Vol. 137, No. 2
JOURNAL OF BACTERIOLOGY, Feb. 1979, p. 811-817 0021-9193/79/02-081 1/07$02.00/0
Oxidation of Carbon Monoxide in Cell Extracts of Pseudomonas carboxydovorans ORTWIN MEYER AND HANS G. SCHLEGEL* Institut fur Mikrobiologie der Universitat, 3400 Gottingen, Federal Republic of Germany Received for publication 26 October 1978
Extracts of aerobically, CO-autotrophically grown cells of Pseudomonas carboxydovorans were shown to catalyze the oxidation of CO to C02 in the presence of methylene blue, pyocyanine, thionine, phenazine methosulfate, or toluylene blue under strictly anaerobic conditions. Viologen dyes and NAD(P)+ were ineffective as electron acceptors. The same extracts catalyzed the oxidation of formate and of hydrogen gas; the spectrum of electron acceptors was identical for the three substrates, CO, formate, and H2. The CO- and the formate-oxidizing activities were found to be soluble enzymes, whereas hydrogenase was membrane bound exclusively. The rates of oxidation of CO, formate, and H2 were measured spectrophotometrically following the reduction of methylene blue. The rate of carbon monoxide oxidation followed simple Michaelis-Menten kinetics; the apparent Km for CO was 45 ,uM. The reaction rate was maximal at pH 7.0, and the temperature dependence followed the Arrhenius equation with an activation energy (AH0) of 35.9 kJ/mol (8.6 kcal/mol). Neither free formate nor hydrogen gas is an intermediate of the CO oxidation reaction. This conclusion is based on the differential sensitivity of the activities of formate dehydrogenase, hydrogenase, and CO dehydrogenase to heat, hypophosphite, chlorate, cyanide, azide, and fluoride as well as on the failure to trap free formate or hydrogen gas in coupled optical assays. These results support the following equation for CO oxidation in P. carboxydovorans: CO H20 C02 + 2 H+ 2eThe CO-oxidizing activity of P. carboxydovorans differed from that of Clostridium pasteurianum by not reducing viologen dyes and by a pH optimum curve that did not show an inflection point. +
+
-*
The first reports on carbon monoxide oxidation by aerobic bacteria date back to the beginning of this century (2, 11, 17, 27). The early literature was critically reviewed by Kistner (14), who succeeded in isolating a hydrogen bacterium named Hydrogenomonas carboxydovorans, capable of oxidizing CO to C02 (15). Since then further aerobic bacteria have been reported to grow on CO as the main carbon and energy source: Seliberia carboxydohydrogena (21, 22, 31), Pseudomonas carboxydoflava (19, 31), Pseudomonasgazotropha (19,31), Comamonas compransoris (19, 31), Achromobacter carboxydus (19, 31), Pseudomonas carboxydovorans (18), the unidentified strains 460 and 461 (D. H. Davis, Ph.D. thesis, University of California, Berkeley, 1967), and the N2-fixing strains S17 and A305 (20). Pseudomonas carboxydovorans, a gram-negative hydrogen bacterium, has been shown to grow aerobically on carbon monoxide as sole
source of carbon and energy (18). Growth on CO was accompanied by the production of carbon dioxide; 16% of the CO was converted into cell carbon as follows (18): 1 02 + 2.19 CO -0 1.83 C02 + 0.36 cell carbon + energy Since little is known about the CO oxidation reaction proper and since there are no reports so far on the enzymic mechanism of the conversion of CO to C02 in aerobic bacteria, it was of interest to study this reaction in cell extracts of P. carboxydovorans. The present work was undertaken to demonstrate the in vitro conversion of CO to C02 by crude or partially fractionated cell extracts of CO-grown P. carboxydovorans and to elucidate some basic properties of the CO-oxidizing system. MATERIALS AND METHODS Chemicals. All chemicals were commercially available. 811
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MEYER AND SCHLEGEL
Organism and cultivation. P. carboxydovorans OM5 (DSM 1227) was grown CO-autotrophically in mineral medium under the conditions described (18). Cells were cultivated in a 10-liter fermentor (Biostat; Braun, Melsungen), harvested in midexponential growth phase, washed twice in 50 mM potassium phosphate buffer (pH 7.0), and stored at -20°C. Preparation of cell extracts. The thawed cells were resuspended in buffer (2 g [wet weight] of cells in about 5 ml of buffer with the addition of 0.4 mg of DNase) and passed through a French pressure cell at 147.1 MPa (1,500 kpond/cm2) or disrupted by sonic oscillation (15 s/ml) with a Braun-Sonic 300 disintegrator (Quigley-Rochester, Rochester, N.Y.) at 0°C. T'he suspension was centrifuged for 2 h at 100,000 x g. The supernatant was referred to as soluble fraction. The sediment was clearly separated into two different layers; the upper layer, which contained the membranes, was suspended in buffer and designated particle fraction. Mixtures of both fractions were called crude extract. Sucrose gradient centrifugation. Discontinuous sucrose gradients were prepared in 13-ml centrifuge tubes by layering sucrose solutions (2 ml) of different concentrations (30 to 80%, wt/vol) on top of one another. Sucrose was dissolved in 50 mM phosphate buffer (pH 7.0). A 1-ml volume of crude extract was layered on the top sucrose layer. The tubes were centrifuged at 30,000 rpm for 20 h in a Christ Vacufuge (Osterode) and fractionated (0.4-ml fractions), and the enzyme activities were determined. Protein determination. The methods of Bradford (4) or Beisenherz et al. (3) were employed for protein estimation. Methylene blue reduction. The reduction of methylene blue with CO, H2, or formate was measured photometrically at 30°C by following the change in absorbance at 615 nm (ejI5 = 37.1 cm2/[.mol) using a Zeiss PM6 photometer in connection with a Goerz Servogor recorder. Solubility of carbon monoxide and hydrogen. At 30°C and atmospheric pressure, 19.4 yl of CO or 16.7 [i of H2 is dissolved in 1 ml of water (13). Preparation of gas mixtures. Gas mixtures of carbon monoxide, hydrogen, and nitrogen were prepared by filling the required quantities of gas into a syringe (100 ml). CO (99.997 and 99.0 vol%), H2 (99.9 vol%-,), N2 (99.99 vol%), 02 (99.995 vol%), and CO2 (99.995 vol%) were obtained from Messer Griesheim GmbH, Dusseldorf, Germany. RESULTS
P. carboxydovorans grew on carbon monoxide (CO-02-N2, 40:5:55) as sole source of carbon and 20 h); growth with H2-02-C02 gas energy (t,1 (90:5:5) was much faster (td = 7 h). CO-oxidizing activity was found in CO-grown cells only; hydrogenase and formate oxidase were also present under these conditions. Localization of enzyme activities. The distribution of hydrogenase, CO-, and formate-oxidizing activities between soluble fraction and the sediment after centrifugation (2 h, 100,000 x g) =
