Vol. 29, No. 2INFECTION AND IMMUNITY, Aug. 1980, p. 551-5600019-9567/80/08-0551/10$02.00/0

Energy Metabolism in Capnocytophaga ochraceaROBERT CALMES,* G. W. RAMBICURE, W. GORMAN, AND THOMAS T. LILLICH

Department of Oral Biology, College of Dentistry, Albert B. Chandler Medical Center, University ofKentucky, Lexington, Kentucky 40536

Among the microflora of the gingival sulcus are members of the genus Capno-cytophaga which have been implicated as possible etiological agents of juvenileperiodontitis and systemic infectious diseases. In this study, the pathway used byC. ochracea strain 25 for generating energy from glucose was investigated. Whengrown in a complex medium supplemented with glucose and NaHCO3, the majorend products formed were acetate (4.6 mmol), succinate (11.0 mmol), pyruvate(4.3 mmol), and oxalacetate (3.6 mmol), and the molar growth yield was 58.Addition of yeast extract to the growth medium caused (i) an increase in acetate(9.2 mmol) and succinate (14.3 mmol), (ii) a decrease in pyruvate (O mmol) andoxalacetate (1.1 mmol), and (iii) the molar growth yield increased to 75. Glucosewas transported by a phosphoenolpyruvate:phosphotransferase system and thencatabolized to phosphoenolpyruvate by enzymes ofthe Embden-Meyerhof-Parnaspathway. No activities were detected for the key enzymes of the Warburg-Dickens, Entner-Douderoff, or hexose phosphoketolase pathways. During growthin the yeast extract-supplemented medium, approximately 37% of the phospho-enolpyruvate carbon was converted to acetate by pyruvate kinase, a pyruvate-decarboxylating enzyme activity, and acetate kinase; the remaining 63% wasconverted to succinate via phosphoenolpyruvate carboxykinase, malate dehydro-genase, fumarate hydratase, and fumarate reductase.

Capnocytophaga is a newly described genusof gliding, anaerobic to microaerophilic, C02-de-pendent, gram-negative oral bacteria consistingof three species: C. ochracea, C. sputigena, andC. gingivalis (28, 50, 55). These capnophilicmembers of the oral microflora (10) have beenimplicated as possible etiological agents of ju-venile periodontitis (34, 35; for reviews, see ref-erences 48 and 49). Recent preliminary reportssuggest that Capnocytophaga sepsis occurs inpatients compromised by hematological or othermalignant disease (3, 18; S. W. Forlenza, M. G.Newman, and U. Blachman, J. Dent. Res.58[Special Issue A]: 1027, 1979); in all of thesestudies, the portal of entry was believed to bethe mouth. Shurin et al. (47) have reported thatboth sonic extracts and spent culture mediumfrom C. sputigena strain 4, isolated from theblood of a patient with a peripheral neutrophildisorder, contained a dialyzable substance whichcaused morphological and functional alterationsin normal neutrophils which mimicked those ofneutrophils from infected patients. In addition,a previously unclassified group of bacteria (des-ignated Center for Disease Control biogroup DF-1) and Bacteroides ochraceus, both isolatedfrom a wide range of human clinical specimens,have recently been shown to be synonymouswith species of Capnocytophaga (36, 56).

Until recently, little was known of the physi-ology of these bacteria. Most of the metabolicstudies were for taxonomic purposes to establishCapnocytophaga as a new genus (28, 50). Sincethen, we have reported that membranes of C.ochracea strain 25 contain cytochromes and pri-mary dehydrogenases that oxidize reduced nic-otinamide adenine dinucleotide (NADH) andsuccinate (29). Kapke et al. (23) have shownphosphoenolpyruvate (PEP) carboxykinase tobe the sole C02-fixing enzyme in C. ochracea,thus establishing the enzymatic basis for theC02-dependent fermentation exhibited by Cap-nocytophaga (28).Because of their possible role as etiological

agents of infectious diseases (3, 18, 34, 35, 47-49;Forlenza et al., J. Dent. Res. 58[Special IssueA]:1027, 1979) and the demonstration of theirperiodontopathic potential as monoinfectants ingnotobiotic rats (21), a careful study was begunof the physiological systems used by Capnocy-tophaga to generate adenosine 5'-triphosphate(ATP) for growth and reproduction. In this com-munication, we describe the pathway for glucosecatabolism and ATP production used by C.ochracea strain 25.

(Part of this work was presented at the AnnualMeeting of the International Association ofDen-tal Research, 5 to 7 June 1980, Osaka, Japan.)

