JOURNAL OF BACTERIOLOGY, Nov. 1969, p. 962-968Copyright 0 1969 American Society for Microbiology

Vol. 100, No. 2Printed in U.S.A.

Enzymes of Intermediary Carbohydrate Metabolismin the Obligate Autotrophs Thiobacillus thioparus

and Thiobacillus neapolitanustEMMETT J. JOHNSON2 AND S. ABRAHAM

Bruce Lyon Memorial Research Laboratory, Children's Hospital Medical Center ofNorthern Californlia,Oakland, California 94609

Received for publication 15 May 1969

Levels of enzymes operative in the Embden-Meyerhof-Parnas (glycolytic) path-way, pentose phosphate cycle, citric acid cycle, and certain other phases of inter-mediary carbohydrate metabolism have been compared in Thiobacillus thioparusand T. neapolitanus. All enzymes of the glycolytic pathway except phosphofructo-kinase were demonstrated in both organisms. There were some striking quantitativedifferences between the two organisms with respect to the activities of the individualenzymes of the glycolytic pathway and the citric acid cycle. Qualitative differenceswere also found: the isocitrate dehydrogenase activity of T. thioparus is strictlynicotinamide adenine dinucleotide phosphate (NADP)-dependent, whereas that ofT. neapolitanus is primarily nicotinamide adenine dinucleotide-dependent, activitywith NADP being low; the glucose-6-phosphate dehydrogenase of T. thioparus isparticulate, whereas that of T. neapolitanus is partly soluble and partly particulate;the 6-phosphogluconate dehydrogenase of T. thioparus is soluble, that of T. neapoli-tanus is partly soluble and partly particulate. All enzymes which function in thecarbon reduction cycle were present at very high levels. In contrast, enzymes whichoperate exclusively in cycles other than the carbon reduction cycle were present atlow levels. Of the enzymes not operative in the carbon reduction cycle that wereexamined, isocitric dehydrogenase had the highest specific activity. Both organismspossessed reduced nicotinamide adenine dinucleotide dehydrogenase activity. Thequalitative and quantitative aspects of the data are discussed in relation to possiblebiochemical explanations of obligate autotrophy.

The existence in obligate autotrophs of meta-bolic pathways for carbohydrate utilization simi-lar to those which exist in heterotrophic organismshas been inferred on the basis of indirect evi-dence, obtained primarily from studies on theassimilation and metabolism of substrates orintermediates associated with these pathways.Very little enzymological evidence bearing on thisquestion is available. Certain enzymes of the tri-carboxylic acid cycle have been detected inThiobacillus thioparus by Cooper (10), and inT. thioparus and T. thiooxidans by Smith et al.(35). Enzymes of the hexosemonophosphatepathway (pentose phosphate cycle) have beendemonstrated in T. ferrooxidans by Gale and

1Presnted in part at the 68th Annual Meeting ofthe AmericanSociety for Microbiology, Detroit, Mich., 5 May 1968.

' Present address: Deprtmet of Microbioloy and Im-munology, Tulane University School of Modicine, New Orleans,La. 70112.

Beck (13). The present report presents quantita-tive measurements of the activities of certainenzymes, notably ones associated with the Emb-den-Meyerhof-Parnas pathway (glycolytic path-way), the pentose phosphate cycle, and the citricacid cycle, in extracts of T. thioparus and T.neapolitanus.

MATERIALS AND METHODS

Conditions for bacterial growth and preparation ofextracts. T. thioparus (ATCC 8158) and T. neapoli-tanus were grown as previously described (26) andharvested during exponential growth. To detectheterotrophic contaminants, samples of liquid cul-tures were plated on tryptose-agar at daily intervalsuntil the time of harvesting. None of the culturesused showed any heterotrophic contamination, asjudged by the absence of growth on these plates after5 days at 30 C.

Approximately 4 g of cells (wet weight) per 10 ml

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of buffer [0.1 M tris(hydroxymethyl)aminomethane(Tris), pH 7.5] was passed through a French pressurecell at 18,000 psi and 2 to 4 C. All subsequent steps inthe treatment of the crude extracts were conducted at2 to 4 C. After treatment with deoxyribonuclease, thecrude extract was freed from coarse fragments andunbroken cells by centrifugation at 4,000 X g for 10min. Separation of crude extracts into soluble andparticulate fractions was performed by centrifugationat 144,000 X g for 120 min, which yielded a super-

natant fluid, designated 144.120, and a pellet, desig-nated 144pI20. The surface of 144p120 was rinsedtwice with 2 ml of 0.1 M Tris buffer (pH 7.5), and thenhomogenized with a Ten Broeck hand homogenizerin 5 ml of the same buffer.

