microbial ecophysiology whey biomethanation ... · 190 chartrain andzeikus by techniques previously...

Vol. 51, No. 1APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Jan. 1986, p. 188-1960099-2240/86/010188-09$02.00/0Copyright © 1986, American Society for Microbiology

Microbial Ecophysiology of Whey Biomethanation: Characterizationof Bacterial Trophic Populations and Prevalent Species in

Continuous CultureM. CHARTRAIN' AND J. G. ZEIKUS2*

Department of Bacteriology, University of Wisconsin, Madison, Wisconsin 53706,1 and Michigan Biotechnology Instituteand the Departments ofBiochemistry and Microbiology, Michigan State University, East Lansing, Michigan 488242

Received 27 June 1985/Accepted 11 October 1985

The organization and species composition of bacterial trophic groups associated with lactose biomethanationwere investigated in a whey-processing chemostat by enumeration, isolation, and general characterizationstudies. The bacteria were spatially organized as free-living forms and as self-immobilized forms appearing inflocs. Three dominant bacterial trophic group populations were present (in most probable number permilliliter) whose species numbers varied with the substrate consumed: hydrolytic, 1010; acetogenic, 107 to 1010;and methanogenic, 106 to 109. The three prevalent species utilizing lactose were identified as Leuconostocmesenteroides, Klebsiella oxytoca, and Clostridium butyricum. Clostridium propionicum and Desulfovibriovulgaris were the dominant lactate-consuming, hydrogen-producing acetogenic bacteria, while D. vulgaris wasthe only significant ethanol-degrading species. Methanosarcina barkeri and Methanothrix soehngenii wereidentified as the dominant acetate-utilizing methanogens, and Methanobacteriumformicicum was the prevalenthydrogen-utilizing methanogen. A microbial food chain is proposed for lactose biomethanation that comprisesmultiple species in three different groups, with the major hydrogen-producing acetogen being a sulfate-reducing species, D. vulgaris, which functioned in the absence of significant levels of environmental sulfate.

The complete degradation of complex organic matter intomethane and carbon dioxide requires the involvement ofvarious microorganisms. The organization of anaerobic bac-teria into different trophic groups that perform specificmetabolic transformations during the degradation of organicmatter has been proposed by various investigators (23, 45,46). The first trophic group is composed of hydrolyticbacteria that degrade complex organic matter into multipleacid and neutral end products. These acids and neutralproducts are further transformed into acetate and hydrogenby the hydrogen-producing acetogens. The presence ofacetogenic bacteria that produce acetate from H2-CO2 aswell as glucose, ethanol, and lactate has been detected indigestors, but their significance in carbon degradation is notestablished (4, 27). Methanogenic bacteria are the terminaltrophic group and utilize acetate, H2-CO2, methanol, andmethylamines as substrates (2, 22, 44). Acetate and H2-CO2are the major methane precursors in sludge digestors (15,34). When sulfate is available, methane production is limitedbecause sulfate-reducing bacteria can outcompetemethanogens for acetate and hydrogen (19, 20, 29, 32, 41).All the bacterial trophic groups involved in anaerobic diges-tion are highly dependent on species metabolic interactions,and inhibition of one group can cause failure of the overallbiomethanation process (31, 36, 44, 45).Enumeration of the bacterial trophic groups and isolation

of the prevalent species involved in anaerobic degradation oforganic matter has been accomplished in some waste treat-ment systems. In sewage sludge, hydrolytic bacterial popu-lations are usually high and generally comprise between 108and 1010 bacteria per ml of sludge (11, 12, 16, 17, 22, 37, 40,45). Studies of bacterial isolations report a large variety ofhydrolytic organisms, with Streptococcus spp., Bacillusspp., Clostridium spp., and various enteric bacteria as

* Corresponding author.

