APPLIED AND ENVIRONMENTAL MICROBIOLOGY, May 1983, P. 1519-15250099-2240/83/051519-07$02.00/0Copyright © 1983, American Society for Microbiology

Vol. 45, No. 5

Hydrogen Metabolism by Decomposing CyanobacterialAggregates in Big Soda Lake, Nevada

RONALD S. OREMLANDU.S. Geological Survey, Menlo Park, California 94025

Received 7 February 1983/Accepted 10 March 1983

Hydrogen production by incubated cyanobacterial epiphytes occurred only inthe dark, was stimulated by C2H2, and was inhibited by 02. Addition of NO3inhibited dark, anaerobic H2 production, whereas the addition of NH4' inhibitedN2 fixation (C2H2 reduction) but not dark H2 production. Aerobically incubatedcyanobacterial aggregates consumed H2, but light-incubated rates (3.6 ,umol of H2g 1 h-1) were statistically equivalent to dark uptake rates (4.8 ,umol of H2 g-1h-1), which were statistically equivalent to dark, anaerobic production rates (2.5to 10 ,umol of H2 g-1 h-1). Production rates of H2 were fourfold higher foraggregates in a more advanced stage of decomposition. Enrichment cultures ofH2-producing fermentative bacteria were recovered from freshly harvested, H2-producing cyanobacterial aggregates. Hydrogen production in these cyanobacter-ial communities appears to be caused by the resident bacterial flora and not by thecyanobacteria. In situ areal estimates of dark H2 production by submergedepiphytes (6.8 ,umol of H2 m2 h 1) were much lower than rates of light-driven N2fixation by the epiphytic cyanobacteria (310 Lmol of C2H4 m-2 h-1).

The abundance of dissolved molecular hydro-gen within the aerobic water column of oceansand lakes often exceeds equilibration levels withthe atmosphere, suggesting an in situ microbialsource (4, 5, 9, 10, 21a, 22, 23). Althoughnumerous types of microorganisms are able toproduce or consume hydrogen or both (underaerobic or anaerobic conditions), production inaerobic, pelagic waters has been attributed tothe activity of N2-fixing cyanobacteria of thegenus Oscillatoria (5, 21a, 22). However, littleexperimental work has been done with free-living, N2-fixing populations of Oscillatoria (orother cyanobacterial genera) to determine theirinvolvement (if any) in the hydrogen cycle ofaquatic environments.

Big Soda Lake is an alkaline (pH 9.7), mero-mictic lake located in western Nevada nearFallon. Aspects of the lake's chemistry andbiology have been published previously (2, 13,18, 20; Y. Kharaka, L. M. Law, W. W. Caroth-ers, and S. W. Robinson, Eos Trans. Am.Geophys. Un. 62:922, 1981; J. E. Cloern andB. E. Cole, Eos Trans. Am. Geophys. Un.63:965,1982; J. E. Cloern, B. E. Cole, and R. S.Oremland, Hydrobiologia, in press). During pe-riods of thermal stratification (April to Decem-ber), the lake's surface waters (upper 20 m) areseverely depleted in dissolved inorganic nitro-gen (<1 ,uM) and pelagic productivity is limited

by nitrogen (2, 20). During this time period,however, dense beds of the submerged macro-phyte Ruppia sp. develop in the lake's littoralzone, and the plant becomes heavily colonizedby epiphytic cyanobacteria, primarily Anabaenasp. (Cloern et al., in press). During the day theseepiphytes evolve considerable amounts of 02bubbles, many of which remain entrapped with-in the epiphyte matrix, thereby giving buoyancy.Uprooted Ruppia-epiphyte aggregates are fre-quently found floating in the lake's pelagic zone.Because dissolved hydrogen (when present) inthe lake exhibits an extremely "patchy" distri-bution (R. S. Oremland, L. Marsh, C. Cul-bertson, and D. J. DesMarais, Eos Trans. Am.Geophys. Un. 62:922, 1981), I theorized thatthese epiphytic cyanobacteria may evolve hy-drogen via their nitrogenase system (7, 12, 14,15), thus providing a source of hydrogen to thelake. However, microscopic examination ofthese cyanobacterial epiphytes revealed notonly heterocystous Anabaena sp. but also alarge, morphologically diverse population ofbacteria. In this paper, I report that these cyano-bacterial communities both produce and con-sume hydrogen under anaerobic and aerobicconditions, respectively. However, hydrogenproduction appears to be caused by the bacterialiving within the aggregates and not by theAnabaena sp.