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MATERIALS AND METHODS

Bacterium, medium, and culture conditions.The bacterium used in this study was C. ochraceastrain 25, the neotype strain for this species (28) whichwas generously provided by S. S. Socransky of ForsythDental Center, Boston, Mass. Members of all threespecies of Capnocytophaga have been shown to havean obligate CO2 requirement for growth (28, 50). Inthe presence of adequate CO2 concentrations, thesebacteria will grow either anaerobically or in the pres-ence of small amounts of oxygen (28, 50). Because ofthe relative indifference of Capnocytophaga sp. tomicroaerophilic conditions and the inconvenience ofusing artificial atmospheres over culture media, a sim-plified method of growing C. ochracea was soughtwhich would yield good crops of cells for enzymologicaland physiological studies. First, because C. ochraceawas somewhat aerotolerant, no special precautionswere taken to ensure anaerobiosis other than to usefreshly prepared medium in tightly capped vesselswith a minimum amount of headspace. Second, theglucose-Trypticase soy broth (BBL Microbiology Sys-tems) in which these bacteria have been routinelygrown (28, 29, 50) could not be used because Trypti-case contained amounts of acetic acid (9; data notshown) which interfered with fermentation productanalyses. To eliminate this problem, the Trypticasecomponent of the medium was replaced by Tryptone(Difco), which resulted in an essentially acetate-freemedium. Third, NaHCO:) was included in the mediumin lieu of an atmosphere containing high levels of CO2as used in previous studies (28, 29, 50). The newmedium contained the following ingredients: 1.7% (wt/vol) Tryptone (Difco), 0.3% (wt/vol) Phytone (BBLMicrobiology Systems), 0.25% (wt/vol) dibasic potas-sium phosphate (pH 7), 0.25% (wt/vol) D-glucose, 0.5%(wt/vol) yeast extract (Difco), and 20 mM NaHCO:;.Unless otherwise specified, this medium was used forall experiments. Routinely, the medium withoutNaHCO:; was prepared freshly for each experimentand sterilized by steam autoclave (121°C, 15 min).After cooling to 39°C, NaHCO:, was added asepticallyfrom a sterile 1 M solution to a final concentration of20 mM. One liter of the complete medium, containedin a 1-liter screw-capped Erlenmeyer flask fitted witha side arm for measurement of growth, was inoculated(30 ml/liter) with a 24-h culture of C. ochracea. Aftercapping, it was incubated statistically at 39°C. Use ofthis medium resulted in the elimination of the lagphase, maximum rates of growth, and large cell crops.Stock cultures of C. ochracea were maintained at 4°Cas stab cultures in 1.5% (wt/vol) agar deeps of themedium described above but without glucose.Growth measurement. Growth of C. ochracea

strain 25 was monitored turbidimetrically at 750 nmvia a sidearm on the culture vessel. Uninoculatedmedium was used as the blank.Fermentation product analysis. Culture super-

natant fluids were quantitatively analyzed for volatileand nonvolatile end products with a Varian 3700 gaschromatograph equipped with a model CDS 111cchromatography data system. Samples obtained atmaximum growth (usually 12 h) were prepared forgas-liquid chromatography by the methods of Holde-

man et al. (20). The following chromatograph param-eters were used to separate and detect the volatile andnonvolatile endproducts. (i) For the thermal conduc-tivity detector (this detector was used in preliminaryexperiments to verify that formate was not producedby C. ochracea) carrier gas (He) flow rate was 40 ml/min, the injector temperature was 200°C, and thedetector temperature was 220°C. The column ovenwas operated isothermally at 115°C for nonvolatilefermentation products; for analysis of volatile fattyacids the following temperature program was used:column oven temperature was held initially at 130°Cfor 6 min followed by a programmed increase in tem-perature at a rate of 10°C/min to 150°C, where thetemperature was held for 10 min. (ii) For the flameionization detector, the carrier gas (N2) flow rate was40 ml/min, the H2 and compressed air flow rates were30 and 300 ml/min, respectively; the injector temper-ature was 200°C; the detector temperature was 220°C;the column oven temperature was isothermal at 150and 130°C, respectively, for volatile and nonvolatileendproducts. The stainless steel columns (183 by 2mm inner diameter; premium grade, Supelco, Belle-fonte, Pa.) were packed with 10% SP-1000/1% phos-phoric acid on 100/120 chromosorb W AW andplugged with phosphoric acid-treated glass wool. Ex-tracts (0.5 to 1.0 IL) were injected, and each componentwas quantified by the integrator of the attendant CDS-111c chromatography data system and compared withknown amounts of the appropriate standard.Harvesting of cells. Cells in the late exponential

phase of growth were harvested by centrifugation andwashed twice with equal volumes of 15 mM KCI.Washed cells were suspended in 50 mM potassiumphosphate buffer (pH 7) or in 50 mM tris(hydroxy-methyl)aminomethane (Tris)-hydrochloride buffer(pH 7.5), depending upon experimental use.Preparation of decryptified cell suspensions.