prevalent species (11-13, 37, 40). Populations of hydrogen-producing acetogens in sludge have been reported (47), andthey are usually isolated as obligate syntrophic associationswith a hydrogen-consuming species (3, 6, 25). Populations ofsulfate-reducing bacteria are often present in sewage sludgeat 104 to 108 organisms per ml, with Desulfovibrio speciesbeing most prevalent (38, 39). Hydrogen-utilizingmethanogens are present in sludge at 108 organisms per ml,with Methanobacterium spp. and Methanospirillum spp.usually present (9, 11, 43, 45). Acetate-utilizing methanogenpopulation levels in sludge are approximately 106 organismsper ml, with Methanosarcina and Methanothrix speciescommonly identified (33, 43, 45, 47).Whey is a by-product of the cheese industry that presents

an important pollution problem (C. D. Parker, Proc. 2nd.Natl. Symp. Food Processing Wastes, Environmental Pro-tection Agency report 12060 FUR 03/71, 1971). The organicportion of liquid whey is mostly composed of lactose, andbiomethanation studies have been realized in pilot plants(C. J. Clanton, P. R. Goodrich, P. A. Lee, and B. D.Backus, Meet. Am. Soc. Agric. Eng. paper 81-6007, 1981).These previous investigations were engineering oriented anddid not deal with the microbiology of whey biomethanation.

Recently, we described a steady-state chemostat systemfor continuous biomethanation of whey at a retention time of100 h and demonstrated the intermediary metabolic route forlactose transformation to methane and carbon dioxide (8).The purpose of the present work was to define the microbialpopulations in this ecosystem in relation to enumeration oftrophic groups, spatial organization, and isolation and iden-tification of prevalent species.

MATERIALS AND METHODS

Chemicals and gases. All chemicals used were of reagentgrade or better and were obtained from Sigma Chemical Co.,

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BACTERIAL POPULATIONS IN WHEY BIOMETHANATION

St. Louis, Mo., or Mallinckrodt, Inc., Paris, Ky. All gasesused were obtained from Matheson Scientific, Inc., Joliet, Ill.Chemostat and physicochemical conditions. The chemostat

had a 1-liter (total capacity) vessel with a 260-ml workingvolume. It was continuously fed with a phosphate basalbuffer (PBB) medium (26) containing 10 g of whey per liter ata retention time of 100 h and was operated at 37°C and pH7.1. A complete description of the system and operatingparameters was given in a separate report (8).

Microscopy. Phase-contrast and epifluorescence observa-tions were made with an Olympus microscope model BH2equipped with a mercury lamp and an automatic-exposurecamera. The autofluorescence of methanogen deazoflavinwas observed with a B (IF-490) excitation filter. Forphotomicrography, liquid samples were removed from thechemostat by syringe with an 18-gauge needle and placed onagar-coated microscope slides. Kodak technical pan film2415 was used for photomicrographs.

Electron microscopic observations were made as follows.Samples were centrifuged for 5 min at 3,000 x g. The pelletwas suspended in a fixation solution consisting of 2.5%glutaraldehyde in 0.1 M cacodylate buffer at pH 7.2 for 1 h.The preparations were stained with 0.015% ruthenium red inthe same buffer. The fixed cells were then embedded in 1.5%ion-agar (Difco Laboratories, Detroit, Mich.) and cut into1-mm3 cubes. The cubes were rinsed three times for 20 mineach time in 0.1 M cacodylate buffer and then placed into1.25% OS04 in 0.1 M cacodylate buffer for 12 h at 4°C. Thepreparations were dehydrated by successive transfers of theblocks in a series of 35, 50, 70, 85, 95, and 100% ethanolsolutions. The material was then washed twice in propyleneoxide for 15 min each time and embedded in Spurr resin (35).Thin sections were cut with a diamond knife on a porterBlum MT-2 ultramicrotome at about 80-nm thickness. Sec-tions were stained for 15 min in saturated uranyl acetate in50% methanol, rinsed, dried, and then stained for 5 min inReynold lead citrate (28). The thin sections were examinedand photographed with a Hitachi UH-11 E transmissionelectron microscope operated at 75 kV.Enumeration techniques. Enumerations were performed

