METABOLISM OF FRUCTOSE-1,6-DIPHOSPHATE AND ACETATE IN ACETOBACTER SUBOXYDANS1

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1 METABOLISM OF FRUCTOSE-1,6-DIPHOSPHATE AND ACETATE IN ACETOBACTER SUBOXYDANS1 PAUL A. KITOS,2 TSOO E. KING, AND VERNON H. CHELDELIN Department of Chemistry and Science Research Institute, Oregon State College, Corvallis, Oregon. Received for publication April 1, 1957 Cell free extracts of Acetobacter suboxydans catalyze the formation of triosephosphate from either glucose or glycerol. The same extracts consume triosephosphate by the aldolase and pentose cycle reactions (Hauge et al., 1955; Kitos, 1956). Although fructose-1, 6-diphosphate (FDP)3 is oxidized by molecular oxygen in the presence of either whole cells or cell extract, other evidence does not indicate that glycolysis operates to a significant degree in this organism (King and Cheldelin, 1954). This paper is concerned with the conversion of FDP to 3-phosphoglyceric acid (3-PGA)3 and the further metabolism of this substrate as far as the products of acetate activation. The conditions under which these glycolytic enzymes function differentiate them from those commonly observed in other cells. MATERIALS AND METHODS Lyophilized cells of A. suboxydans ATCC strain 621 were prepared as reported previously (King and Cheldelin, 1954). For the preparation of cell free extracts, 2.5 g of lyophilized cells were suspended in 50 ml of ice-cold 0.01 molar trimethylolaminomethane (Tris)3 buffer at ph 8.2 and subjected to sonic 1 Supported by grants from the Nutrition Foundation, Inc., and the Division of Research Grants and Fellowships, National Institutes of Health, U. S. Public Health Service. Published with the approval of the Monographs Publications Committee, Research paper No. 318, School of Science, Department of Chemistry. 2 Taken from the dissertation for the degree of Doctor of Philosophy of Paul A. Kitos, Oregon State College, Present address, E. I. dupont de Nemours Co., Newark, Delaware. 3The following abbreviations are used throughout this paper: FDP = fructose-1,6-diphosphate; 3-PGA = 3-phosphoglyceric acid; Tris = trimethylolaminomethane, CoA = Coenzyme A; DPN and TPN = di- and triphosphopyridine nucleotide; ATP = adenosine triphosphate. disintegration according to a method employed previously (King and Cheldelin, 1956). The cofactors in the cell extract were removed by dialysis for 18 hr at 1 to 3 C, against a suspension of Dowex-50 in the hydrogen form. The protein precipitated within the dialysis bag4 and was collected by centrifuging at 2000 X G. The precipitate was redissolved in 0.01 molar Tris buffer at ph 8.2 and brought to the original volume of cell extract. The small amount of residue which was formed was removed by low speed centrifugation. The supernatant solution was clear, slightly pink and generally less active than the crude cell free extract. Unless otherwise stated, the chemicals used were obtained commercially and used without further purification. These were obtained as follows: Coenzyme A (CoA)3, approximately 80 per cent pure, from either Pabst laboratories or Sigma Chemical Co.; adenosine triphosphate (ATP),3 94 per cenit pure with respect to acid hydrolyzable phosphate, from Pabst laboratories; FDP, about 68 per cent pure, was obtained from Schwarz laboratories. It was purified by Dowex-1 chromatography according to the method of Khym and Cohn (1953). Sodium ethylenediamine tetraacetic acid (versene) was obtained from the Versene Corporation. Acetate (1-C'4-labeled) was purchased from Tracerlab, Inc. Acethydroxamic acid was determined by the method of Lipmann and Tuttle (1945). The system of Chou and Lipmann (1952) was used for the preparation of pigeon liver acetate-activating enzyme. Citric acid was measured by the 4 Protein precipitation during dialysis was observed only in crude cell free extract under the conditions described. The precipitate was not formed in partially purified preparations. The amount of precipitate as well as the final ph depended upon the salt concentrations in the original extracts. The removal of cofactors by this treatment was found to be more complete in the crude extracts than in partially purified fractions. 565

