Department of Microbiology and Enzymology, Delft University of Technology, Julianalaan 67, NL-2628 BC Delft, The Netherlands1
Author for correspondence: Johannis A. Duine. Tel: +31 15 2785051. Fax: +31 15 2782355. e-mail: j.a.duine{at}stm.tudelft.nl
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ABSTRACT |
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Keywords: nicotinoprotein alcohol dehydrogenase, Rhodococcus erythropolis
Abbreviations: MNO, methanol:NDMA oxidoreductase; NDMA, N,N-dimethyl-4-nitrosoaniline; np-ADH, nicotinoprotein alcohol dehydrogenase
The EMBL accession number for the sequence reported in this paper is P81747.
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INTRODUCTION |
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MNO (EC 1 . 1 . 99 . ) is a member of the group of nicotinoprotein alcohol dehydrogenases, i.e. enzymes which contain tightly bound, non-exchangeable NAD(P)(H) in the active site acting as cofactor instead of coenzyme. MNO was detected in Amycolatopsis methanolica NCIB 11946 (grown on methanol, ethanol, butan-1-ol, hexan-1-ol or acetate) and Mycobacterium gastri MB 19 (grown on methanol, propan-1-ol, propan-2-ol or glycerol) (Bystrykh et al., 1993a , b
; van Ophem et al., 1993
), i.e. Gram-positive, methylotrophic actinomycetes closely related to (non-methylotrophic) rhodococci. It is a 490500 kDa homodecameric class III alcohol dehydrogenase, each subunit containing a tightly, but not covalently bound NADP(H) cofactor (Bystrykh et al., 1993a
). Since no transfer of reducing equivalents takes place to externally added NAD(P), MNO is not active in the common assays for NAD(P)-dependent alcohol dehydrogenases. Its alcohol dehydrogenase activity can be demonstrated, however, by using the artificial electron acceptor NDMA. In such an assay, the enzyme acts as a coenzyme-independent alcohol: NDMA oxidoreductase and obeys a ping-pong mechanism. Moreover, MNO catalyses the dismutation of formaldehyde and the NADH-dependent reduction of aldehydes (Bystrykh et al., 1993b
). Gene-disruption mutants of A. methanolica lacking a functionally active gene encoding MNO were unable to grow on methanol, ethanol, propan-1-ol or butan-1-ol as sole carbon source (Hektor & Dijkhuizen, 1996
; Hektor, 1997
). It seems, therefore, that MNO is crucial for oxidation of these C1C4 alcohols.
Surprisingly, apart from MNO, A. methanolica (grown on methanol, ethanol, butan-1-ol, hexan-1-ol or acetate) contains a second nicotinoprotein alcohol:NDMA oxidoreductase (EC 1 . 1 . 99 . ) (van Ophem et al., 1993 ) for which the name nicotinoprotein alcohol dehydrogenase (np-ADH) has been proposed (Piersma et al., 1998
). The enzyme is quite different from MNO with respect to protein structure and catalytic performance. np-ADH is a homotri- or tetrameric class I alcohol dehydrogenase of 120 kDa, each 39 kDa subunit containing firmly, non-covalently bound NADH as cofactor. It catalyses the NAD(P)-independent oxidation of various primary alcohols (but not methanol) with the concomitant reduction of NDMA, according to a ping-pong mechanism. However, in contrast to MNO it catalyses neither the dismutation of formaldehyde nor the NADH-dependent reduction of aldehydes. Despite the fact that MNO and np-ADH are simultaneously induced during growth on certain alcohols (methanol, ethanol, butan-1-ol or hexan-1-ol), and ethanol, propan-1-ol and butan-1-ol are good substrates for both enzymes in vitro, the physiological role of MNO in A. methanolica cannot be taken over by np-ADH, as judged from the inability of mutants lacking the MNO-encoding gene to grow on these alcohols. Since such studies using mutants lacking a functionally active gene encoding np-ADH have not been performed so far, the physiological role of np-ADH remains to be elucidated.
During our earlier work on novel types of alcohol dehydrogenases involved in the kinetic resolution of racemic alcohols (Geerlof et al., 1994 ), strong indications were obtained that a nicotinoprotein alcohol dehydrogenase found in certain rhodococci is involved in resolving enantiomeric mixtures of alcohols. To study this in more detail, R. erythropolis DSM 1069 was chosen as a model organism. From these studies it appeared that this strain is able to produce MNO (characterization to be published elsewhere) as well as np-ADH. Here we report on the induction, purification and characterization of np-ADH.
