Alkane Oxidation by a Particulate Preparation from Candida

Transcription

1 JOURNAL OF BACTERIOLOGY, June 1971, p Copyright i 1971 American Society for Microbiology Vol. 106, No. 3 Printed in U.S.A. Alkane Oxidation by a Particulate Preparation from Candida CHAO-MIN LIU AND MARVIN J. JOHNSON Department of Biochemistrv, University of Wisconsin, Madison, Wisconsin Received for publication 5 February 1971 The oxidation of decane by a cell-free particulate preparation from Candida intermedia was studied. Decane is oxidized to decanoate via decanol and decanaldehyde. Oxidation of decane to decanol requires molecular oxygen. Decanol is oxidized to decanaldehyde by a nicotinamide adenine dinucleotide-linked dehydrogenase differing greatly in specificity from ordinary yeast alcohol dehydrogenase. Decanaldehyde is oxidized to decanoate by a nicotinamide adenine dinucleotidelinked dehydrogenase that oxidizes long-chain aldehydes but not short-chain aldehydes. The enzymes that oxidize decane, decanol, and decanaldehyde are all induced when decane is present in the medium. These enzymes are apparently located in the cell membrane. The initial reaction in oxidation of alkanes by bacteria has been shown by Peterson et al. (6, 7) to be hydroxylation of the terminal methyl group by an oxygenase. It is to be expected that oxidation of alkanes by yeasts should involve a similar initial reaction. lizuka et al. (2) concluded that, in Candida, alkanes are oxidized anaerobically to I-alkenes by nicotinamide adenine dinucleotide (NAD). It has been pointed out (3) that such a route is thermodynamically unfeasible. It was our intention in the work described here to determine whether the oxidation of alkanes by Candida proceeded via oxygenase hydroxylation and to determine the location within the cell of the alkane-oxidizing system. A particulate enzyme preparation was obtained that oxidized decane to decanoic acid. Fractionation of the preparation showed that the enzyme system was contained in the cytoplasmic membrane, not in the mitochondria. The enzyme preparation required molecular oxygen to oxidize decane. It also oxidized decanol and decanaldehyde to decanoic acid with NAD. Radioactivitytrapping experiments showed that decanol and decanaldehyde were intermediates in decane oxidation. The entire membrane-bound enzyme system was induced by decane. MATERIALS AND METHODS Organisms. C. intermedia NRRL-Y6328-1, originally isolated from soil with hexadecane as carbon source (5), was grown on decane in the following medium (in grams per liter): NH4H2PO4, 5.0; KH2PO4, 0.7; MgSO4-7H2O, 0.4; CaCl2 2H2O, 0.1; NaCl, 0.1; ZnSO4 5H2O, 4 x 10-4; MnSO4 H2O, 6 x 10-4; 830 FeCI3, 4 x 10 4; CuSO4-5H2O, 2.5 x The ph was adjusted to 5.5 with NaOH. A 1-mI amount of decane and 8 mg of yeast extract (Difco) per 100 ml of medium were added just before inoculation. With this medium and the conditions described below, the generation time was 7.2 hr. The cell yield on decane was found to be approximately 0.6 g of dry cells per g of decane in an experiment run with a 2-liter fermentor. Other yeasts listed in Table 3 were also grown in the same manner, except that various carbon sources were used as required. The generation time for C. intermedia growing on glycerol in a shaken flask was about 8 hr. C. utilis (UW no. 3), Candida sp. (E. Mc. no. 80), and Saccharomyces carisbergensis (ATCC no. 9080) were obtained from Elizabeth McCoy of the Department of Bacteriology; C. pseudotropicalis (NRRL Y- 83) was obtained from D. A. Stuiber of the Department of Food Science. Particulate cell-free preparation. The yeast was grown at 30 C on a rotary shaker in a 2-liter flask containing 500 ml of medium. Cells were harvested by centrifugation during exponential growth and washed with cold distilled water. The ph of the culture at harvest was about 3.5. The amount of cells obtained was about I g (dry weight). The cells so prepared could be stored in a freezer (-20 C) for months without losing decaneoxidizing activity. To obtain a cell-free particulate preparation, the washed cell paste was suspended in an equal volume of 0.02 M potassium phosphate buffer (ph 7.0) containing 5 x 10-4 M dithiothreitol and 0.25 M sucrose, and was treated in a French press at 15,000 psi twice at 0 to 5 C. Cell debris and unbroken cells were removed by 10- min centrifugation at 800 x g. Centrifugation of the supernatant at 10,000 x g for 30 min yielded a particulate fraction that oxidized decane to decanoic acid. The supernatant fraction did not oxidize decane. Higher activity was found in the cell debris fraction, but this