on 1-ml samples taken from the chemostat operating understeady-state conditions for 2 months or for 14 volumeturnovers. Samples were removed by syringe with an 18-gauge needle and immediately injected into a 26-ml pressuretube (Bellco Glass, Inc., Vineland, N.J.) containing 9 ml ofprereduced PBB medium and 10 to 15 glass beads of 0.5 mmin diameter and sealed with a black bung. The tube wasplaced on a Vortex mixer for 1 min, and the bacterial flocswere disrupted. A similar treatment was applied to the twosuccessive decimal dilution tubes. Enumerations of lactose-degrading bacteria showed an increase of approximately100-fold when the glass bead disruption procedure was used.Except when specified, the enumerations were performed bythe most probable number technique, with three tubes perdilution (1).

Lactose-utilizing bacteria were enumerated in 26-ml pres-sure tubes containing 9 ml of APT basal medium (Difco) and10 mM lactose. The tubes were incubated for 1 week at 37°C,and the results were interpreted on the basis of a 0.2-unitincrease in the optical density at 660 nm when comparedwith a noninoculated tube.

Sulfate-reducing bacteria utilizing lactate or ethanol wereenumerated in pressure tubes containing 9 ml of PBB me-dium, 20 mM carbon substrate and electron donor, 20 mMFeSO4, 0.05% yeast extract, and 0.02% sodium ascorbate asreducing agent. Sulfate-reducing bacteria utilizing hydrogen

were enumerated in a similar medium but with 1.8 atm (182.3kPa) of H2-CO2 (80:20) in the head space as electron source.The tubes were incubated at 37°C for 2 weeks, and theresults were interpreted on the basis of formation of an Fe2Sblack precipitate.

Syntrophic bacterial populations utilizing lactate, ethanol,propionate, and succinate were enumerated in 26-ml pres-sure tubes containing 9 ml of PBB medium, 20 mM respec-tive energy source, and 1 ml of a Methanobacteriumformicicum culture (optical density at 660 nm, 0.3) as thehydrogen-consuming partner. A series of controls were usedthat contained the same components but omitted the energysource. The most probable number technique tubes wereincubated at 37°C for a month, and the results were inter-preted on the basis of an increase in methane production(>0.5%) between the controls and the enumeration tubes forthe same dilution.The homoacetogenic bacteria (or hydrogen-consuming

acetogenic bacteria) were enumerated by plate count on themedium described by Braun et al. (4). The plates wereincubated inside a pressure paint pot that contained anatmosphere of H2-CO2 at 20 lb/in2 for a period of 3 weeks at37°C. The results were interpreted on the basis of a change inthe pH indicator color around the acid-producing colonies.Methanogenic bacteria utilizing acetate or formate were

enumerated in 26-ml pressure tubes containing 9 ml of PBBmedium, 80 mM respective substrate, and 0.05% yeastextract. Methanogens forming methane from hydrogen andcarbon dioxide were enumerated in pressure tubes contain-ing 9 ml of PBB medium and 0.05% yeast extract andpressurized with 1.8 atm (182.3 kPa) of H2-CO2 (80:20). Thetubes were incubated at 37°C for a month, and the resultswere interpreted on the basis of the presence of methane inthe head phase of the tube (>0.5% of CH4).

Isolation and identification of species. All the followingtechniques were performed under strict anaerobic conditionseither with an anaerobic glove bag or by utilizing syringes forall transfers in pressure tubes. The isolations were made bydilution from the positive end-dilution tube. Dilutions (0.1ml) were plated on the appropriate enrichment mediumsolidified with 1.5% purified agar (Difco). The plates wereincubated inside the anaerobic glove bag except for thehydrogen-utilizing bacteria, which were incubated inside apressure jar with H2-CO2 at 20 lb/in2. All media for theisolation of methanogenic bacteria contained 1,200 U ofpenicillin G (Sigma) per ml. Plates used for isolation ofacetate-utilizing methanogens were incubated inside ananaerobic jar containing filter paper soaked with 2.5% Na2S.