1519

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

APPL. ENVIRON. MICROBIOL.

MATERIALS AND METHODS

Flask experiments. Experiments were conductedduring July through November, when plants and epi-phytes were abundant. Freshly harvested, Anabaena-colonized Ruppia sp. stems were cut into small sec-tions (5 to 8 cm in length), and several sections wereplaced into 125-ml Erlenmeyer flasks which contained50 ml of Big Soda Lake surface water (salinity = 2.7g/liter; 0.5 g of plant dry weight per flask). Flasks weresealed under air with sleeved rubber septa and incu-bated statically (at 12°C), either exposed to ambientsunlight or in darkness (covered with two layers ofaluminum foil). Acetylene (15 kPa) and NH4Cl (100mg) were added to selected flasks by syringe injection.Flasks were vigorously handshaken (15 s) at samplingintervals to facilitate gas-phase equilibration. Subsam-ples (2.25 ml) of the gas phases were removed by glasssyringes and stored in 2-ml nonsterile Vacutainers(Becton, Dickinson & Co.). The Vacutainers werenext injected with 0.25 ml of a 15% NaCl solution andstored inverted to retard diffusion of trace gasesthrough the stopper. Analysis of H2 was performed bythermal conductivity gas chromatography (Ar carrier),and hydrocarbons were measured by flame-ionizationgas chromatography as described previously (6, 17).All analyses were performed within 1 week of collec-tion of samples at Big Soda Lake.

In another series of flask experiments, the thick(diameter, 1 to 3 cm) epiphytic cyanobacterial aggre-gates (CBAs) were separated from the Ruppia sp.stems (diameter, 0.3 cm) by gentle shaking in a bucket.To achieve a uniform suspension, the dissociatedCBAs were subjected to moderate hand-stirring withlake surface water (500 ml), followed by repetitivepassages (15 X) in and out of a wide-bore (0.5 cm), 50-ml plastic syringe. The resulting suspension had theappearance and consistency of pea soup. The suspen-sion was dispensed (50 ml) into 125-ml Erlenmeyerflasks (1 to 1.8 g of dry weight per flask), and the flaskswere sealed as described previously. Selected flaskswere flushed with N2 (flow, 150 ml min-1 for 5 min),injected with NH4Cl (100 mg), C2H2 (15 kPa), H2 (2kPa), NaNO3 (42.5 mg), or glucose (200 mg), or sealedwith HgCl2 (500 mg) as indicated. In some experi-ments, older CBAs (as evidenced by a brownish color,more fluid texture, and putrescent odor) found floatingwith uprooted Ruppia sp. were tested for their abilityto form H2. Flasks were incubated with reciprocalshaking (150 rpm) in the light or dark (foil-covered) atroom temperature (18 to 20°C). Headspace subsamples(0.25 or 0.5 ml) were stored in Glaspak syringes(Becton, Dickinson & Co.), with the needle endsinserted into large rubber bungs, until they wereanalyzed (<1.5 h after each sampling). Gases werequantified by gas chromatography as outlined previ-ously. At the end of the experiment, flask contentswere collected on filter paper (Whatman no. 1), driedat 50°C, and weighed.

Bacterial enrichment cultures. Enrichment culturesof anaerobic bacteria capable of H2 formation fromcyanobacterial cell material were recovered from theepiphytic CBAs of Big Soda Lake. Frozen (150 ml)Anabaena sp. aggregates were thawed and boiled in150 ml of tap water for 30 min. The resulting suspen-sion was centrifuged and decanted twice (7,500 x g)and then passed through a 1-,um filter (Millex; Milli-

pore Corp.) to remove particulates. The clear, amber-colored algal extract was mixed with Big Soda Lakesurface water (7:3, lake water-extract). The resultingmedium was dispensed (10 ml) into culture tubes (18by 150 mm; Bellco Co.), sealed with recessed butylrubber stoppers (no. 1; A. H. Thomas Co.), flushedwith N2 (120 ml min-1 for 2.5 min), placed in a tubepress, and sterilized by autoclaving (15 lb/in2, 20 min).Cultures were inoculated with 0.15 ml of dark-incubat-ed (N2 atmosphere), dispersed CBAs that had pro-duced H2 (see above). Inoculations were made at thetermination of the flask experiment (6 h of incubation).Tubes were flushed with sterile N2 (100 ml min-' for 4min) and incubated at 18°C with gentle shaking. Bacte-rial growth was measured at 550 nm (Spectronic 21colorimeter), with a sterile medium blank. Productionof H2 was measured by gas chromotography as out-lined above. Cultures were maintained by transfers(0.4 ml) at 2-week intervals.Chamber experiments. Estimates of in situ rates of

