Degradation of Tetrahydrofurfuryl Alcohol by Ralstonia eutropha Is ...

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dent aldehyde dehydrogenase. Polyethylene glycol (PEG), a xenobiotic compound, contains an ether in addition to the alcohol function, as in THFA. Dye-linked ...
APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Dec. 1997, p. 4891–4898 0099-2240/97/$04.0010 Copyright © 1997, American Society for Microbiology

Vol. 63, No. 12

Degradation of Tetrahydrofurfuryl Alcohol by Ralstonia eutropha Is Initiated by an Inducible Pyrroloquinoline Quinone-Dependent Alcohol Dehydrogenase ¨ DER, GRIT ZARNT, THOMAS SCHRA

AND

JAN R. ANDREESEN*

Institut fu ¨r Mikrobiologie, Martin-Luther-Universita ¨t Halle, Halle, Germany Received 2 June 1997/Accepted 7 October 1997

An organism tentatively identified as Ralstonia eutropha was isolated from enrichment cultures containing tetrahydrofurfuryl alcohol (THFA) as the sole source of carbon and energy. The strain was able to tolerate up to 200 mM THFA in mineral salt medium. The degradation was initiated by an inducible ferricyanidedependent alcohol dehydrogenase (ADH) which was detected in the soluble fraction of cell extracts. The enzyme catalyzed the oxidation of THFA to the corresponding tetrahydrofuran-2-carboxylic acid. Studies with npentanol as the substrate revealed that the corresponding aldehyde was released as a free intermediate. The enzyme was purified 211-fold to apparent homogeneity and could be identified as a quinohemoprotein containing one pyrroloquinoline quinone and one covalently bound heme c per monomer. It was a monomer of 73 kDa and had an isoelectric point of 9.1. A broad substrate spectrum was obtained for the enzyme, which converted different primary alcohols, starting from C2 compounds, secondary alcohols, diols, polyethylene glycol 6000, and aldehydes, including formaldehyde. A sequence identity of 65% with a quinohemoprotein ADH from Comamonas testosteroni was found by comparing 36 N-terminal amino acids. The ferricyanide-dependent ADH activity was induced during growth on different alcohols except ethanol. In addition to this activity, an NAD-dependent ADH was present depending on the alcohol used as the carbon source. oxidize PEG to its corresponding carboxylic acid (21, 47). The first enzyme of this kind was a ferricyanide-dependent ADH characterized as a quinohemoprotein isolated from Comamonas testosteroni as apoenzyme that contains pyrroloquinoline quinone (PQQ) and heme c in the holoenzyme (14). A similar quinohemoprotein that also converted PEG was isolated from Rhodopseudomonas acidophila (47). A strain of P. putida was found to be able to induce three distinct PQQ-dependent ADHs during growth on different alcohols (40). Ralstonia eutropha (formerly Alcaligenes eutrophus) strains were previously shown to induce different NAD(P)-dependent ADHs when grown on different alcohols (29, 33, 46). However, no ferricyanide-dependent ADH activity was reported for that organism. We now report that a strain identified as R. eutropha was able to grow on THFA and that the degradation of THFA was initiated by an inducible quinohemoprotein ADH designated tetrahydrofurfuryl alcohol dehydrogenase (THFA-DH). The enzyme was purified to apparent homogeneity, and the basic properties, showing some deviations from those of quinohemoprotein ADHs previously isolated (10, 40, 47), were determined.

Tetrahydrofurfuryl alcohol (THFA) is a xenobiotic cyclic ether which is frequently used as an organic solvent in industry. Furthermore, THFA is released in large amounts, e.g., during synthesis of different industrial products such as furan resins. However, no information is available on the bacterial degradation of this compound. THFA is a poor substrate for the tetrahydrofuran (THF)-degrading Rhodococcus ruber 219 (5). Microorganisms screened for utilization of several O-heterocyclic compounds, like furfuryl alcohol, furfural, furan-2-carboxylate, or THF, were not able to grow on THFA as the sole source of carbon and energy (24, 44). Many reports about bacteria able to degrade heterocyclic compounds exist. The initial reaction can be catalyzed by quite different classes of enzymes (1, 5, 6, 12, 20). In the case of THF, a monooxygenase reaction catalyzing an initial hydroxylation adjacent to the heteroatom has been proposed (8). A monooxygenase was also responsible for initial hydroxylation of the electron-rich compound pyrrole-2-carboxylate (17). Furthermore, molybdenum-containing dehydrogenases play an important role in the degradation of aromatic heterocyclic compounds, such as furan-2-carboxylate, nicotinate, or isonicotinate (23, 25, 32, 37). In the case of the degradation of furfuryl alcohol by Pseudomonas putida, the oxidation of furfuryl alcohol to furfural is initiated by an NAD-dependent alcohol dehydrogenase (ADH) (23), whereas the further oxidation of the aldehyde to the corresponding carboxylic acid is carried out by a dichlorophenol-indophenol (DCPIP)-dependent aldehyde dehydrogenase. Polyethylene glycol (PEG), a xenobiotic compound, contains an ether in addition to the alcohol function, as in THFA. Dye-linked ADHs were isolated from different bacteria able to

MATERIALS AND METHODS Chemicals. THFA, 2,6-DCPIP, dithiothreitol, PMS, nitroblue tetrazolium chloride, ferricyanide (potassium hexacyanoferrate), procion red, and the marker proteins for sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE) were purchased from Sigma (Deisenhofen, Germany). All other chemicals and PQQ were received from Fluka (Heidelberg, Germany). The chemicals were at least of analytical-reagent grade. SP-Sepharose and butyl-Sepharose were obtained from Pharmacia (Freiburg, Germany). Materials and marker proteins for isoelectric focusing were from Serva (Heidelberg, Germany). The capillary columns and the equipment for gas chromatography (GC) were purchased from Seitz Chromatographie Produkte (Weiterstadt, Germany), Hewlett Packard (Waldbronn, Germany), or Shimadzu (Duisburg, Germany). Type strains. The type strains used for identification of the isolated THFAdegrading bacterium were received from the following sources: Serratia marcescens was from a strain collection of our institute, Pseudomonas fluorescens was from the former Institute of Microbiology and Experimental Therapy, Jena, Germany (IMET 10619), and Agrobacterium tumefaciens was from the Deutsche