Chemicals. All chemicals were obtained from com-mercial sources and were either reagent grade or of thehighest purity available. Adenosine triphosphate(ATP) was purcahsed from Pabst Laboratories,Milwaukee, Wis. Nicotinamide adenine dinucleotidephosphate (NADP) was purchased from either Cal-biochem, Los Angeles, Calif., or Boehringer Mann-heim Corp., New York, N.Y. Glucose-6-phosphatedehydrogenase, glucose-phosphate isomerase, pyru-vate kinase, glyceraldehydephosphate dehydrogenase,triosephosphate isomerase, c-glycerophosphate dehy-drogenase, 6-phosphogluconate, glucose-l-phosphate,and phosphoenolpyruvate were purchased from Boeh-ringer Mannheim Corp., New York, N.Y. Glucose-6-phosphate, fructose-1,6-diphosphate, nicotinamideadenine dinucleotide [oxidized (NAD) and reduced(NADH)], adenosine diphosphate (ADP), isocitrate,dihydroxyacetone phosphate, and ethylenediamine-tetraacetate (EDTA) were purchased from SigmaChemical Co., St. Louis, Mo. Lactate dehydrogenase,malate dehydrogenase, cis-aconitate, coenzyme A(CoA), oxalacetate, malate, and 3-phosphoglyceratewere purchased from Calbiochem. Potassium citratewas purchased from Mallinckrodt Chemical Works,St. Louis, Mo.; potassium pyruvate, from NutritionalBiochemical Corp., Cleveland, Ohio; and 5,5'-dithio-bis-(2 nitrobenzoic acid), from Aldrich Chemical Co.,Milwaukee, Wis. Acetyl-CoA was prepared accordingto the method of Stadtman (37) and fructose-6-phos-phate (glucose-6-phosphate-free), according to themethod of Borrebaek and Abraham (6).Enzyme assays. All enzyme assays, unless otherwise

noted, were performed with the 144,120 fraction.Enzyme activities were determined by following thechanges in optical density of the reaction mixtures at30 C with a Gilford automatic recording spectro-photometer. In all enzyme assays, concentrations ofsubstrates and of added enzymes (where needed) wereat least 10 times those required to yield maximalactivities. Absorbancy changes were measured withreference to reaction mixtures, devoid either of sub-strates or of coenzymes. The reactions were started byadditions of substrates, after a brief period of preincu-bation of the enzyme in the reaction mixture. Depend-ing upon the assay, 0.01 to 0.1 ml of the enzyme-containing fraction was used. Measurements of initialvelocities were made under conditions in whichkinetics were zero-order, and activity was proportionalto enzyme concentration.

Specific activities are reported as nanomoles eitherof pyridine nucleotide oxidized or reduced or of sub-strate converted to product, per milligram of proteinper minute. The following extinction coefficients wereused in the calculations: reduced NADP (NADPH)and NADH (340 nm), 6.22 X 103 liter X mole-1 Xcm-, (16); cis-aconitate (240 nm), 3.54 X 101liter Xmole-1 X cm-' (31); fumarate (240 nm), 2.11 X 103liter X mole-1 X cm-1 (31). Protein was determinedby the method of Gornall et al. (14) or that of Lowryet al. (24).