J. BACTERIOL.
is presented in Table 1. A total of 61% of the COoxidizing activity and 44% of the formate-oxidizing activity were found in the soluble fraction, but 96% of hydrogenase activity was recovered in the sediment. The percentages of CO-oxidizing activity in the soluble and the particle fraction turned out to be nearly equal. Because this observation suggested the existence of both a soluble and a particulate CO-oxidizing activity, the distribution of enzyme activities was examined by sucrose gradient centrifugation. The localization of hydrogenase, CO-oxidizing activity, formate-oxidizing activity, and NADH oxidase in CO-grown cells of P. carboxydovorans was studied by fractionation of crude extracts prepared by sonic disruption (Fig. 1A) or with a French pressure cell (Fig. 1B) in discontinuous sucrose gradients. The almost exact coincidence of activity profiles of hydrogenase and NADH oxidase (Fig. 1A and B) unambiguously disproved the existence of a soluble hydrogenase. Both the CO- and the formate-oxidizing activities were localized in the 30% sucrose layer, which contained only a few small particles. The particles of sonic and French press extracts behaved differently during sucrose gradient centrifugation, as indicated in the distribution pattern of hydrogenase activity. When sonic extracts were used, the hydrogenase activity profiles exhibited pronounced peaks in the 60 and 80% sucrose layers (Fig. 1A). In contrast, hydrogenase of French press extracts was bound to smaller and apparently much more homogeneous particles localized in the 50% sucrose layer TABLE 1. Distribution of specific CO-, formate-, and H2-oxidizing activities after centrifugation of a crude sonic extract of P. carboxydovorans" Oxidizing activity (nmol min-' mg-') in: Substance oxidized Supernatant 99 91 41
Pellet
CO 64 Formate 116 876 H2 0.91 ml of 50 mM buffer KH2PO4-KOH (pH 'Assay: 7.0); 0.05 ml of 2 mM glucose in buffer; 0.02 ml of 2.5 mM methylene blue in buffer; 0.01 ml of mix (1 U of glucose oxidase + 1 U of catalase in buffer); serumstoppered cuvettes were flushed with the following gases for at least 5 min: CO (CO-oxidizing activity), H2 (H2-oxidizing activity), N2 in the presence of 0.1 ml of sodium formate (in buffer) in the presence of 0.81 ml of 50 mM KH2PO4-KOH buffer (pH 7.0) (formateoxidizing activity); start with 10 1l of extract (supernatant after 2 h, 100,000 x g) of CO-grown cells of P. carboxydovorans and 1 to 8 mg of soluble protein or 4 to 14 mg of particle protein per ml; reduction of methylene blue was measured at 615 nm, 30°C.
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gradient centrifugation, both the CO- and the formate-oxidizing activities must be due to soluble enzymes. Electron acceptors. Crude extracts of COgrown P. carboxydovorans readily catalyzed the oxidation of carbon monoxide with basic dyes of the thiazine group such as methylene blue or thionine (Lauth's violet) and with dichlorophenolindophenol, pyocyanine, and toluylene blue (Table 2). Dyes with a potential lower than -100 mV [NAD(P)+, the viologens] were not reduced. Polarographic measurements (Yellow Springs electrode) showed that the reduction of 02 with CO was catalyzed by the particle fraction exclusively, but not by the soluble fraction (2 h, 100,000 x g). For hydrogenase and the formateTABLE 2. Reduction of electron acceptors with carbon monoxide catalyzed by the 100,000 x g supernatant of P. carboxydovorans" Electron acceptor
Carbon monoxide Methyl viologen Benzyl viologen Neutral red NADP+, NAD+ FAD+, FMN+c Riboflavin Indigo (tri-, tetra-)sulfonates Vitamin K, Vitamin K1 Pyocyanine Methylene blue Thionine (Lauth's violet) Phenazine methosulfate Toluylene blue 0
5
10
15
20 25 Fraction number
30
FIG. 1. Distribution patterns of the CO-, formate-, H2-, and NADH-oxidizing enzyme activities of crude extracts of P. carboxydovorans after sucrose density centrifugation. (A) Sonic extract; (B) French pressure cell extract. After cell disintegration the crude extract was layered on top of a 30 to 80% sucrose density gradient and centrifuged for 20 h at 30,000 rpm; NADH oxidase activity was measured according to Aggag and Schlegel (1). All other enzyme assays are described in the legend to Table 1. Symbols: CO (0), formate (0), H2 (U), and NADH (O).
with a shoulder in the 60% sucrose layer (Fig. 1B). From these results the conclusion is drawn that the P. carboxydovorans hydrogenase is a membrane-bound enzyme exclusively. In contrast, according to results obtained by sucrose
(mV)
-540 -440 -359 -320 -320 -219 -298 -111 (-70, -30) -50 -44 -34 +11 +70
CO oxidation rates ( of rate with methylene blue)5
0 0 0 0 0 0 0 3 7 83 100 30
+80
70
+110
88
+217 20 Dichlorophenolindophenol +245 0 Cytochrome c +429 0 Ferricyanide +816 0 Oxygen "Assay: 1.1 rml of 50 mM KH2PO4-KOH buffer (pH 7.0); 0.1 ml of mix (1 U of glucose oxidase + 1 U of catalase); 0.5 ml of 2 mM glucose; 0.3 ml of soluble fraction (100,000 x g, 2 h, 3.9 mg of soluble protein per ml) of P. carboxydovorans in the main compartment; 0.2 ml of KOH (20%) in the central cup; 0.3 ml of 7.5 mM electron acceptor in buffer (water-insoluble vitamins in ethanol); atmosphere 100% CO; start with electron acceptor.
'CO uptake rates were manometrically measured at 30°C; the rate with methylene blue was 124 nmol
min-' mg-'. 'FADW, Oxidized flavine adenine dinucleotide; FMN+, oxidized riboflavine 5'-phosphate.
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oxidizing activity an electron acceptor spectrum was found identical to that of the CO-oxidizing activity. Therefore, the reduction of methylene blue followed spectrophotometrically at 615 nm was used as a standard assay in the following studies. pH dependence. Maximum methylene blue reduction rates with CO were found in the range of pH 6.5 to 7.2 (Fig. 2). The pH curve shows no inflection point and is quite different from that reported for CO oxidation catalyzed by extracts of Clostridium pasteurianum (24) or Methanobacterium thermoautotrophicum (5). Temperature dependence. Maximum rates of methylene blue reduction with CO were found at 65°C (Fig. 3); this value is relatively high for an enzyme functioning in a mesophilic bacterium and may be considered a hint for the involvement of a transition metal. The Arrhenius plot revealed an activation energy (AH0) of 35.9 kJ/mol (8.6 kcal/mol). Stoichiometry of methylene blue reduction with CO. Due to the low apparent Km of 45 [zM CO, the clearly exergonic reduction of methylene blue (E"' = +11 mV) with carbon monoxide (Et') = -540 mV) proceeded to complete oxidation of the substrate. Thus the stoichiometry of the reaction could be determined by adding limiting amounts of CO and by measuring the amount of methylene blue reduced.
Methylene blue was reduced by carbon monoxide at a 1:1 ratio. This indicates that CO oxidation provides two electrons and carbon dioxide is the reaction product. Inactivation by heat. The CO-oxidizing activity was relatively heat stable (Fig. 4). Portions of 5 ml of the soluble fraction were incubated in a water bath (80°C); at intervals, samples were taken and cooled in ice. After 2 min of exposure to 80°C, the formate-oxidizing activity disappeared, whereas 40% of the CO-oxidizing activity remained. This indicates that the formate-oxi-
>1
z
0
40
30
20
10
50
60
70
80
(OC) FIG. 3. Temperature dependence of the CO-oxidizing activity. Methylene blue reduction as described in the footnote to Table 1. Temperature
0o3 -100 -0
E u') 0 .2 (D
50I
-0
Z O.1i
I 10
0 i
0 0
3
4
5
6
7
8
pH
FIG. 2. pH dependence of the rate of methylene blue reduction with CO. Assay: buffers as indicated; methylene blue reduction as described in the footnote to Table 1. Symbols: (-) KH2PO4-NaOH, 50 mM; (O) Tris-maleate, 50 mM; (A) citrate-Na2HPO4, 100 mM.