* Corresponding author. Mailing address: Institut fu ¨r Mikrobiologie, Universita¨t Halle, Kurt-Mothes-Str. 3, D-06099 Halle, Germany. Phone: 49-345-5526350. Fax: 49-345-5527010. E-mail: j.andreesen @mikrobiologie.uni-halle.de. 4891

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Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany (DSM 30204). Organism and growth conditions. R. eutropha Bo (DSM 11098) was isolated from soil by using enrichment cultures with THFA as the sole source of carbon and energy. The strain was grown in baffled 2-liter flasks containing 700 ml of mineral medium. The mineral salt medium was prepared as described previously (24). After sterilization, 100 mM potassium phosphate buffer (pH 8.0) and 30 mM THFA were added separately. The flasks were inoculated (5%, vol/vol) and incubated at 130 rpm on a rotary shaker for 24 h at 28 to 30°C. The cells were harvested at the end of the exponential growth phase by centrifugation at 8,500 3 g at 4°C. The wet cell paste was washed once with ice-cold 0.9% NaCl and stored at 220°C until used. Identification of the microorganism. For the determination of physiological and biochemical properties, the strain was grown in 5-ml aliquots of mineral salt medium in 20-ml culture tubes under the conditions described above. The substrate was added separately to a final concentration of 10 mM, and growth was monitored at 600 nm. Quinones were extracted from freeze-dried cells as described by Lechner et al. (28) and separated by high-pressure liquid chromatography (HPLC) using a LiChrospher 100-RP 18 100-Å column (250 by 4 mm; Machery-Nagel, Dueren, Germany), and a mobile phase of methanol–i-propanol (4:1, vol/vol) at a flow rate of 1 ml/min. The ubiquinones were identified by comparing their retention times (tRs) to those extracted from following reference strains: Serratia marcescens (ubiquinone 8 [Q8]), Pseudomonas fluorescens (Q9), and Agrobacterium tumefaciens (Q10). The fatty acid pattern was analyzed according to the method of Va¨isa¨nen and Salkinoja-Salonen (41), based on GC of whole-cell fatty acid methyl esters, with an HP 5898 A Microbial Identification System (Hewlett Packard Company, Avondale, Pa.) and the MIS System Software 3.9 (Microbial ID, Newark, Del.). Enzyme assay. All enzyme assays were performed in a final volume of 1.5 ml at 30°C. The standard assay for determination of THFA-DH activity contained 1 mM ferricyanide–0.5 mM THFA–5 mM CaCl2 in 50 mM Tris-HCl, pH 8.0. The reaction was started by the addition of THFA, and initial rates were derived from ferricyanide reduction following the decrease in absorbance at 405 nm (ε 5 958 M21 cm21). One unit of specific activity was defined as the reduction of 1 mmol of ferricyanide per min per mg of protein under the specific conditions of the assay. The activity of the NAD-dependent ADH was measured in 50 mM TrisHCl, pH 8.0, containing 2 mM NAD, 2 mM semicarbazide, and 5 mM substrate at 340 nm (ε 5 6,300 M21 cm21). One unit of specific activity was defined as the reduction of 1 mmol of NAD per min per mg of protein. Malate dehydrogenase (35) and alkaline phosphatase (4) activities were determined as described previously. The protein content was determined by the method of Bradford (9), with bovine serum albumin as a standard. Purification of THFA-DH. Wet cell paste (264 g) was suspended in 132 ml of 20 mM Tris-HCl, pH 7.8, containing 5 mM CaCl2 (buffer A). After the addition of 3 ml of Benzonase (Merck, Darmstadt, Germany) per 10-ml suspension and 10 mM phenylmethanesulfonyl fluoride, the cells were disrupted by using a French press at 110 MPa. Cell debris was removed by two succeeding centrifugations of the resulting extract at 30,000 3 g for 30 min at 4°C. The obtained supernatant was diluted eightfold with 10 mM Tris-HCl, pH 7.8, containing 3 mM CaCl2 (buffer B) and applied at a flow rate of 1.5 ml/min to an SP-Sepharose column (bed volume, 14.5 by 5 cm) previously equilibrated with buffer B. The column was washed with buffer B, and the THFA-DH activity was eluted by a linear gradient from buffer B to 0.2 M KCl in buffer B at a flow rate of 2 ml/min. The pooled fractions containing THFA-DH activity were brought to a final concentration of 1 M (NH4)2SO4 by adding the solid salt with constant stirring. Precipitated protein was removed by centrifugation for 20 min at 30,000 3 g and 4°C. The resulting supernatant was applied at a flow rate of 1.5 ml/min to a butyl-Sepharose column (bed volume, 4.1 by 5 cm) equilibrated with buffer A containing 1 M (NH4)2SO4. The bound protein was eluted with a linear gradient from buffer A containing 1 M (NH4)2SO4 to buffer A. Under these conditions, THFA-DH activity eluted at 0.75 M (NH4)2SO4. Pooled fractions containing THFA-DH activity were dialyzed for 20 h against buffer B and subsequently loaded at a flow rate of 1 ml/min on a procion red affinity column (bed volume, 4.6 by 5 cm) previously equilibrated with buffer B. Elution of THFA-DH activity was achieved by a linear gradient from buffer B to 0.2 M KCl in buffer B. The resulting pool was concentrated by chromatography on a butyl-Sepharose column under the conditions described above, except that THFA-DH activity was eluted with buffer B. Desalting of the THFA-DH pool was achieved by using a G-25 column equilibrated with buffer A. The purified protein was stored at 280°C. Localization experiments. Localization experiments were performed by osmotic shock according to the method of Koemen (22). Fresh THFA-grown cells of R. eutropha Bo were harvested at the exponential growth phase (optical density at 436 nm 5 8) by centrifugation (12,000 3 g, 4°C, 15 min). The pellet was washed twice with 20 mM Tris-HCl, pH 8.0, and resuspended in 10 ml of 50 mM Tris-HCl, pH 7.8, containing 0.5 M saccharose and 5 mM EDTA at room temperature (optical density at 436 nm 5 100). After the addition of 20 mg of lysozyme per g wet weight, the solution was stirred for 20 min at room temperature. Subsequently, the suspension was centrifuged at 10,000 3 g (4°C, 8 min), the supernatant (lysozyme fraction) was removed, and the pellet was immediately resuspended in 10 ml of ice-cold MgSO4 buffer (20 mM Tris-HCl, pH 7.8,