After incubation, the colonies were picked with a sterileglass capillary made from a Pasteur pipette, and the cellswere inoculated into a 26-ml pressure tube containing 2.5 mlof the appropriate medium. Purity of the strains was che9kedafter growth in liquid medium by microscopy and by streak-ing on a plate of the appropriate medium and observingcolony homogeneity. Species identification of the isolatedstrains was performed by the dichotomous tests described inthe Bergey manuals (7, 18) and in authoritative reviews onmethanogens (2, 21, 44). General tests (7, 18) employed forspecies identification included catalase, oxidase, urease,substrate range, and spore location.Mass balance experiments. Mass balance experiments

were performed in 26-ml pressure tubes which contained 10ml of the indicated medium and an atmosphere of nitrogenand were sealed with a black bung. After inoculation andsubsequent incubation at 37°C, substrate consumption, endproduct formation, and biomass production were quantified

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190 CHARTRAIN AND ZEIKUS

by techniques previously reported (8). Carbon dioxide,acetate, butyrate, propionate, ethanol, and acetoin werequantified by standard gas chromatographic procedures.Lactate and formate were detected by high-pressure liquidchromatographic techniques (8). Lactose was measured byenzymatic analysis with a lactose assay kit (BoehringerMannheim Biochemicals, Indianapolis, Ind.). Dry weightwas measured by filtration of a known volume through amembrane filter (pore size, 0.22 ,um; Millipore Corp.,Bedford, Mass.) that was dried at 60°C until a constantweight was reached. The amount of cellular carbon (C) wascalculated (in millimoles) by the following formula: C =(milligrams [dry weight] of cells x 0.451)/12. The electroncomposition of the cell was calculated with the value of 4.21electrons per mol of cellular carbon (10).

RESULTS

Microscopic analysis. Microscopic examination of thewhey-processing chemostat revealed that the microbial com-position was highly heterogeneous, with free-living bacterialforms and numerous self-immobilized bacteria in flocs.Protists and eucaryotic nematodes were not observed.

Figure 1 shows representative photomicrographs of thewhey-processing chemostat bacterial population. Figure 1Ashows the morphological heterogeneity of the free-livingbacterial population, which comprised cocci in short chains,curved rods, and various rod-shaped bacteria. Figure 1Bshows a typical floc. Samples from the chemostat containednumerous flocs, and their sizes varied from 1 to 20 ,um indiameter. Figure 1C shows a floc observed under UVepifluorescence. High numbers of these fluorescent, irregu-larly shaped rods indicative of Methanobacterium for-micicum were observed within these flocs. Fluorescentspirillum and sarcina-shaped bacteria were also observedwithin the flocs but in much lower numbers (data notshown).

Figure 2 shows a typical low-power electron photo-micrograph of a thin-sectioned floc which was stained withruthenium red to reveal glycocalyx matrix material. Theseultrastructural data support the size and architectural fea-tures described for the Leuconostoc, Klebsiella,Desulfovibrio, Methanobacterium, Methanosarcina, andMethanothrix species which were isolated and which arecharacterized below.

Trophic-group analysis. The different bacterial trophicgroups in the chemostat were enumerated to assess theprevalence of specific metabolic groups and species. Table 1shows the approximate number of bacterial trophic groupsas enumerated on separate occasions. Indicative of stability,the number of organisms obtained in the whey-processingchemostat varied less than 1 order of magnitude betweensampling times separated by 2 weeks. The trophic grouppopulation numbers observed in decreasing order of domi-nance were lactose hydrolytics at 1010, hydrogen-producingacetogens at 108 to 1010, and methanogens at 106 to 109.Notably, sulfate-reducing bacteria which degraded lactateand ethanol were as numerous as hydrogen-producingacetogenic populations enumerated as syntrophic bacteria.

