of naphthalene, 1-naphthol, indene, and indole by Rhodococcus sp. strain NCIMB 12038. Appl. Environ. Microbiol. 63:151-155.[Abstract]
7
7. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254.[CrossRef][Medline]
8
8. Brahmia, O., and C. Richard. 2005. Photochemical transformation of 1-naphthol in aerated aqueous solution. Photochem. Photobiol. Sci. 4:454-458.[CrossRef][Medline]
9
9. Cerniglia, C. E., and D. T. Gibson. 1977. Metabolism of naphthalene by Cunninghamella elegans. Appl. Environ. Microbiol. 34:363-370.[Abstract/Free Full Text]
10
10. Detmer, K., and V. Massey. 1984. Effect of monovalent anions on the mechanism of phenol hydroxylase. J. Biol. Chem. 259:11265-11272.[Abstract/Free Full Text]
11
11. Detmer, K., and V. Massey. 1985. Effect of substrate and pH on the oxidative half-reaction of phenol hydroxylase. J. Biol. Chem. 260:5998-6005.[Abstract/Free Full Text]
12
12. Eppink, M. H., E. Cammaart, D. Van Wassenaar, W. J. Middelhoven, and W. J. van Berkel. 2000. Purification and properties of hydroquinone hydroxylase, a FAD-dependent monooxygenase involved in the catabolism of 4-hydroxybenzoate in Candida parapsilosis CBS604. Eur. J. Biochem. 267:6832-6840.[Medline]
13
13. Gibson, D. T., and R. E. Parales. 2000. Aromatic hydrocarbon dioxygenases in environmental biotechnology. Curr. Opin. Biotechnol. 11:236-243.[CrossRef][Medline]
14
14. Gosselin, R. E., H. C. Hodge, R. P. Smith, and M. N. Gleason. 1976. Clinical toxicology of commercial products. Williams and Wilkins, Baltimore, MD.
15
15. Haigler, B. E., W. C. Suen, and J. C. Spain. 1996. Purification and sequence analysis of 4-methyl-5-nitrocatechol oxygenase from Burkholderia sp. strain DNT. J. Bacteriol. 178:6019-6024.[Abstract/Free Full Text]
16
16. Hayaishi, O., M. Katagiri, and S. Rothberg. 1955. Mechanism of the pyrocatechase reaction. J. Am. Chem. Soc. 77:5450-5451.
17
17. Hodgson, E., and P. E. Levi. 1996. Pesticides: an important but underused model for the environmental health sciences. Environ. Health Perspect. 104(Suppl. 1):97-106.
18
18. Hosokawa, K., and R. Y. Stanier. 1966. Crystallization and properties of p-hydroxybenzoate hydroxylase from Pseudomonas putida. J. Biol. Chem. 241:2453-2460.[Abstract/Free Full Text]
19
19. Husain, M., B. Entsch, D. P. Ballou, V. Massey, and P. J. Chapman. 1980. Fluoride elimination from substrates in hydroxylation reactions catalyzed by p-hydroxybenzoate hydroxylase. J. Biol. Chem. 255:4189-4197.[Abstract/Free Full Text]
20
20. Kirchner, U., A. H. Westphal, R. Muller, and W. J. van Berkel. 2003. Phenol hydroxylase from Bacillus thermoglucosidasius A7, a two-protein component monooxygenase with a dual role for FAD. J. Biol. Chem. 278:47545-47553.[Abstract/Free Full Text]
21
21. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.[CrossRef][Medline]
22
22. Locher, H. H., T. Leisinger, and A. M. Cook. 1991. 4-Toluene sulfonate methyl-monooxygenase from Comamonas testosteroni T-2: purification and some properties of the oxygenase component. J. Bacteriol. 173:3741-3748.[Abstract/Free Full Text]
23
23. Malmstrom, B. G. 1982. Enzymology of oxygen. Annu. Rev. Biochem. 51:21-59.[CrossRef][Medline]
24
24. Mason, H. S., W. L. Folks, and L. Peterson. 1955. Oxygen transfer and electron transport by the phenolase complex. J. Am. Chem. Soc. 77:2914-2915.
25
25. Mishra, V., R. Lal, and Srinivasan. 2001. Enzymes and operons mediating xenobiotic degradation in bacteria. Crit. Rev. Microbiol. 27:133-166.[CrossRef][Medline]
26
26. Miura, R. 2001. Versatility and specificity in flavoenzymes: control mechanisms of flavin reactivity. Chem. Rec. 1:183-194.[CrossRef][Medline]
27
27. Moonen, M. J. H., M. W. Fraaije, I. M. C. M. Rietjens, C. Laane, and W. J. H. van Berkel. 2006. Flavoenzymes-catalyzed oxygenations and oxidations of phenolic compounds. Adv. Synth. Catal. 344:1023-1035.
