TORI M. HOEHLER,1 MARY HOGAN,4 LINDA L. JAHNKE,1 RICHARD M. KELLER,5
SCOTT R. MILLER,6 LESLIE E. PRUFERT-BEBOUT,1 CHRIS RALEIGH,7
MICHAEL ROTHROCK,3 and KENDRA TURK4
ABSTRACT
Photosynthetic microbial mat communities were obtained from marine hypersaline salternponds, maintained in a greenhouse facility, and examined for the effects of salinity varia-tions. Because these microbial mats are considered to be useful analogs of ancient marinecommunities, they offer insights about evolutionary events during the .3 billion year timeinterval wherein mats co-evolved with Earth’s lithosphere and atmosphere. Although photo-synthetic mats can be highly dynamic and exhibit extremely high activity, the mats in the pre-sent study have been maintained for .1 year with relatively minor changes. The major groupsof microorganisms, as assayed using microscopic, genetic, and biomarker methodologies, areessentially the same as those in the original field samples. Field and greenhouse mats weresimilar with respect to rates of exchange of oxygen and dissolved inorganic carbon across themat–water interface, both during the day and at night. Field and greenhouse mats exhibitedsimilar rates of efflux of methane and hydrogen. Manipulations of salinity in the water over-lying the mats produced changes in the community that strongly resemble those observed inthe field. A collaboratory testbed and an array of automated features are being developed tosupport remote scientific experimentation with the assistance of intelligent software agents.This facility will permit teams of investigators the opportunity to explore ancient environ-mental conditions that are rare or absent today but that might have influenced the early evo-lution of these photosynthetic ecosystems. Key Words: Microbial mat—Biogeochemistry—Biomarkers. Astrobiology 2, 383–402.
383
1Exobiology Branch and 5Computational Sciences Division, NASA Ames Research Center, Moffett Field, Califor-nia.
2SETI Institute, Mountain View, California.3Department of Microbiology, Arizona State University, Tempe, Arizona.4Institute of Marine Sciences, University of California, Santa Cruz, California.6Department of Genetics, North Carolina State University, Raleigh, North Carolina.7Montana State University, Bozeman, Montana.
INTRODUCTION
MODERN MICROBIAL MATS are thought to be ex-tant representatives of Earth’s most ancient
ecosystems (Walter, 1976). Geochemical evidenceof the existence of photosynthetic microbial mats,and their mineralized counterparts, stromatolites,has been identified in rocks as old as 3.0 Ga(Beukes and Lowe, 1989). As a living repositoryof genetic, physiological, isotopic, and biogeo-chemical information on the co-evolution of aplanet and the only known biosphere, modernmicrobial mats are invaluable objects of study.Modern microbial mat studies have provided im-portant insights on rates of biological activity(Revsbech et al., 1983; Canfield and Des Marais,1993; Des Marais, 1995), genetic diversity (Wardet al., 1990; Garcia-Pichel et al., 1998; Nübel et al.,2001), stable isotopic fractionation (Schidlowski,1988; Des Marais and Canfield, 1994), and organic(Boon, 1984; Ward et al., 1985) and atmospheric(Visscher and Van Gemerden, 1991; Visscher andKiene, 1994; Hoehler et al., 2001) biomarkers, aswell as minerals (Reid et al., 2000), that have beenused to interpret the fossil record of these com-munities over geologic time.
Photosynthetic microbial mats are self-sustain-ing, complete ecosystems in which light energyabsorbed over a diel (24-h) cycle drives the syn-thesis of spatially organized, diverse biomass. Mi-croorganisms with tightly coupled metabolismsin the mat catalyze transformations of carbon, ni-trogen, sulfur, and metals. Radiant energy fromthe sun sustains oxygenic and anoxygenic pho-tosynthesis, which in turn provides chemical en-ergy (as organic photosynthates and oxygen) tothe rest of the community. When oxygenic pho-tosynthesis ceases at night, the upper layers of themat become highly reduced and sulfidic (Jør-gensen et al., 1979). Counteracting gradients ofoxygen and sulfide shape the chemical environ-ment and provide daily-contrasting microenvi-ronments separated on a scale of a few millime-ters (Revsbech et al., 1983; Revsbech andJørgensen, 1986). While photosynthetic bacteriadominate the biomass and productivity of themat, many aspects of the ecosystem’s emergentbehavior may ultimately depend on the associ-ated nonphotosynthetic microbial communities,including the anaerobes. Additionally, transfor-mation of photosynthetic productivity by the mi-crobial community may contribute diagnostic“biosignature” gases that could represent search
targets for remote spectroscopic life detection ef-forts [e.g., Terrestrial Planet Finder (Des Maraiset al., 2002)]. To understand the overall structureand function of mat communities, it is thus criti-cal to determine the nature and extent of the in-teractions between phototrophic and nonphoto-synthetic microorganisms, including anaerobicmicroorganisms.