I
t
1
2
3
4 5 6 Time (min)
7
FIG. 4. Differentiation of CO-oxidizing activity and formate-oxidizing activity of the soluble fraction by exposure to 80°C. The soluble fraction of the crude sonic extract (2 h at 100,000 x g) was incubated at 80°C, samples were taken at intervals, and the rate of methylene blue reduction was measured with CO (-) or formate (0) as described in the footnote to Table 1.
CO OXIDATION IN P. CARBOXYDOVORANS
VOL. 137, 1979
dizing activity is not involved in the CO oxidation reaction. Effects of inhibitors. Hypophosphite (NaH2PO2), chlorate (NaCl03), cyanide (NaCN), and azide (NaN3) are known as metalcomplexing agents and used as inhibitors of formate dehydrogenases. At high inhibitor concentrations, the formate-oxidizing activity as well as the CO-oxidizing activity was very low (Table 3). However, the formate-oxidizing activity was subject to a much stronger inhibition than the CO-oxidizing activity. In the presence of fluoride ([NaF] > 300 mM) the CO-oxidizing activity was scarcely affected, whereas the formate-oxidizing activity was almost completely suppressed (Fig. 5). Kinetics of methylene blue reduction with CO. The reduction of methylene blue by the soluble fraction of the sonic extract with CO proceeded linearly with time (0 to 3 min), and the rate was proportional to the amount of protein added in the range measurable photometrically (E615 < 2.5). The dependence of the methylene blue reduction rate on the substrate concentration (CO) followed simple Michaelis-Menten kinetics: plots of 1/v versus 1/[S] were linear. The Km turned out to be 45 ,aM CO. Exclusion of hydrogen gas and free formate as intermediates of the CO oxidation reaction. Cells of P. carboxydovorans growing CO-autotrophically contain hydrogenase and formate oxidase (18). Thus, the oxidation of carbon monoxide to carbon dioxide and hydrogen gas (CO + H20 -' CO2 + H2; AGO' = -20.1 kJ [-4.8 kcal]) (25) and the hydratation to free formate (CO + H20-- HCOO- + H+; AGO = -16.3 kJ [-3.9 kcal]) (25), which is afterwards oxidized to C02, have to be considered as alternate possibilities of CO oxidation; both reactions are thermodynamically possible. If hydrogen gas is formed during the oxidation of CO it should be possible to couple this reaction to the reduction of NAD+ catalyzed by the
815
soluble NAD-specific hydrogenase of Alcaligenes eutrophus (Fig. 6). After addition of this hydrogenase to the reaction mixture, however, NAD+ reduction was not detectable (Fig. 6, B). Controls performed by the addition of small amounts of hydrogen-saturated buffer (Fig. 6, C and D) or of methylene blue (Fig. 6, E) showed that the hydrogenase remained still active in the reaction mixture and that the CO-oxidizing activity was still capable of oxidizing CO. A similar assay was used to couple the formation of formate with the reduction of methyl viologen catalyzed by extracts of Clostridium cylindrosporum (Fig. 7). However, the dye was 100
.0 p
p
-
50
10 0
100
[NaF](mM)
500
FIG. 5. Differentiation of the CO-oxidizing activity and formate-oxidizing activity of the soluble fraction by inhibition by fluoride. The rate of methylene blue reduction was measured as described in the footnote to Table 1.
TABLE 3. Effect of metal-complexing agents on the CO- and the formate-oxidizing activities" Inhibition (%)
Inhibitor (100 mM)
CO oxidation
Formate oxidation
11 Hypophosphite (NaH2PO2) 89.4 Chlorate (NaCI03) 0 64.1 64.1 100 Cyanide (NaCN) Azide (NaN3) 92.2 100 Fluoride (NaF) 2.3 81.7 " Assay: the CO-oxidizing activity and the formateoxidizing activity were assayed according to the legend to Table 1, but in the presence of 100 mM inhibitor.
20
Ti,me (min
FIG. 6. Experiment to detect hydrogen gas as a possible intermediate of CO oxidation. Assay: 0.91 ml of 50 mM KH2PO4-KOH buffer (pH 7.0); 10 ,ul of mix (1 U of glucose oxidase + 1 U of catalase); 50 ILI of 2 mMglucose + 30 LM NAD+; serum-stoppered cuvettes were flushed with CO. Arrows: A, addition of 20 u1 of crude bacterial extract of P. carboxydouorans; B, 20 fLl of NAD+-specific hydrogenase; C, 100 /1l of H2saturated buffer; D, 50 u1 of H2-saturated buffer (so far measured at 365 nm); E, 10I l of 2.5 mM methylene blue (measured at 615 nm).
816
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MEYER AND SCHLEGEL
not reduced; addition of formate d( emonstrated that the extract was still capable c)f catalyzing the reduction of methyl viologen w,ith formate. The results clearly indicate that ne ither hydrogen gas nor free formate is an inte rmediate of the carbon monoxide oxidation reac tion. Other properties of the CO ox idation reaction. The CO-oxidizing activity w. as only present in CO-grown cells of P. carbo)xydoL'orans and was very sensitive to alterat.ions of the growth conditions; it was not lost wihen extracts were aerobically dialyzed for 24 h ag ainst 50 mM phosphate buffer (pH 7.0), whereas tthe formateoxidizing activity was completely destroyed. Ethylenediaminetetraacetic acid (5'0 mM) was without effect on the CO-oxidizing aictivity. The reduction of methylene blue with CO started immediately after addition of solulble fraction; however, the reaction mediated by the particle fraction sometimes started after a laLg of up to 15 min.
The CO-oxidizing activity was as,sayed under strictly anaerobic conditions, indicat ing that molecular oxygen is not involved and the second oxygen atom in the CO2 produceci is derived from water; thus, CO oxidation is riot due to a monooxygenase.
DISCUSSION During aerobic growth of P. carbo rydoLtorans with carbon monoxide as the sol e source of carbon and energy, CO is oxidized to CO2 (18). CO oxidation was also shown to occutr in extracts of CO-grown cells. Different electroin acceptors such as pyocyanine, methylene bluie, thionine,
A
B
c
c |
2_____
Time (min)2
FIc. 7. Experiment to detect formate as apossible intermiediate of CO oxidation. Assay: c(omponents as in the legend to Fig. 6, but in the pres;ence of 5 [aM W methyl ciologen instead of NAD+. At-rr 9)VS: A, addition of 10 til of crude bacterial extract cif P. carboxydocorans; B, 10 pl of craude extract off C. cylindro-
sporum containing about 0.15 U of foaimq7ntprP h>vr1r)t ltlCt (XUctlY(l genase; C. .30 yl of 5 mM formate.