N2 fixation (C2H2 reduction) and H2 production bysubmerged, epiphytized Ruppia sp. beds were ob-tained by conducting experiments in Plexiglas cham-bers (Fig. 1). The chamber design (volume, 4 liters)allowed for insertion of the wider, tapered-base cylin-drical section (19 by 13 cm) into the sediments to a

B

FIG. 1. Design of benthic metabolic chambers forareal quantification of in situ N2 fixation and H2production. Chambers were constructed of Plexiglas(0.6-cm thickness).

1520 OREMLAND

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

HYDROGEN METABOLISM IN BIG SODA LAKE

>4 - 1.6-

0 1.2 -

EE

04 c,4 0~~.8-

0.4

00 1 2 3 0 1 2 3

Hours Hours

FIG. 2. N2 fixation and H2 production by cyano-bacterial communities epiphytic on Ruppia sp. plantsincubated in unshaken flasks during October 1981. (A)N2 fixation (C2H2 reduction) by epiphytes incubated inthe light (0), in the dark (0), and in the light withNH42+ (A). (B) H2 production by flasks incubated inthe light (A), in the dark (A), in the light with C2H2(0), and in the dark with C2H2 (0). All points repre-sent the mean of three experimental flasks, and barsindicate ±1 standard deviation. Flasks were shakenbriefly (15 s) before sampling.

depth of 3 cm. The chambers were held in place withlead diving weights (2.7 Kg). The smaller top cylindri-cal section (15.5 by 6.5 cm) was partially filled with agas mixture containing a Kr internal standard. Afteraddition of the gas mixtures to the chambers, all portswere sealed (except in C2H2 experiments, in whichone port was left open for 0.5 h to allow for waterreplacement of dissolved C2H2), and the gas phase wasbubbled several times through the liquid phase (tofacilitate equilibration) by means of a hand pump. Thisprocedure was followed immediately before each sam-pling. Initial gas phase compositions and volumeswere as follows: for N2-fixation experiments, air (200ml) + C2H2 (200 ml) + Kr (6 ml); for H2 production,N2 (400 ml) + Kr (6 ml). Replicate light and dark(covered with two layers of duct tape) chambers weredeployed in the littoral zone (depth, 1 m). Samples (6ml) of the chamber gas phases were withdrawn bysyringe (Glaspak, 10 ml; Becton, Dickinson & Co.)and were stored in 4-ml test tubes (10 by 75 mm) sealedwith size 000 black rubber stoppers. The tubes wereentirely filled with gas sample by displacement ofdistilled water with gas injected from the Glaspaksyringes. Tubes were stored inverted with 0.2 ml ofdistilled water overlying the internal portion of thestopper (to prevent leakage). Analysis of samples wasperformed within 12 h of sampling by proceduresoutlined previously. Sediments associated with theRuppia sp. beds contained gas bubbles. Analysis ofthe bubbles failed to detect any H2 or C2H4; however,CH4 levels varied from a trace (0.1%) to a majorcomponent (40%o by volume). Chambers were placedin a region of sediment where gas ebullition appearedto be low and was dominated by daytime photosyn-thetic 02 production by the plant/algal community.

FIG. 3. Production of H2 by dispersed cyanobac-terial aggregates incubated in the dark with constantshaking. Materials were incubated under N2 (0), air(A), or N2 with HgCl2 (A). In addition, dispersedcyanobacterial aggregates in a more advanced stage ofdecay were incubated in the dark under N2 (0). Allpoints represent the mean of three experimentalflasks, and bars indicate ±1 standard deviation (exceptat 2 h for older CBAs). Only one control flask withHgCl2 was run. Experiments were conducted duringOctober 1982.