Compared with their historical distributionand abundance, microbial mats are today con-fined to a relatively restricted number of habitats.Well-developed modern photosynthetic mats oc-cur in both high-salinity and high-temperatureenvironments. This restricted distribution isthought to be due primarily to the higher rates ofgrazing in more moderate modern environments(Garret, 1970), although alternative hypotheseshave been forwarded. In any event, the limitedgeographical distribution of mats on the present-day Earth also serves to limit their usefulness asanalogs of similar ecosystems growing in morediverse ancient environments. Environments thatare not well represented on the modern Earth, butthat have been extremely important over evolu-tionary time, include marine environments hav-ing low concentrations of sulfate and/or free oxy-gen (Holland, 1984; Canfield and Teske, 1996).
The effects on microbial processes of variousenvironmental parameters important in Earth’spast (such as sulfate and oxygen concentrations)may be studied in culture. However, the emer-gent properties of complex microbial ecosystemsare not necessarily predictable, or understand-able, on the basis of these types of experiments(Wilson and Botkin, 1990). Either some microbialprocesses do not occur in culture [e.g., anaerobicmethane oxidation (Reeburgh, 1980) and sulfatereduction under aerobic conditions (Canfield andDes Marais, 1991)], or they occur at rates vastlydifferent than rates observed in nature. In addi-tion, relatively few (,1%) of the total number ofmicrobes present in nature are available in cul-ture (Ward et al., 1990; Amann et al., 1995). A mi-crobial mat is a highly complex assemblage of organisms possessing many different modes ofmetabolism, all of which are interacting with eachother at some level in beneficial and/or compet-itive ways. The end products of the metabolismof one group of organisms frequently constitutethe substrates of another. These considerationsargue for an alternative approach in which theentire microbial ecosystem is subjected to exper-imental manipulation.
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To attain several key objectives in astrobiology,we must understand better the mechanisms andrates at which trace gases are produced and con-sumed by microbial ecosystems. As dominantcomponents of our biosphere for at least 2 billionyears of its .3.5 billion year history, microbialmats played a pivotal role in shaping the com-position of Earth’s early atmosphere, including itseventual oxygenation. NASA’s present searchstrategy for the detection of life on extrasolarplanets includes the observation and interpreta-tion of spectral features of biogenic gases in re-mote atmospheres (Des Marais et al., 2002). How-ever, the current strategy relies principally uponthe detection of O2, and thus might preclude thedetection of life that might resemble our own pre-oxygenated biosphere for as much as 2 billionyears of its early existence. Clearly, the rates ofproduction and consumption of reduced gases bymicrobial communities over geologic time mustbe better constrained to enhance our search forlife elsewhere. The use of simulation facilities,such as the greenhouse described here, will aug-ment that endeavor.
We report here our attempts to maintain intactmodern photosynthetic microbial mats under ap-proximate in situ conditions, and to monitor theresponses of these mats to experimental manipu-lations. We have constructed a facility in whichphotosynthetic microbial mats can be maintainedover long periods of time, using natural illumi-nation and realistic water flow and other envi-ronmental conditions. We describe here the re-sults of our first experiment, in which we usedthis facility to manipulate the salinity of wateroverlying the mats. As variations in salinity alsooccur in the natural environment where thesemats were collected, we are able to assess the de-gree to which our experimental facility can re-produce naturally occurring phenomena. Futureuses of this experimental system, in which con-ditions not presently found in the modern envi-ronment will be simulated, are discussed.
MATERIALS AND METHODS
Field site
Microbial mats were collected in salterns man-aged by the salt-producing company Exporta-dora de Sal. S.A. de C.V., located on the PacificOcean side of the Peninsula of Baja California Sur,
Mexico. This field site was previously describedin detail (Des Marais, 1995). Briefly, lagoon wa-ter, having a salinity of ,40‰ (parts per thou-sand), is pumped through a series of concentrat-ing ponds, and then into crystallizing ponds. Thesalinity of the water is gradually raised until itbecomes saturated with respect to sodium chlo-ride, at which point the brine is pumped out ofthe crystallizing pond, and the salt is harvested.Extensive and well-developed microbial mats oc-cur in concentrating areas in which the salinity isbetween ,65‰ and 130‰.