Enzymes were measured according to well-estab-lished methods or modifications of currently usedtechniques, as indicated below: glucokinase [ATP:D-glucose-6-phosphotransferase, EC 2.7.1.2 (1, 5,11)]; fructokinase [ATP: D-fructose-6-phosphotrans-ferase, EC 2.7.1.4 (1, 5, 11)]; mannokinase [ATP:D-mannose-6-phosphotransferase, EC 2.7.1.7 (1)];phosphoglucomutase [a-D-glucose-1, 6-diphosphate:a-D-glucose-1-phosphate phosphotransferase, EC2.7.5.1 (3)]; glucosephosphate isomerase [D-glucose-6-phosphate ketol-isomerase, EC 5.3.1.9 (3)]; man-nosephosphate isomerase [D-mannose-6-phosphateketol-isomerase, EC 5.3.1.8 (1, 2)]; phospho-fructokinase [ATP:D-fructose-6-phosphate 1-phos-photransferase, EC 2.7.1.11 (3)]; aldolase [fructose-1,6-diphosphate D-glyceraldehyde-3-phosphate-lyase,EC 4.1.2.13 (3)]; glyceraldehydephosphate dehydro-genase [D-glyceraldehyde-3-phosphate: NAD oxido-reductase (phosphorylating), EC 1.2.1.12 (23)];phosphoglycerate kinase [ATP: 3-phospho-D-glycer-ate 1-phosphotransferase, EC 2.7.1.3 (8)]; pyruvatekinase [ATP: pyruvate phosphotransferase, EC2.7.1.40 (9)]; citrate synthase [citrate oxaloacetate-lyase (CoA-acetylating) (36)]; aconitase [citrate (iso-citrate) hydro-lyase, EC 4.2.1.3 (30)]; isocitrate de-hydrogenase, NADP-linked [threo-D.-isocitrate:NADP oxidoreductase (decarboxylating), EC1.1.1.42] and NAD-linked [threo-D.-isocitrate: NADoxidoreductase (decarboxylating), EC 1.1.1.41 (27)];fumarase [L-malate-hydro-lyase, EC 4.2.1.2 (25) ];malate dehydrogenase [L-malate: NAD oxidoreduc-tase, EC 1 .1 .1.37 (28)]; malic enzyme [L-malate:NADP oxidoreductase (decarboxylating), EC1.1.1.40 (29)]; glucose-6-phosphate dehydrogenase[D-glucose-6-phosphate: NADP oxidoreductase, EC1.1.49 (22)]; 6-phosphogluconate dehydrogenase[6-phospho-D-gluconate: NADP oxidoreductase (de-carboxylating), EC 1.1.1.44 (17)]; ribosephosphateisomerase [D-ribose-5-phosphate ketol-isomerase, EC5.3.1.6 (4)]; reduced NAD dehydrogenase [NADH:(acceptor) oxidoreductase, EC 1.6.99.3] was meas-ured by following the decrease in optical density at340 mm in a system that contained 150 ,umoles ofglycylglycine buffer, pH 7.5, 0.4 ,umole of NADH,and crude extract in a total volume of 2 ml; andglycerol kinase [ATP: glycerol phosphotransferase,EC 2.7.1.30 (7)].

RESULTSAll values reported for specific activities are

average values, determined for at least four inde-pendently prepared extracts. Variation was never

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more than 5% for different extracts for any

given enzyme activity.Enzymes of the Embden-Meyerhof-Parnas path-

way. The activities of some enzymes of the Emb-den-Meyerhof-Parnas pathway in T. thioparusand T. neapolitanus are shown in Table 1. Bothglucokinase and fructokinase were demonstrated.Glucokinase activity was considerably higher inT. neapolitanus than in T. thioparus. Fructokinaseactivity, however, was higher in T. thioparus thanin T. neapolitanus. No mannokinase activitycould be demonstrated in extracts of either or-ganism.The level of phosphoglucomutase in T. neapoli-

tanus was double that in T. thioparus, but bothorganisms had similar levels of glucosephosphateisomerase and mannosephosphate isomerase. Nophosphofructokinase could be demonstrated ineither organism. Aldolase was present in bothorganisms. The specific activities of glyceralde-hydephosphate dehydrogenase and phospho-glycerate kinase were very high in both organisms,exceeding 1,500 in T. thioparus and 800 in T.neapolitanus. The level of pyruvate kinase inT. neapolitanus was almost double that in T.thioparus.Enzymes of the citric acid cycle. The specific

activities of some enzymes of the citric acid cyclein T. thioparus and T. neapolitanus are shown inTable 2. The specific activity of citrate synthase(condensing enzyme) in T. thioparus was aboutfour times as high as in T. neapolitanus, and theaconitase activity of T. thioparus was also some-

TABLE 1. Relative activities of enzymes of theEmbden-Meyerhof-Parnas pathway in the obligateautotrophs T. thioparus and T. neapolitanus"

Specific activity

EnzymeT. T.

thioparus eapolitanus

Glucokinase............... 4.1 58.1

Fructokinase.. 9.8 3.3Mannokinase .............. NAb NAbPhosphoglucomutase ....... 13.4 28.7Glucosephosphate isomer-

ase.................... 22.1 22.4

Mannosephosphate isom-erase .................... 17.4 21.8

Phosphofructokinase....... NAb NAbAldolase................... 25.4 19.3

Glyceraldehydephosphatedehydrogenase.......... . 1,768 896

Phosphoglycerate kinase.. 1,515 835

Pyruvate kinase............ 30.0 52.2

a See text for enzyme assays and expression ofactivities.

b No measurable activity.