28
28. Nam, J. W., H. Nojiri, T. Yoshida, H. Habe, H. Yamane, and T. Omori. 2001. New classification system for oxygenase components involved in ring-hydroxylating oxygenations. Biosci. Biotechnol. Biochem. 65:254-263.[CrossRef][Medline]
29
29. Neujahr, H. Y. 1991. Phenol hydroxylase, p. 65-85. In F. Muller (ed.), Chemistry and biochemistry of flavo enzymes, vol. 2. CRC Press, Boca Raton, FL.
30
30. Ohta, Y., and D. W. Ribbons. 1976. Bacterial metabolism of resorcinylic compounds: purification and properties of orcinol hydroxylase and resorcinol hydroxylase from Pseudomonas putida ORC. Eur. J. Biochem. 61:259-269.[Medline]
31
31. Pessione, E., S. Divari, E. Griva, M. Cavaletto, G. L. Rossi, G. Gilardi, and C. Giunta. 1999. Phenol hydroxylase from Acinetobacter radioresistens is a multicomponent enzyme. Purification and characterization of the reductase moiety. Eur. J. Biochem. 265:549-555.[Medline]
32
32. Prabhu, Y., and P. S. Phale. 2003. Biodegradation of phenanthrene by Pseudomonas sp. strain PP2: novel metabolic pathway, role of biosurfactant and cell surface hydrophobicity in hydrocarbon assimilation. Appl. Microbiol. Biotechnol. 61:342-351.[Medline]
33
33. Rajagopal, B. S., V. R. Rao, G. Nagendraappa, and N. Sethunathan. 1984. Metabolism of carbaryl and carbofuran by soil enrichment and bacterial cultures. Can. J. Microbiol. 30:1458-1466.
34
34. Rao, N. A., S. P. Felton, F. M. Huennekens, and B. Mackler. 1963. Flavin mononucleotide: the coenzyme of reduced diphosphopyridine nucleotide dehydrogenase. J. Biol. Chem. 238:449-455.[Free Full Text]
35
35. Suske, W. A., M. Held, A. Schmid, T. Fleischmann, M. G. Wubbolts, and H. P. Kohler. 1997. Purification and characterization of 2-hydroxybiphenyl 3-monooxygenase, a novel NADH-dependent, FAD-containing aromatic hydroxylase from Pseudomonas azelaica HBP1. J. Biol. Chem. 272:24257-24265.[Abstract/Free Full Text]
36
36. Swetha, V. P., and P. S. Phale. 2005. Metabolism of carbaryl via 1,2-dihydroxynaphthalene by soil isolates Pseudomonas sp. strains C4, C5, and C6. Appl. Environ. Microbiol. 71:5951-5956.[Abstract/Free Full Text]
37
37. van Berkel, W. J., and F. Muller. 1991. Flavin-dependent monooxygenases with special reference to p-hydroxybenzoate hydroxylase, p. 1-29. In F. Muller (ed.), Chemistry and biochemistry of flavo enzymes, vol. 2. CRC Press, Inc., Boca Raton, FL.
38
38. van Berkel, W. J., N. M. Kamerbeek, and M. W. Fraaije. 2006. Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J. Biotechnol. 124:670-689.[CrossRef][Medline]
39
39. van Berkel, W. J., and W. J. Van Den Tweel. 1991. Purification and characterisation of 3-hydroxyphenylacetate 6-hydroxylase: a novel FAD-dependent monooxygenase from a Flavobacterium species. Eur. J. Biochem. 201:585-592.[Medline]
40
40. van der Bolt, F. J., R. H. van den Heuvel, J. Vervoort, and W. J. van Berkel. 1997. 19F NMR study on the regiospecificity of hydroxylation of tetrafluoro-4-hydroxybenzoate by wild-type and Y385F p-hydroxybenzoate hydroxylase: evidence for a consecutive oxygenolytic dehalogenation mechanism. Biochemistry 36:14192-14201.[CrossRef][Medline]
41
41. Vontor, T. S. J., and M. Vecera. 1972. Kinetics and mechanism of hydrolysis of 1-naphthyl,N-methyl carbamate and NN-dimethyl carbamates. Collect. Czech. Chem. Commun. 37:2183-2196.
42
42. Walker, N., N. F. Janes, J. R. Spokes, and P. van Berkum. 1975. Degradation of 1-naphthol by a soil pseudomonad. J. Appl. Bacteriol. 39:281-286.[Medline]
43
43. Whitby, L. G. 1953. A new method for preparing flavin-adenine dinucleotide. Biochem. J. 54:437-442.[Medline]
44
44. Xun, L., E. Topp, and C. S. Orser. 1992. Diverse substrate range of a Flavobacterium pentachlorophenol hydroxylase and reaction stoichiometries. J. Bacteriol. 174:2898-2902.[Abstract/Free Full Text]

---

Journal of Bacteriology, April 2007, p. 2660-2666, Vol. 189, No. 7
0021-9193/07/$08.00+0     doi:10.1128/JB.01418-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via Google Scholar Google Scholar Articles by Swetha, V. P. Articles by Phale, P. S. Search for Related Content PubMed PubMed Citation Articles by Swetha, V. P. Articles by Phale, P. S.