APPL. ENVIRON. MICROBIOL. containing 5 mM MgSO4). After incubation for 10 min on ice, the suspension was centrifuged (10,000 3 g, 4°C, 10 min) and the resulting supernatant (MgSO4 fraction) was carefully removed. The pellet (cytoplasmic fraction) was resuspended in 2 ml of 50 mM Tris-HCl, pH 8.0, containing 5 mM CaCl2. The cells were disrupted with a French press under the conditions described above. The protein in the lysozyme and MgSO4 fractions was precipitated by the addition of 85% (NH4)2SO4, and both fractions were subsequently dialyzed against 50 mM Tris-HCl, pH 8.0, containing 5 mM CaCl2 at 4°C. Spectral measurements. The steady-state kinetic analysis and UV-visible (UV/ VIS) absorption spectra of the enzyme or the cofactors were recorded with an Uvikon 930 spectrophotometer (Kontron) at 30°C. An absorption coefficient of ε 5 25,400 M21 cm21 at 249 nm and pH 4 was used for quantitative determination of PQQ (42). Electrophoresis. Homogeneity of the purified enzyme was analyzed under denaturing conditions by SDS-PAGE according to the method of Laemmli (27), with 12.5% polyacrylamide gels. Electrophoresis was carried out at constant current of 25 mA and maximal voltage. Gels were stained either with Coomassie brilliant blue G-250 (45) or with silver stain (7), and heme staining was performed with 3,39,5,59-tetramethylbenzidine (39). Determination of cofactors. The heme cofactor was detected as described by Bartsch (3). The content of heme c was estimated by addition of an equal volume of alkaline pyridine to samples containing THFA-DH or cytochrome c, respectively. After reduction by sodium dithionite, the ferrohemochrome spectrum of the samples was recorded. The amount of heme c was calculated by using an extinction coefficient of 25,800 M21 cm21 at 550 nm. The PQQ cofactor was separated from the enzyme by the SDS method as described by Van der Meer et al. (42). Subsequently, the cofactor was analyzed by HPLC by applying it to an RP-C18 Daltosil column (100 Å; 120 by 4 mm; Serva) equilibrated with 0.1% trifluoroacetic acid in aqua dest (aqua destillata) at a flow rate of 1 ml/min. PQQ was eluted by a linear gradient of increasing concentration of acetonitrile (0 to 40%, vol/vol) in 0.1% trifluoroacetic acid in aqua dest. The elution profile was recorded at both 215 and 254 nm. Determination of reaction products. To determine the stoichiometry of substrate conversion, 28 nM THFA-DH, 1 mM THFA, and 2 mM ferricyanide were incubated under standard assay conditions as described above. Aliquots of 200 ml were removed at several times, and the reaction was stopped by the addition of 0.8 ml of 20% H2SO4. Precipitated protein was removed by centrifugation (40,000 3 g, 4°C), and the amount of remaining THFA was determined by GC using a capillary column (BGB-1701; Seitz Chromatographie Produkte). Determination of reaction products and intermediates was carried out by incubating (i) 85 nM THFA-DH in 50 mM Tris-HCl, pH 8.0, containing 5 mM CaCl2, 2 mM pentanol, and 9 mM ferricyanide and (ii) 282 nM THFA-DH in 75 mM MOPS (morpholinepropanesulfonic acid)-NaOH, pH 8.0, containing 5 mM CaCl2, 5 mM ethanol, and 22 mM ferricyanide, respectively, at room temperature. The reactions were started by addition of the substrate, and aliquots of 200 ml were removed every 2 min and prepared as described above. The amount of substrate consumed or product formed was determined with a GC capillary column (HP-Innowax; Hewlett Packard) under the conditions described above. Determination of molecular weight and isoelectric point. The molecular weight of the subunit was determined by using SDS gels with a 10-to-27.5% polyacrylamide gradient. Mixtures 4 and 5 (Serva) were used as marker proteins. Alternatively, the molecular weight of the subunit was determined by MALDI (matrix-assisted-laser desorption-ionization) mass spectrometry, which was performed on a reflectron type time-of-flight mass spectrometer (REFLEX; BrukerFranzen Analytik, Bremen, Germany). Gel filtration chromatography and native gradient PAGE were used for determination of the native molecular weight. A fast-performance liquid chromatography column (300 by 10 mm; FPLC Superdex 200 column; Pharmacia) or a Hiload Superdex 200 column (600 by 16 mm; Pharmacia) was equilibrated with 0.02 M Tris-HCl, pH 8.0, containing 0.15 M KCl and 0.01 M CaCl2 or with 0.15 M Tris-HCl, pH 8.0, containing 0.01 M CaCl2, respectively. THFA-DH was applied to the column at a flow rate of 0.25 ml/min. Native linear gradient PAGE (4 to 27.5% polyacrylamide) was performed as described previously (6). Isoelectric focusing was carried out as described by Robertson et al. (33), with ampholytes in the pH range of 3 to 10. The isoelectric point of THFA-DH was determined by using a marker kit from Serva (pH 3 to 10). N-terminal sequence analysis. Purified THFA-DH was desalted by HPLC using a Nucleosil 500-5 C3-PPN RP column (125 by 4 mm; Machery-Nagel). The column was equilibrated with 0.08% trifluoroacetic acid in aqua dest (solvent A). A gradient from solvent A to 80% acetonitrile in solvent A was developed for elution of the enzyme. The THFA-DH obtained was concentrated and applied to an Applied Biosystems 476 A Protein Sequencer. The sequence analysis was carried out according to the instruction manual.