RESULTSFlask experiments. Nitrogenase activity was

evident when pieces of the epiphyte-colonizedRuppia sp. plants were subjected to the C2H2-reduction assay (air, unshaken). Ethylene pro-duction was immediate and linear for light-incu-bated flasks (rate, 1.7 ,umol of C2H4 g-1 h-1),and production was inhibited by NH4' and bydark incubation (Fig. 2A). No C2H4 was formedin flasks (light or dark) incubated without C2H2(not shown). By contrast, H2 production oc-curred only in dark-incubated flasks (rate, 0.13,umol of H2 g-1 h-1) and H2 production rateswere stimulated by C2H2 (0.53 ,umol of H2 g-1h-1), whereas no production occurred in light-incubated flasks (with or without C2H2; Fig.2B).Dark incubation of dispersed CBAs under N2

with constant shaking resulted in the immediateand linear production of H2 (rate, 2.5 ,umol of H2g-1 h-1), and production was entirely inhibitedby 02 or by Hg2+ (Fig. 3). Addition of glucoseprovided no initial stimulation (2.4 ,umol of H2g-1 h-1 for 4.5 h); however, production ratesincreased after 4.5 h (4.4 p.mol of H2 g 1 h 1).This dark, anaerobic production of H2 was unaf-fected by addition of NH4' but was stimulated

VOL. 45, 1983 1521

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

APPL. ENVIRON. MICROBIOL.

TABLE 1. Consumption of H2 by aerobicallyincubated dispersed CBAsa

iLmol of H2 g-' dry wt Lmol of H2Time (h) per flaskb:

Lightc Darkc light + HgC12

0 66.9 (9.3) 59.6 (8.9) 53.3 (5.8)1.25 63.6 (5.1) 51.8 (8.1) 53.2 (1.1)3.25 55.7 (6.5) 45.6 (6.5) 53.9 (2.2)8.00 39.2 (6.4) 20.9 (2.5) 52.4 (0.8)a Flasks were incubated in the light or dark with

constant reciprocal shaking. Results represent themean of three experimental flasks, and the numbers inparentheses indicate 1 standard deviation. Dry weightswere 0.8 ± 0.06 g for light-incubated flasks and 0.96 ±0.04 g for dark-incubated flasks. Approximately 60,umol of H2 was added to flasks initially.

b Dry weights were not obtained for HgCI2-contain-ing flasks.

c Light H2 uptake rate = 3.6 ,umol g-1 h-1.d Dark H2 uptake rate = 4.8 ,umol g-1 h-1.

by C2H2 (data not shown). Incubation of dis-persed CBAs in the light under air resulted in alinear rate of C?H2 reduction to C2H4 (13.4 ,umolof C2H4 g 1 h- ), but H2 was not produced (withor without C2H2; data not shown). In addition,CBAs in a more advanced stage of decay (foundfloating free with uprooted, dying Ruppia sp.)produced H2 (dark, N2 atmosphere) at rates thatwere fourfold higher (10 FLmol of H2 g-1 h1)than material harvested from rooted macro-phytes (Fig. 3). Addition of NaNO3 (500 ,umolper flask) to dispersed CBAs (dark, unshaken,N2 atmosphere) caused a fivefold inhibition ofH2 production rates (e.g., 46 ,umol of H2 perflask h-1 without NO3- versus 8.8 ,umol of H2per flask h-1 with NO3-).When dispersed CBAs were incubated under

air plus H2, a linear rate of H2 disappearancewas observed (Table 1). Uptake of H2 in the light(rate, 3.6 ,umol of H2 g-1 h-1) was statisticallyequivalent to the dark H2 uptake rate (4.8 ,umolof H2 g-1 h-1). No H2 uptake occurred in Hg2+-killed controls (light incubated).Enrichment cultures. Inoculation of culture

media with 0.15 ml of the contents of one of the

TABLE 2. Growth and H2 production by ananaerobic bacterial enrichment culture.