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METHODS |
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Cultivation and induction experiments.
Rhodococcus erythropolis DSM 1069 was precultivated on a medium containing 2% (w/v) D-glucose, 1% (w/v) bactopeptone, 0·5% (w/v) NaCl and 0·05% (w/v) yeast extract in a rotary shaker incubator (200 r.p.m.) for 48 h at 30 °C. For induction studies, 25 ml precultures were used to inoculate 2 l Erlenmeyer flasks, containing 0·5 l mineral salt medium (Eggeling & Sahm, 1984 ), 0·05% (w/v) yeast extract, and growth substrate (see legend of Table 1
). The flasks were shaken (at 200 r.p.m., at 30 °C) and the cells were harvested in the late exponential growth phase (after approximately 48 h, except for growth on citronellol, which required 168 h) by centrifugation (20 min at 16300 g, at 4 °C) and washed twice with 10 mM potassium phosphate buffer, pH 7·0, at 4 °C. An aliquot of cell paste (typically 12 g) was resuspended in an equal volume of the same buffer, sonicated (using an MSE sonicator for 4x10 s on ice, at high power, amplitude 2, 10 µm peak to peak), and centrifuged (30 min at 20000 g, at 4 °C). The supernatant was used as the cell-free extract and assayed for various alcohol dehydrogenase activities.
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Enzyme purification.
All purification steps were performed at room temperature unless indicated otherwise. The fractions were assayed for the presence of np-ADH, MNO or NAD(P)-dependent benzyl alcohol dehydrogenase activities. Thawed cell paste (50 g) was resuspended in an equal volume of 10 mM potassium phosphate buffer, pH 7·0, at 4 °C. After adding bovine DNase I, the suspension was passed three times through a French pressure cell under a pressure of 27·6 MPa. Centrifugation (60 min at 16300 g, at 4 °C) yielded the cell-free extract. The extract was dialysed for 1 h against 1 M potassium phosphate buffer, pH 7·0, at 4 °C, and centrifuged (20 min at 16300 g, at 4 °C). Aliquots (10 ml) of the supernatant were applied to a 40 ml Phenyl-Sepharose FF hydrophobic interaction column (Pharmacia Biotech) equilibrated with 1 M potassium phosphate buffer, pH 7·0, at 4 °C. A linear gradient of potassium phosphate (1 M10 mM, pH 7·0) was applied for 45 min at a flow rate of 2 ml min-1. Fractions showing np-ADH activity (eluting in the final part of the gradient) were concentrated by centrifugation (3000 g) using a 30 kDa cutoff filter (Centriprep 30, Amicon). Concentrate obtained from five separate runs on Phenyl-Sepharose was applied to a Poros HQ (1x10 cm) anion-exchange column (Boehringer Mannheim) equilibrated with 20 mM potassium phosphate buffer, pH 7·2. A linear gradient of potassium chloride (01·5 M) in the same buffer was applied for 60 min at a flow rate of 2 ml min-1. The np-ADH activity, eluting at 35 mM KCl, was concentrated as above and applied in 200 µl portions to a Superdex 200 (1x30 cm) gel-filtration column (Pharmacia) equilibrated with 10 mM potassium phosphate buffer, pH 7·0, containing 0·2 M KCl, at a flow rate of 0·5 ml min-1. Fractions with np-ADH activity from 10 separate runs were pooled and stored frozen at -80 °C.
Enzyme assays.