RESULTS Morphological and physiological characterization of the THFA-degrading strain. The THFA-degrading microorganism isolated from soil was characterized as a gram-negative, motile, short rod. Q8 was determined to be the main isoprenoid qui-

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TABLE 1. Induction of different ADHs during growth of R. eutropha Bo on different alcohols Sp act (mU/mg) in crude extracts assayed witha: Growth substrate

b

THFA FeCN

THFA Furfuryl alcohol Ethanol Pentanol 2-Pentanol

160 30 0 60 320

Furfuryl alcohol NAD c

0 0 0 0 0

Ethanol

Pentanol

2-Pentanol

1,4-Butandiol

FeCN

NAD

FeCN

NAD

FeCN

NAD

FeCN

NAD

FeCN

NAD

130 10 0 50 240

30 10d 210 110 120

40 0 0 0 220

60 40 380 180 160

260 40 0 120 490

70 30 370 210 200

300 50 0 150 570

20 20 180 210 100

170 30 0 60 320

0 0 50 20 30

a

Activities were determined as described in Materials and Methods. R. eutropha was grown at 30°C and 170 rpm in a mineral medium containing 30 mM substrate. Cell extracts were obtained as indicated in Materials and Methods. 0, no detectable activity under the conditions used. d Furfuryl alcohol (0.5 mM) was used in the enzyme assay. b c

none. The strain was not able to utilize carbohydrates such as glucose, mannose, arabinose, or mannitol as sole sources of carbon and energy. No growth was observed on L-serine, L-lysine, glycine, d-aminovalerate, maleate, acetamide, and glycerol. Compounds like DL-valine, acetate, lactate, 2-oxoglutarate, succinate, citrate, n-butyrate, gluconate, caproate, phenylacetate, nicotinate, benzoate, ethanol, n-pentanol, or 2-hexanol were utilized as growth substrates. Application of different test systems, such as API 20 NE, determination of the main isoprenoid quinone, and analysis of the fatty acid pattern led to the organism’s identification as R. eutropha (formerly A. eutrophus [46]). We were interested in whether the isolated microorganism could use other heterocyclic compounds as sole sources of carbon and energy. Besides utilizing THFA and its corresponding acid, the strain was able to utilize different substituted aromatic O- and N-heterocyclic compounds, e.g., furfuryl alcohol, furan-2-carboxylate, pyrrole-2-carboxaldehyde, pyrrole2-carboxylate, and nicotinate. However, no growth was observed on the unsubstituted aromatic or saturated heterocyclic compounds furan, tetrahydrofuran, morpholine, 1,4-dioxane, pyrrolidinone, piperidine-2-carboxylate, and THF amine. Induction of different ADH activities. A ferricyanide-dependent THFA-DH activity could be detected after growth of R. eutropha Bo on a defined mineral medium containing up to 200 mM THFA as the sole source of carbon and energy. A maximal specific activity of 0.16 U/mg was obtained in crude extracts when cells were harvested at the end of the exponential growth phase. This activity was inducible, for it was not detected in cells grown on, e.g., citrate. However, when R. eutropha Bo was grown on different alcohols, such as pentanol, a ferricyanidedependent ADH activity was detected in the corresponding crude extracts (Table 1). This activity was quite low in furfuryl alcohol-grown cells and could not be detected in cells grown on ethanol. It was previously reported that strains of R. eutropha can express an NAD-dependent ADH when grown on aliphatic alcohols (36). These results were confirmed during our investigations, in which NAD-dependent ADH activities were detected preferentially in crude extracts from cells grown on aliphatic alcohols (Table 1). The induction level of this activity was quite low if the two heterocyclic alcohols served as substrates. It also became obvious from the data presented in Table 1 that THFA was not converted by the NAD-dependent ADH expressed during growth on different alcohols. In contrast to C. testosteroni (14), addition of PQQ to the growth medium had no effect on the lag phase or on the growth rate of R. eutropha Bo and did not influence the ferricyanidedependent THFA-DH activity in crude extracts.