Time H2 (4mol A55(h) per tube)0 0.00 0.0654 0.02 0.075

28 2.37 0.14052 10.06 0.120a A550, Absorbancy at 550 nm.

dark/N2 flasks (Fig. 3; inoculated at the end ofthe 6-h experiment) resulted in anaerobic bacte-rial growth and H2 production (Table 2). Similarresults were obtained when 0.15 ml of the mate-rial in a more advanced stage of decomposition(Fig. 3) was inoculated into identical media.Both enrichments were subsequently carriedwith transfers at 2-week intervals (dark, 28°C,unshaken). Growth (as indicated by increasedabsorbance at 550 nm) was essentially the sameas shown in Table 2; however, H2 productionrates increased fourfold, but final H2 concentra-tions achieved after 48 h were similar (12 ,umolof H2 per tube). Microscopic examination re-vealed a bacterial population of diverse mor-phologies (e.g., rods, cocci, spirilla, etc.). Nocyanobacterial cells were observed. Addition ofglucose (10 mM) to algal extract media stimulat-ed bacterial growth (absorbance at 550 nm =0.53 at 25 h) and H2 production (35 ,umol of H2per tube at 25 h). Addition of C2H2 (2.5 ml) to aculture did not enhance H2 production as com-pared with a culture incubated without C2H2(after 17 h of incubation, both cultures producedabout 11 ,umol of H2 per tube).Chamber experiments. Replicate light-incubat-

ed chambers reduced C2H2 to C2H4 but did notevolve H2 (Fig. 4A). The estimated mean rate ofC2H4 production was 310 jxmol of C2H4 m 2h-1. This value was obtained by assuming alinear production rate and multiplying the 5-hheadspace C2H4 concentrations (116 and 123,umol ml-1) by the final measured headspacevolumes (230 and 235 ml). The Bunsen coeffi-cient at 15°C for C2H4 (16) was applied to theequations of Flett et al. (8) to estimate dissolvedC2H4, which was calculated to be 69% of head-space C2H4. The chamber base area (293 cm2)was then extrapolated to a meter-squared basis.Dark chambers produced 16-fold less C2H4 thanlight chambers; however, increases in head-space H2 were observed during the incubation(Fig. 4B). Dark production rates of C2H4 wereestimated to be 20 ,mol of C2H4 m-2 h-1.Chambers incubated without C2H2 in the gas

phase (N2 + Kr atmosphere) did not accumulatesignificant levels of C2H4. Replicate dark-incu-bated chambers produced H2 without a lag;production rates were linear for 4 h (6.8 ,umol ofH2 m-2 h-1) and continued overnight (Fig. 5).By contrast, light-incubated chambers did notaccumulate H2 during the first 4 h of incubation.Significant levels of H2 were noted in "light"chambers only after an overnight (dark) incuba-tion period- Methane also increased in the gasphases of chambers (linear for the first 4 h), andmean areal production rates for dark chambers(32.2 pumol of CH4 m2 h1) were about seven-fold greater than light-chamber rates (4.8 ,umolof CH4 m-2 h-1).

1522 OREMLAND

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

HYDROGEN METABOLISM IN BIG SODA LAKE 1523

A B S

120-12

E- E~~~~~~~-~- 10

E=r~2H.j.-i~8

40~~~~~

1 2 3 4 5 1 2 3 4

Hours

FIG. 4. N2 fixation (C2H2 reduction), H2 produc-tion, and internal standard Kr concentrations for ben-thic chambers incubated in Big Soda Lake duringOctober 1982. Chambers were deployed at 11 a.m. ona partly cloudy day. (A) Replicate chambers incubatedin the light: levels of gas-phase C2H4 (0, 0), H2 (°,*), and Kr (A, A). (B) Replicate chambers incubatedin the dark: levels of gas-phase C2H4 (0, 0), H2 (°,U), and Kr (A, A). Initial gas phases of all chambersconsisted of air (200 ml), C2H2 (200 ml), and Kr (6 ml).

DISCUSSION

A considerable amount of research has beendevoted to the study of hydrogen metabolism(production and consumption) in cyanobacteriafor the ultimate purpose of biological fuel pro-duction. These studies have been conductedwith pure cultures, isolated heterocysts, andvarious hydrogenase enzyme systems. By con-trast, there have been few investigations ofhydrogen metabolism in nature in environmentsother than anaerobic sediments, rumens, orroot-nodule associations. Although the potentialfor aerobic cyanobacterial hydrogen productionexists, this has not as yet been demonstrated tooccur in natural habitats (5). Interpretation ofresults obtained with free-living cyanobacterialpopulations can be clouded by possible contri-bution(s) to hydrogen metabolism made by othermicrobes (e.g., bacteria) present within thecyanobacterial matrix. Thus, the approach takenin this study was to attempt to experimentally"dissect" the Big Soda Lake cyanobacterialaggregates and, if possible, assign the responsi-bility for hydrogen metabolism to a particulargroup of microorganisms.