TABLE 2. Relative activities ofenzymes of the citricacid cycle in the obligate autotrophs T. thioparus

and T. neapolitanusf

Specific activityEnzyme Substrate orEnzyme ~~cofactor T. T. nea-

thioparus politanus

Citrate synthase Acetyl-CoA, 40.7 10.9oxalacetate

Aconitase Citrate 26.0 NMbAconitase Isocitrate 12.0 NMIsocitrate dehy- NADP 204.4 11.5drogenase

Isocitrate dehy- NAD NAC 203. 1drogenase

Isocitrate dehy- NADP, cis- 32.0 13.0drogenase aconitate

Isocitrate dehy- NADP, citrate 17.0 1.7drogenase

Fumarase Fumarate 79.6 63.7Fumarase Malate 39.9 42.3Malate dehydro- Malate, NAD 136.4 14.5

genaseMalate dehydro- Oxalacetate, 223.5 NMgenase NADH

Malic enzyme NADP 26.8 24.0

a See text for enzyme assays and expression ofactivities.

b Not measured.c No measurable activity.

what greater than that of T. neapolitanus. Anextremely active isocitrate dehydrogenase, spe-cific for NADP, was demonstrated in T. thio-parus. No evidence for an NAD-specific dehydro-genase could be found. In contrast, NADP-linked isocitrate dehydrogenase activity in T.neapolitanus was very low, NAD being by far themost effective acceptor for the reaction in thisspecies. Similar amounts of fumarase weredemonstrated in extracts of T. thioparus andT. neapolitanus. A high level of malate dehydro-genase activity was present in extracts of T.thioparus; the level was much lower in T. neapoli-tanus. Malic enzyme had approximately the samespecific activity in both organisms.Enzymes of the pentose phosphate cycle. As

shown in Table 3, glucose-6-phosphate dehydro-genase was present at a much lower level in T.thioparus than in T. neapolitanus. Glucose-6-phos-phate dehydrogenase activity of T. thioparus wasparticulate, being localized in 144pl20; that ofT. neapolitanus was distributed between particu-late (144p120) and soluble (144,120) fractions.The 6-phosphogluconate dehydrogenase activityof T. thioparus occurred in the soluble (144,120)fraction. Although most of the 6-phosphoglu-conate dehydrogenase of T. neapolitanus was in

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TABLE 3. Relative activities of enzymes of the pentose phosphate cycle (hexose monophosphateshunt) in the obligate autotrophs T. thioparus and T. neapolitanusa

Enzyme

Specific activity

T. thioparus T. neapolitanus

Crude extract 144p120O Crude extract 144,120 144pl20

Glucose-6-phosphatedehydrogenase .. 1.8 4.1 18.2 9.7 9.3

6-Phosphogluconatedehydrogenase .... 3.2 9.3 - 13.0 17.9 2.5

Ribosephosphateisomerase.......... 4,570 - - 5,050 -

a See text for enzyme assays and expression of activities.bThe supernatant fraction after centrifugation of the crude extract at 144,000 X g for 120 min.c The particulate fraction after centrifugation of the crude extract at 144,000 X g for 120 min.

the soluble fraction, a small amount of activitywas detected in the particulate fraction. Ribose-phosphate isomerase activity was present at highand similar levels in extracts of both organisms.

Miscellaneous enzymes. As shown in Table 4,the specific activity of NADH dehydrogenase inT. thioparus was almost four times as high as

that in T. neapolitanus. The extracts of bothspecies oxidized NADPH at much lower rates.No attempt was made to determine whether thisactivity reflected the presence of an NADPHoxidase or an NADPH-NAD transhydrogenase.

DISCUSSIONAlthough it seems probable a priori that the

central pathways of intermediary metabolism inautotrophic organisms are for the most partsimilar to those in heterotrophic organisms, verylittle direct evidence bearing on this question isavailable. A number of possible explanations ofthe biochemical basis of autotrophy still remainopen (18, 21, 35). One possible explanation forthe failure of obligate autotrophs to grow at theexpense of organic compounds is that they lackone or more of the essential enzymes required forthe metabolism of such compounds. Some evi-dence in favor of this explanation for certainphotoautotrophs has been presented (33, 35).Detailed enzymatic analyses of obligate auto-trophs are required before any firm conclusionsabout the relationship between enzyme deficien-cies and obligate autotrophy can be reached.For the obligate sulfur-oxidizing autotrophs T.thioparus and T. neapolitanus, only two reportson the presence of certain enzymes of the citricacid cycle have appeared (10, 35).The data presented here show both quantitative

and qualitative differences with respect to theenzymes of the glycolytic pathway, the citric acid

cycle, and the pentose phosphate cycle betweenT. thioparus and T. neapolitanus. One notableobservation was that neither organism containedmeasurable phosphofructokinase. Phosphofruc-tokinase also has not been detected in Nitrocystisoceanus (39). Since most phosphofructokinasesappear to be allosteric proteins [inhibited by ci-trate and high concentrations of ATP, and acti-vated by adenosine monophosphate (AMP)], it isdifficult to be certain that optimal conditions forthe assay of any given phosphofructokinase havebeen employed. However, no phosphofructo-kinase activity could be demonstrated in crudeextracts of T. neapolitanus with ATP concentra-tions ranging from 0.5 to 5 mM. It is thereforepossible that the absence (or, at best, the presenceof extremely low levels) of this enzyme is charac-teristic of this organism. If so, there is not anoperational glycolytic pathway in T. neapolitanus.The glyceraldehyde phosphate dehydrogenase

and phosphoglycerate kinase activities of T.thioparus and T. neapolitanus were over 100 times