All measurements were performed in duplicate at 20 °C. np-ADH activity was measured by following the reduction rate of NDMA at 440 nm with benzyl alcohol as electron donor. The reaction mixture contained (total volume 1 ml): benzyl alcohol, 20 µM; NDMA, 28 µM; potassium phosphate buffer, pH 7·0, 10 mM. The reaction was started by the addition of an appropriate amount of enzyme. Specific activities were calculated using a molar absorption coefficient for NDMA of 35400 M-1 cm-1 at 440 nm (Dunn & Bernard, 1971 ). In order to determine whether extracts exhibited NDMA-dependent alcohol oxidation activity for the alcohol used as growth substrate, benzyl alcohol in the assay was replaced by this alcohol. The optimal concentration of the latter (as indicated by the values in parentheses in Table 1
), yielding an apparent maximal rate, was determined by varying each substrate in the range from 5 µM to 25 mM (or up to saturation in the case of alcohols with low solubility). MNO activity was measured in 50 mM potassium phosphate buffer, pH 6·3, by following the reduction of NDMA (28 µM, final concentration), starting the reaction by the addition of methanol (25 mM, final concentration) (Bystrykh et al., 1993b
). NAD(P)-dependent alcohol dehydrogenase activities were determined in 0·1 M sodium pyrophosphate buffer, pH 9·0 (Duine et al., 1984
) by measuring the formation rate of NAD(P)H at 340 nm in the presence of the alcohol used as growth substrate (apparent maximal rates were determined as indicated above). Assays for aldehyde dismutase activity were carried out in a 1·5 ml reaction vessel containing (final concentrations) 50 mM potassium phosphate buffer, pH 7·0, aldehyde (formaldehyde, 5 mM; acetaldehyde, 2 mM; propionaldehyde, 0·1 mM; methylglyoxal, 5 mM; benzaldehyde, 5 mM; phenylacetaldehyde, 0·2 mM) and an appropriate amount of enzyme (typically 0·67 µM) in a total volume of 1·0 ml. The reaction was started by the addition of enzyme. Samples (100 µl) were taken at regular intervals and the reaction was stopped by the addition of 2 µl formic acid (100%) [in the case of formaldehyde, aqueous acetic acid (10%) was used instead]. After centrifugation (2 min at 20000 g), the remaining substrate and the products were determined by injecting 1·0 µl of supernatant into the HP-Innowax GC-column (see below). However, in the case of benzaldehyde and phenylacetaldehyde, analysis was done according to the HPLC method described below for the veratryl alcohol/methylglyoxal interconversion. Formaldehyde solutions were prepared by heating paraformaldehyde in water for 8 h at 100 °C.
Substrate specificity.
Values for V'max, K'm and K'i of the NDMA-linked dehydrogenase activity of np-ADH were determined by varying the concentration of the alcohol or aldehyde substrate (in the range from 1 µM to 200 mM, or up to saturation) in the assay mixture at a constant concentration of NDMA (28 µM). The apparent kinetic parameter values of the alcohol:carbonyl oxidoreductase activity of np-ADH were determined by varying the concentration of the aldehyde or ketone substrate (from 1 µM up to saturation) at a fixed concentration of veratryl alcohol (1 mM) and measuring the initial rates of veratraldehyde formation at 310 nm (310=9300 M-1 cm-1). The steady-state kinetic data were analysed by non-linear regression using an equation derived from MichaelisMenten kinetics (with a single substrate), including a substrate-inhibition term (Cornish-Bowden, 1995
), using Igor Pro software (WaveMetrics) on a Power Macintosh G3 computer (Apple).
Interconversion of the veratryl alcohol/methylglyoxal combination.
The np-ADH-catalysed oxidation of veratryl alcohol to veratraldehyde with the concomitant reduction of methylglyoxal was carried out at 20 °C in a quartz cuvette. The reaction mixture (total volume 2 ml) consisted of 10 mM potassium phosphate buffer, pH 7·0, 0·15 mM veratryl alcohol, 1·0 mM methylglyoxal and 0·1 µM np-ADH. The progression of the conversion with time was monitored by measuring veratraldehyde formation at 310 nm. Samples (25 µl) were taken at regular intervals and the reaction was quenched by adding 50 µl of the HPLC column eluent (aqueous 30%, v/v, acetonitrile containing 1%, v/v, acetic acid), and centrifuging (2 min at 20000 g). An aliquot of the supernatant (20 µl) was applied to a Nova-pak C18 reversed-phase column (150x3·9 mm, Waters) followed by isocratic elution at a flow rate of 0·8 ml min-1 (Piersma, 1998 ). The eluate was monitored at 236 nm, 278 nm and 310 nm using a UV/VIS photodiode-array detector (Hewlett Packard, model 1040 A). Baseline separation could be achieved with this setup for veratryl alcohol, veratraldehyde and veratric acid, allowing reliable integration of the peak areas of the compounds using Chemstation chromatography software (Hewlett Packard). Quantitative data were obtained by using calibration curves of the authentic compounds chromatographed in the same way.
Molecular mass determinations.