Production or consumption of hydrogen ob-served in these experiments was an actual envi-ronmental occurrence and not an artifact of theincubation conditions. This was borne out by theobservations that both production and uptake ofhydrogen did not display lag phases and thatrates were linear during relatively short (a fewhours) incubations in either flasks (Fig. 2B and3; Table 1) or in situ chambers (Fig. 5). In

HoursFIG. 5. H2 production by replicate dark- and light-

incubated chambers deployed in Big Soda Lake duringOctober 1982. Gas phase consisted of N2 (400 ml) plusKr (6 ml). Kr values decreased by 30% during theincubation. Chambers were deployed at 3 p.m. on apartly cloudy day. Chambers were incubated in thelight (A, 0) and dark (A, 0). After overnight (19 h)incubation, final H2 levels (nmol ml-') were: A, 21; 0,6.2; A, 1.8; 0, not determined.

addition, hydrogen production and uptake wasblocked by Hg2+, thereby indicating its biologi-cal nature (Fig. 3; Table 1).Hydrogen production by the CBAs did not

appear to be a function of the Anabaena sp.hydrogenase or nitrogenase system. No H2 pro-duction occurred in the light, and productionwas only observed in the dark (Fig. 2B, 3, and5). Hydrogen production by the nitrogenasesystem of Anabaena cylindrica is a light-depen-dent phenomenon (7, 12, 14, 15, 19). Further-more, although NH4' inhibited N2 fixation bythe CBAs (Fig. 2A), it did not inhibit dark H2production. In addition, C2H2 did not inhibit(but instead stimulated) dark H2 production(Fig. 2B). If dark H2 production were due tonitrogenase, then C2H2 should act as an electronsink and less H2 should be formed in its presencethan in its absence (19, 21). Additionally, aero-bic photoproduction of H2 by A. cylindricarequires either C2H2 or CO to block uptakehydrogenases (7, 12, 14, 15). However, no H2production was observed by CBAs incubatedaerobically in the light with C2H2 (Fig. 2A).Finally, dark H2 production by anoxygeniccyanobacteria requires a supply of reductant,such as sodium dithionite, reduced methyl violo-gen, or, perhaps, H2S (3). None of these sub-stances was added to the incubation flasks, andthe flasks had no sulfide odor at the end of theexperiment. Thus, the results obtained for H2production by Big Soda Lake CBAs do not fitthe pattern expected for nitrogenase- or hydro-genase-linked H2 evolution by cyanobacteria.

Instead, evidence indicates that the H2 pro-

VOL. 45, 1983

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

1524 OREMLAND

duction observed in these experiments was aconsequence of the bacterial fermentation ratherthan cyanobacterial activity. Hydrogen produc-tion occurred only in the dark (Fig. 2B) andrequired anaerobic conditions (Fig. 3). Inhibi-tion of H2 production in the light (Fig. 2A) wastherefore probably caused by the production ofphotosynthetic 02 by the cyanobacteria. Theproduction of H2 in the dark under air occurredin unshaken flasks (Fig. 2B), and these condi-tions approximate actual in situ conditions expe-rienced by the CBAs at night. Thus, constantrapid shaking under air prevents the establish-ment of anaerobic microniches within the cyano-bacterial matrix. Additional evidence for bacte-rial involvement includes the higher rates of H2production associated with CBAs in a moreadvanced stage of decomposition (Fig. 3) andthe recovery of anaerobic enrichment culturescapable of growth on and H2 production fromcyanobacterial extract (Table 2). Finally, addi-tion of nitrate to dispersed aggregates caused afivefold inhibition of H2 production. This sug-gests potential coupling of bacterial electrontransport to dissimilatory nitrate reduction ordenitrification in lieu of hydrogenase (when ni-trate is available), as occurs in decomposingperiphyton (25, 26). However, production ofN20 did not occur during incubation of CBAs inthe presence of C2H2 with or without addednitrate (data not shown).The stimulation of dark H2 production by