Purification of THFA-DH. THFA-DH from R. eutropha Bo was purified to apparent homogeneity by a three-step procedure (Table 2). The homogeneous enzyme was enriched 211fold and had a specific activity of 43.6 U/mg with THFA as a substrate. According to the purification factor and the yield, it can be calculated that THFA-DH represents about 0.5% of the total soluble protein. A single band was obtained when the purified enzyme was analyzed by SDS-PAGE (Fig. 1). The stability of THFA-DH activity could be improved by the addition of 5 mM CaCl2 during purification. Addition of PQQ (0.01 to 1 mM) had no positive effect on the enzyme activity during purification. No significant loss in activity was observed when the enzyme was stored for prolonged periods in 20 mM TrisHCl, pH 8.0, containing 5 mM CaCl2 at 280°C. Properties of THFA-DH. The subunit molecular mass of THFA-DH was determined to be 73.6 kDa by SDS-polyacrylamide gel-gradient PAGE and 73.4 kDa by mass spectrometry. During gel filtration experiments, an apparent strong interaction with the column matrix was observed on the FPLC Superdex 200 column, even at various salt concentrations. Therefore, a Hiload Superdex 200 gel filtration column was used. Under these conditions THFA-DH activity corresponding to a molecular mass of 79.5 kDa eluted from the column, indicating a monomeric structure. No protein band was obtained when the homogeneous enzyme was applied to the system routinely used for nondenaturing gradient-PAGE, possibly due to its still positive net charge. The pH optimum of THFA-DH activity with THFA (0.5 mM) as the substrate was between pH 8.5 and 9.0 at 30°C, and the highest stability was also obtained in this pH range. A temperature optimum for THFA-DH activity was found between 40 and 50°C with 50 mM potassium phosphate buffer. As observed during purification of THFA-DH, the enzyme was tightly bound to the cation exchanger SP-Sepharose at pH

TABLE 2. Purification of ferricyanide-dependent THFA-DH from R. eutropha Boa Purification

Protein (mg)

Total activity (U)

Sp act (U/mg)

Yield (%)

Purification (fold)

Cell extract SP-Sepharose Butyl-Sepharose Procion red

11,252 585 73 21

2,250 3,042 1,876 834

0.2 5.2 25.7 39.7

100 135 84 37

1 28 137 211

a Enzyme activity was determined in 50 mM Tris-HCl, pH 8.2, containing 5 mM CaCl2, 1 mM ferricyanide, and 0.5 mM THFA at 30°C.

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FIG. 1. SDS–12.5% PAGE and Coomassie blue stain of THFA-DH from R. eutropha after different purification steps. Lane 1, marker proteins; lane 2, crude extract (20 mg); lane 3, SP-Sepharose pool (12.5 mg); lane 4, butyl-Sepharose pool (3.3 mg); lane 5, homogeneous THFA-DH after chromatography on procion red (0.5 mg).

7.8, indicating the strongly basic character of the protein. This was confirmed by the results obtained during gel filtration chromatography and native gradient PAGE experiments. In accordance with these data, the isoelectric point of THFA-DH was determined to be 9.1 by isoelectric focusing. Cofactor content. The cofactor isolated from THFA-DH revealed the same tR as PQQ after HPLC separation (tR 5 12.3 min). However, a second peak at tR 5 15.8 min (data not shown) was observed with the isolated cofactor and commercially available PQQ. Because trifluoroacetic acid (0.1% in aqua dest) was used as the solvent, PQQ might have undergone a spontaneous autohydrogenation to PQQH2 at low pH values, as suggested previously (42). The isolated cofactor showed the same spectrum as PQQ when analyzed by UV/VIS spectroscopy. The amount of isolated PQQ was calculated by the absorption coefficient to represent 4.25 nmol of PQQ from 4.6 nmol of homogeneous protein (73.5 kDa), corresponding to a molar ratio of THFA-DH to PQQ of 1 to 0.93. This indicated that no significant amount of PQQ was lost during purification and that native THFA-DH contained one molecule of PQQ per monomer. The heme cofactor could not be extracted with 2-butanone, indicating that it was covalently bound to the protein. This was confirmed by the observation that only a single band at 73.6 kDa occurred after SDS-PAGE and heme staining of homogeneous THFA-DH. Addition of sodium dithionite resulted in a typical ferrohemochrome spectrum, showing absorption maxima at 414, 520, and 550 nm. A total of 2.5 nmol of heme c was detected from 2.3 nmol of purified protein, indicating a molar ratio of 1.07 to 1. Thus, THFA-DH contained both heme c and PQQ in a stoichiometric ratio to the monomer. Spectral properties of THFA-DH. The purified, homogeneous THFA-DH had a red-brownish color and showed the typical UV/VIS absorption spectrum of reduced cytochrome c. Absorption maxima of the reduced enzyme were determined at 419, 523, and 552 nm (Fig. 2). Quite similar absorption spectra were obtained for the holoenzymes of quinohemoprotein ADHs isolated from C. testosteroni and P. putida (14, 40). The ratio of A280/A416 was 1.07 for THFA-DH of R. eutropha Bo, compared to 1.21 reported for the ADH of C. testosteroni (10). Addition of 100 mM ferricyanide to THFA-DH (2.8 mM) resulted in the oxidation of the heme c cofactor as monitored by recording the absorption spectra between 200 and 600 nm. As observed for the ADH from C. testosteroni and P. putida, THFA-DH tended to persist in the reduced form. A high

APPL. ENVIRON. MICROBIOL.

excess of ferricyanide was required to achieve complete oxidation. The oxidized protein could be reduced by addition of THFA (0.5 mM) (Fig. 2), showing that the heme c cofactor was involved in substrate oxidation. Activators and inhibitors. Addition of PQQ to the homogeneous enzyme had no effect on the specific activity. However, the specific activity decreased drastically when the protein was dialyzed for 20 h at 4°C against 20 mM Tris-HCl, pH 8.0, containing 2 mM EDTA. It could be completely restored when saturating amounts of PQQ (50 mM) and CaCl2 (5 mM) were added to the enzyme assay. These results confirmed the assumption that bivalent cations, especially Ca21 ions, are involved in the binding process of PQQ (14). Saturation kinetics were obtained at calcium ion concentrations of about 3 mM (data not shown). Calcium ions could be replaced by magnesium ions without a significant change in specific activity. However, cobalt ions (3 mM) and manganese ions (3 mM) led to complete inhibition of THFA-DH when added to the enzyme assay. A further 2.5-fold increase in enzyme activity could be observed in the presence of ammonium sulfate (15 mM) with THFA or n-pentanol as a substrate. Other monovalent ions, such as sodium or potassium, were without any effect. Substrate spectrum and electron acceptor usage. THFADH exhibited a broad substrate spectrum and was able to oxidize primary, secondary, and cyclic alcohols, diols, PEG 6000, and aldehydes (Table 3). The activities obtained for different substrates were compared to the maximal activity determined for THFA. Although R. eutropha Bo was grown on THFA, the purified THFA-DH showed the highest activity with 2-hexanol under the conditions used (Table 3). No activity was obtained with glycerol and glucose as substrates, which is in agreement with the observation that the strain was not able to grow on these substances. Although methanol did not serve as a substrate for THFA-DH, a distinct activity was detected for the corresponding aldehyde, formaldehyde (Table 3). Interestingly, THFA-DH from R. eutropha Bo was able to con-