2 566 KITOS, KING, AND CHELDELIN [VOL. 74 acetic anhydride-pyridine technique (Saffran and Denstedt, 1948). Inorganic and organic phosphates were determined by the methods described by Le Page t(1949). 3-Phosphoglyceric acid was determined as 3 hr stable phosphate according to Le Page (1949). (Thus any compound with this property would be recorded in these experiments as 3- PGA.) Fructose and FDP were assayed by the resorcinol method of Roe (Le Page, 1949). Nonspecific sugar tests were carried out by the anthrone method (Morris, 1948). The colorimetric procedure of Barker and Summerson (1941) for lactic acid and the colorimetric estimation of pyruvic acid as the 2,4-dinitrophenylhydrazone (Lardy, 1947) were used. Spectrophotometric measurements of DPN and TPN reduction were carried out in a Beckman model B spectrophotometer with 1 cm2 cuvette. Tetrazolium (triphenyltetrazolium chloride) reduction experiments were conducted as before (Hauge et al., 1955). In the radiochemical experiments, direct plating was carried out on stainless steel planchets of 25 or 31 mm diameter. The samples were counted under standard conditions with a Geiger tube mounted on a shielded sample holder, connected to a Tracerlab 64 utility scaler. The counts per min were corrected for background and for self absorption wherever possible. RESULTS Fructose diphosphate dissimilation. Oxidation of FDP in the crude cell extract was strongly stimulated by TPN. However, when the oxidation was carried out in the presence of versene or when a Dowex-50 treated extract was used, the reactions became DPN specific. Addition of M'IgK to the Dowex treated enzymes restored the ability to utilize TPN (data not shown). It appeared from this behavior that versene and Dowex-50 were removing 1MIg from the system, thereby preventing the conversion of FDP to fructose-6-phosphate, and hence blocking pentose cycle operation. The DPN specific oxidation of FDP in the absence of the pentose cycle suggested a conversion to triosephosphate and subsequent triosephosphate dehydrogenase activity. Since aldolase and triosephosphate isomerase function in this organism (Hauge et al., 1955), FDP serves as a ready source of glyceraldehyde-3-phosphate. The DPN-specific oxidation of triosephosphate was enhanced at values above ph 8.2, having an optimum near 9.0. The light absorption at 340 m, was not due to the dihydroxyacetone- DPN complex that has been shown to form at high ph values (King and Cheldelin, 1954; Burton and Kaplan, 1953). This was shown by experiments in which tetrazolium was used as the ultimate hydrogen acceptor. Under these conditions the tetrazolium reduction was mediated by DPN but not by TPN. The DPN-specific oxidation of FDP required either arsenate or orthophosphate. Absence of these cosubstrates resulted in no nucleotide reduction (figure 1). Iodoacetate completely inhibited the oxidation reaction. The accumulation of 3-PGA, determined as 3 hr stable P, was approximately equivalent to the amount of FDP oxidized, as shown in table 1. Since the Dowex-50 treated cell extract was free of Mg, use of this preparation eliminated the need for fluoride to inhibit enolase activity and trap 3-PGA. Anaerobically, FDP was converted to a product that responded positively to the Barker-Summerson lactic acid test. However, the same color reaction was given aerobically; under these conditions lactate is rapidly converted to acetate m. 0.6 E 0 to, C) o. 2rnoles As04 no AsO-.-~~ TIME, MINUTES. Figure 1. Dependence upon arsenate of the DPN-specific FDP oxidation in Acetobacter suboxydans. Spectrophotometric measurements express DPN reduction at 340 ma at room temperature. Each vessel contained Tris, ph 9.0, 80,umoles; versene, 108 Mmoles; DPN, 1 /Amole; FDP, 0.88,umole; A. suboxydans Dowex-50 treated sonic cell free extract, 0.2 ml; and water to 2 ml volume. Blank corrections were made.