-
or phenazine methosulfate were readily reduced by carbon monoxide, whereas oxidized flavine adenine dinucleotide, oxidized riboflavine 5'phosphate, cytochrome c, and the pyridine nucleotides turned out to be ineffective. This supports the assumption that the electrons are delivered at the level of succinate dehydrogenase and that a quinone is possibly the physiological electron acceptor. Centrifugation of extracts in sucrose gradients characterized hydrogenase as a membranebound enzyme, whereas the CO- and the formate-oxidizing activities were soluble or only slightly attached to the membrane; their distribution at a 1:1 ratio between the soluble and particle fraction after centrifugation may be due to partial adsorption of the soluble enzyme to the particles. Dehydrogenation of CO is probably due to a separate enzyme and does not involve formate dehydrogenase or hydrogenase activity. This conclusion is based on the differential sensitivity of the enzyme activities to heat and metal-complexing inhibitors such as NaF, as well as on the failure to trap free formate or hydrogen gas in coupled optical assays. The basic reaction actually used in P. carboxydou,orans is CO + H2O -* CO2 + 2 Ht + 2e The reaction serves two different functions: (i) to provide electrons for energy generation and reductive syntheses and (ii) to produce CO2 for the reductive pentose phosphate cycle. The COoxidizing system thus serves catabolic functions; it is inducible and is only formed during growth on CO. Hydrogenase is inducible and is only present in cells grown with hydrogen (H2,-02CO2) or with pyruvate, whereas the formateoxidizing activity is constitutive and is present in cells grown with CO (CO-O0), H2 (H2-02GCO,), acetate, or pyruvate. In contrast, the physiological function of COoxidizing activity in methanogenic bacteria (5, 7, 8, 16, 23), sulfate-reducing bacteria (28, 29, 30), photosynthetic bacteria (26), and clostridia (9, 1)0, 24) is far from clear. In these bacteria the CO-oxidizing activity is constitutive. It may be due to the nonspecific activities of enzymes fulfilling quite different physiological functions, e.g. in the case of methane monooxygenase (2, 6). Furthermore, the CO-oxidizing activities of the anaerobic bacteria have been shown to reduce methylene blue only slightly, but viologen dyes with high rates, whereas extracts of P. carboxvdo'orans are not capable of oxidizing CO with vriologen dyes as electron acceptors. ACKNOWLEDGMENTS (KOLDINT We thiank K. Schneider for providing SOIlIble hydrogenase
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CO OXIDATION IN P. CARBOXYDOVORANS
of A. eutrophus and R. Wagner for providing extracts of C. cylindrosporum. LITERATURE CITED 1. Aggag, M., and H. G. Schlegel. 1973. Studies on a grampositive hydrogen bacterium, Nocardia opaca lb. III. Purification, stability and some properties of the soluble hydrogen dehydrogenase. Arch. Microbiol. 100:25-39. 2. Beijerinck, M., and A. van Delden. 1903. Uber eine farblose Bakterie, deren Kohlenstoffnahrung aus der atmospharischen Luft herruhrt. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. 2 10:33-47. 3. Beisenherz, G., H. J. Bolze, T. Bucher, R. Czok, K. H. Garbade, E. Meyer-Arendt, and G. Pfleiderer. 1953. Diphosphofructose-Aldolase, PhosphoglyceraldehydDehydrogenase, Milchsaure-Dehydrogenase, Glycerophosphat-Dehydrogenase und Pyruvat-Kinase aus Kaninchenmuskulatur in einem Arbeitsgang. Z. Naturforsch. 8b:555-577. 4. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254. 5. Daniels, L., G. Fuchs, R. K. Thauer, and J. G. Zeikus. 1977. Carbon monoxide oxidation by methanogenic bacteria. J. Bacteriol. 132:118-126. 6. Ferenci, T. 1974. Carbon monoxide stimulated respiration in methane-utilizing bacteria. FEBS Lett. 41:94-98. 7. Fischer, F., R. Lieske, and K. Winzer. 1931. Biologische Gasreaktionen. I. Mitt. Die Umsetzungen des Kohlenoxyds. Biochem. Z. 236:247-267. 8. Fischer, F., R. Lieske, and K. Winzer. 1932. Uber die Bildung von Essigsaure bei der biologischen Umsetzung von Kohlenoxyd und Kohlensaure mit Wasserstoff zu Methan. Biochem. Z. 245:2-12. 9. Fuchs, G., G. Andress, and R. K. Thauer. 1975. COoxidation by anaerobic bacteria: indications for the involvement of a vitamin B12 compound, p. 231-236. In H. G. Schlegel, G. Gottschalk, and N. Pfennig (ed.), Microbial production and utilization of gases. E. Goltze KG, Gottingen. 10. Fuchs, G., U. Schnitker, and R. K. Thauer. 1974. Carbon monoxide oxidation by growing cultures of Clostridium pasteurianum. Eur. J. Biochem. 49:111-115. 11. Hasemann, W. 1927. Zersetzung von Leuchtgas und Kohlenoxyd durch Bakterien. Biochem. Z. 184:147-171. 12. Hubley, J. H., J. R. Milton, and J. F. Wilkinson. 1974. The oxidation of carbon monoxide by methane-oxidizing bacteria. Arch. Microbiol. 95:365-368. 13. Lax, E. 1967. D'Ans-Lax: Taschenbuch fur Chemiker und Physiker, vol. 1, p. 1-1205. Springer-Verlag, Berlin. 14. Kistner, A. 1953. On a bacterium oxidizing carbon monoxide. Proc. K. Ned. Akad. Wet. Ser. C 56:443-450.
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15. Kistner, A. 1954. Conditions determining the oxidation of carbon monoxide by Hydrogenomonas carboxydovorans. Proc. K. Ned. Akad. Wet. Ser. C 57:186-195. 16. Kluyver, A. J., and C. G. Schnellen. 1947. On the fermentation of carbon monoxide by pure cultures of methane bacteria. Arch. Biochem. 14:57-70. 17. Lantzsch, K. 1922. Actinomyces oligocarbophilus (Bacillus oligocarbophilus Beij.), sein Formwechsel und seine Physiologie. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. 2 57:309-319. 18. Meyer, O., and H. G. Schlegel. 1978. Reisolation of the carbon monoxide utilizing hydrogen bacterium Pseudomonas carboxydovorans (Kistner) comb. nov. Arch. Microbiol. 118:35-43. 19. Nozhevnikova, A. N., and G. A. Zavarzin. 1974. On the taxonomy of CO-oxidizing gram-negative bacteria. Izv. Akad. Nauk. SSSR Ser. Biol. 3:436-440. 20. Ooyama, J., and T. Shinohara. 1971. Simultaneous fixation of CO and N, in the presence of H2 and O0 by a bacterium. Rep. Ferment. Res. Inst. 40:1-5. 21. Sanjieva, E. U., and G. A. Zavarzin. 1971. Oxidation of carbon monoxide by Seliberia carboxydohydrogena. Dokl. Akad. Nauk. SSSR 196:956-958. 22. Savelieva, N. D., and A. N. Nozhevnikova. 1972. Autotrophic growth of Seliberia carboxydohydrogena during oxidation of hydrogen and carbon monoxide. Mikrobiologiya 41:813-817. 23. Stephenson, M., and L. H. Stickland. 1933. The bacterial formation of methane by the reduction of one carbon compounds by molecular hydrogen. Biochem. J. 27:1517-1527. 24. Thauer, R. K., G. Fuchs, B. Kaufer, and U. Schnitker. 1974. Carbon monoxide oxidation in cell-free extracts of Clostridium pasteurianum. Eur. J. Biochem. 45:343349. 25. Thauer, R. K., K. Jungermann, and K. Decker. 1977. Energy conservation in chemotrophic bacteria. Bacteriol. Rev. 41:100-180. 26. Uffen, R. L. 1976. Anaerobic growth of a Rhodopseudomonas species in the dark with carbon monoxide as sole carbon and energy substrate. Proc. Natl. Acad. Sci. U.S.A. 73:3298-3302. 27. Wehmer, C. 1926. Biochemische Zersetzung des Kohlenoxyds. Ber. DtLsch. Chem. Ges. 59:887-890. 28. Yagi, T. 1958. Enzymic oxidation of carbon monoxide. Biochim. Biophys. Acta 30:194-195. 29. Yagi, T. 1959. Enzymic oxidation of carbon monoxide. II. J. Biochem. (Tokyo) 46:949-955. 30. Yagi, T., and N. Tamiya. 1962. Enzymic oxidation of carbon monoxide. III. Reversibility. Biochim. Biophys. Acta 65:508-509. 31. Zavarzin, G. A., and A. N. Nozhevnikova. 1977. Aerobic carboxydobacteria. Microb. Ecol. 3:305-326.