The subunit molecular mass of np-ADH was determined by SDS-PAGE (Laemmli, 1970 ) using 825% polyacrylamide gradient gels on a Phast-System (Pharmacia) according to the manufacturers instructions. A low-molecular-mass marker kit (Pharmacia) was used for calibration and Coomassie brilliant blue R250 for protein staining. The molecular mass of native np-ADH was determined by gel filtration on a Superdex 200 (30x1 cm) column equilibrated with 10 mM potassium phosphate buffer, pH 7·0, containing 0·2 M KCl, at a flow rate of 0·5 ml min-1. Blue dextran (2000 kDa), thyroglobulin (669 kDa), ferritin (440 kDa), catalase (232 kDa), aldolase (158 kDa) and bovine serum albumin (BSA, 67 kDa) were used as molecular mass references.
Protein determination.
Protein concentrations were estimated using the bicinchoninic acid/CuSO4 method (Pierce) with desalted BSA as a standard (Smith et al., 1985 ).
N-terminal sequence analysis.
The N-terminal amino acid sequence of the enzyme (the blotted band obtained with SDS-PAGE) was determined by automated Edman degradation using a Procise 494 sequenator (Applied Biosystems Division, Perkin Elmer) running in pulsed-liquid mode.
Determination of bound NADH.
The enzyme-bound chromophore was dissociated from the enzyme using a slightly modified urea-extraction procedure (Bystrykh et al., 1993a ). For that purpose, an aliquot of enzyme in 0·1 M Tris/HCl, pH 8·5, was brought to 6 M urea by adding a saturated urea solution in the same buffer. The mixture was heated for 2 min at 85 °C. After cooling and centrifugation (5 min at 20000 g), the supernatant was applied to a MonoQ (0·5x5 cm) column equilibrated with 10 mM Tris/HCl, pH 8·5, containing 6 M urea. Elution was done at a flow rate of 1 ml min-1 with a linear gradient of 01 M KCl in the same buffer. The eluate was monitored with the UV/VIS photodiode-array detector. The resulting elution profile and the absorption spectra of the compounds corresponding to the peaks in the chromatogram were compared with those of urea-treated authentic NAD, NADH, NADP and NADPH.
Fluorescence spectroscopy.
Fluorescence spectra were obtained at room temperature using a Shimadzu spectrofluorimeter, model RF-5001PC. Fluorescence emission spectra were recorded from 350 to 600 nm using a fixed excitation wavelength of 334 nm (bandwidth 3 nm). Fluorescence excitation spectra were obtained by scanning from 250 to 400 nm and recording the fluorescence emission at a fixed wavelength of 430 nm (bandwidth 3 nm).
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RESULTS |
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Purification and structural properties
The induction studies indicated that benzyl alcohol as growth substrate gives rise to the highest level of NDMA-dependent alcohol dehydrogenase activity without inducing MNO or NAD(P)-dependent alcohol dehydrogenase activities. Therefore, this alcohol was chosen as a growth substrate to produce cells as starting material for the purification of np-ADH. The data for the purification protocol of a typical batch are presented in Table 2. SDS-PAGE of the final preparation revealed a single band corresponding to an Mr of 38000 (±5%). Gel-filtration chromatography of native enzyme yielded a single peak, which corresponded to an Mr of 150000 (±10%). Since overlaying of the chromatograms taken at 205, 280 and 330 nm showed a single, symmetrical peak with the same retention time, it is concluded that the final enzyme preparation was homogeneous. A single N-terminal amino acid sequence was detected (Fig. 1
), showing high sequence identity to that of np-ADH from A. methanolica (van Ophem et al., 1993
), i.e. 77% identical amino acid residues out of 22. This suggested that the enzyme purified is an np-ADH which has, according to the data given above, a homotetrameric structure.
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From the apparent kinetic parameter values in Table 3(a, b
), it follows that variation in the specificity constant values is mainly caused by the K'm values, not by the V'max values. The best substrates are primary alcohols containing a phenyl group (compare methanol and ethanol with benzyl alcohol and 2-phenylethanol) or an aliphatic chain of a certain length (pentan-1-ol up to heptan-1-ol). Substitution of the phenyl ring (with hydroxyl and/or methoxy groups) results in less favourable substrates, unless the aliphatic chain is enlarged somewhat. Branching of the aliphatic chain also gives improvement (compare butan-1-ol with 3-methylbutan-1-ol). Introduction of a double bond has a negative effect (compare propan-1-ol with 2-propen-1-ol) but this can be compensated by substitution with a phenyl group (compare 2-propen-1-ol with cinnamyl alcohol).