3 1957] FRUCTOSE-1,6-DIPHOSPHATE AND ACETATE IN A. SUBOXYDANS 567 TABLE 1 Stoichiometry of fructose-i,6-diphosphate (FDP) oxidation by a diphosphopyridine nucleotide-specific Acetobacter suboxydans enzyme FDP Added Enzyme Tetrazolium FDP 3-PGA* Added Reduced Consumed Produced j.moles ml pmoles pmoles Amoles The reaction was carried out in vacuo in 3" test tubes at 37 C. Each vessel contained Tris, ph 9.0, 30,moles; versene, 40.5,Amoles; arsenate, 1,umole; DPN, 0.1 /mole; triphenyltetrazolium chloride, 10,umoles; and water to 1 ml. Enzyme = Dowex-50 treated sonic digested cell extract. Appropriate blanks were run containing no FDP. Time, 140 min. The reactions were stopped by mixing the contents of each tube with 1 ml of 12% trichloracetic acid and 3 ml H20, then centrifuging. Two ml aliquots of the supernatant solution were used for 3 hr stable P determinations; 1 ml for total P after H2SO4 digestion; and 1 ml for residual FDP determination. * Phosphoglyceric acid. TABLE 2 Cofactors for the acetylation of coenzyme A by cell free extracts of Acetobacter suboxydans Ingredient Missing Acethydroxamic Acid Formed Crude cell etat,umoles Dowex-50 treated cell extract,umoles 1. None Enzyme CoA ATP KOAc...ọ MgCl Glutathione KF Complete system: A. suboxydans cell free extract (enzyme), 10 mg protein; acetate, 10 jumoles; glutathione, 20 jamoles; MgCl2, 7.5,umoles; CoA, 0.4,mole; KF, 45,moles; ATP, 10 jamoles; NH20H 500 jamoles; Tris buffer, ph 8, 50 Mmoles. Total volume = 2 ml. Time = 1 hr at 30 C. (experiments not shown). The color was found to be due to dihydroxyacetone, and therefore the anaerobic production of lactate from FDP could not be claimed. TABLE 3 Citrate formation by pigeon liver enzyme and Acetobacter suboxydans cell free extract A. sub- Pigeon Acetate Pyruvate ATP oxydans Liver Con- Citrate Cell densing Formed Extract Enzyme,umoles IAmoles,umoles ml ml,umoles Experiments were carried out in 3" test tubes at 37 C. Time, 135 min. In addition to the variables tested each vessel contained Tris, ph 8.2, 100 iamoles; glutathione, 5,umoles; MgC12, 5,umoles; CoA, 0.5,umole; ATP, 0.5,mole; oxalacetate, 10,moles; and water to 1 ml. The reaction was stopped by the addition of 1 ml of 10 per cent trichloracetic acid and the protein was removed by centrifugation. One ml of the supernatant layer was tested for citrate. TABLE 4 Arylamine acetylation by reconstituted enzyme systems Pigeon Liver Enzyme Acelobacter susboxydans Amid A40 Acetate A-60 Acetyl Extrat Acetylated activating acceptor xtrac paminoedi ml ml ml "moles Experiments were carried out in 3" test tubes at 37 C. Time = 60 min. In addition to the variables stated, each vessel contained Tris buffer, ph 8.2, 200 inmoles; glutathione, 10 Mmoles; MgCl2, 10 jsmoles; p-aminobenzoic acid, 2 j,moles; CoA, 1 j,mole; ATP, 10 jamoles; and water to 2 ml. A. suboxydans cell free extract, Dowex-50 treated, contained 12.6 mg protein per ml. When 3-PGA was introduced into the cell free system, TPN reduction was observed after a lag of 10 to 20 min, provided that Mg+-+ and ADP were added. However, the products of the oxidation were not identified.