TABLE 4. Relative activities of miscellaneousenzymes in the obligate autotrophs T. thioparus

and T. neapolitanusa

Specific activity

EnzymeT. T.

thipoarus neapolitanus

Reduced NAD dehydro-genaseb.................. 40.4 11.2

Reduced NADP dehydro-genase....... 1.8 2.8

Glycerol kinase......... 19.9 3.1

a See text for enzyme assays and expression ofactivities.

b Measured on crude extract only.

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those reported for T. ferrooxidans (13), and theglyceraldehyde phosphate dehydrogenase activitywas 50 to 100 times that reported for N. oceanus(39). Both of these enzymes are essential catalystsof the carbon reduction cycle, and it is difficult tounderstand how N. oceanus can grow autotroph-ically at the rates which have been measured, ifits glyceraldehyde phosphate dehydrogenase ac-tivity is really as low as reported (39). Conceiv-ably, this enzyme may be NADP-dependent inN. oceanus; its activity was measured with NAD(39).Although D-3-phosphoglyceric acid mutase and

enolase were not assayed, there is indirect evi-dence from other work (26) that both enzymesare present in T. thioparus and T. neapolitanus.Chromatography and autoradiography of prod-ucts produced by C01-fixing systems from T.neapolitanus (26) and T, thioparus (E. J. Johnson,unpublished observations) revealed that, although3-phosphoglyceric acid is the predominant prod-uct, measurable amounts of pbosphoenolpyruvicacid are also produced. Both phosphoglyceratephosphomutase and enolase are required for theconversion of 3-phosphoglycerate to phospho-enolpyruvate. In addiion, there is evidence thatorganisms with a complete carbon reductioncycle, such as T. thioparus and T. neapolitanus,possess a triose phosphate isomerase (12). Indi-rect evidence indicates that enzymes involved inthe metabolism of pyruvate, such as pyruvateoxidase, are present in both organisms: cell sus-pensions can oxidize pyruvate and incorporatepyruvate carbon into trichloroacetic acid-solublecompounds, nucleic acids, lipids, and proteins(20). Accordingly, all enzymes of the,glycolyticcycle except phosphofructokinase appear to bepresent in both of these organisms. Unless thephosphofructokinase of these organisms hasunusual properties which prevent'its demonstra-tion by standard assay methods, it must be con-cluded that the lack of this enzyme makes im-possible a utilization of hexoses by the glycolyticpathway. However, hexosephosphate might bemetabolized by a pathway which by-passes thisapparent lesion: for example, fructose-6-phos-phate could be converted to triose phosphatethrough the reactions of the carbon reductioncycle.Cooper (10) demonstated isocitrate dehydro-

genase, aconitase, and malate dehydrogenase inextracts prepared from T. thioparus. He could notdetect isocitratase, and did not obtain conclusiveevidence for the presence of fumarase and citratesynthase. We have been able to demonstratefumarase, malate dehydrogenase, and malic en-zyme in both T. thioparus and T. neapolitanus.Suggestive evidence for the presence of succinate

dehydrogenase was also presented by Cooper(10), and the fact that a-ketoglutarate was pro-duced from isocitrate suggested the presence ofoxalosuccinate decarboxylase as well as isocitricdehydrogenase. In addition, the presence of iso-citrate dehydrogenase, succinate dehydrogenase,and malate dehydrogenase has been shown inT. thiooxidans and in a different strain of T.thioparus; however, reduced NAD dehydrogenase(NADH oxidase) and a-ketoglutarate dehydro-genase could not be demonstrated in these or-ganisms (35). We have shown the presence ofcitrate synthase and aconitase in both T. thioparusand T. neapolitanus. It has been suggested that adeficiency of aconitase may provide the biochem-ical basis for photoautotrophy in the genusChlorobium, as this enzyme could not be demon-strated in the anaerobic obligate photoautotrophC. thiosulfatophilum (33). However, aconitase isclearly present in the obligate chemoautotrophs.