FIG. 2. UV/VIS spectra of purified THFA-DH from R. eutropha Bo. Absorption spectra of the purified THFA-DH (2.8 mM) were recorded in 50 mM Tris-HCl, pH 8.0, containing 5 mM CaCl2 at 25°C. Shown are spectra for reduced THFA-DH (——), oxidized THFA-DH after the addition of 100 mM potassium ), and reduced THFA-DH obtained by the addition of THFA ferricyanide ( (500 mM) to the oxidized enzyme (– – –).

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TABLE 3. Substrate specificity of THFA-DHa Substrate

V9maxb (%)

Primary alcohols Methanol .......................................................................................... 0 Ethanol ............................................................................................. 175 Propanol........................................................................................... 166 n-Butanol ......................................................................................... 166 n-Pentanol........................................................................................ 176 n-Hexanol......................................................................................... 129 Secondary alcohols 2-Butanol.......................................................................................... 65 2-Pentanol ........................................................................................ 220 2-Hexanol......................................................................................... 254 Aldehydes Formaldehyde.................................................................................. 125 Acetaldehyde ................................................................................... 168 Butanal ............................................................................................. 150 Pentanal............................................................................................ 137 Furfural ............................................................................................ 44 Cyclic alcohols THF-2-alcohol ................................................................................. 100 THF-3-alcohol ................................................................................. 119 3-Hydroxy-THF ............................................................................... 45 Furfuryl alcohol............................................................................... 86 Diols and polyols 1,2-Ethandiol ................................................................................... 5 1,2-Butandiol ................................................................................... 89 1,3-Butandiol ................................................................................... 140 1,4-Butandiol ................................................................................... 138 PEG 6000......................................................................................... 70 Glycerol ............................................................................................ 0 Glucose............................................................................................. 0 a Enzyme activities were determined in 20 mM MOPS-NaOH, pH 8.0, containing 5 mM CaCl2, with 1 mM ferricyanide, various concentrations of substrate, and 18.8 nM THFA-DH at 30°C. b The relative V9max, measured at saturating substrate concentrations, is given as a percentage of the activity obtained with THFA (43.6 U/mg 5 100%).

vert PEG 6000, in contrast to the two similar quinohemoprotein ADHs isolated from P. putida (40). As observed in cell extracts, the highest THFA-DH activity was determined with ferricyanide as the electron acceptor (Table 4). Addition of PMS to ferricyanide showed no significant increase of the specific activity. The enzyme transferred electrons to cytochrome c at a very low rate, but this could be significantly increased by using PMS as a mediator. A much TABLE 4. Activity of THFA-DH with different electron acceptors Electron acceptor

Activitya (%)

Potassium ferricyanide ...................................................................... 100 Potassium ferricyanide-PMS ............................................................ 105 DCPIP................................................................................................. 6 DCPIP-PMS ....................................................................................... 17 Cytochrome c ..................................................................................... 3 Cytochrome c-PMS ........................................................................... 82 Othersb ................................................................................................ 0 a The activities were determined in 50 mM Tris-HCl, pH 8.2, containing 5 mM CaCl2, 0.5 mM THFA, and 18.8 nM THFA-DH. The artificial electron acceptors were used at concentrations as follows: ferricyanide, 1 mM (set to 100%); DCPIP, 0.2 mM; PMS, 70 mM; and cytochrome c, 20 mM. b NAD(P) alone and nitroblue tetrazolium with or without PMS.

FIG. 3. Detection of reaction products during conversion of n-pentanol (A) and ethanol (B) by THFA-DH. A solution of 2 mM pentanol–9 mM ferricyanide–50 mM Tris-HCl (pH 8.0) containing 5 mM CaCl2 and 85 nM THFA-DH (A) and a solution of 5 mM ethanol–22 mM ferricyanide–75 mM MOPS-NaOH (pH 8.0) containing 5 mM CaCl2, and 282 nM THFA-DH (B) were incubated with continuous stirring at room temperature. Aliquots of 200 ml were removed every 2 min and analyzed by GC as described in Materials and Methods. The results for alcohol (F), aldehyde (E), and carboxylic acid (}) detection are shown.

lower stimulation was observed if PMS was added to the twoelectron acceptor 2,6-DCPIP. THFA-DH did not react with nitroblue tetrazolium chloride (with or without PMS), NAD, or NADP as the electron acceptor. Intermediates and products. The stoichiometry of the electron transfer reaction catalyzed by THFA-DH was determined by conversion experiments with THFA as a substrate and ferricyanide as an electron acceptor under the conditions described in Materials and Methods. These experiments showed that the oxidation of 1 nmol of THFA was accompanied with the reduction of 4 nmol of ferricyanide (data not shown). From these data it could be suggested that THFA-DH converted the alcohol beyond the aldehyde level to its corresponding carboxylic acid. In order to determine whether the alcohol is directly oxidized to the carboxylic acid or whether the corresponding aldehyde is released from the enzyme as an intermediate, both substrate and reaction products were analyzed by GC during enzymatic conversion. THFA could not be used as a substrate during these experiments, because the corresponding aldehyde is not commercially available and no supposed intermediate could be identified by GC. Pentanol was chosen because it was one of the best substrates for THFA-DH (Table 3). When 2 mM n-pentanol and 9 mM ferricyanide were incubated with 85 nM THFA-DH, pentanal was formed as a transient intermediate (Fig. 3). The oxidation of pentanal was initiated after the