FIG. 1. Phase-contrast photomicrographs of the whey-processing chemostat bacterial population. The bar represents 10p.m. (A) Free-living bacterial population; (B) bacteria immobilizedwithin a floc; and (C) bacterial floc under UV light, indicating evendistribution of epifluorescent methanogenic rods.

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FIG. 2. Electron photomicrograph of a thin section of ruthenium red-stained floc material from the whey-processing chemostat. The barrepresents 1 ,um. Mb, Methanosarcina barkeri; Dv, D. vulgaris; Ms, Methanothrix soehngenii, Mf, Methanobacteriumformicicum; and Lm,L. mesenteroides.

Microscopic analysis indicated that the bacteria present inthe end-dilution tubes for both of these enumerations had thesame morphology. Enumerations for butyrate-, propionate-,and succinate-degrading bacteria revealed that these popu-

lations were at the assay detection limits (103/ml). Hydrogen-consuming acetogenic populations were lower than popula-tions of methanogens.

Prevalent-species analysis. The prevalent species in thechemostat ecosystem were isolated from end-dilution tubesused to enumerate the trophic groups. Three different spe-

cies of hydrolytic bacteria that consumed lactose were

isolated and identified as Leuconostoc mesenteroides, Kleb-siella oxytoca, and Clostridium butyricum. Figure 3 showsphase-contrast photomicrographs of these species. The firstspecies was a gram-positive coccus in short chains, and itgrew aerobically and anaerobically, was devoid of catalase,and fermented glucose into lactate and smaller amounts ofethanol and acetate. It was identified as L. mesenteroides on

the basis of these observations and additional standardbiochemical identification tests (7). The second species was

a strict anaerobe, and it was a gram-positive, sporeformingrod that fermented glucose to butyrate, acetate ethanol,hydrogen, and CO2. It was identified as C. butyricum on thebasis of these observations and additional biochemical tests(7). The third species was a facultative anaerobe, gram-

TABLE 1. Trophic-group populations in a steady-statewhey-processing chemostata

MPN seriesbBacterial group

1 2

I (Lactose consuming hydrolytic) 2.3 x 1010 4.6 x 1010II (Acetogenic)H2 producing, lactate consuming 1.5 x 108 4.6 x 108H2 producing, ethanol consuming 2.4 x 109 1.1 x 1010H2 consuming 1.9 x 107 1.1 x 107

III (Methanogenic)H2 consuming 1.5 x 108 1.1 x l09Formate consuming 2.8 x 107 9.3 x 108Acetate consuming 7.5 x 106 4.3 x 106

IV (Sulfate reducing)H2 consuming 2.4 x 109 9.3 x 108Lactate consuming 2.4 x 109 1.5 x 109Ethanol consuming 2.3 x 108 4.6 x 109a The enumerations were performed when the chemostat was operating at a

retention time of 100 h at 37°C and pH 7.1 and was fed with PBB mediumcontaining 10 g of whey per liter. The enumerations were done by the mostprobable number technique, with three tubes per dilution except for theH2-consuming acetogens, which were enumerated on plates.

b MPN, Most probable number.

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FIG. 4. Phase-contrast photomicrographs of the prevalent hy-drogen-producing acetogens isolated from the whey-processing che-mostat. The bar represents 10 ,um. (A) D. vulgaris; (B) C.propionicum.

negative rod which was catalase positive and oxidase nega-tive and fermented glucose to acetate, ethanol, H2, lactate,and acetoin. On the basis of these observations and addi-tional biochemical tests (18), it was identified as K. oxytoca.Table 2 compares the lactose fermentation balance of thethree different hydrolytic strains.Hydrogen-producing acetogens which degraded lactate or

ethanol were enriched in syntrophic cultures that containedMethanobacterium formicicum as the hydrogen-consumingpartner. Microscopic observations of these lactate

FIG. 3. Phase-contrast photomicrographs of the prevalent lac-tose-hydrolytic strains isolated from the whey-processing chemo-stat. The bar represents 10 pum. (A) L. mesenteroides; (B) C.butyricum; and (C) K. oxytoca.