C2H2 was an unexpected, yet consistent resultof these experiments. The mechanism(s) bywhich this occurs is beyond the scope of thispaper, but perhaps takes place at the bacteriallevel, possibly by inhibition of metalloenzymesas is believed to occur in methylotrophs (1).However, C2H2 did not stimulate H2 productionby the enrichment culture recovered from theCBAs. Thus, either the C2H2 stimulation iscaused by microbes other than the ones presentin the enrichment, or the culture conditions ofthe enrichment somehow preclude the C2H2effect. Aerobic photoproduction of H2 by A.cylindrica requires C2H2, CO, or both (7, 12, 14,15). Acetylene also inhibits aerobic uptake hy-drogenases of diazotrophic bacteria (24), but notof Anabaena sp. strain 7120 (11). Thus, themolecular basis of the C2H2 requirement forphotoproduction of H2 by heterocystous cyano-bacteria is also unknown.Aerobic uptake of H2 by the CBAs may have

been caused by consumption by bacteria (e.g.Alcaligenes), by uptake hydrogenases associat-ed with heterocysts, or both (11). Results of thechamber experiments indicate that net dark H2production rates (Fig. 5) are low (6.8 ,umol of H2m-2 h-1), especially as compared to the light-driven rates of C2H2 reduction (Fig. 4A), which

were much higher (310 ,umol of C2H4 m-2 h-1).Because potential rates of anaerobic H2 produc-tion and aerobic H2 consumption are of equalmagnitude (Fig. 3; Table 1), the net amount ofH2 released by these cyanobacterial communi-ties will be governed by the abundance of 02within the matrix. Oxygen both inhibits H2 pro-duction by fermentative bacteria and enablesheterocysts and H2-oxidizing bacteria to con-sume any H2 produced. These two effects of 02will therefore tend to keep in situ net H2 produc-tion rates low and well below the gross H2production rates observed in flask experiments.The results presented in this paper suggest that arapid shift from aerobic to anaerobic environ-ments occurs within these matrices as a functionof diurnal photosynthetic activity and respira-tion. Hydrogen production by fermentative bac-teria associated with decomposing algal materialmay be of importance in providing H2 to this andother aquatic environments, such as brackishponds (M. Scranton, P. A. Loud, and P. C.Novelli, Eos Trans. Am. Geophys. Un. 63:990,1982) and oceanic surface waters (S. J. Schroppand J. R. Schwarz, Abstr. Annu. Meet. Am.Soc. Microbiol. 1982, N93, p. 193).

ACKNOWLEDGMENTS

I am grateful to C. Culbertson for technical assistance, and Ithank R. L. Smith, B. F. Taylor, and M. Firestone for helpfuldiscussions and manuscript review.

LITERATURE CITED1. Anthony, C. 1982. The biochemistry of methylotrophs.

Academic Press, Inc., New York.2. Axler, R. P., R. M. Gersberg, and L. J. Paulson. 1978.

Primary productivity in Big Soda Lake, Nevada. GreatBasin Nat. 38:187-192.

3. Belkin, S., and E. Padan. 1978. Hydrogen metabolism inthe facultative anoxygenic cyanobacteria (blue-green al-gae) Oscillatoria limnetica and Aphanotece halophytica.Arch. Microbiol. 116:109-111.

4. Bullister, J. H., N. L. Guinasso, Jr., and D. R. Schink.1982. Dissolved hydrogen, carbon monoxide, and meth-ane at the CEPEX site. J. Geophys. Res. 87:2022-2034.

5. Conrad, R., M. Aragno, and W. Seiler. 1983. Productionand consumption of hydrogen in a eutrophic lake. Appl.Environ. Microbiol. 45:502-510.

6. Culbertson, C. W., A. J. B. Zehnder, and R. S. Orem-land. 1981. Anaerobic oxidation of acetylene by estuarinesediments and enrichment cultures. Appl. Environ. Mi-crobiol. 41:396-403.

7. Daday, A., R. A. Platz, and G. D. Smith. 1977. Anaerobicand aerobic hydrogen gas formation by the blue-green algaAnabaena cylindrica. Appl. Environ. Microbiol. 34:478-483.

8. Flett, R. J., R. D. Hamilton, and N. E. R. Campbell. 1976.Aquatic acetylene-reduction techniques: solutions to sev-eral problems. Can. J. Microbiol. 22:43-51.