In agreement with Cooper (10), we found thatthe isocitrate dehydrogenase of T. thioparus isstrictly NADP-dependent. However, this enzymein T. neapolitanus is predominantly NAD-depend-ent. The specific activities of the isocitrate dehy-drogenases from both organisms are considerablyhigher than those reported for any of the auto-trophs studied by Smith et al. (35).

It appears, therefore, that all the enzymes ofthe citric acid cycle with the exception of a-keto-glutarate dehydrogenase, which was not measuredeither by Cooper (10) or by us, have been demon-strated in T. thioparus. a-Ketoglutarate dehydro-genase could not be demonstrated in extracts ofthe two strict autotrophic thiobacilli studied bySmith et al. (35).The ribosephosphate isomerase activity was

very high and was about the same in both or-ganisms. Since transketolase, transaldolase, andphosphoketoepimerase are known to participatein the carbon reduction cycle (12), which theseorganisms possess, it appears that a completepentose phosphate cycle can operate in bothT. thioparus and T. neapolitanus. It is clear thatall enzymes which participate in the carbon reduc-tion cycle are present at very high levels. In con-trast, those enzymes which operate exclusively incycles other than carbon reduction are present atmuch lower levels.

Other enzymes operative in the metabolism ofheterotrophic organisms have also been demon-strated in extracts of T. thioparus and T. neapoli-tanus. There were quantitative differences betweenthe two organisms in several instances, e.g., therewas almost four times .as much reduced NADdehydrogenase and over six times as much glyc-erol kinase in T. thioparus as in T. neapolitanus.

It is clear that there are differences, both quali-

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tative and quantitative, between these two or-ganisms in enzymatic constitution. This is un-doubtedly reflected in the differences in theirabilities to assimilate and incorporate organiccompounds into cellular components (20). Fur-ther, these organisms differ in other respects.Jackson et al. (19) recently reported values for themean deoxyribonucleic acid base composition(moles per cent guanine plus cytosine) of 62(strain NCIB 8370) and 66 (strain NCIB 8349)for T. thioparus, and a value of 56 for T. neapoli-tanus. Although these two organisms are generallyconsidered closely related, the available informa-tion accordingly suggests that this may not betrue.Some of our findings have a bearing on the basis

of obligate autotrophy. Smith et al. (35) presentedevidence that the lack of reduced NAD dehydro-genase provides the biochemical basis for obligateautotrophy in the strains of T. thioparus andT. thiooxidans examined by them. They furtherstated that obligate autotrophs probably evolvedfrom heterotrophic ancestors by elimination ofthis enzyme, which plays a key role in hetero-trophic metabolism. Their conclusions do not,however, hold for all obligate chemosyntheticautotrophic bacteria, or even for all members ofthe genus Thiobacillus. The evidence presentedhere shows that one strain of T. thioparus (ATCC8158) contains a fairly active NADH dehydro-genase. Furthermore, the specific activity of theNADH dehydrogenase of T. neapolitanus re-ported in this paper is similar to that reported forfacultative heterotrophs by Smith et al. (35),whereas the specific activity of this enzyme inT. thioparus is even higher. Hempfling and Vish-niac (15) also found NADH dehydrogenase inT. neapolitanus, and could demonstrate oxygenuptake with NADH as election donor. Thesefindings have recently been confirmed by Tru-dinger and Kelly (38). There is no doubt, there-fore, that this obligate chemoautotroph canmediate a transfer of electrons from NADH tooxygen. Thus, if the obligate autotrophs studiedin this present investigation fail to couple thebreakdown of organic substrates with the genera-tion of ATP, it is not because they lack NADHoxidase, but presumably because this enzymesystem is an uncoupled one.

If coupling mechanisms are absent, there wouldbe only one conceivable mechanism for theobligate autotrophs to produce ATP from the me-tabolism of organic substrates: substrate phos-phorylation during glycolysis. However, if phos-phofructokinase is absent, as our data suggest,glycolysis may not be possible. As a consequenceof the inability to couple phosphorylation to theoxidation of NADH, together with a block in the

glycolytic pathway prior to the triose phosphatestage (which would eliminate the possibility of asubstrate level phosphorylation, at least in thecase of certain substrates), the production ofATP from the metabolism of certain organic com-pounds would be prevented. These defects mayprovide a possible biochemical basis for obligatechemoautotrophy in these organisms. Alterna-tively, very specific control mechanisms involvingATP, ADP, AMP, and reduced and oxidizedpyridine nucleotides may explain their obligatechemoautotrophy. In any case, as suggested in anearlier report (20), all autotrophs may not havethe same biochemical basis for that autotrophy.It should be noted that some organisms previouslyconsidered to be obligate autotrophs have nowbeen grown heterotrophically (32, 34). The factremains, however, that some bacteria still behaveas obligate autotrophs despite all efforts to growthem as heterotrophs.