For transport assays, the membranes of washed cellswere perturbed as previously described (8) with tolu-ene-acetone (1:4, vol/vol).

Preparation of cell-free extracts. Washed cellsfrom a 2-liter culture were disrupted by sonication aspreviously described (29). Cellular debris was removedby centrifugation at 40,000 x g for 30 min at 4°C. Theresulting supernatant fluid was designated as the crudecell-free extract.

PEP:glucose PT assay. The method of Schach-tele (45) was modified and used to measure the for-mation of ['4C]2-deoxyglucose (DG)-phosphate from['4C]DG. Use ofDG as a substrate for the PEP:glucosephosphotransferase (PT) system was based on theinability of bacterial hexokinases to catalyze the ATP-dependent phosphorylation of this analog (13,41). Thereaction mixture contained in a total volume of 1 ml:["4C]DG, 1 mM (1 ,uCi/,umol); PEP, 2 mM; MgCl2, 5mM; NaF, 10 mM; and potassium phosphate buffer(pH 7), 50 mM. The reaction was started by addingdecryptified cells (5 mg of bacterial protein) and in-cubating at 37°C. After 20 min, triplicate 50-,ul portionsof the reaction mixture were transferred directly to2.5-cm disks of Whatman DE-81 diethylaminoethyl(DEAE)-cellulose paper which were immediatelyplaced in deionized, distilled water (at least 20 ml/disk) to elute unphosphorylated ['4CIDG. After 15

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min, the water was decanted, replaced with an equalvolume, and allowed to stand an additional 15 min.The DEAE-cellulose disks, to which ['4C]DG-phos-phate was bound, were removed, dried under a heatlamp, transferred to minicounting vials containing 3.5ml of Aquasol, and counted in a Packard Tri-Carbliquid scintillation spectrometer. Under these condi-tions, the rate of ['4C]DG-phosphate synthesis waslinear with respect to time and cell concentration.Data were expressed as nanomoles of ['4C]DG-phos-phate formed per minute per milligram of proteincorrected for spurious retention of nonphosphorylated['4C]DG to the DEAE-cellulose disks.

Alkaline phosphatase treatment. To determinewhether phosphorylation of ['4C]DG was the basis forits retention to DEAE-cellulose, the PT system assaywas modified slightly. The assay was performed in 50mM Tris-hydrochloride buffer (pH 7.0). After the 20-min reaction period, the reaction was stopped by heat-ing at 100'C for 5 min. The pH was quickly adjustedto 8.2 with 0.1 N KOH, 1 mg of highly purified alkalinephosphatase was added; and the reaction mixture wasincubated for 30 min at 370C. After readjustment ofthe reaction mixture to pH 7 with 0.1 N HCI, thetransfer, washing, and counting of samples were asdescribed above for the transport assay.Enzyme assays. The following enzymes were as-

sayed spectrophotometrically in 50 mM Tris-hydro-chloride buffer (pH 7.5) at saturating substrate con-centrations with crude cell-free extracts. Hexokinase(EC 2.7.1.1) was assayed by the method of Pilkis (38).Phosphofructokinase (EC 2.7.1.11) activity was meas-ured as described by Kemerer et al. (24). Fructosediphosphate aldolase (EC 4.1.2.18) activity was deter-mined by the method of Lebherz and Rutter (27). Theassay for enolase (EC 4.2.1.11) was that of Spring andWold (51). Pyruvate kinase (EC 2.7.1.40) was assayedby the method of Tuominen and Bernlohr (54). Lac-tate dehydrogenase (EC 1.1.1.27) was assayed as de-scribed by Wittenberger (58). Glucose 6-phosphatedehydrogenase (EC 1.1.1.49) activity was measuredusing the method of Olive and Levy (37). The assay of6-phosphogluconate dehydrogenase (EC 1.1.1.43) wasby the method of Bridges and Wittenberger (5). Themethod used to assay transaldolase (EC 2.2.1.2) wasthat of Tsolas and Joris (53). The 2-keto-3-deoxy-6-phosphogluconate aldolase (EC 4.1.2.14) assay wascarried out as described by Hammerstedt et al. (17).Malate dehydrogenase (EC 1.1.1.37) was assayed asdescribed by Murphey and Kitto (33). Fumarate hy-dratase (EC 4.2.1.2) activity was measured in themanner of Hill and Bradshaw (19). Fumarate reduc-tase was assayed with the method of Kroger et al. (26).Pyruvate decarboxylating activity was assayed bymonitoring the release of ['4C]C02 from [1_-4C]pyru-vate by crude cell-free extracts as described by Kresze(25). Acetate kinase (EC 2.7.2.1) activity was deter-mined by the method of Kahane and Muhlrad (22) incell-free extracts partially purified as follows: ice-cold10% (wt/vol) streptomycin sulfate was added drop-wise, with stirring, to the crude cell-free extract to afinal concentration of 1% (wt/vol). After standing inice for 30 min, the solution was centrifuged at 20,000x g for 20 min, and the pellet was discarded. Finelyground crystalline ammonium sulfate was added with