2 VOL. 106, 1971 ALKANE OXIDATION 831 fraction was not used because it was contaminated by unbroken cells. The particulate fraction [containing at least three enzymes: decane-oxidizing enzyme(s), decanol dehydrogenase, and decanaldehyde dehydrogenasel was washed with the same buffer (containing no sucrose) and was routinely used for study. One gram of cells (dry basis) yjelded about 60 mg of particulate fraction, containing about 37 mg of protein as determined by the method of Lowry et al. (4) with bovine serum albumin as standard. Enzyme assays. Because the observed rate of conversion of decanol to decanoate was always much greater than the rate of formation of decanol from decane, the formation of radioactive decanoate from decane-l-'4c could be used as a measure of enzyme activity. For routine assay, nmoles of decane-l- '4C (1.1 x 106 dpm) dissolved in 95% ethanol (5 uliters) was incubated for I hr at 30 C with the enzyme preparation and 90 Mmoles of glycine-naoh buffer (ph 10.0) in a total volume of I ml. Ethyl alcohol did not affect the enzyme activity appreciably at concentrations below 0.4 M. Under these assay conditions, the amount of radioactive decanoate formed (up to about 3 nmoles) was proportional to the amount of cell-free preparation added. It was found that activity was greatest at ph 10 even though the decanol and decanaldehyde dehydrogenases had a ph optimum close to 9. Apparently the oxygenase system, which was rate-limiting, was most active at ph 10. The rate of decanoate formation did not increase when the reaction mixture was continuously shaken during the assay, showing that the reaction was not limited by the oxygen transfer. After incubation, the mixture was acidified with 0.1 ml of 15% sulfuric acid and extracted twice with an equal volume of ether. The ether extracts were combined and extracted with 0.1 M KOH. Samples of the KOH solution were chromatographed on paper strips, which were scanned for radioactivity. Decanol and decanaldehyde dehydrogenase activities were determined by following the reduction of NAD at 340 nm with a Beckman DB spectrophotometer used with a recorder. In the routine assay, 2.0 ml of tris(hydroxymethyl)aminomethane-hydrochloride buffer (0.01 M), ph 8.9, 2 umoles of NAD in distilled water, and 0.1 ml of enzyme preparation were combined in both reference and sample cuvettes. Water was added to a final volume of 3.0 ml. The reaction was started by adding aqueous suspension of substrate (5 x 10-4 M, final concentration) to the sample cuvette. One unit of dehydrogenase activity was defined as the amount causing the reduction of I umole of NAD per min. Specific activity was expressed as units per milligram of protein. Initial velocity was taken as a measure of enzyme activity. Since the solubility of decanol and decanaldehyde in water are very low, being of the order of 10-5 M at room temperature, substrates to be used were sonically dispersed in distilled water immediately before use. Chromatography. Paper chromatography was carried out by the ascending method on Whatman no. I filter paper with the following solvent systems: solvent system 1, 100 ml of p-dioxane plus I ml of 1.5 N aqueous NH3; solvent system 2, 100 ml of 95% ethanol plus I ml of concentrated (28%) NHs; solvent system 3, 100 ml of 95% ethanol plus I ml of 88% HCOOH. Radioactivity measurements. Decane-1-'4C was determined with a liquid scintillation spectrometer (Packard Instrument Co., Inc.). Radioactivity in decanoic acid formed by the enzyme reaction was determined by the following