ACKNOWLEDGMENTS

The excellent technical assistance of Rosalie French is gratefullyacknowledged. We thank Edith Engleson and Mona Ingelstromfor their fine technical assistance with various analyses.

This investigation was supported by Contract NAS-2-3901with the National Aeronautics and Space Administration, AmesResearch Center, Moffett Field, Calif., and by Public HealthService grant FR-5467 from the National Institutes of Health.

LITERATURE CITED

1. Abraham, S., B. Borrebaek, and I. L. Chaikoff. 1964. Effectof dietary carbohydrate on glucokinase and mannokinaseactivities of various rat tissues. J. Nutr. 83:273-288.

2. Abraham, S., W. M. Fitch, and I. L. Chaikoff. 1961. Mannosemetabolism and the demonstration of mannokinase andphosphomannoisomerase activities in the lactating ratmammary gland. Arch. Biochem. Biophys. 93:278-282.

3. Abraham, S., L. Kopelovich, P. R. Kerkof, and I. L. Chaikoff.1965. Metabolic characteristics of preparations of isolatedsheep thyroid cells. I. Activity levels of enzymes concernedwith glycolysis and the tricarboxylic acid cycle. Endocri-nology 76:178-190.

4. Axelrod, B. 1955. Pentose phosphate isomerase, p. 363-366.In S. P. Colowick and N. 0. Kaplan (ed.), Methods inenzymology, vol. 1. Academic Press Inc., New York.

5. Blumenthal, M. D., S. Abraham, and I. L. Chaikoff. 1964.Dietary control of liver glucokinase activity in the normalrat. Arch. Biochem. Biophys. 104:211-224.

6. Borrebaek, B., S. Abraham, and I. L. Chaikoff. 1964. En-zymatic removal of glucose-6-phosphate from fructose-6-phosphate preparations. Anal. Biochem. 8:367-372.

7. Bublitz, C., and 0. Wieland. 1962. Glycerokinase, p. 354-361.In S. P. Colowick and N. 0. Kaplan (ed.), Methods inenzymology, vol. 5. Academic Press Inc., New York.

8. BUicher, T. 1955. Phosphoglycerate kinase from brewer's yeast,p. 415-422. In S. P. Colowick and N. 0. Kaplan (ed.),Methods in enzymology, vol. 1. Academic Press Inc., NewYork.

9. BUcher, T., and G. Pfleiderer. 1955. Pyruvate kinase frommuscle, p. 435-440. In S. P. Colowick and N. 0. Kaplan(ed.), Methods in enzymology, vol. 1. Academic Press Inc.,New York.

10. Cooper, R. C. 1964. Evidence for the presence of certaintricarboxylic acid cycle enzymes in Thiobacillus thioparus.J. Bacteriol. 88:624-629.

11. Dipietro, D. L., and S. Weinhouse. 1960. Hepatic glucokinase

VOL. 100, 1969 967

on February 14, 2020 by guest

http://jb.asm.org/

Dow

nloaded from

JOHNSON AND ABRAHAM

in the fed, fasted and alloxan-diabetic rat. J. Biol. Chem.235:2542-2545.

12. Elsden, S. R. 1962. Photosynthesi and lithotrophic carbondioxide fixation, p. 1-40. In I. C. Gunsalus and R. Y.Stanier (ed.), The bacteria, vol. 3, Biosynthesi. AcademicPress Inc., New York.

13. Gale, N. L., and J. V. Beck. 1967. Evidence for the Calvincycle and hexose monophosphate pathway in T7hobacillusferrooxidans. J. Bacteriol. 94:1052-1059.

14. Gornall, A. C., C. J. Bardawill, and M. M. David. 1949.Determination of senm proteins by mean of the biuretreaction. J. Biol. Chem. 177-.751-766.

15. Hempfling, W. P., and W. Vihniac. 1965. Oxidative phos-phorylation in extracts of Thiobacillus X. Biochem. Z. 342:272-287.

16. Horecker, B. L., and A. Komberg. 1948. The extinction coeffi-cients of the reduced band of pyridine nucleotides. J. Biol.Chem. 175:385-390.

17. Horecker, B. L., and P. Z. Smyrniotis. 1955. 6-Phosphoglu-conic dehydrogenase, p. 323-327. In S. P. Colowick and N.0. Kaplan (ed.), Methods in ezymrology, voL 1. AcademicPress Inc., New York.