stirring to the supernatant fluid obtained above until40% (wt/vol) saturation was obtained; after standingin ice for 30 min, the precipitate was removed bycentrifugation at 20,000 x g for 20 min and discarded.Additional ammonium sulfate was added with stirringuntil 70% (wt/vol) saturation was obtained, then theprecipitate was sedimented as above, and the super-natant fluid was discarded. The precipitate was dis-solved in 10 ml of 50 mM Tris-hydrochloride (pH 7.5)containing 1 mM dithiothreitol and dialyzed for 16 hagainst 4 liters of the same buffer. The resulting prep-aration, the 40 to 70% ammonium sulfate precipitate,is referred to as the partially purified acetate kinase.The specific activities of all enzymes are expressed

as nanomoles per minute per milligram of proteinunless otherwise specified.

Protein estimations. The method of Lowry et al.(30) was used to estimate the protein content of de-cryptified cells and cell-free extracts with egg whitelysozyme as the standard.

Chemicals. All biochemicals, substrates, and en-zymes were purchased from Sigma Chemical Co.Tryptone and yeast extract were obtained from Difco,and Phytone was purchased from BBL MicrobiologySystems. All radiochemicals and Aquasol were ob-tained from New England Nuclear Corp. All otherchemicals were of analytical grade and were purchasedfrom Fisher Scientific Co., except for "enzyme-grade"ammonium sulfate, which was obtained from Schwarz-Mann.

RESULTS

Growth of C. ochracea. Cells grown in com-plex medium containing glucose, NaHCO3, andyeast extract had a generation time of 2.4 h. Themaximum absorbance at 750 nm was 1.04, whichcorresponded to a cell crop of 1.04 g (dry wt) ofcells per liter of medium. During growth, the pHof the medium decreased from 7.0 to 5.7. Cul-tures grown under strict anaerobic conditionswith an atmosphere of 100% CO2 or with 20mMNaHCO3 had the same growth characteristics asthe aerobic culture supplemented with NaHCO3.Optimum growth was obtained when theNaHCO3 concentration was 20 mM, a value sim-ilar to the [S]o.!, for HCO3 of 10 mM observedby Kapke et al. (23) for PEP carboxykinase. Theaddition of yeast extract to the medium causeda 15% increase in the cell crop; as will be dis-cussed in the next section, the addition of yeastextract also caused shifts in both the kinds andamounts of glucose fermentation products.Analysis of fermentation products and

determination of Yg1.. Qualitative analysesof glucose fermentation products formed at max-imum growth were performed initially to obtainsome insight to the catabolic pathway(s) usedby C. ochracea to generate energy for growth.In addition to acetate and succinate, the majorend products reported by others (28, 50), pyru-vate and oxalacetate, were also formed in mod-

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erate amounts. The presence of these metabo-lites suggested that they might be intermediatesin the glucose catabolic pathways to the majorend products. Experiments designed to obtainmore abundant growth of C. ochracea throughsupplementation of the medium with yeast ex-tract supported this hypothesis. Although thetotal amount of all products formed duringgrowth in either the presence or absence of yeastextract was essentially the same (24.6 and 23.5mmol, respectively), differences in the quantityof individual products were observed (Table 1).A careful quantitative analysis of fermentationproducts revealed that cells which were culturedin the absence of yeast extract formed acetate(4.6 mmol), pyruvate (4.3 mmol), oxalacetate(3.6 mmol), and succinate (11.0 mmol), whereasduring growth in the presence of yeast extract,no pyruvate accumulated and oxalacetate accu-mulation was reduced by two-thirds. These in-termediate products were replaced stoichiomet-rically (within 95%) by acetate and succinate.Inclusion of yeast extract in the growth mediumalso caused the molar growth yield on glucose,i.e., the YgiucOse (4), to increase from 57.9 to 75.3.If these Y,,,,ose values are used in conjunctionwith the mean value for YATP of 10.5 determinedby Bauchop and Elsden (4), the theoretical num-ber of moles ofATP formed during fermentationof 1 mol of glucose can be calculated. In thepresence of yeast extract, 7.2 mol of ATP wasgenerated per mol of glucose, whereas only 5.5mol of ATP was formed when yeast extract wasomitted. These values, together with the Ygiucosevalues, suggest that energy transduction wasmore efficient when C. ochracea was grown withyeast extract supplementation than when cellswere cultured in its absence.PEP:glucose PT system characteristics.