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TABLE 2. Lactose fermentation balances of the prevalent hydrolytic species isolated from the whey biomethanation chemostata

Amt of end product formed (mmolU100 mmol of lactose) Product recoveryOrganism

Lactate Acetate Ethanol Butyrate Formate Acetoin CO2 H2 Cells Carbon Electrons

L. mesenteroides 246 43 15 31 2 x 10-4 186 91 86C. butyricum 56 158 78 55 199 270 219 106 112K. oxytoca 46 73 157 160 49 97 8 x 10-3 98 104 97

a The bacteria were grown in 26-ml pressure tubes containing an atmosphere of nitrogen and sealed with a butyl rubber bung. The tubes contained 10 ml ofreduced TYE medium (44) with 5 mM lactose and were incubated at 37°C until a constant optical density at 660 nm was reached.

cocultures indicated two kinds of hydrogen-producingacetogens, a gram-negative curved rod and a sporeforming,gram-positive straight rod. Only one morphotype of hydro-gen-producing acetogen which degraded ethanol was ob-served, and it was a gram-negative curved rod. The lactateand ethanol consumers were isolated by transferring thecultures often to eliminate the slower-growing methanogenin the presence or absence of added sulfate. Figure 4 showsphase-contrast photomicrographs of the two lactate-degrading species isolated. The gram-negative curved rodwas strictly anaerobic and reduced sulfate as electron accep-

tor. It utilized lactate, ethanol, and hydrogen as electronsources and was identified as Desulfovibrio vulgaris (7). Thesporeforming gram-positive rod was a strict anaerobe, itfermented lactate into acetate, propionate, and H2-CO2, andit was identified as Clostridium propionicum (7). Theethanol-degrading species possessed the same features asthe isolated curved rod which degraded lactate and wasidentified as D. vulgaris. The organisms that were isolatedfrom sulfate-reducing enrichments which degraded lactate orethanol were also identified as D. vulgaris.

Table 3 compares the fermentation balances for the hy-drogen-producing acetogens isolated from the chemostat. C.propionicum fermented lactate into propionate, acetate,hydrogen, and CO2. In the presence of sulfate, D. vulgarisfermented lactate into acetate, H2S, small amounts ofethanol, and traces of hydrogen. Acetate, H2S, and traces ofhydrogen were the products of ethanol degradation by thisspecies.The prevalent hydrogen- or formate-consuming methano-

genic species isolated was a short rod which formed chainsand flocs and did not grow at 60°C. This strain was identifiedas Methanobacterium formicicum (2, 21, 44). Two differentacetate-utilizing methanogens were isolated. A sarcina thatused acetate, H2-CO2, and methanol as methane precursorsand did not grow at 60°C was identified as Methanosarcinabarkeri (2, 21, 44). A long, filamentous, fat rod with squareends that grew on acetate but not on H2-CO2 was identifiedas Methanothrix soehngenii (14, 45). Figure 5 shows phase-contrast photomicrographs of these three methanogenic spe-

cies, which were isolated from the whey-processing chemo-stat.