4 568 KITOS, KING, AND CHELDELIN [VOL. 74 v 0.6_ x 0 o < UOO.2 O._ (no w 'I )L npjmoles Figure 2. Effect of reactant concentrations on the enzymatic formation of acetyl CoA. Measurements depict acethydroxamic acid formation at 30 C for 60 min. Complete system contained: Tris, ph 8.2, 100 Mmoles; potassium acetate, 100,moles; glutathione, 10,umoles; NH20H, 500,Amoles; MgCl2, 10 MAmoles; Acetobacter suboxydans Dowax-50 treated cell free extract (enzyme); protein, 1.2 mg; and water to 2 ml volume. TABLE 5 Fate of C14 from CH3C1400H administered to resting cells of Acetobacter suboxydans CH3C'400H Added Radioactivity Distribution Experiment NO. Total Respiratory CO2 Medium Cells _Moles _ pm Cpm % of Total Cpm % of Total Cpm % of Total X X X X 10' X Each vessel contained 250 mg dry wt of fresh A suboxydans cells in a 500 ml 3 necked standard taper flask. Composition of medium, 0.4 per cent monopotassium phosphate, ph 6.0. Volume = 100 ml. CO2 free air was bubbled through the medium for 60 min before CH3C'400Na was added to the vessels. Experiment 1: 6 MAmoles CH3C'400Na, specific activity 5,c/Mmole. Experiment 2: 200,umoles CH3C1400Na, specific activity c//Amole. In both experiments 250,moles glucose were added after 2 hr. The metabolic CO2 was trapped by sparging through 0.5 N NaOH. The NaOH traps were replaced periodically and the CO2 was plated as BaCO,, weighed and counted. The activation of acetate. A. suboxydans converts lactate, pyruvate, ethanol, and acetaldehyde to acetate and acetoin (King and Cheldelin, 1954). Although acetate is not oxidized either by whole cells, cell free extracts, or cell homogenates, it may be mobilized to form acetyl CoA (Kitos et al., 1955). Treatment of the cell extract with Dowex-50 removes various endogenous cofactors so that acetate activation is completely dependent upon ATP, Mg++ and CoA (table 2). Glutathione presumably functions to maintain the CoA in the reduced state, consequently its effect is only stimulatory. The inhibition by fluoride, particularly with the Dowex-50 treated enzyme, has been attributed to the effect of the CoA acetylating enzyme (Aisenberg and Potter, 1955). Fluoride was introduced to inhibit ATPase. Lipoic acid in amounts up to 50m,ug per ml exerted no effect on the acetylation of CoA when the Dowex-50 treated enzyme was employed. Experiments in which alumina treated cell free extracts were used likewise elicited no response to added lipoic acid. The alumina treatment reduced the lipoic acid content of the extracts to 50 per cent of its normal level (King et al., 1956). The requirements for other components of the acetylation system are qualitatively the same as those described for other tissues (Lynen and Ochoa, 1953). The effect of the concentration of reactants upon the acetylation of CoA is illustrated in figures 2A, B and C. The optimum concentrations of each of these components are 5 X 10-2 molar acetate, 5 X 10-3 molar Mg++, 5 X 10-3 molar ATP and approx 5 X 10-4 molar CoA. The ph optimum lies between 7.8 and 9.0. At ph 7.4,

5 1957] FRUCTOSE-1,6-DIPHOSPHATE AND ACETATE IN A. SUBOXYDANS 569 that used by Rao (1955), the activity was only about 2 of the optimum value. Inorganic phosphate stimulates the formation of acetyl CoA in cell free systems. The nature of the stimulation is not clear. The fact that cell free extracts of A. suboxydans catalyze the formation of acetyl CoA suggests that the active acyl group may subsequently be transferred to appropriate acceptors. However, attempts to detect the presence of the condensing enzyme were unsuccessful. Only when the acceptor fraction from pigeon liver was added to the otherwise complete system was any citrate formed (table 3). A. suboxydans cell free extract was likewise unable to acetylate arylamines. However, when the cell extract was coupled with the A-60 pigeon liver enzyme fraction of Chou and Lipmann (1952), which contained the acceptor enzyme for acetylation of sulfonamides (but no acetate activating enzyme), acetylation of p- aminobenzoic acid, sulfanilimide or 4-aminoazobenzene (Handschumacher et al., 1951) could be realized (table 4). When the pigeon liver A-40 and A-60 fractions (the donor and acceptor enzymes) were combined, acetylation resulted in a manner similar to that from the combined A. suboxydans cell extract and pigeon liver enzyme. However, combined pigeon liver A-40 and A. suboxydans cell extract were inactive in this respect. Thus, there is no evidence for the occurrence of either the condensing enzyme or the arylamine acetylating enzyme in the cell free extracts of A. suboxydans. The stimulation by ATP of citrate formation from pyruvate suggests I4-3 03k~~ x \ m medium u) 2 00 z ^ - 01 glucose o added 0 I_ /cells _o..