Decryptified cells of C. ochracea strain 25 trans-ported the nonmetabolizable glucose analog 2-DG by using a system having the characteristicsof a PEP:PT system as shown in Table 2. In thepresence of PEP and Mg2", the phosphorylationof ["C]DG was stimulated 22-fold; deletion ofPEP or Mg2" from the reaction mixture causedreductions in ['4C]DG-phosphate synthesis of 95and 80%, respectively. Addition of glucose at 10times the concentration of ['4C]DG present inthe reaction inhibited phosphorylation of['4C]DG by 95%, indicating that both glucoseand its analog competed for the same transportsystem. Of the three phosphate donors tested,only PEP and its immediate metabolic precur-sor, 2-phosphoglycerate, were capable of stimu-lating ['4C]DG-phosphate formation; no stimu-lation was observed with ATP. For 2-phospho-glycerate to serve as the energy source, it was

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TABLE 2. PEP:glucose PT system in C. ochraceastrain 25a

Assay condition Sp acthComplete 44Heated cells' <1Minus PEP .. ................. 2Minus Mg2e . .. .. 9Plus 10 mM glucose .. 2Plus ATPd . 1Plus 2-phosphoglycerate 5Plus 2-phosphoglycerate, minus NaF 31Plus alkaline phosphatase 2

a Decryptified cells were prepared from stationary-phase cells. Experimental conditions were as describedin the text. Each reaction contained 5 mg of bacterialprotein.

b Nanomoles of ['4C]DG-phosphate formed per min-ute per milligram of protein.

'Contains cells heated at 100'C for 10 min.dATP or 2-phosphoglycerate was substituted for

PEP as indicated; final concentration was 1 mM.

necessary to delete NaF from the reaction. Thisobservation suggests that the enolase of C.ochracea, which catalyzes the conversion of 2-phosphoglycerate to PEP, is sensitive to NaF asin other oral bacteria (16). Treatment of thereaction mixture with purified alkaline phospha-tase after the PEP:PT assay, but before transferof aliquots of the reaction to the DEAE-cellulosedisks, essentially eliminated retention of all ra-dioactive material from the disks. This would beexpected if the anionic ['4C]DG-phosphate wascleaved by alkaline phosphatase to form thenon-ionic ['4C]DG plus inorganic phosphate.Evidence for glucose catabolism via the

Embden-Meyerhof-Parnas pathway. Anal-ysis of key enzymes (12) was used for initialdetermination of the catabolic pathway used byC. ochracea strain 25 for dissimilation of glucose(Table 3). No activities were detected for thekey enzymes of the Warburg-Dickens pathway,i.e., glucose-6-phosphate dehydrogenase, 6-phos-phogluconate dehydrogenase, and transaldolase,thereby negating the presence of the Warburg-Dickens pathway. Similarly, the absence of glu-cose 6-phosphate dehydrogenase and 6-phos-phogluconate dehydrogenase negated operationof the glucose phosphoketolase pathway sinceentry into this pathway is via glucose 6-phos-phate or 6-phosphogluconate. The lack of thepreceding two dehydrogenases and 2-keto-3-deoxy-6-phosphogluconate aldolase indicatedthe absence of the Entner-Douderoff pathway.Activities were detected for phosphofructoki-nase and fructose diphosphate aldolase, bothkey enzymes of the Embden-Meyerhof-Parnaspathway, as well as for hexokinase, glucose phos-phate isomerase, enolase, and pyruvate kinase

(Table 4). No lactate dehydrogenase activity wasdetected when measured in either the oxidativeor reductive direction, or in the presence orabsence of fructose-1,6-diphosphate, a metabolicintermediate which has been shown to act as apositive effector of lactate dehydrogenases inother oral bacteria (6, 7). The absence of lactatedehydrogenase was not suprising, since no lac-tate was observed during the analysis of fermen-tation products (Table 1). The results of theenzyme analysis indicated that glucose wasprobably catabolized to pyruvate by the Emb-den-Meyerhof-Parnas pathway in C. ochraceastrain 25.Fate of PEP. The occurrence of a branch

point in the dissimilatory pathway at PEP wasindicated by (i) the fermentation product anal-ysis as well as the existence of a stoichiometricrelationship between pyruvate and acetate andbetween oxalacetate and succinate (Table 1);and (ii) the presence of PEP carboxykinase (23)and succinic dehydrogenase (29). Therefore, asearch was made for possible enzymes in the

TABLE 3. Specific activities of key enzymes of majorglucose catabolic pathways in C. ochracea strain

25"Key enzyme Pathwayh Sp act"

Phosphofructokinase EMP 67Fructose diphosphate aldolase 89Glucose 6-phosphate WD 0dehydrogenase

6-Phosphogluconate 0dehydrogenase

Transaldolase 02-Keto-3-deoxy-6- ED 0

phosphogluconate aldolase

"Cells were grown to stationary phase, cell-freeextracts were prepared, and enzymes were analyzed asdescribed in the text.

h EMP, Embden-Meyerhof-Parnas; WD, Warburg-Dickens; ED, Entner-Douderoff.