In many cases, severe substrate inhibition was observed (often in the submillimolar range), hampering the determination of kinetic parameter values, especially when there were small differences in K'm and K'i values. This indicates that in such cases the substrate binds nearly equally well to the oxidized and the reduced enzyme forms.
Conversion experiments
For reasons of analytical simplicity, most enzyme-catalysed conversion experiments were carried out with the couple veratryl alcohol/methylglyoxal. As shown in Fig. 3, a small amount of np-ADH catalysed the complete oxidation of veratryl alcohol into veratraldehyde with a stoichiometric amount of methylglyoxal as electron acceptor, in line with the fact that no veratric acid was found in the final reaction mixture. Complete conversion was also achieved using the couple veratraldehyde/ethanol, although the reaction proceeded approximately 10-fold slower than that for the couple veratryl alcohol/methylglyoxal. These observations indicate that np-ADH is stable for several hours in mixtures of alcohols and aldehydes during catalysis. Rapid, irreversible inactivation of np-ADH was observed (typically within a few minutes) when performing alcohol oxidation in the presence of NDMA (not shown). This phenomenon could be partly suppressed by adding DTT (or other thiols) to the assay mixture. Incubating the enzyme with solely NDMA, an alcohol, or an aldehyde did not lead to any inactivation (unpublished results). It seems, therefore, that the inactivation of np-ADH in the assay mixture is caused by products generated by the reduction of NDMA, an observation which has also been made in the case of horse liver alcohol dehydrogenase (Dunn & Bernard, 1971
).
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DISCUSSION |
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In view of the induction of np-ADH by many alcohols, as deduced from the activity exhibited by extracts in the assay with benzyl alcohol and NDMA, the absence or low activity of other alcohol dehydrogenases, and the broad substrate specificity of np-ADH, the enzyme may be the general alcohol-oxidizing enzyme for R. erythropolis DSM 1069. Since its assay is not standard practice, the significance of np-ADH may be even wider, as it may have been overlooked so far in other organisms. Accordingly, a search in the databases was made to see whether structural evidence could be found. It appeared that the N-terminal amino acid sequence of the AdhD gene product of Mycobacterium tuberculosis H37Rv (Cole et al., 1998 ), which was designated by the authors as a putative alcohol dehydrogenase, has 73% identical amino acids, as shown in Fig. 1
. Also, a significant sequence identity (51%) was observed with a hypothetical alcohol dehydrogenase from R. rhodochrous NCIMB 13064, whose corresponding gene occurs in a dehalogenation operon (Kulakova et al., 1997
), and with the AdhB gene product (46%) from M. tuberculosis (not shown). Recently, the complete amino acid sequence of np-ADH from A. methanolica has been determined and structural modelling was carried out using this sequence and the known three-dimensional structure of horse liver alcohol dehydrogenase (see Piersma, 1998
). Taking the differences with respect to structural and catalytic aspects into account, this provided a number of amino acid residues which can be used as markers to distinguish between the two types of alcohol dehydrogenase (Table 4
). Applying this to the sequences found in the databases, it appears that characteristic amino acid residues for np-ADH are present, most of them in the gene product of the adhD gene of M. tuberculosis and fewer in those of the hypothetical alcohol dehydrogenase from R. rhodochrous and the AdhB gene product of M. tuberculosis (Table 4
). It can be anticipated, therefore, that np-ADHs may also be found in other nocardioform actinomycetes.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Ashraf, W. & Murrell, J. C. (1992). Genetic, biochemical and immunological evidence for the involvement of two alcohol dehydrogenases in the metabolism of propane by Rhodococcus rhodochrous PNKb1.Arch Microbiol 157, 488-492.