JOURNAL OF BACTERIOLOGY, Feb. 1979, p. 811-817 0021-9193/79/02-081 1/07$02.00/0
Oxidation of Carbon Monoxide in Cell Extracts of Pseudomonas carboxydovorans ORTWIN MEYER AND HANS G. SCHLEGEL* Institut fur Mikrobiologie der Universitat, 3400 Gottingen, Federal Republic of Germany Received for publication 26 October 1978
Extracts of aerobically, CO-autotrophically grown cells of Pseudomonas carboxydovorans were shown to catalyze the oxidation of CO to C02 in the presence of methylene blue, pyocyanine, thionine, phenazine methosulfate, or toluylene blue under strictly anaerobic conditions. Viologen dyes and NAD(P)+ were ineffective as electron acceptors. The same extracts catalyzed the oxidation of formate and of hydrogen gas; the spectrum of electron acceptors was identical for the three substrates, CO, formate, and H2. The CO- and the formate-oxidizing activities were found to be soluble enzymes, whereas hydrogenase was membrane bound exclusively. The rates of oxidation of CO, formate, and H2 were measured spectrophotometrically following the reduction of methylene blue. The rate of carbon monoxide oxidation followed simple Michaelis-Menten kinetics; the apparent Km for CO was 45 ,uM. The reaction rate was maximal at pH 7.0, and the temperature dependence followed the Arrhenius equation with an activation energy (AH0) of 35.9 kJ/mol (8.6 kcal/mol). Neither free formate nor hydrogen gas is an intermediate of the CO oxidation reaction. This conclusion is based on the differential sensitivity of the activities of formate dehydrogenase, hydrogenase, and CO dehydrogenase to heat, hypophosphite, chlorate, cyanide, azide, and fluoride as well as on the failure to trap free formate or hydrogen gas in coupled optical assays. These results support the following equation for CO oxidation in P. carboxydovorans: CO H20 C02 + 2 H+ 2eThe CO-oxidizing activity of P. carboxydovorans differed from that of Clostridium pasteurianum by not reducing viologen dyes and by a pH optimum curve that did not show an inflection point. +
+
-*
The first reports on carbon monoxide oxidation by aerobic bacteria date back to the beginning of this century (2, 11, 17, 27). The early literature was critically reviewed by Kistner (14), who succeeded in isolating a hydrogen bacterium named Hydrogenomonas carboxydovorans, capable of oxidizing CO to C02 (15). Since then further aerobic bacteria have been reported to grow on CO as the main carbon and energy source: Seliberia carboxydohydrogena (21, 22, 31), Pseudomonas carboxydoflava (19, 31), Pseudomonasgazotropha (19,31), Comamonas compransoris (19, 31), Achromobacter carboxydus (19, 31), Pseudomonas carboxydovorans (18), the unidentified strains 460 and 461 (D. H. Davis, Ph.D. thesis, University of California, Berkeley, 1967), and the N2-fixing strains S17 and A305 (20). Pseudomonas carboxydovorans, a gram-negative hydrogen bacterium, has been shown to grow aerobically on carbon monoxide as sole
source of carbon and energy (18). Growth on CO was accompanied by the production of carbon dioxide; 16% of the CO was converted into cell carbon as follows (18): 1 02 + 2.19 CO -0 1.83 C02 + 0.36 cell carbon + energy Since little is known about the CO oxidation reaction proper and since there are no reports so far on the enzymic mechanism of the conversion of CO to C02 in aerobic bacteria, it was of interest to study this reaction in cell extracts of P. carboxydovorans. The present work was undertaken to demonstrate the in vitro conversion of CO to C02 by crude or partially fractionated cell extracts of CO-grown P. carboxydovorans and to elucidate some basic properties of the CO-oxidizing system. MATERIALS AND METHODS Chemicals. All chemicals were commercially available. 811
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MEYER AND SCHLEGEL
Organism and cultivation. P. carboxydovorans OM5 (DSM 1227) was grown CO-autotrophically in mineral medium under the conditions described (18). Cells were cultivated in a 10-liter fermentor (Biostat; Braun, Melsungen), harvested in midexponential growth phase, washed twice in 50 mM potassium phosphate buffer (pH 7.0), and stored at -20°C. Preparation of cell extracts. The thawed cells were resuspended in buffer (2 g [wet weight] of cells in about 5 ml of buffer with the addition of 0.4 mg of DNase) and passed through a French pressure cell at 147.1 MPa (1,500 kpond/cm2) or disrupted by sonic oscillation (15 s/ml) with a Braun-Sonic 300 disintegrator (Quigley-Rochester, Rochester, N.Y.) at 0°C. T'he suspension was centrifuged for 2 h at 100,000 x g. The supernatant was referred to as soluble fraction. The sediment was clearly separated into two different layers; the upper layer, which contained the membranes, was suspended in buffer and designated particle fraction. Mixtures of both fractions were called crude extract. Sucrose gradient centrifugation. Discontinuous sucrose gradients were prepared in 13-ml centrifuge tubes by layering sucrose solutions (2 ml) of different concentrations (30 to 80%, wt/vol) on top of one another. Sucrose was dissolved in 50 mM phosphate buffer (pH 7.0). A 1-ml volume of crude extract was layered on the top sucrose layer. The tubes were centrifuged at 30,000 rpm for 20 h in a Christ Vacufuge (Osterode) and fractionated (0.4-ml fractions), and the enzyme activities were determined. Protein determination. The methods of Bradford (4) or Beisenherz et al. (3) were employed for protein estimation. Methylene blue reduction. The reduction of methylene blue with CO, H2, or formate was measured photometrically at 30°C by following the change in absorbance at 615 nm (ejI5 = 37.1 cm2/[.mol) using a Zeiss PM6 photometer in connection with a Goerz Servogor recorder. Solubility of carbon monoxide and hydrogen. At 30°C and atmospheric pressure, 19.4 yl of CO or 16.7 [i of H2 is dissolved in 1 ml of water (13). Preparation of gas mixtures. Gas mixtures of carbon monoxide, hydrogen, and nitrogen were prepared by filling the required quantities of gas into a syringe (100 ml). CO (99.997 and 99.0 vol%), H2 (99.9 vol%-,), N2 (99.99 vol%), 02 (99.995 vol%), and CO2 (99.995 vol%) were obtained from Messer Griesheim GmbH, Dusseldorf, Germany. RESULTS
P. carboxydovorans grew on carbon monoxide (CO-02-N2, 40:5:55) as sole source of carbon and 20 h); growth with H2-02-C02 gas energy (t,1 (90:5:5) was much faster (td = 7 h). CO-oxidizing activity was found in CO-grown cells only; hydrogenase and formate oxidase were also present under these conditions. Localization of enzyme activities. The distribution of hydrogenase, CO-, and formate-oxidizing activities between soluble fraction and the sediment after centrifugation (2 h, 100,000 x g) =