Nanomoles per minute per milligram of protein.

TABLE 4. Specific activities of enzymes of theEmbden-Meyerhof-Parnas pathway in C. ochracea

strain 25'

Enzyme Sp act'Hexokinase 523

Glucose phosphate isomerase 93Phosphofructokinase 67Fructose diphosphate aldolase 89Enolase ... .... 78Pyruvate kinase 50Lactate dehydrogenase 0

" Cells were grown to stationary phase, cell-freeextracts were prepared, and enzymes were assayed asdescribed in the text.

h Nanomoles per minute per milligram of protein.

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terminal pathways from PEP to acetate andsuccinate.As shown in Table 4, conversion of PEP to

pyruvate was catalyzed by pyruvate kinase. Py-ruvate was then catabolized to acetate by apyruvate-decarboxylating enzyme and acetatekinase (Table 5). No phosphotransacetylase (EC2.3.1.8) activity was detected with several con-ditions of assay.The pyruvate-decarboxylating enzyme ex-

hibited a dependence upon thiamine pyrophos-phate (TPP); in the absence to TPP, decarbox-ylation of pyruvate decreased by 70%. The activ-ity of the enzyme was enhanced by the inclusionof the sulfhydryl-protecting reagent dithiothrei-tol in the assay. The dependence of the pyru-vate-decarboxylating enzyme upon TPP and di-thiothreitol suggested that the activity meas-ured may be that of the pyruvate dehydrogenasemultienzyme complex (25); however, attemptsto measure this lipoate-dependent enzyme com-plex with the dismutation assay (40) and a two-step variation (39) of the assay were not suc-cessful. Therefore, it is uncertain at presentwhether the pyruvate-decarboxylating activitywas due to the pyruvate dehydrogenase complexor to other TPP-dependent pyruvate-decarbox-ylating enzymes (11, 15, 57).

Acetate kinase was detected in high levels incrude cell-free extracts prepared from C. ochra-cea (data not shown). Because of the presenceof high endogenous NADPH-generating activitywhich interfered with the assay, it was necessaryto fractionate the extract with ammonium sul-fate. The 40 to 70% (wt/vol) ammonium sulfatefraction contained most of the acetate kinaseactivity and had <10% of the protein in the

TABLE 5. Conversion ofpyruvate to acetate by C.ochracea strain 25"

Enzyme Sp act'Pyruvate decarboxylating activityComplete 2.3 (100)CMinus dithiothreitol 2.0 (87)Minus TPP 0.7 (30)

Acetate kinaseComplete 15 (100)Minus acetyl-phosphate 2 (13)Minus ADP 0 (0)"Cells were grown to stationary phase, cell-free

extracts were prepared, and enzymes were assayed asdescribed in the text.

bPyruvate decarboxylating activity, nanomoles of['4C]C02 released per minute per milligram of protein;acetate kinase, micromoles of NADPH formed perminute per milligram of protein.

'Parenthetic values are the percentage of the "com-plete" reaction.

crude cell-free extract. As shown in Table 5, theNADPH formed by the enzyme couple, with thepartially purified acetate kinase, was dependentupon acetyl-phosphate as the substrate and ac-counted for 87% of the total NADPH generatedin the reaction mixture during conversion ofacetyl-phosphate to acetate. Acetate kinase ex-hibited an absolute dependence upon its othersubstrate, adenosine diphosphate (ADP), thusproducing 1 mol of ATP per mol of acetateformed.The conversion of PEP to succinate was me-

diated by PEP carboxykinase (23), malate de-hydrogenase, fumarate hydratase, and fumaratereductase as shown in Table 6 and Fig. 1. Theratio of the specific activities of malate dehydro-genase to fumarate reductase was approximately10, suggesting that the assay conditions for fu-marate reductase were not optimal, or that theelectron donor was not NADH, or both.

DISCUSSIONThe purpose of this study was to describe the

metabolic pathways used by C. ochracea strain25 to generate ATP from glucose. As shown inFig. 1, C. ochracea has evolved a complement ofenzymes and a transport mechanism that effi-ciently conserves energy from glucose dissimi-lation. Glucose was transported across the cellmembrane by a PEP:glucose PT system (Table2) forming glucose 6-phosphate, which was ca-tabolized to PEP.