DISCUSSIONThe bacterial flora present inside the whey-processing

chemostat was organized into two major components, self-immobilized cells in flocs and free-living cells. A complexfood chain comprising three distinct trophic groups wasresponsible for lactose biomethanation. The prevalent spe-cies of lactose hydrolyzers, lactate- and ethanol-degradingacetogens, and methanogens isolated from the chemostatwere previously described species of facultative and obligateanaerobes common to a variety of other ecosystems.The mixed bacterial floc observed in this chemostat eco-

system was interesting because this phenomenon is usuallymore common to up-flow fixed bed reactors used inanaerobic wastewater treatment systems (30). The flocscomprised different bacterial species from all three majortrophic groups. Apparently even with good mixing, shortretention time, and a readily degradable soluble substrate,methanogens and nonmethanogens like to grow as consortia.Thiele et al. reported the physiological importance of flocformation in facilitating metabolic communication betweenspecies via juxtapositioning of hydrogen-producing and hy-drogen-consuming syntrophic partners (J. Thiele, M.Chartrain, and J. G. Zeikus, unpublished results).The specific trophic group population levels in this whey

biomethanation ecosystem were what was expected for anactive anaerobic digestion process. Lactose-degrading bac-terial populations were high, as is any microbial populationenumerated in sewage sludge contact digestors (11, 12, 16,17, 22, 37, 40). Hydrogen-producing acetogens that degradedlactate and ethanol were more numerous than those whichdegraded propionate, butyrate, and succinate. These resultssupport our previous 14C tracer studies, which demonstratedthat lactate and ethanol were the major intermediary metab-olites formed during lactose biomethanation by the whey-processing chemostat, with butyrate and propionate onlyformed at background levels when the chemostat was oper-ating at a retention time of 100 h (8). The methanogenic

TABLE 3. Carbon fermentation balances for the prevalent hydrogen-producing acetogens isolated fromthe whey biomethanation chemostat

Amt of end product formed (mmol/100 mmol of substrate) Product recovery (%)Organism Substrate

Propionate Acetate Ethanol CO2 H2 H2S Cells Carbon Electrons

C. propionicuma Lactate 63 23 27 1 13 92 93D. vulgarish Lactate 71 17 61 3 x 1O-3 35 27 115 97

Ethanol 90 1 1 x 1O-3 38 28 118 94a Cells were grown in a 26-ml pressure tube containing PBB medium with 100 mM lactate. The tubes were incubated at 37°C, and end products were measured

after a constant optical density at 660 nm was reached.b Cells were grown in a 26-ml pressure tube containing PBB medium, 40 mM respective substrate, and 40 mM sodium sulfate. The tubes were incubated at 37°C,

and end products were measured after a constant optical density was reached.

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populations utilizing hydrogen and formate were within therange reported for sewage sludge digestors (12, 33). Themethanogens which degraded acetate were present at lowerlevels, probably because the two species identified grew inclumps or long filaments which were not significantly dis-rupted by the enumeration methods used.

Characterization of the lactose-hydrolyzing bacteria asspecies of enteric bacteria, Clostridium, and lactics is notsurprising, as these bacteria have been isolated from sewagesludge digestors (11-13, 22, 40). Leuconostoc and Klebsiellaspecies are common to milk products and may be naturallyassociated with nonsterile whey. K. oxytoca and L.mesenteroides are exopolysaccharide formers and may bemajor contributors to floc formation in this wheybiomethanation ecosystem. The coexistence at high popula-tion densities of three species which utilize lactose underenergy-limited conditions raises interesting questions of spe-cies competition for energy source and survival which willbe addressed in a separate report (manuscript in prepara-tion).

Notably, the prevalent hydrogen-producing acetogenswhich degraded lactate and ethanol in this ecosystem werenot obligate proton-reducing acetogenic species but were D.vulgaris and C. propionicum. Desulfovibrio spp. must begrowing syntrophically via interspecies hydrogen transfer (5,24), because sulfate was absent in these ecosystems. There-fore, this result provides the first evidence that in a naturallyoccurring process, sulfate-reducing bacteria function in car-bon degradation as syntrophs in the absence of sulfate.These findings extend the classic coculture discoveries ofBryant and co-workers (5, 24) which first established thisphenomenon in artificial cocultures. Although C.propionicum was isolated as a prevalent lactate-degradingspecies, propionate was not a major metabolite in thewhey-processing chemostat. Thus, it is assumed that C.propionicum was growing in the chemostat by coupling H2transfer to methanogens. In a separate report (manuscript inpreparation), we have shown that the metabolism of C.propionicum is altered via interspecies hydrogen transfertowards the production of more acetate and less propionatewhen grown in the presence of Methanobacteriumformicicum.