J. BACTERIOL.
is presented in Table 1. A total of 61% of the COoxidizing activity and 44% of the formate-oxidizing activity were found in the soluble fraction, but 96% of hydrogenase activity was recovered in the sediment. The percentages of CO-oxidizing activity in the soluble and the particle fraction turned out to be nearly equal. Because this observation suggested the existence of both a soluble and a particulate CO-oxidizing activity, the distribution of enzyme activities was examined by sucrose gradient centrifugation. The localization of hydrogenase, CO-oxidizing activity, formate-oxidizing activity, and NADH oxidase in CO-grown cells of P. carboxydovorans was studied by fractionation of crude extracts prepared by sonic disruption (Fig. 1A) or with a French pressure cell (Fig. 1B) in discontinuous sucrose gradients. The almost exact coincidence of activity profiles of hydrogenase and NADH oxidase (Fig. 1A and B) unambiguously disproved the existence of a soluble hydrogenase. Both the CO- and the formate-oxidizing activities were localized in the 30% sucrose layer, which contained only a few small particles. The particles of sonic and French press extracts behaved differently during sucrose gradient centrifugation, as indicated in the distribution pattern of hydrogenase activity. When sonic extracts were used, the hydrogenase activity profiles exhibited pronounced peaks in the 60 and 80% sucrose layers (Fig. 1A). In contrast, hydrogenase of French press extracts was bound to smaller and apparently much more homogeneous particles localized in the 50% sucrose layer TABLE 1. Distribution of specific CO-, formate-, and H2-oxidizing activities after centrifugation of a crude sonic extract of P. carboxydovorans" Oxidizing activity (nmol min-' mg-') in: Substance oxidized Supernatant 99 91 41
Pellet
CO 64 Formate 116 876 H2 0.91 ml of 50 mM buffer KH2PO4-KOH (pH 'Assay: 7.0); 0.05 ml of 2 mM glucose in buffer; 0.02 ml of 2.5 mM methylene blue in buffer; 0.01 ml of mix (1 U of glucose oxidase + 1 U of catalase in buffer); serumstoppered cuvettes were flushed with the following gases for at least 5 min: CO (CO-oxidizing activity), H2 (H2-oxidizing activity), N2 in the presence of 0.1 ml of sodium formate (in buffer) in the presence of 0.81 ml of 50 mM KH2PO4-KOH buffer (pH 7.0) (formateoxidizing activity); start with 10 1l of extract (supernatant after 2 h, 100,000 x g) of CO-grown cells of P. carboxydovorans and 1 to 8 mg of soluble protein or 4 to 14 mg of particle protein per ml; reduction of methylene blue was measured at 615 nm, 30°C.
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gradient centrifugation, both the CO- and the formate-oxidizing activities must be due to soluble enzymes. Electron acceptors. Crude extracts of COgrown P. carboxydovorans readily catalyzed the oxidation of carbon monoxide with basic dyes of the thiazine group such as methylene blue or thionine (Lauth's violet) and with dichlorophenolindophenol, pyocyanine, and toluylene blue (Table 2). Dyes with a potential lower than -100 mV [NAD(P)+, the viologens] were not reduced. Polarographic measurements (Yellow Springs electrode) showed that the reduction of 02 with CO was catalyzed by the particle fraction exclusively, but not by the soluble fraction (2 h, 100,000 x g). For hydrogenase and the formateTABLE 2. Reduction of electron acceptors with carbon monoxide catalyzed by the 100,000 x g supernatant of P. carboxydovorans" Electron acceptor
Carbon monoxide Methyl viologen Benzyl viologen Neutral red NADP+, NAD+ FAD+, FMN+c Riboflavin Indigo (tri-, tetra-)sulfonates Vitamin K, Vitamin K1 Pyocyanine Methylene blue Thionine (Lauth's violet) Phenazine methosulfate Toluylene blue 0
5
10
15
20 25 Fraction number
30
FIG. 1. Distribution patterns of the CO-, formate-, H2-, and NADH-oxidizing enzyme activities of crude extracts of P. carboxydovorans after sucrose density centrifugation. (A) Sonic extract; (B) French pressure cell extract. After cell disintegration the crude extract was layered on top of a 30 to 80% sucrose density gradient and centrifuged for 20 h at 30,000 rpm; NADH oxidase activity was measured according to Aggag and Schlegel (1). All other enzyme assays are described in the legend to Table 1. Symbols: CO (0), formate (0), H2 (U), and NADH (O).
with a shoulder in the 60% sucrose layer (Fig. 1B). From these results the conclusion is drawn that the P. carboxydovorans hydrogenase is a membrane-bound enzyme exclusively. In contrast, according to results obtained by sucrose
(mV)
-540 -440 -359 -320 -320 -219 -298 -111 (-70, -30) -50 -44 -34 +11 +70
CO oxidation rates ( of rate with methylene blue)5
0 0 0 0 0 0 0 3 7 83 100 30
+80
70
+110
88
+217 20 Dichlorophenolindophenol +245 0 Cytochrome c +429 0 Ferricyanide +816 0 Oxygen "Assay: 1.1 rml of 50 mM KH2PO4-KOH buffer (pH 7.0); 0.1 ml of mix (1 U of glucose oxidase + 1 U of catalase); 0.5 ml of 2 mM glucose; 0.3 ml of soluble fraction (100,000 x g, 2 h, 3.9 mg of soluble protein per ml) of P. carboxydovorans in the main compartment; 0.2 ml of KOH (20%) in the central cup; 0.3 ml of 7.5 mM electron acceptor in buffer (water-insoluble vitamins in ethanol); atmosphere 100% CO; start with electron acceptor.
'CO uptake rates were manometrically measured at 30°C; the rate with methylene blue was 124 nmol
min-' mg-'. 'FADW, Oxidized flavine adenine dinucleotide; FMN+, oxidized riboflavine 5'-phosphate.
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oxidizing activity an electron acceptor spectrum was found identical to that of the CO-oxidizing activity. Therefore, the reduction of methylene blue followed spectrophotometrically at 615 nm was used as a standard assay in the following studies. pH dependence. Maximum methylene blue reduction rates with CO were found in the range of pH 6.5 to 7.2 (Fig. 2). The pH curve shows no inflection point and is quite different from that reported for CO oxidation catalyzed by extracts of Clostridium pasteurianum (24) or Methanobacterium thermoautotrophicum (5). Temperature dependence. Maximum rates of methylene blue reduction with CO were found at 65°C (Fig. 3); this value is relatively high for an enzyme functioning in a mesophilic bacterium and may be considered a hint for the involvement of a transition metal. The Arrhenius plot revealed an activation energy (AH0) of 35.9 kJ/mol (8.6 kcal/mol). Stoichiometry of methylene blue reduction with CO. Due to the low apparent Km of 45 [zM CO, the clearly exergonic reduction of methylene blue (E"' = +11 mV) with carbon monoxide (Et') = -540 mV) proceeded to complete oxidation of the substrate. Thus the stoichiometry of the reaction could be determined by adding limiting amounts of CO and by measuring the amount of methylene blue reduced.
Methylene blue was reduced by carbon monoxide at a 1:1 ratio. This indicates that CO oxidation provides two electrons and carbon dioxide is the reaction product. Inactivation by heat. The CO-oxidizing activity was relatively heat stable (Fig. 4). Portions of 5 ml of the soluble fraction were incubated in a water bath (80°C); at intervals, samples were taken and cooled in ice. After 2 min of exposure to 80°C, the formate-oxidizing activity disappeared, whereas 40% of the CO-oxidizing activity remained. This indicates that the formate-oxi-
>1
z
0
40
30
20
10
50
60
70
80
(OC) FIG. 3. Temperature dependence of the CO-oxidizing activity. Methylene blue reduction as described in the footnote to Table 1. Temperature
0o3 -100 -0
E u') 0 .2 (D
50I
-0
Z O.1i
I 10
0 i
0 0
3
4
5
6
7
8
pH
FIG. 2. pH dependence of the rate of methylene blue reduction with CO. Assay: buffers as indicated; methylene blue reduction as described in the footnote to Table 1. Symbols: (-) KH2PO4-NaOH, 50 mM; (O) Tris-maleate, 50 mM; (A) citrate-Na2HPO4, 100 mM.