Assays for the key enzymes (12) of the War-burg-Dickens, Entner-Douderoff, and glucosephosphoketolase pathways were negative (Table3). It is possible, however, that the key enzymesof these pathways were inactivated during prep-aration of cell-free extracts; thus, to prove thatthe Embden-Meyerhof-Parnas pathway is theexclusive sequence for primary glucose dissimi-lation in C. ochracea, it will be necessary totrace the distribution of specifically labeled glu-

TABLE 6. Conversion ofPEP to succinate by C.ochracea strain 25"

Enzyme Sp act'PEP carboxykinasec 5.4Malate dehydrogenase 91Fumarate hydratase 56Fumarate reductase 9

"Cells were grown to stationary phase, cell-freeextracts were prepared, and malate dehydrogenaseand fumarate hydratase were assayed as described inthe text.bNanomoles per minute per milligram of protein;

for PEP carboxykinase, micromoles per minute permilligram of protein.

' Reference 23.

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ENERGY METABOLISM IN C. OCHRACEA 557

GlucoseOral milieu

PTS Cell ~~membraneaTPEP Cytoplasm

Glucose b Glucos-S-P

t ~ ~~ic Pyruv tot Fructooe4-PATP ,4 d

Fructose S,6-dIP

1NAD+ Glyceraldehyde-3-P Dlhydroxyacetono-P

NADH2 -' ATP

2- P-Glycorate

To Glucose PTS PEP

CO" 'ATP - -j ATP

NADH2 Oxalactate Pyruvate

NAD+ kh--- O

Malate Acetyl-P

/1Fumarate ATPam

Succlnate Acetate

FIG. 1. Pathway from glucose to succinate and acetate in C. ochracea. Note that this figure does not showthe stoichiometry of the catabolic sequence. Enzymes which have been demonstrated (--) are (a) PEP:glucosePTsystem, (b) hexokinase, (c) glucose 6-phosphate isomerase, (d)phosphofructokinase, (e) fructose diphosphatealdolase, (f) enolase, (g) pyruvate decarboxylating activity, (i) acetate kinase, (j) PEP carboxykinase (23), (k)malate dehydrogenase, (I) fumarate hydratase, and (m) fumarate reductase; enzymatic reactions not yetdetermined (--).

cose carbons to their respective endproducts. Atthis time we tentatively conclude that glucose iscatabolized to pyruvate by the Embden-Meyer-hof-Parnas pathway. Romano et al. (42) haveprovided indirect evidence supporting this con-clusion by showing that fermentative bacteria,which use the PEP:PT system for sugar trans-port, dissimilate glucose by the Embden-Mey-erhof-Parnas pathway.Energy was conserved or generated in C.

ochracea at several points during catabolism ofglucose to acetate and succinate. The uptake ofglucose by the PEP:PT system conserves energywhich can be used elsewhere for biosynthesis(43, 44). Energy was generated in the Embden-Meyerhof-Parnas pathway by phosphoglyceratekinase (EC 2.7.2.3) and pyruvate kinase. Of the2 mol of PEP formed per mol of glucose in C.ochracea, 1 mol was used by the PEP:glucosePT system to transport another mole of glucoseacross the cell membrane; the other was catab-

olized disproportionately to acetate (37%) andsuccinate (63%) (Table 1). Regardless of thebranch into which PEP was shuttled, ATP wasformed-by PEP carboxykinase (23) on the pathto succinate or by pyruvate kinase and acetatekinase on the acetate leg (Tables 4 and 5). En-ergywise it was more advantageous for PEP tobe channeled to acetate since two ATP-gener-ating steps were demonstrated for that leg of theterminal pathway: pyruvate kinase and acetatekinase. In addition, the pyruvate generated fromPEP during operation of the PEP:glucose PTsystem can be further metabolized to acetate,thus gaining another ATP at acetate kinase. Theremaining portion of glucose carbon (as PEP)was channeled to succinate, with ATP beinggenerated in that leg of the pathway at PEPcarboxykinase (23).The above energy-yielding pathway in C.

ochracea is somewhat like the pathways of glu-cose catabolism reported for Cytophaga succin-

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icans (1) and Bacteroides succinogenes (32) inthat glucose is incompletely oxidized in a C02-dependent fermentation. There are, however, atleast two important differences between each ofthese species and C. ochracea. (i) Both B. suc-cinogenes (46) and C. succinicians (2) produceformic, acetic, and succinic acids from glucose.C. ochracea forms only acetate and succinate(50 and the present study). (ii) Although allthree species have C02-dependent fermenta-tions, B. succinogenes (32) and C. succinicians(1) fix C02 with a guanosine 5'-diphosphate-re-quiring enzyme which forms guanosine 5'-tri-phosphate as a product of C02 fixation; further-more, C. succinicians may have a second uni-dentified C02-fixing enzyme that does not re-quire a phosphate acceptor (1). On the otherhand, the sole C02-fixing enzyme in C. ochraceais an ADP-dependent PEP carboxykinase whichsynthesizes ATP as a product (23).