Microbial mats were collected on May 27, 2000from two localities: Area 4, near the dike sepa-rating Area 4 from Area 5 (27°41.34509N,113°55.02709W), and Area 5, near the dike sepa-rating Area 5 from Area 6 (27°44.41409N,113°54.79509W). At the time of the collection, thesalinities of Area 4 and Area 5 were ,90‰ and,120‰, respectively. Microbial mats in both ofthese locations are ,5 cm thick and well lami-nated. The dominant cyanobacterium of the matcommunity is Microcoleus chthonoplastes. Sectionsof mat ,20 3 25 cm were cut and removed fromthe bottom of the concentrating pond by diversusing metal spatulas and were immediatelyplaced into tight-fitting black acrylic trays. In thisway, exposure of the deeper anaerobic layers ofthe mats to air and light was minimized. Matswere covered with relatively high-salinity water(180‰) overnight in an effort to slow overallmetabolic rates for transport. The trays contain-ing the mats were then transported by vans backto our laboratory in larger plastic trays coveredby tight-fitting plastic film. In this way, the matswere kept moist but not covered with water, andwere exposed to some natural light over the ,48h required for the relocation. Although six matsfrom Area 5 and 12 mats from Area 4 were main-tained in the greenhouse throughout the entireexperiment, results from only the Area 4 matswill be presented here.
Greenhouse facility
Upon arrival at Ames Research Center, themats were transferred to a greenhouse modifiedfor these experiments by replacing the originalglass with UV-transparent OP-4 acrylic (trans-mission in the UV-B, UV-A, and visible ,90% inthe greenhouse). Mats were placed onto a spe-cially built table (Fig. 1) having six clear acrylicflow boxes (150 3 22 cm), each flow box holding
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FIG. 1
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three trays of mat. The depth of the water in theflow boxes can be adjusted with standpipes. Forthis experiment, the depth of the water overlyingthe mats was ,3 cm; therefore the total volumeof the water in each flow box at any one time was,10 L.
Brine used for the experiment was collectedfrom one of the higher-salinity (,130‰) concen-trating areas that had retained its original com-plement of seawater solutes, and diluted withdeionized water to the appropriate salinity. Matswere arbitrarily assigned to one of two salinitytreatments on the table. Three of the flow boxesrecirculated brine collected from the field site,which was diluted to the in situ salinity of 90‰(hereafter referred to as the NORMAL salinitytreatment). The other three flow boxes recircu-lated brine having an elevated salinity of 120‰(hereafter referred to as the HIGH salinity treat-ment). Water was recirculated from a single reser-voir holding 60 L of brine through the three in-terconnected flow boxes that constituted each ofthe two salinity treatments.
Environmental control
The flow boxes assigned to each of the salinitytreatments were interspersed on the table, so thatthe treatments alternated (e.g., NORMAL, HIGH,NORMAL, etc.). Salinity was maintained at theselevels over the course of the experiment with ad-ditions of deionized water to replace losses dueto evaporation. Temperature control of the air inthe greenhouse was achieved with a combinationof an evaporative cooler and propane heater. Precise temperature control of the water being re-circulated over the mats was achieved by circu-lating temperature-controlled water through tita-nium heat-exchanging coils submerged in thereservoirs. Normal in situ daily water columntemperature variations were simulated in thegreenhouse by (1) controlling the temperature in-crease (attributable to solar heating) during thedaytime to stay below the maximum temperatureobserved in situ and (2) turning off temperaturecontrol at night to allow the water temperaturein the flow boxes to decrease slowly with the de-crease in greenhouse air temperature. This simu-lation of diel temperature variations, while not always completely faithful to the precise near-si-nusoidal pattern of temperature change observedin situ (owing to the simplicity of the apparatus),reproduced the normal range of temperatures ob-
served in situ and the approximate duration oftime at each temperature. The similarity betweengreenhouse and in situ temperatures was docu-mented by an extensive set of measurements per-formed the year after this experiment was run.However, since no changes had been made to theapparatus since the time of the experiments re-ported here, this comparison should be valid forthe period of time discussed here. Using sub-mersible data loggers (StowAway TidbiT, OnsetComputer Corp., Bourne, MA), recording at 10-min intervals from June 2001 through September2001, the average temperature in the salterns was22.8 6 1.99°C (mean 6 SD, for n 5 16,914),whereas water temperatures in the greenhouseover the same interval of time were 20.3 6 2.62°C(n 5 16,191). Water column salinity in the flowboxes was determined using a refractometer(Delta model, Bellingham 1 Stanley, TunbridgeWells, UK), which was corrected for temperatureeffects at each reading. Light [as photosyntheti-cally available radiation (PAR)] was monitoredand averaged at 10-min intervals, using a datalogger coupled to a terrestrial quantum sensor(LI-1000 and LI-192SA, LI-COR, Inc., Lincoln,NE). An underwater light sensor (LI 192SA, LI-COR, Inc.) was used to measure irradiance at themat surface in situ on several field trips. The irradiance at the mat surface, expressed as a per-centage of the irradiance incident upon the sur-face of the pond, is comparable with measure-ments of irradiance in the greenhouse.