methods. A 5-inch (ca cm) wide paper chromatogram (routinely developed with solvent system 2) of the KOH extract from enzyme assay was cut into 1-inch (2.54-cm) strips, and each strip was scanned for radioactivity by a Vanguard model 880 automatic chromatogram scanner. For radioactivity measurement on thin-layer plates, silica gel from an area (I by 3 cm) of the chromatogram plate was transferred to a counting vial and counted with a liquid scintillation counter. Decane-l-14C was purchased from Nuclear-Chicago Corp. It was dissolved in 95% ethanol and used without addition of carrier. Decanol-1-"4C was obtained from Mallinckrodt/Nuclear, St. Louis, Mo. It was dissolved in 35% aqueous solution of t-butanol and used directly for enzyme assay. RESULTS Properties of the decane-oxidizing system. Aerobic incubation of radioactive decane with cell-free enzyme preparation under the conditions described above produced only one radioactive compound in the KOH extract. This was characterized as decanoic acid by paper chromatographic comparison with authentic decanoic acid, with the three solvent systems described, and by preparation of the p-bromophenacyl derivative. The derivative was repeatedly crystallized without significant change in specific activity. Formation of decanoate was used as an assay for decane-oxidizing activity. Molecular oxygen was found to be essential for the reaction. In an experiment in which 520 Ag of extract protein was incubated for 60 min, 9.6 nmoles of decanoate was formed in the presence of oxygen, but only 0.4 nmoles was formed when oxygen was excluded. Oxygen could not be replaced by ferricyanide, 2,6-dichloroindophenol, flavine adenine dinucleotide, flavine mononucleotide, or methylene blue. The activity was inhibited by sodium azide and by potassium cyanide (98 and 87%, respectively, at 0.01 M). Since (as shown below) the oxidation proceeds via decanol and decanaldehyde, and since these compounds are oxidized anaerobically, only the oxidation of decane to decanol requires molecular oxygen. The decane-oxidizing enzyme system is apparently located in the cell membrane. Mitochondria prepared by the use of snail enzyme (Helicase, obtained from Industrie Biologique Francaise) had no activity. Lysed mitochondria were also inactive. Formation of decanol and decanaldehyde from decane. During decane oxidation to decanoate,

3 832 LIU AND JOHNSON J. BACTERIOL. no detectable decanol or decanaldehyde accumulated. However, when unlabeled decanol or decanaldehyde was present in the reaction mixture at 10-3 M when decane-1-'4c was being oxidized, little radioactive decanoate was formed, but radioactivity accumulated in the pool of added intermediate. Unlabeled decanol and decanaldehyde were readily oxidized by the preparation. In one experiment, enzyme preparation (0.37 mg of protein) was incubated with nmoles of decane-1-14c (1.1 x 106 dpm) in the presence of unlabeled decanol (5 gmoles) and 90,imoles of glycine-naoh buffer (ph 10.0) for 75 min at 30 C in a total volume of I ml and then extracted directly with ether. Ether extracts from 10 such incubation tubes were combined and used to prepare the 3,5-dinitrobenzoate ester of decanol, which was subjected to thin-layer chromatography with silica gel as an adsorbent. The 3,5-dinitrobenzoate ester was located on the chromatogram either by ultraviolet light or by spraying with a 1% solution of a-naphthylamine in ethanol. Co-chromatography with the authentic derivative and measurement of radioactive distribution on the chromatograms (silica gel as an adsorbent) with two solvent systems (benzenepentane, 1: 1, and benzene-ethyl acetate, 95:5) indicated that "C-labeled decanol was formed enzymatically from decane-1-14c and trapped by the unlabeled decanol. Enzymatic oxidation of radioactive decane to decanaldehyde was demonstrated in a similar manner. The 2,4-dinitrophenylhydrazone of decanaldehyde was prepared and shown to be radioactive by similar procedures. Oxidation of decanol to decanaldehyde and decanoic acid. The same particulate fraction that catalyzed