18. Ida, S., and M. Alexander. 1965. Permeability of Nitrobacteragilis to organic compounds. J. Bacteriol. 90:151-156.

19. Jackson, J. F., D. J. M. Moriarty, and D. J. D. Nicholas.1968. Deoxyribonucleic acid base composition and tax-onomy of thiobacil1i and nitrifying bacteria. J. Gen. Micro-biol. 53:53-60.

20. Johnson, E. J., and S. Abraham. 1969. Assimilation andmetabolism of exogenous organic compounds by the strictautotrophs Thiobacilus thloparus and Thlobacilus neapoli-tanus. J. Bacteriol. 97:1198-1208.

21. Kelly, D. P. 1967. Problems of the autotrophic microorga-nisms. Sci. Progr. 55:35-51.

22. Kornberg, A., and B. L. Horecker. 1955. Glucose--phosphatedehydrogenase, p. 323-327. In S. P. Colowick and N. 0.

Kaplan (ed.), Methods in enzymology, vol. 1. AcademicPress Inc., New York.

23. Krebs, E. G. 1955. Glyceraldehyde-3-phosphate dehydro-genase from yeast, p. 407-411. In S. P. Colowick and N. 0.

Kaplan (ed.), Methods in enzymology, vol. 1. AcademicPress Inc., New York.

24. Lowry, 0. H., N. J. Rosebrough, A. L. Farr, and R. J. Ran-dall. 1961. Protein measurement with the Folin phenolreagent. J. Biol. Chem. 193:265-275.

25. Massey, V. 1955. Fumamse, p. 729-735. In S. P. Colowick andN. 0. Kaplan (ed.), Methods in enzymology, vol. 1.Academic Press Inc., New York.

26. Mayeux, J. V., and E. J. Johnson. 1967. Effect of adenosinemonophosphate, adenosine diphosphate, and reduced nico-tinamide adenine dinucleotide on adenosine triphosphate-dependent carbon dioxide fixation in the autotroph Thio-bacillus neapolitanus. J. Bacteriol. 94:409-414.

27. Ochoa, S. 1955. Isocitric dehydrogenase system (TPN) frompig heart, p. 699-704. In S. P. Colowick and N. 0. Kaplan(ed.), Methods in enzymology, vol. 1. Academic Press Inc.,New York.

28. Ochoa, S. 1955. Malic dehydrogenase from pig heart, p. 735-739. In S. P. Colowick and N. 0. Kaplan (ed.), Methods inenzymology, vol. 1. Academic Press Inc., New York.

29. Ochoa, S. 1955. "Malic" enzyme, p. 739-753. In S. P. Colo-wick and N. 0. Kaplan (ed.), Methods in enzymology, vol.1. Academic Press Inc., New York.

30. Racker, E. 1950. Spectrophotometric measurements of theenzymatic formation of fumaric and cis-aconitic acids.Biochim. Biophys. Acta 4:211-214.

31. Rauen, H. M. 1956. Biochemisches Taschenbuch, p. 988.

Springer Verlag, Berlin.32. Shafia, F., and R. F. Wilkinson, Jr. 1969. Growth of Ferro-

bacillus ferrooxidans on organic matter. J. Bacteriol. 97:256-260.

33. Smillie, R. M., and W. R. Evans. 1963. Biochemical basis forthe obligate photoautotrophy of green bacteria of the genus

Chloroblum, p. 151-159. In H. Gest, A. San Pietro, and L. P.Vernon (ed.), Bacterial photosynthesis. The Antioch Press,Yellow Springs, Ohio.

34. Smith, A. J., and D. S. Hoare. 1968. Acetate assimilation byNitrobacter agilis in relation to its "obligate autotrophy."J. Bacteriol. 95:844-855.

35. Smith, A. J., J. London, and R. Y. Stanier. 1967. Biochemicalbasis of obligate autotrophy in blue-green algae and thio-bacilli. J. Bacteriol. 94:972-983.

36. Srere, P. A., H. Brazil, and L. Gonen. 1963. The citrate con-

densing enzyme of pigeon breast muscle and moth flightmuscle. Acta Chem. Scand. 17:S129-S134.

37. Stadtman, E. R. 1957. Preparation and assay of acyl coenzymeA and other thiol esters; use of hydroxylamine, p. 931-941.In S. P. Colowick and N. 0. Kaplan (ed.), Methods inenzymology, vol. 3. Academic Press Inc., New York.

38. Trudinger, P. A., and D. P. Kelly. 1968. Reduced nicotinamideadenine dinucleotide oxidation by Thiobacillus neapolitanusand Thiobacillus strain C. J. Bacteriol. 95:1962-1963.

39. Williams, P. J. leB., and S. W. Watson. 1968. Autotrophy inNitrosocystis oceanus. J. Bacteriol. 96:1640-1648.

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