Bell, K. S., Philp, J. C., Aw, D. W. J. & Christofi, N. (1998). The genus Rhodococcus.J Appl Microbiol 85, 195-210.[Medline]
Bystrykh, L. V., Vonck, J., van Bruggen, E. F. J., van Beeumen, J., Samyn, B., Govorukhina, N. I., Arfman, N., Duine, J. A. & Dijkhuizen, L. (1993a). Electron microscopic analysis and structural characterization of novel NADP(H)-containing methanol:N,N'-dimethyl-4-nitrosoaniline oxidoreductases from the Gram-positive methylotrophic bacteria Amycolatopsis methanolica and Mycobacterium gastri MB19.J Bacteriol 175, 1814-1822.[Abstract]
Bystrykh, L. V., Govorukhina, N. I., van Ophem, P. W., Hektor, H. J., Dijkhuizen, L. & Duine, J. A. (1993b). Formaldehyde dismutase activities in Gram-positive bacteria oxidizing methanol.J Gen Microbiol 139, 1979-1985.
Bystrykh, L. V., Govorukhina, N. I., Dijkhuizen, L. & Duine, J. A. (1997). Tetrazolium dye-linked alcohol dehydrogenase of the methylotrophic actinomycete Amycolatopsis methanolica is a three-component complex.Eur J Biochem 247, 280-287.[Abstract]
Cole, S. T., Brosch, R., Parkhill, J. & 38 other authors (1998). Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393, 537544.[Medline]
Cornish-Bowden, A. (1995). Fundamentals of Enzyme Kinetics, 2nd edn. London: Portland Press.
Duine, J. A., Frank, J. & Berkhout, M. P. J. (1984). NAD-dependent, PQQ-containing methanol dehydrogenase: a bacterial dehydrogenase in a multienzyme complex.FEBS Lett 168, 217-221.[Medline]
Dunn, M. F. & Bernard, S. A. (1971). Rapid kinetic evidence for adduct formation between the substrate p-nitroso-N,N-dimethylaniline and reduced nicotinamide-adenine dinucleotide during enzymic reduction.Biochemistry 10, 4569-4575.[Medline]
Eggeling, L. & Sahm, H. (1984). An unusual formaldehyde oxidizing system in Rhodococcus erythropolis grown on compounds containing methyl groups.FEMS Microbiol Lett 25, 253-257.
Eggeling, L. & Sahm, H. (1985). The formaldehyde dehydrogenase of Rhodococcus erythropolis, a trimeric enzyme requiring a cofactor and active with alcohols.Eur J Biochem 150, 129-134.[Abstract]
Eklund, H., Müller-Wille, P. & Horjales, E. (1990). Comparison of three classes of human liver alcohol dehydrogenase. Emphasis on different substrate binding pockets.Eur J Biochem 193, 303-310.[Abstract]
Finnerty, W. R. (1992). The biology and genetics of the genus Rhodococcus.Annu Rev Microbiol 46, 193-218.[Medline]
Geerlof, A., van Tol, J. B. A., Jongejan, J. A. & Duine, J. A. (1994). Enantioselective conversion of the racemic C3-alcohol synthons, glycidol (2,3-epoxy-1-propanol), and solketal (2,2-dimethyl-4-(hydroxymethyl)-1,3-dioxolane) by quinohaemoprotein alcohol dehydrogenases and bacteria containing such enzymes.Biosci Biotechnol Biochem 58, 1028-1036.
Hektor, H. J. (1997). Physiology and biochemistry of primary alcohol oxidation in the Gram-positive bacteria Amycolatopsis methanolica and Bacillus methanolicus. PhD thesis, Groningen State University.
Hektor, H. J. & Dijkhuizen, L. (1996). Mutational analysis of primary alcohol metabolism in the methylotrophic actinomycete Amycolatopsis methanolica.FEMS Microbiol Lett 144, 73-79.
Jaeger, E. (1988). Purification of coniferyl alcohol dehydrogenase from Rhodococcus erythropolis.Methods Enzymol 161, 301-306.
Jaeger, E., Eggeling, L. & Sahm, H. (1981). Partial purification and characterization of a coniferyl alcohol dehydrogenase from Rhodococcus erythropolis.Curr Microbiol 6, 333-336.
Kersten, P. J., Stephens, S. K. & Kirk, T. K. (1990). Glyoxal oxidase and the extracellular peroxidases of Phanerochaete chrysosporium. In Biotechnology in Pulp and Paper Manufacture, pp. 457-463. Edited by T. K. Kirk & H.-M. Chang. Stoneham, MA: Butterworth-Heinemann.
Krier, F., Kreit, J. & Millière, J. B. (1998). Characterization of partially purified alcohol dehydrogenase from Rhodococcus sp. strain GK1.Lett Appl Microbiol 26, 283-287.