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TABLE 5. Alignment of the N-terminal amino acid sequences of THFA-DH from R. eutropha Bo and the quinohemoprotein ADH from C. testosteroni (38)a Organism

Sequence

R. eutropha Bo.................. AADAAA---RVDGAAIRANEAGTPNWPSYGLDYA C. testosteroni.............. TGPAAQAAAAVQRVDGDFIRANAARTPDWPT-GVDYA a

Identical amino acids are indicated in boldface letters.

pentanol concentration had decreased to about 10% of the initial amount, which is in agreement with the corresponding kinetic data for these substrates (Km [pentanol] 5 0.012 mM; Km [pentanal] 5 0.061 mM). If ethanol (5 mM) was incubated with 22 mM ferricyanide and 282 nM THFA-DH, the corresponding aldehyde was detected in a very low concentration, about 0.02 mM (Fig. 3), which is in agreement with the Km values obtained for ethanol and acetaldehyde, 3.1 and 0.075 mM, respectively. From these results it can be concluded at least for pentanol and ethanol that the in vitro oxidation of an alcohol to its corresponding carboxylic acid by THFA-DH was a two-step reaction with the aldehyde as a true intermediate. N-terminal amino acid sequence. The N-terminal amino acid sequence obtained for THFA-DH from R. eutropha Bo revealed a high similarity to the quinohemoprotein ADH from C. testosteroni (38). An identity of 65% and a similarity of 80% were determined by comparing the 36 N-terminal amino acids (Table 5). No similarity to the sequence of any other quinoprotein was found. However, an alignment of the complete amino acid sequence of ADH from C. testosteroni with those of other quinoproteins (38) showed that there is generally a low similarity at the N-terminal part of these proteins. Localization of THFA-DH. Genetic data indicated that quinohemoprotein ADH of C. testosteroni contains a leader peptide for translocation of the protein into the periplasm (38). Therefore, experiments were performed to get information about the localization of THFA-DH from R. eutropha Bo. Malate dehydrogenase and alkaline phosphatase were used as marker enzymes for the cytoplasm and periplasm, respectively. However, the activity of alkaline phosphatase was very low in crude extract from cells grown on THFA. After separation of the different fractions, no alkaline phosphatase activity could be detected. The periplasmic fraction contained only 9.6% of the malate dehydrogenase activity but contained 45% of the THFA-DH activity. Thus, it might be assumed that the THFADH is localized to about 50% in the periplasm. DISCUSSION The enrichment of microorganisms able to utilize THFA as a sole source of carbon and energy led to the isolation of a strain identified as R. eutropha (DSM 11098). This is the first report on the degradation of THFA by a microorganism. The initial oxidation of THFA is catalyzed by an NAD-independent dye-linked ADH which oxidizes THFA to its corresponding carboxylic acid. This is significantly different from the breakdown of the analogous aromatic compound furfuryl alcohol by P. putida, in which the alcohol group is first oxidized to the aldehyde level by an NAD-dependent ADH. Furfural is then converted to furan-2-carboxylate by a second dye-linked dehydrogenase (23). THFA-DH from R. eutropha Bo was able to convert both furfuryl alcohol and furfural in addition to THFA. It is interesting that during growth of R. eutropha Bo on ethanol or furfuryl alcohol as substrate, no or only a negligible, low ferricyanide-dependent activity was detected. However, a high induction of the latter activity was obtained after growth

on 2-pentanol even if ethanol or furfuryl alcohol was used as electron donor. If one compares the relation of the ferricyanide-dependent activities induced after growth on different alcohols, it might be speculated that the organism synthesizes different ADHs of this type, showing a deviating specificity at least for ethanol. Thus, a situation similar to that reported for P. putida, which formed three different dye-linked ADHs depending on the growth conditions (40), seems to exist. However, the organism was also able to express an NAD-dependent ADH activity during growth on different alcohols. It was previously shown that R. eutropha strains were able to induce different NAD(P)-dependent ADHs depending on the growth conditions used (19, 26, 29). Jendrossek et al. (18) described R. eutropha mutants able to use ethanol as a sole source of carbon and energy in contrast to the wild-type strains N9A and H16. Under conditions of restricted oxygen supply, the wildtype strains produced an unspecific NAD(P)-dependent ADH as a typical fermentation enzyme (36). Madyastha and Gururaja (29) isolated a secondary ADH from a strain of R. eutropha unable to oxidize short-chain alcohols like ethanol or ipropanol. Our investigations indicated that R. eutropha Bo grown on different alcohols was not able to produce an NADdependent ADH to convert THFA. This was in agreement with the observation that no or only a low NAD-dependent ADH activity was present in THFA-grown cells. A quite similar induction level was obtained for furfuryl alcohol-grown cells, although the organism expressed an NAD-dependent ADH able to convert this substrate after growth on, e.g., ethanol. The highest NAD-dependent ADH activity was detected in cells grown on ethanol. The isolation and characterization of the different ADHs present in R. eutropha Bo are necessary to obtain more information on the relation between the alcohol used as growth substrate and the induction pattern of different ADHs. The characterization of the purified THFA-DH of R. eutropha Bo indicated that the enzyme is a quinoprotein ADH of group II (30), being a monomer of about 73 kDa and containing one PQQ and one covalently bound heme c. During recent years, quinoproteins of this group were isolated from C. testosteroni, P. putida, a Pseudomonas sp., and R. acidophila (10, 16, 40, 47). The quinohemoprotein ADH from R. eutropha Bo exhibited the highest similarity to the corresponding protein from C. testosteroni regarding its isoelectric point, molecular mass, and N-terminal amino acid sequence (38, 47). However, C. testosteroni produces only the apoenzyme without the PQQ cofactor (14), whereas R. eutropha Bo synthesized the holoenzyme. It was supposed that the former organism can grow slowly on alcohols by producing an NAD-dependent ADH, as was detected in R. eutropha H16 and Bo (14, 36). The presence of PQQ improves growth of C. testosteroni significantly, as was also observed for other PQQ-dependent proteins, such as glucose dehydrogenase from Escherichia coli (14, 15). In natural habitats, PQQ is often provided by other bacteria found to produce the coenzyme in large amounts and excrete it into the environment (34, 43). A different situation seems to exist in P. putida, in which three distinct quinoprotein ADHs were induced, depending on the alcohol used as the growth substrate (40). All three enzymes were isolated as holoenzymes, and two of them belong to the group II quinoprotein ADHs (30, 40). However, the isoelectric points of these enzymes were determined to be 6.5 and 5.0, which are much lower than that reported for the quinohemoprotein ADH from C. testosteroni (about 9.0 [47]) and that determined for THFA-DH from R. eutropha Bo in this study (9.1). PEG 6000 was no substrate for the enzymes isolated from P. putida but was converted by the enzymes of C. testosteroni and R. eutropha Bo (40, 47). Very