I
t
1
2
3
4 5 6 Time (min)
7
FIG. 4. Differentiation of CO-oxidizing activity and formate-oxidizing activity of the soluble fraction by exposure to 80°C. The soluble fraction of the crude sonic extract (2 h at 100,000 x g) was incubated at 80°C, samples were taken at intervals, and the rate of methylene blue reduction was measured with CO (-) or formate (0) as described in the footnote to Table 1.
CO OXIDATION IN P. CARBOXYDOVORANS
VOL. 137, 1979
dizing activity is not involved in the CO oxidation reaction. Effects of inhibitors. Hypophosphite (NaH2PO2), chlorate (NaCl03), cyanide (NaCN), and azide (NaN3) are known as metalcomplexing agents and used as inhibitors of formate dehydrogenases. At high inhibitor concentrations, the formate-oxidizing activity as well as the CO-oxidizing activity was very low (Table 3). However, the formate-oxidizing activity was subject to a much stronger inhibition than the CO-oxidizing activity. In the presence of fluoride ([NaF] > 300 mM) the CO-oxidizing activity was scarcely affected, whereas the formate-oxidizing activity was almost completely suppressed (Fig. 5). Kinetics of methylene blue reduction with CO. The reduction of methylene blue by the soluble fraction of the sonic extract with CO proceeded linearly with time (0 to 3 min), and the rate was proportional to the amount of protein added in the range measurable photometrically (E615 < 2.5). The dependence of the methylene blue reduction rate on the substrate concentration (CO) followed simple Michaelis-Menten kinetics: plots of 1/v versus 1/[S] were linear. The Km turned out to be 45 ,aM CO. Exclusion of hydrogen gas and free formate as intermediates of the CO oxidation reaction. Cells of P. carboxydovorans growing CO-autotrophically contain hydrogenase and formate oxidase (18). Thus, the oxidation of carbon monoxide to carbon dioxide and hydrogen gas (CO + H20 -' CO2 + H2; AGO' = -20.1 kJ [-4.8 kcal]) (25) and the hydratation to free formate (CO + H20-- HCOO- + H+; AGO = -16.3 kJ [-3.9 kcal]) (25), which is afterwards oxidized to C02, have to be considered as alternate possibilities of CO oxidation; both reactions are thermodynamically possible. If hydrogen gas is formed during the oxidation of CO it should be possible to couple this reaction to the reduction of NAD+ catalyzed by the
815
soluble NAD-specific hydrogenase of Alcaligenes eutrophus (Fig. 6). After addition of this hydrogenase to the reaction mixture, however, NAD+ reduction was not detectable (Fig. 6, B). Controls performed by the addition of small amounts of hydrogen-saturated buffer (Fig. 6, C and D) or of methylene blue (Fig. 6, E) showed that the hydrogenase remained still active in the reaction mixture and that the CO-oxidizing activity was still capable of oxidizing CO. A similar assay was used to couple the formation of formate with the reduction of methyl viologen catalyzed by extracts of Clostridium cylindrosporum (Fig. 7). However, the dye was 100
.0 p
p
-
50
10 0
100
[NaF](mM)
500
FIG. 5. Differentiation of the CO-oxidizing activity and formate-oxidizing activity of the soluble fraction by inhibition by fluoride. The rate of methylene blue reduction was measured as described in the footnote to Table 1.
TABLE 3. Effect of metal-complexing agents on the CO- and the formate-oxidizing activities" Inhibition (%)
Inhibitor (100 mM)
CO oxidation
Formate oxidation
11 Hypophosphite (NaH2PO2) 89.4 Chlorate (NaCI03) 0 64.1 64.1 100 Cyanide (NaCN) Azide (NaN3) 92.2 100 Fluoride (NaF) 2.3 81.7 " Assay: the CO-oxidizing activity and the formateoxidizing activity were assayed according to the legend to Table 1, but in the presence of 100 mM inhibitor.
20
Ti,me (min
FIG. 6. Experiment to detect hydrogen gas as a possible intermediate of CO oxidation. Assay: 0.91 ml of 50 mM KH2PO4-KOH buffer (pH 7.0); 10 ,ul of mix (1 U of glucose oxidase + 1 U of catalase); 50 ILI of 2 mMglucose + 30 LM NAD+; serum-stoppered cuvettes were flushed with CO. Arrows: A, addition of 20 u1 of crude bacterial extract of P. carboxydouorans; B, 20 fLl of NAD+-specific hydrogenase; C, 100 /1l of H2saturated buffer; D, 50 u1 of H2-saturated buffer (so far measured at 365 nm); E, 10I l of 2.5 mM methylene blue (measured at 615 nm).
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MEYER AND SCHLEGEL
not reduced; addition of formate d( emonstrated that the extract was still capable c)f catalyzing the reduction of methyl viologen w,ith formate. The results clearly indicate that ne ither hydrogen gas nor free formate is an inte rmediate of the carbon monoxide oxidation reac tion. Other properties of the CO ox idation reaction. The CO-oxidizing activity w. as only present in CO-grown cells of P. carbo)xydoL'orans and was very sensitive to alterat.ions of the growth conditions; it was not lost wihen extracts were aerobically dialyzed for 24 h ag ainst 50 mM phosphate buffer (pH 7.0), whereas tthe formateoxidizing activity was completely destroyed. Ethylenediaminetetraacetic acid (5'0 mM) was without effect on the CO-oxidizing aictivity. The reduction of methylene blue with CO started immediately after addition of solulble fraction; however, the reaction mediated by the particle fraction sometimes started after a laLg of up to 15 min.
The CO-oxidizing activity was as,sayed under strictly anaerobic conditions, indicat ing that molecular oxygen is not involved and the second oxygen atom in the CO2 produceci is derived from water; thus, CO oxidation is riot due to a monooxygenase.
DISCUSSION During aerobic growth of P. carbo rydoLtorans with carbon monoxide as the sol e source of carbon and energy, CO is oxidized to CO2 (18). CO oxidation was also shown to occutr in extracts of CO-grown cells. Different electroin acceptors such as pyocyanine, methylene bluie, thionine,
A
B
c
c |
2_____
Time (min)2
FIc. 7. Experiment to detect formate as apossible intermiediate of CO oxidation. Assay: c(omponents as in the legend to Fig. 6, but in the pres;ence of 5 [aM W methyl ciologen instead of NAD+. At-rr 9)VS: A, addition of 10 til of crude bacterial extract cif P. carboxydocorans; B, 10 pl of craude extract off C. cylindro-
sporum containing about 0.15 U of foaimq7ntprP h>vr1r)t ltlCt (XUctlY(l genase; C. .30 yl of 5 mM formate.