- o I I TIME, HOURS. Figure S. CH3C'400H incorporation into Acetobacter suboxydans. Experimental conditions as in table TABLE 6 Distribution of C14 in cell fractions after exposure to CH3C'400H Neutral Lipid Fraction Acid Fraction Alkaline ether extraction Extraction of cell Total hydrolyzate Cpm in Cells Whole cells, Cell hydrolyzate, Fraction 3 fraction 1 fraction 2 C % of Cpm % of C % of Total Total Cpm Total 762,00094,000i , , An aliquot of cells from experiment 1, table 5, was centrifuged from the medium, washed in 35 ml H20 and resuspended in 20 ml H20. An aliquot of the cell suspension was made alkaline with NaOH and extracted overnight with ether in a Kutscher-Steudel liquid-liquid extractor (fraction 1). The ether was evaporated and an aliquot of the oily residue was counted by direct plating. The cell suspension from the first extraction was then hydrolyzed in 1 N HCl for 2 hr at 100 C. The solution was made alkaline with NaOH and reextracted with ether for additional neutral fatsoluble material (fraction 2). The hydrolyzate was then made acidic and extracted further with ether to remove fatty acids (fraction 3). that the latter compound is first converted to free acetate before acetyl CoA is formed. In an attempt to detect the participation of administered acetate in the metabolism of A. suboxydans, resting cells were treated with CH3C'4- OOH at two levels of specific activity. The respiratory CO2, medium and cells were then examined for radioactivity. The results are summarized in table 5. In both experiments the CO2 production was negligible, and there was no significant appearance of C'4 in the CO2. After the experiment had proceeded for 2 hr, unlabeled glucose was added to the flask containing acetate of high specific activity (experiment 2, table 5). This resulted in a rapid increase in CO2 production but the C'4 content was still negligible throughout the 10 hr course of the experiment. Analysis of the medium and cells revealed no incorporation of acetate into the cells and no decrease in medium radioactivity until the point of glucose addition to the reaction vessels (figure 3). Thereafter the acetate in the medium decreased promptly to about 50 per cent of its original concentration and the cellular radioactivity increased correspondingly. When 200

6 570 KITOS, KING, AND CHELDELIN [VOL. 74,umoles of acetate of specific activity tic/,umole were used (representing 1/200 dilution of radioactivity compared to the previous experiment) no net incorporation or medium depletion was observed. Examination of the cells revealed that small but measurable quantities of isotope had been incorporated, principally into the lipid fraction (table 6). As shown in figure 3 this entry of acetate occurred only after the administration of glucose. In may be concluded that acetate is incorporated in extremely low concentrations into respiring cells of A. suboxydans, but only in the presence of an added energy source. This is in accordance with the fact that acetate may be activated only in the presence of added ATP. The slight incorporation of acetate, even in the presence of glucose, makes it unlikely that lipid synthesis by this route is a major process in this organism. DISCUSSION From the observations at least two of the reactions characteristic of glycolysis, namely, aldolase and glyceraldehyde-3-phosphate dehydrogenase operate in A. suboxydans under the special conditions tested. However, the conversion of 3-PGA to acetate could not be demonstrated. Indeed this seems unlikely in either resting or proliferating cultures in view of the negligible accumulation of the expected reaction products; triosephosphate has only a transient existence when exposed to cell extract, yet unless the system is deficient in Mg+ this substrate disappears predominantly via the pentose cycle. As