the formation of radioactive decanoic acid from decane-1-14c also oxidized decanol to decanoic acid. Figure I shows the time course of decanol oxidation. Radioactive decanol-14c was used for the assay. The results indicate that I mole of decanol was oxidized, presumably via decanaldehyde, to I mole of decanoic acid with concomitant reduction of two equivalents of NAD. Decanaldehyde was barely detectable in the reaction mixture, probably because of its low steady-state concentration in the reaction sequence. Decanoic acid was identified as the final product of decanol oxidation by the methods already described, and decanaldehyde was established as an intermediate by a radioactivity-trapping experiment like that already described. The particulate enzyme preparation was able to oxidize either decanol or decanaldehyde as substrate with NAD (not with nicotinamide adenine dinucleotide phosphate). Dehydrogenase activity was found in the particles that could be brought down completely by centrifuging the French-pressed cell suspension at 105,000 x g for 1 hr, as well as in the usual 10,000 x g fraction. Most of the studies on dehydrogenase properties were done with the fraction between 10,000 x g (30 min) and 105,000 x g (1 hr), which was found to have a higher specific activity. The dehydrogenase activity was completely inhibited by sulfhydryl reagents such as p-chloromercurobenzoate at 3.3 x 10-' M. The optimum ph for both alcohol and aldehyde dehydrogenation was 8.0 to 9.0. The observed substrate specificity is shown in Tables I and 2. It will be noted that short-chain alcohols and aldehydes were not oxidized. Ordinary yeast alcohol (8) or aldehyde (1) dehydrogenases attack long-chain substrates more slowly than short-chain substrates. M ichaelis constants }_0.2 I LJ 0 I a 12 MINUTES DGNOL NADH DECANOATE DECANALDEHYDE FIG. 1. Time course of decanol oxidation. The reaction mixture contained 200 Amoles of tris(hydroxymethyl)aminomethane-hydrochloride buffer, ph 7.5, 0.958,umole of decanol-l-'4c (specific activity, 2.61 mci/mmole, dissolved in 0.05 ml of 35% aqueous solution of t-butanol), and 12,moles of NA D and enzyme preparation (320,ug of protein) in a total volume of 3.5 ml. Assay was carried out at 30 C. Enzyme was used to start the reaction. A zero-time control was run with the same amount of boiled enzyme. Reduced NAD formation was measured spectrophotometrically. For estimation of decanol-l-'4c oxidation, a 0.5-mI sample of the reaction mixture was added to a I by 15 cm test tube containing 0.1 ml of 0.2 N H2SO4. Two micromoles each of unlabeled decanol, decanaldehyde, and decanoate were then added, and the mixture was adjusted to ph 10.5 with 0.2 N KOH. ft was then extracted with ether. Decanaldehyde was separated from decanol in the ether extract by shaking with 3 ml of aqueous solution of 10-- I M hydrazinobenzene sulfonate at ph 5.0. Decanoate in the aqueous layer after the first ether extraction was extracted with ether after acidification and reextracted with aqueous KOH. The resultant fractions were then suitably diluted and counted for radioactivity in a liquid scintillation counter with dioxane as solvent.

4 VOL. 106, 1971 ALKANE OXIDATION 833 were determined to be 3.85 x 10-6 M for decanol and 1.76 and 10-6 M for decanaldehyde [tris(hydroxymethyl)aminomethane buffer, ph 8.9]. Induction of the decane-oxidizing system. The specific activity of enzyme preparations made from cells growing on lactose was low. When TABLE 1. Relative dehydrogenation rates of alcoholsa Relative Alcohol (Coc) velocity (decanol = 100) Ethanol < 0.05 Butanol < 0.05 Pentanol Hexanol Octanol Decanol...h-b 100 Undecanol III Hexadecanol Octanol Heptanol a The preparation used had been sedimented at 105,000 x g for I hr. In some cases, 1.7% t-butanol was present to aid solution of the substrate. At the concentration used, t-butanol was not inhibitory and was not oxidized by the preparation. Reaction conditions were as described in Materials and Methods. bspecific activity of the preparations used varied from 0.08 to 0.4 units per mg of