Kulakova, A. N., Larkin, M. J. & Kulakov, L. A. (1997). The plasmid-located haloalkane dehalogenase gene from Rhodococcus rhodochrous NCIMB 13064.Microbiology 143, 109-115.[Abstract]
Laemmli, U. K. (1970). Cleavage of structural proteins during assembly of the head of bacteriophage T4.Nature 227, 680-685.[Medline]
Ludwig, B., Akundi, A. & Kendall, K. (1995). A long-chain secondary alcohol dehydrogenase from Rhodococcus erythropolis ATCC 4277. Appl Environ Microbiol 61, 3729-3733.
Misset-Smits, M., van Ophem, P. W., Sakuda, S. & Duine, J. A. (1997). Mycothiol, 1-O-(2'-[N-acetyl-L-cysteinyl]amino-2'-deoxy--D-glucopyranosyl)-D-myo-inositol, is the factor of NAD/factor-dependent formaldehyde dehydrogenase.FEBS Lett 409, 221-222.[Medline]
Nagy, I., Verheijen, S., de Schrijver, A., van Damme, J., Proost, P., Schoofs, G., Vanderleyden, J. & de Mot, R. (1995). Characterization of the Rhodococcus sp. NI86/21 gene encoding alcohol:N,N'-dimethyl-4-nitrosoaniline oxidoreductase inducible by atrazine and thiocarbamate herbicides.Arch Microbiol 163, 439-446.[Medline]
van Ophem, P. W. & Duine, J. A. (1994). NAD- and co-substrate (GSH or factor)-dependent formaldehyde dehydrogenases from methylotrophic microorganisms act as a class III alcohol dehydrogenase.FEMS Microbiol Lett 116, 87-94.
van Ophem, P. W., van Beeumen, J. & Duine, J. A. (1992). NAD-linked, factor-dependent formaldehyde dehydrogenase or trimeric, zinc-containing, long-chain alcohol dehydrogenase from Amycolatopsis methanolica.Eur J Biochem 206, 511-518.[Abstract]
van Ophem, P. W., van Beeumen, J. & Duine, J. A. (1993). Nicotinoprotein (NAD(P)-containing) alcohol/aldehyde oxidoreductases. Purification and characterization of a novel type from Amycolatopsis methanolica.Eur J Biochem 212, 819-826.[Abstract]
Peters, J., Zelinski, T. & Kula, M.-R. (1992). Studies on the distribution and regulation of microbial keto ester reductases.Appl Microbiol Biotechnol 38, 334-340.
Peters, J., Zelinski, T., Minuth, T. & Kula, M.-R. (1993). Synthetic applications of the carbonyl-reductases isolated from Candida parapsilosis and Rhodococcus erythropolis.Tetrahedron Asymm 4, 1683-1692.
Piersma, S. R. (1998). Structure and catalytic mechanism of nicotinoprotein alcohol dehydrogenases from Amycolatopsis methanolica. PhD Thesis, Delft University of Technology.
Piersma, S. R., Visser, A. J. W. G., de Vries, S. & Duine, J. A. (1998). Optical spectroscopy of nicotinoprotein alcohol dehydrogenase from Amycolatopsis methanolica: a comparison with horse liver alcohol dehydrogenase and UDP-galactose epimerase.Biochemistry 37, 3068-3077.[Medline]
de Schrijver, A., Nagy, I., Schoofs, G., Proost, P., Vanderleyden, J., van Pée, K.-H. & de Mot, R. (1997). Thiocarbamate herbicide-inducible nonheme haloperoxidase of Rhodococcus erythropolis NI86/21.Appl Environ Microbiol 63, 1911-1916.[Abstract]
Smith, P. K., Krohn, R. I., Hermanson, G. T. & 7 other authors (1985). Measurement of protein using bicinchoninic acid. Anal Biochem 150, 7685.[Medline]
Warhurst, A. M. & Fewson, C. A. (1994). Biotransformations catalyzed by the genus Rhodococcus.Crit Rev Biotechnol 14, 29-73.[Medline]
Zelinski, T., Peters, J. & Kula, M.-R. (1994). Purification and characterization of a novel carbonyl reductase isolated from Rhodococcus erythropolis.J Biotechnol 33, 283-292.[Medline]
Received 16 August 1999;
revised 2 November 1999;
accepted 23 December 1999.
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