9. Herr, F. L., and W. R. Barger. 1978. Molecular hydrogenin the near-surface atmosphere and dissolved in waters ofthe tropical North Atlantic. J. Geophys. Res. 83:6199-6205.

10. Herr, F. L., M. I. Scranton, and W. R. Barger. 1981.Dissolved hydrogen in the Norwegian Sea: mesoscalesurface variability and deep water distribution. Deep-SeaRes. 28:1001-1016.

APPL. ENVIRON. MICROBIOL.

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

HYDROGEN METABOLISM IN BIG SODA LAKE 1525

11. Houchins, J. P., and R. H. Burris. 1981. Light and darkreactions of the uptake hydrogenase in Anabaena 7120.Plant Physiol. 68:712-716.

12. Houchins, J. P., and R. H. Burris. 1981. Physiologicalreactions of the reversible hydrogenase from Anabaena7120. Plant Physiol. 68:717-721.

13. Kimmel, B. L., R. M. Gersberg, L. J. Paulson, R. P.Axler, and C. R. Goldman. 1978. Recent changes in themeromictic status of Big Soda Lake, Nevada. Limnol.Oceanogr. 23:1021-1025.

14. Lambert, G. R., A. Daday, and G. D. Smith. 1979. Effectsof ammonium ions, oxygen, carbon monoxide, and acety-lene on anaerobic and aerobic hydrogen formation byAnabaena cylindrica B629. Appl. Environ. Microbiol.38:521-529.

15. Lanbert, G. R., A. Daday, and G. D. Smith. 1979. Dura-tion of hydrogen formation by Anabaena cylindrica B629in atmospheres of argon, air, and nitrogen. Appl. Environ.Microbiol. 38:530-536.

16. Lange, N. A. 1967. Handbook of chemistry, 10th ed.Handbook Publishers, Sandusky, Ohio.

17. Oremland, R. S. 1981. Microbial formation of ethane inanoxic estuarine sediments. Appl. Environ. Microbiol.42:122-129.

18. Oremland, R. S., L. Marsh, and D. J. Des Marais. 1982.Methanogenesis in Big Soda Lake, Nevada: an alkaline,moderately hypersaline desert lake. Appl. Environ. Mi-crobiol. 43:462-468.

19. Peterson, R. B., and R. H. Burris. 1978. Hydrogen metab-

olism in isolated heterocysts of Anabaena 7120. Arch.Microbiol. 116:125-132.

20. Priscu, J. C., R. P. Axler, R. G. Carlton, J. E. Reuter,P. A. Arneson, and C. R. Goldman. 1982. Vertical profilesof primary productivity, biomass and physico-chemicalproperties in meromictic Big Soda Lake, Nevada, U.S.A.Hydrobiologia 96:113-120.

21. River-Ortiz, J. M., and R. H. Burris. 1975. Interactionsamong substrates and inhibitors of nitrogenase. J. Bacteri-ol. 123:537-545.

21a.Scranton, M. I. 1983. The role of the cyanobacteriumOscillatoria (Trichodesmium) thiebautii in the marinehydrogen cycle. Mar. Ecol. Prog. Ser. 11:79-87.

22. Scranton, M. I., M. M. Jones, and F. L. Herr. 1982.Distribution and variability of dissolved hydrogen in theMediterranean Sea. J. Mar. Res. 40:873-891.

23. Sester, P. J., J. L. Bullister, E. C. Frank, N. L. Guinasso,Jr., and D. R. Schink. 1982. Relationships between re-duced gases, nutrients and fluorescence in surface watersoff Baja California. Deep-Sea Res. 29:1203-1216.

24. Smith, L. A., S. Hill, and M. G. Yates. 1976. Inhibition byacetylene of conventional hydrogenase in nitrogen-fixingbacteria. Nature (London) 262:209-210.

25. Trlska, F. J., and R. S. Oremland. 1981. Denitrificationassociated with periphyton communities. Appl. Environ.Microbiol. 42:745-748.

26. Ventullo, R. M., and J. J. Rowe. 1982. Denitrificationpotential of epilithic communities in lotic environment.Curr. Microbiol. 7:29-33.

VOL. 45, 1983

on Novem

ber 24, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from