-
or phenazine methosulfate were readily reduced by carbon monoxide, whereas oxidized flavine adenine dinucleotide, oxidized riboflavine 5'phosphate, cytochrome c, and the pyridine nucleotides turned out to be ineffective. This supports the assumption that the electrons are delivered at the level of succinate dehydrogenase and that a quinone is possibly the physiological electron acceptor. Centrifugation of extracts in sucrose gradients characterized hydrogenase as a membranebound enzyme, whereas the CO- and the formate-oxidizing activities were soluble or only slightly attached to the membrane; their distribution at a 1:1 ratio between the soluble and particle fraction after centrifugation may be due to partial adsorption of the soluble enzyme to the particles. Dehydrogenation of CO is probably due to a separate enzyme and does not involve formate dehydrogenase or hydrogenase activity. This conclusion is based on the differential sensitivity of the enzyme activities to heat and metal-complexing inhibitors such as NaF, as well as on the failure to trap free formate or hydrogen gas in coupled optical assays. The basic reaction actually used in P. carboxydou,orans is CO + H2O -* CO2 + 2 Ht + 2e The reaction serves two different functions: (i) to provide electrons for energy generation and reductive syntheses and (ii) to produce CO2 for the reductive pentose phosphate cycle. The COoxidizing system thus serves catabolic functions; it is inducible and is only formed during growth on CO. Hydrogenase is inducible and is only present in cells grown with hydrogen (H2,-02CO2) or with pyruvate, whereas the formateoxidizing activity is constitutive and is present in cells grown with CO (CO-O0), H2 (H2-02GCO,), acetate, or pyruvate. In contrast, the physiological function of COoxidizing activity in methanogenic bacteria (5, 7, 8, 16, 23), sulfate-reducing bacteria (28, 29, 30), photosynthetic bacteria (26), and clostridia (9, 1)0, 24) is far from clear. In these bacteria the CO-oxidizing activity is constitutive. It may be due to the nonspecific activities of enzymes fulfilling quite different physiological functions, e.g. in the case of methane monooxygenase (2, 6). Furthermore, the CO-oxidizing activities of the anaerobic bacteria have been shown to reduce methylene blue only slightly, but viologen dyes with high rates, whereas extracts of P. carboxvdo'orans are not capable of oxidizing CO with vriologen dyes as electron acceptors. ACKNOWLEDGMENTS (KOLDINT We thiank K. Schneider for providing SOIlIble hydrogenase
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CO OXIDATION IN P. CARBOXYDOVORANS
of A. eutrophus and R. Wagner for providing extracts of C. cylindrosporum. LITERATURE CITED 1. Aggag, M., and H. G. Schlegel. 1973. Studies on a grampositive hydrogen bacterium, Nocardia opaca lb. III. Purification, stability and some properties of the soluble hydrogen dehydrogenase. Arch. Microbiol. 100:25-39. 2. Beijerinck, M., and A. van Delden. 1903. Uber eine farblose Bakterie, deren Kohlenstoffnahrung aus der atmospharischen Luft herruhrt. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. 2 10:33-47. 3. Beisenherz, G., H. J. Bolze, T. Bucher, R. Czok, K. H. Garbade, E. Meyer-Arendt, and G. Pfleiderer. 1953. Diphosphofructose-Aldolase, PhosphoglyceraldehydDehydrogenase, Milchsaure-Dehydrogenase, Glycerophosphat-Dehydrogenase und Pyruvat-Kinase aus Kaninchenmuskulatur in einem Arbeitsgang. Z. Naturforsch. 8b:555-577. 4. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254. 5. Daniels, L., G. Fuchs, R. K. Thauer, and J. G. Zeikus. 1977. Carbon monoxide oxidation by methanogenic bacteria. J. Bacteriol. 132:118-126. 6. Ferenci, T. 1974. Carbon monoxide stimulated respiration in methane-utilizing bacteria. FEBS Lett. 41:94-98. 7. Fischer, F., R. Lieske, and K. Winzer. 1931. Biologische Gasreaktionen. I. Mitt. Die Umsetzungen des Kohlenoxyds. Biochem. Z. 236:247-267. 8. Fischer, F., R. Lieske, and K. Winzer. 1932. Uber die Bildung von Essigsaure bei der biologischen Umsetzung von Kohlenoxyd und Kohlensaure mit Wasserstoff zu Methan. Biochem. Z. 245:2-12. 9. Fuchs, G., G. Andress, and R. K. Thauer. 1975. COoxidation by anaerobic bacteria: indications for the involvement of a vitamin B12 compound, p. 231-236. In H. G. Schlegel, G. Gottschalk, and N. Pfennig (ed.), Microbial production and utilization of gases. E. Goltze KG, Gottingen. 10. Fuchs, G., U. Schnitker, and R. K. Thauer. 1974. Carbon monoxide oxidation by growing cultures of Clostridium pasteurianum. Eur. J. Biochem. 49:111-115. 11. Hasemann, W. 1927. Zersetzung von Leuchtgas und Kohlenoxyd durch Bakterien. Biochem. Z. 184:147-171. 12. Hubley, J. H., J. R. Milton, and J. F. Wilkinson. 1974. The oxidation of carbon monoxide by methane-oxidizing bacteria. Arch. Microbiol. 95:365-368. 13. Lax, E. 1967. D'Ans-Lax: Taschenbuch fur Chemiker und Physiker, vol. 1, p. 1-1205. Springer-Verlag, Berlin. 14. Kistner, A. 1953. On a bacterium oxidizing carbon monoxide. Proc. K. Ned. Akad. Wet. Ser. C 56:443-450.
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15. Kistner, A. 1954. Conditions determining the oxidation of carbon monoxide by Hydrogenomonas carboxydovorans. Proc. K. Ned. Akad. Wet. Ser. C 57:186-195. 16. Kluyver, A. J., and C. G. Schnellen. 1947. On the fermentation of carbon monoxide by pure cultures of methane bacteria. Arch. Biochem. 14:57-70. 17. Lantzsch, K. 1922. Actinomyces oligocarbophilus (Bacillus oligocarbophilus Beij.), sein Formwechsel und seine Physiologie. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. 2 57:309-319. 18. Meyer, O., and H. G. Schlegel. 1978. Reisolation of the carbon monoxide utilizing hydrogen bacterium Pseudomonas carboxydovorans (Kistner) comb. nov. Arch. Microbiol. 118:35-43. 19. Nozhevnikova, A. N., and G. A. Zavarzin. 1974. On the taxonomy of CO-oxidizing gram-negative bacteria. Izv. Akad. Nauk. SSSR Ser. Biol. 3:436-440. 20. Ooyama, J., and T. Shinohara. 1971. Simultaneous fixation of CO and N, in the presence of H2 and O0 by a bacterium. Rep. Ferment. Res. Inst. 40:1-5. 21. Sanjieva, E. U., and G. A. Zavarzin. 1971. Oxidation of carbon monoxide by Seliberia carboxydohydrogena. Dokl. Akad. Nauk. SSSR 196:956-958. 22. Savelieva, N. D., and A. N. Nozhevnikova. 1972. Autotrophic growth of Seliberia carboxydohydrogena during oxidation of hydrogen and carbon monoxide. Mikrobiologiya 41:813-817. 23. Stephenson, M., and L. H. Stickland. 1933. The bacterial formation of methane by the reduction of one carbon compounds by molecular hydrogen. Biochem. J. 27:1517-1527. 24. Thauer, R. K., G. Fuchs, B. Kaufer, and U. Schnitker. 1974. Carbon monoxide oxidation in cell-free extracts of Clostridium pasteurianum. Eur. J. Biochem. 45:343349. 25. Thauer, R. K., K. Jungermann, and K. Decker. 1977. Energy conservation in chemotrophic bacteria. Bacteriol. Rev. 41:100-180. 26. Uffen, R. L. 1976. Anaerobic growth of a Rhodopseudomonas species in the dark with carbon monoxide as sole carbon and energy substrate. Proc. Natl. Acad. Sci. U.S.A. 73:3298-3302. 27. Wehmer, C. 1926. Biochemische Zersetzung des Kohlenoxyds. Ber. DtLsch. Chem. Ges. 59:887-890. 28. Yagi, T. 1958. Enzymic oxidation of carbon monoxide. Biochim. Biophys. Acta 30:194-195. 29. Yagi, T. 1959. Enzymic oxidation of carbon monoxide. II. J. Biochem. (Tokyo) 46:949-955. 30. Yagi, T., and N. Tamiya. 1962. Enzymic oxidation of carbon monoxide. III. Reversibility. Biochim. Biophys. Acta 65:508-509. 31. Zavarzin, G. A., and A. N. Nozhevnikova. 1977. Aerobic carboxydobacteria. Microb. Ecol. 3:305-326.