protein. For each substrate, the velocity is reported in terms of the velocity with decanol at the same concentration. such cells were transferred to a growth medium containing decane (0.01 M), the enzyme activity rapidly increased. Induction was complete after 2 to 4 hr after a cell weight increase of roughly 30%. Data on activity of enzyme preparations for a number of organisms and the effect of decane induction on the activity are shown in Table 3. It is apparent that the decane-oxidizing oxygenase is inducible and that the alcohol and aldehyde dehydrogenases are also induced. The dehydrogenases which are not cytoplasmic and whose specificity is directed toward high-molecularweight substrates appear to be parts of a membrane-bound hydrocarbon oxidation system. Comparison of the activity of our particulate preparations with that of the intact cell shows that only about 0.1 % of the decane-oxidizing ac- TABLE 2. Relative dehydrogenation rates of aldehydesa Aldehvde Relative rate A-dehyde (decanaldehyde= 100) C2 to C7 aldehydes... <0.1 Octanaldehyde Decanaldehyde Dodecanaldehyde TABLE 3. Decane-oxidizing enzyme system in veastsa a The substrate concentration was 5 x 10-4 M in all cases. The activity of the preparation used (centrifuged at 105,000 x g for I hr) on decanaldehyde was 0.07 unit per mg of protein. Enzyme activity' Decane-l- 4C Growth on oxidized' Decanol Decanaldehyde Organism hexadecane dehydrogenase dehydrogenase Control Decane- Control Decaneinduced inducedinue Candida intermedia (NRRL Y ) Candida sp (E.Mc No. 80) Candida utilis < < (UW no. 3) Candida pseudotropicalis <0.1 <0.1 <0. <0. I (NRRL-Y83) Saccharomyces carlsbergensis <0. I <0. I <0.1 <0. I (ATCC no. 9080) a Yeasts were grown on glycerol as their sole source of carbon, except that C. pseudotropicalis and Candida sp. E.Mc. no. 80 were grown on glucose because of their extremely slow growth on glycerol. Induction by decane was carried out as follows. A growing culture (about 1.5 g, dry weight, per liter) was divided into two portions, to one of which 0.01 m decane was added. Induction was complete in about 6 hr. Decane-oxidizing enzyme activity was assayed as described in Materials and Methods, except that the total volume of the assay mixture was reduced to 0.5 ml, containing nmoles of decane-j- 4C and 0.4 mole of 0.1 M glycine-naoh buffer, ph Intact cells were used. Total KOH-extractable acidic products formed in I hr were taken as a measure of n-decane-oxidizing enzyme activity. Dehydrogenase activities were determined as described. Expressed as micromoles of decane-1-'4c oxidized per hour per gram of cells. Expressed as units per gram of protein.

5 834 LIU AND JOHNSON J. BACTERIOL. tivity of the cell is recovered in our preparations, but the preparations have an activity on decanol and decanaldehyde of the same order of magnitude as that of the intact cell. ACKNOWLEDGMENTS This investigation was supported by Public Health Service grant Al from the National Institute of Allergy and Infectious Diseases. LITERATURE CITED 1. Black, S Yeast aldehyde dehydrogenase. Arch. Biochem. Biophys. 34: lizuka, H., M. lida, Y. Unami, and Y. Hoshino n- Decane dehydrogenation by a cell-free extract of Candida rugosa. Z. Allg. Mikrobiol. 8: Johnson, M. J Utilization of hydrocarbons by microorganisms. Chem. Ind. (London) Lowry, 0. H., J. J. Rosebrough, A. L. Farr, and R. J. Randall Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193: Miller, T. L., S. Lie, and M. J. Johnson Growth of a yeast on normal alkanes. Biotech. Bioeng. 6: Peterson, J. A., and M. J. Coon Enzymatic w-oxidation. Ill. Purification and properties of rubredoxin, a component of the w-hydroxylation system of Pseudomonas oleovorans. J. Biol. Chem. 243: Peterson, J. A., M. Kusunose, E. Kusinose, and M. J. Coon Enzymatic o-oxidation. 11. Function of rubredoxin as the electron carrier in e-hydroxylation. J. Biol. Chem. 242: Van Eys, J., and N. 0. Kaplan Yeast alcohol dehydrogenase. III. Relation of alcohol structure to activity. J. Amer. Chem. Soc. 79: Downloaded from on April 2, 2019 by guest