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recently, a quinohemoprotein ADH with an isoelectric point of 6.0 was isolated from R. acidophila, and it showed a broad substrate spectrum, also converting PEG (47). In general, the substrate spectrum of all group II quinoprotein ADHs described is very broad but restricted to the conversion of alcohols and aldehydes (14, 40, 47). Lupanine hydroxylase from a Pseudomonas strain is the only exception known so far. This enzyme catalyzes the initial hydroxylation of lupanine by a dehydrogenation of a C-N bond followed by a subsequent hydration (16). The conversion experiments performed with the THFA-DH from R. eutropha Bo suggested that the in vitro oxidation of the alcohols to their corresponding carboxylic acids is a two-step reaction. As shown at least for ethanol and n-pentanol, the aldehyde was released from the enzyme as an intermediate. Similar results were obtained for the PQQ-dependent ADH from C. testosteroni, and it was proposed that both alcohol and aldehyde compete for the same active site of the enzyme (11). However, for other substrates the possibility that the alcohol is oxidized directly to its corresponding carboxylic acid without any intermediate cannot be excluded. The high sequence homology obtained for the N termini of THFA-DH from R. eutropha Bo and the quinohemoprotein ADH from C. testosteroni indicated that these enzymes might be closely related. This assumption is supported by the fact that both organisms belong to the b-group of proteobacteria (31). Analysis of the quinohemoprotein ADH gene of C. testosteroni showed that the protein contains a leader peptide for its translocation into the periplasm (38). It was also reported that different quinoprotein methanol dehydrogenases are localized in the periplasm, transferring electrons to specific cytochromes in their vicinity (2, 13). Our data indicate that the THFA-DH of R. eutropha Bo is probably a periplasmic enzyme, too. The complete sequence of the gene encoding THFA-DH might lead to more information about the localization and the structural similarities to other quinoproteins. ACKNOWLEDGMENTS We thank H. Ebner (Institut fu ¨r Mikrobiologie, Go ¨ttingen, Germany) for providing enrichment cultures, P. Ru ¨cknagel and T. Pfeifer (Max-Plank-Gesellschaft Forschungsstelle Enzymologie der Proteinfaltung, Halle, Germany) for N-terminal amino acid sequence analysis and mass spectrometry experiments, respectively, and U. Lechner of our institute for providing data on the fatty acid pattern of the organism. This work was partly supported by grants of the Forschungsfo ¨rderung des Landes Sachsen-Anhalt and Fonds der Chemischen Industrie. REFERENCES 1. Aislabie, J., A. K. B. H. Hurst, S. Rothenburger, and R. M. Atlas. 1990. Microbial degradation of quinoline and methylquinoline. Appl. Environ. Microbiol. 56:345–351. 2. Anthony, C. 1996. Quinoprotein-catalysed reactions. Biochem. J. 320:697– 711. 3. Bartsch, R. G. 1971. Cytochromes: bacterial. Methods Enzymol. 23:344–363. 4. Bergmeyer, H. U., M. Grassl, and H. E. Walter. 1983. Phosphatase, alkaline, p. 269–270. In H. U. Bergmeyer (ed.), Methods of enzymatic analysis, 3rd ed., vol. 2. VCH, Weinheim, Germany. 5. Bernhardt, D., and H. Diekmann. 1991. Degradation of dioxane, tetrahydrofuran and other cyclic ethers by an environmental Rhodococcus strain. Appl. Microbiol. Biotechnol. 36:120–123. 6. Blaschke, M., A. Kretzer, C. Scha ¨fer, M. Nagel, and J. R. Andreesen. 1991. Molybdenum-dependent degradation of quinoline by Pseudomonas putida Chin IK and other aerobic bacteria. Arch. Microbiol. 155:164–169. 7. Blum, H., H. Beier, and H. J. Gross. 1987. Improved silver staining of plant proteins, RNA, and DNA in polyacrylamide gels. Electrophoresis 8:93–99. 8. Bock, C., R. M. Kroppenstedt, and H. Diekmann. 1996. Degradation and bioconversion of aliphatic and aromatic hydrocarbons by Rhodococcus ruber 219. Appl. Microbiol. Biotechnol. 45:408–410.

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