a matter of fact even under the most favorable experimental conditions the oxidation of glyceraldehyde-3-phosphate in the cell extract is much slower than the slowest step in the pentose cycle (phosphogluconate oxidation; Hauge et al., 1955). Thus the contribution of triosephosphate dehydrogenase to the metabolism of this organism under normal conditions is still to be determined. One or more functions of CoA in A. suboxydans must be vital since pantothenic acid is an essential component of the medium. However, the role of CoA acetylation remains obscure except for its part in lipid synthesis. Neither the oxidation of acetate or of lipid material is likely by conventional means since at no time did significant radioactivity from carboxyl labeled acetate appear in the metabolic gases. Experiments with radioactive glucose, glycerol and other substrates are in progress to determine the relative contributions of glycolysis, the pentose cycle and other pathways to total carbohydrate dissimilation in this organism, both in the presence and absence of adequate nitrogen sources. SUMMARY Fructose-1,6-diphosphate is dissimilated by a diphosphopyridine nucleotide specific oxidation to 3-phosphoglyceric acid in Acetobacter suboxydans cell free extracts. This proceeds in the absence of Mg++, a condition which prevents entrance of fructose diphosphate into the pentose cycle. 3-Phosphoglyceric acid induces enzymatic pyridine nucleotide reduction in presence of Mg+ and adenosine diphosphate. Cell free extracts of the microorganism catalyze the formation of acetyl coenzyme A but not the formation of citrate or acetyl arylamines. Acetate is incorporated in small amounts into cell lipid fractions, only in the presence of an energy source. REFERENCES AISENBERG, A. C. AND POTTER, V. R Effect of fluoride and dinitrophenol on acetate activation. Federation Proc., 14, 171. BARKER, S. B. AND SUMMERSON, W. H The colorimetric determination of lactic acid in biological material. J. Biol. Chem., 138, BURTON, R. M. AND KAPLAN, N A DPN-specific glycerol dehydrogenase from Aerobacter aerogenes. J. Am. Chem. Soc., 75, CHOU, T. C. AND LIPMANN, F Separation of acetyl transfer enzymes in pigeon liver extract. J. Biol. Chem., 196, HANDSCHUMACHER, R. E., MUELLER, G. C., AND STRONG, F. M An improved enzymatic assay for coenzyme A. J. Biol. Chem., 189, HAUGE, J. G., KING, T. E., AND CHELDELIN, V. H Oxidation of dihydroxacetone via the pentose cycle in Acetobacter suboxydans. J. Biol. Chem., 214, KHYM, J. X. AND COHN, W. E The separation of sugar phosphates by ion exchange with the use of the borate complex. J. Am. Chem. Soc., 75, KING, T. E. AND CHELDELIN, V. H Oxidations in Acetobacter suboxydans. Biochim. et Biophys. Acta, 14,

7 1957] FRUCTOSE-1,6-DIPHOSPHATE AND ACETATE IN A. SUBOXYDANS 571 KING, T. E. AND CHELDELIN, V. H Glucose oxidation and cytochromes in solubilized particulate fractions of Acetobacter suboxydans. J. Biol. Chem., 224, KING, T. E., KAWASAKI, E. H., AND CHELDELIN, V. H Tricarboxylic acid cycle activity in Acetobacter pasteurianum. J. Bacteriol., 72, KITOS, P. A Terminal oxidation pathways in Acetobacter suboxydans. Ph.D. Thesis, Oregon State College. EITOS, P. A., KING, T. E., AMBROSE, J. A., AND CHELDELIN, V. H Acetate activation in Acetobacter suboxydans. Federation Proc., 14, LARDY, H. A In Manometric techniques and tissue metabolism, pp , 2nd ed. Edited by W. W. Umbreit, R. H. Burris, and J. F. Stauffer. Burgess Publishing Co., Minneapolis, Minn. LE PAGE, G. A In Manometric techniques and tissue metabolism, pp , 2nd ed. Edited by W. W. Umbreit, R. H. Burris, and J. F. Stauffer. Burgess Publishing Co., Minneapolis, Minn. LIPMANN, F. AND TUTTLE, L. C A specific micromethod for the determination of acyl phosphates. J. Biol. Chem., 159, LYNEN, F. AND OCHOA, S Enzymes of fatty acid metabolism. Biochim. et Biophys. Acta, 12, MORRIS. D. L Quantitativedetermination of carbohydrates with Dreywood's anthrone reagent. Science, 107, RAO, N. R. Ragbavendra 1955 Pyruvate and acetate metabolism in Acetobacter suboxydans and Acetobacter aceti. Doctoral Dissertation, University of Illinois, Urbana. SAFFRAN, M. AND DENSTEDT, 0. F A rapid method for the determination of citric acid. J. Biol. Chem., 175, Downloaded from on September 6, 2018 by guest