Methanobacterium formicicum was the dominant hydro-gen- and formate-utilizing methanogen isolated from thechemostat, but other hydrogen-consuming methanogenssuch as Methanospirillum spp. and Methanosarcina spp.were present at lower numbers, suggesting that competitionfor hydrogen occurs between species. The dominant acetate-utilizing methanogens isolated from the chemostat at a 100-hretention time were Methanosarcina barkeri andMethanothrix soehngenii. An explanation for substrate com-petition between these species for acetate was recentlysuggested (47) on the basis that Methanosarcina spp. growrapidly, with lower substrate affinity, and predominate athigh dilution rate, whereas Methanothrix spp. grow moreslowly, with higher substrate affinity, and predominate atlow dilution rate. In the present ecosystem, the dilution ratewas moderate and the acetate concentration was very low,enabling both species to grow.

FIG. 5. Phase-contrast photomicrographs of the prevalentmethanogens isolated from the whey-processing chemostat. The barrepresents 10 ,um. (A) Methanobacterium formicicum; (B)Methanosarcina barkeri; and (C) Methanothrix soehngenii.



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I. Hydrolytic bacteria

Clostridiun butyricun

Klebsiella oxytoca

Leuoxnostoc mesenteroides

II. Acetogenic bacteria

ACETATEDesulfovibrio vulgaris

Clostridiun propionicum

III. Methanogenic bacteria

Methanobacterium formicicum

Methanosarcina barker i

Methanothr ix soehngeni i

FIG. 6. The microbial food chain responsible for biomethanation of lactose in three distinct but simultaneous trophic phases. The heavyarrows indicate the major flux of carbon and electrons during biomethanation.

The present study extends the model proposed (8) forcarbon and electron flow during lactose biomethanation inthe whey-processing chemostat ecosystem by identifying thespecific species accounting for lactose hydrolysis,acetogenesis from lactate and ethanol, and methanogenesisfrom acetate plus H2-CO2. A microbial food chain for lactosebiomethanation which integrates the major route of carbonand electron flow and the specific species composition isshown in Fig. 6.

ACKNOWLEDGMENTSThis research was supported by awards from the Institute Pasteur,

Societe Lyonnaise des Eaux, Le Pec, France, and the MichiganBiotechnology Institute.

LITERATURE CITED

1. American Public Health Association. 1965. Standard methods forthe examination of water and wastewater, 12th ed. AmericanPublic Health Association, Washington, D.C.

2. Balch, W. E., G. E. Fox, L. J. Magrum, C. R. Woese, and R. S.Wolfe. 1979. Methanogens: reevaluation of a unique biologicalgroup. Microbiol. Rev. 43:260-296.

3. Boone, D. R., and M. P. Bryant. 1980. Propionate-degradingbacterium, Syntrophobacter wolinii sp. nov. gen. nov., frommethanogenic ecosystems. Appl. Environ. Microbiol. 40:626-632.

4. Braun, M., S. Schobert, and G. Gottschalk. 1979. Enumerationof bacteria forming acetate from H2 and CO2 in anaerobichabitats. Arch. Microbiol. 120:201-204.

5. Bryant, M. P., L. L. Campbell, C. A. Reddy, and M. R. Crabill.1977. Growth of Desulfovibrio in lactate or ethanol media low insulfate in association with H2-utilizing methanogenic bacteria.Appl. Environ. Microbiol. 33:1162-1169.

6. Bryant, M. P., E. E. Wolin, M. J. Wolin, and R. S. Wolfe. 1967.M. omelianski, a symbotic association of two species of bacte-ria. Arch. Microbiol. 59:20-21.

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