It was possible to determine the hypotheticalmoles of ATP produced from the catabolism of1 mol of glucose to acetate and succinate byusing the data in Table 1 and the catabolicsequence shown in Fig. 1. In yeast extract-sup-plemented cultures, the net amount of ATPsynthesized was 3.37 mol/mol of glucose. Toobtain this sum, it was necessary to considerthat ATP was generated (i) through catabolismof the pyruvate formed from PEP in the opera-tion of the PEP:glucose PT system, and (ii) byphosphoglycerate kinase, pyruvate kinase, ace-tate kinase, and PEP carboxykinase. The theo-retical moles of ATP calculated by usingBauchop and Elsden's mean value of YATP of10.5 (4) and the Ygiuose for the "plus-yeast ex-tract" culture presented in Table 1 was 7.2 molof ATP per mol of glucose catabolized. Thisvalue was 3.8 mol of ATP higher than expectedfrom experimental data. In the "minus-yeastextract" culture the theoretical value was 5.5mol of ATP per mol of glucose, a value 2.3 molhigher than was expected from the ATP-gener-ating sites shown in Fig. 1, suggesting that C.ochracea possessed other, as yet unknown, stepsfor ATP synthesis. The reason for the increasein ATP calculated for the yeast extract-supple-mented medium may be as follows: Withoutyeast extract, the medium was deficient in oneor more vitamins, cofactors, or metabolic inter-mediates needed to catalyze the conversion ofglucose metabolites (i.e., pyruvate and oxalace-tate) which accumulated in cultures lackingyeast extract (Table 1). The addition of yeastextract allowed the conversion of pyruvate andoxalacetate to proceed to acetate and succinate,respectively, with the generation of more ATPat acetate kinase and possibly at other sites.

There are several alternatives for the addi-tional ATP-generating sites necessary to recon-cile the differences between the experimentaldata and theoretical calculations. Since C.ochracea strain 25 contains primary dehydro-genases and cytochromes (29) and forms succi-nate as an end product, we explored possibilitieswhich involve a cytochrome-linked reduction offumarate to succinate. First, there may be acytochrome linkage between the acetate andsuccinate-yielding branches of PEP metabolismsimilar to the one reported in B. succinogenes(32). Thus, electrons generated during the TPP-dependent decarboxylation of pyruvate (Table5) could be transferred to a flavoprotein, then tothe cytochromes, which in turn were oxidized byfumarate with the concomitant generation ofATP. A second possibility for ATP productioncould involve the production of H2 during thedecarboxylation of pyruvate which, togetherwith a cytochrome, would be used to reducefumarate to succinate (14). Production of H2during pyruvate decarboxylation would also ex-plain our failure to detect the reduced fermen-tation products required to achieve a redox bal-ance in the culture in the face of carbon recov-eries approaching 100%. A third alternative forincreased ATP production in C. ochracea is acytochrome-linked reduction of fumarate byNADH, the most common electron donor forfumarate reductases in bacteria (52). In supportof this alternative, we have previously reportedthat purified C. ochracea membranes containedNADH dehydrogenase and cytochromes whichwere reduced by NADH (29). In the presentstudy we have presented evidence for an NADH-fumarate reductase activity in membrane frac-tions of C. ochracea (Table 6). Furthermore,cytochrome b has been recently detected inmembranes of C. gingivalis which could be re-duced with NADH and oxidized with fumarate(D. G. Longley and E. R. Leadbetter, Abstr.Annu. Meet. Am. Soc. Microbiol., 1980, K12, p.128).The coupling ofenergy generation to fumarate

reduction has been inferred by the presence ofcytochromes and high molar growth yields inmany bacteria (14), as well as in several speciesof the closely related genus, Bacteroides (forreview, see reference 31). The previously re-ported cytochromes (29) and the high molargrowth yields obtained for C. ochracea in thepresent study add to these inferences. The cy-tochrome-linked pathway would augment theATP generated by substrate-level phosphoryla-tions in other parts of the catabolic pathway forglucose.Although the exact role of Capnocytophaga

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sp. in the etiology of juvenile periodontitis andcertain systemic infections remains uncertain,these newly described gliding bacteria probablycontribute to the overall pathogenic potential ofthe microflora in the oral ecosytem. As the phys-iology of the Capnocytophaga is more closelydefined, its role in oral and systemic diseasesshould emerge.

ACKNOWLEDGMENTS

We thank Raymond B. Bridges and Albert T. Brown fortheir many helpful discussions and May C. Fu for her experttechnical assistance. We are also indebted to Connie Thaxtonfor her diligence in preparation of the typescript.

This investigation was made possible through grants fromthe University of Kentucky Research Foundation (7900069)(R.C.) and the College of Dentistry Research Committee (R.C.and T.T.L.).

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