Greenhouse oxygen microelectrode measurements
Oxygen concentrations within the microbialmats were monitored at various times through-out the course of the experiment using oxygenmicroelectrodes. We used Clark-type microelec-trodes incorporating guard cathodes (model 737-GC, Diamond General Development Corp., AnnArbor, MI). In the initial phase of the experiment,microelectrodes were positioned using motorizedmicromaniupulators mounted to bases placedacross the flow boxes. Halfway through the ex-periment, the microsensors were transitioned toa positioning system using a robotic xyz posi-tioning system installed over the flow boxes (Fig.1). In all cases, the positioning was controlled by,and data were acquired with, custom softwarewritten in the LabVIEW programming environ-ment (National Instruments Corp., Austin, TX).
In general, the electrodes were advanced
A MICROBIAL MAT GREENHOUSE COLLABORATORY 387
within the mat at 100-mm steps. Oxygen micro-electrode signal output was calibrated to oxy-gen concentrations using a two-point calibra-tion. Because the water circulating through theflow boxes was constantly aerated through theaction of the pumps and through contact withthe atmosphere across a large surface area ofwater, the electrode current at any point in thewater overlying the mats was taken to be equalto the current produced in air-saturated waterat that particular temperature and salinity. Theexact value of this oxygen concentration canthen be identified using published values (Sher-wood et al., 1991). The second point in the cali-bration, the output of the electrode at an oxy-gen concentration of 0, was provided by theasymptotic minimum of electrode currentwithin the permanently anoxic lower parts ofthe mat.
In situ microelectrode measurements
Measurements of in situ concentrations of oxy-gen within the microbial mat were made usinga diver-operated microprofiler (Unisense,Århus, Denmark) at 100-mm intervals. Calibra-tions of the oxygen electrode were performed asdescribed for greenhouse microelectrodes(above). As the ponds are well mixed duringwindy periods of the day, electrode readings inthe well-mixed portions of the water columnabove the mat surface were taken to representair-saturated values at a given temperature andsalinity. In the morning, bottom water can below in oxygen and, therefore, was not used forelectrode calibrations; rather, readings closer tothe air–water interface were used to calibrate theelectrodes. In all cases, measurements of the as-ymptotic minimum in electrode current withinthe permanently anoxic lower parts of the matwere used as the zero oxygen concentration cal-ibration point.
Community composition analyses
Light microscopy. Microbial community compo-sition was followed throughout the course of theexperiment using a variety of techniques. First-order observations about the community weremade using light microscopy (Nikon MicrophotFX/A, Nikon USA). We did not employ quanti-tative microscopy techniques, and so these ob-servations will be referred to as “nonquantita-tive” microscope observations.
Cyanobacterial 16S rRNA gene fingerprints bypolymerase chain reaction (PCR)/denaturing gradientgel electrophoresis (DGGE). Cores (10 mm in di-ameter) were taken from the microbial mats onSeptember 15, 2000, and the photic zone (top 3mm) was aseptically removed. The samples werehomogenized in a Tenbroeck tissue grinder(VWR Scientific Products, Brisbane, CA). Cell ly-sis and DNA extraction were performed as pre-viously described (Nübel et al., 1997). In short, thehomogenized samples were frozen and thawedrepeatedly, followed by incubation with 1%sodium dodecyl sulfate and proteinase K. DNAextraction was performed using hexadecyl-methylammonium bromide in conjunction withphenol/chloroform/isoamyl alcohol (25:24:1 byvolume), followed by isopropyl alcohol precipi-tation. Cyanobacterial/plastid-specific PCR am-plification was carried out using the oligonu-cleotide primers CYA359F (with a 40-nucleotideGC-rich sequence at the 59 end) and CYA805R,which were used for specific amplification of 16SrRNA genes from cyanobacteria and plastids(Nübel et al., 1997). Each 50-mL PCR mixture con-tained 5 mL of 103 Takara Ex Taq™ PCR buffer(PanVera Corp., Madison, WI), 4 mL of Takara de-oxynucleotide triphosphate mixture (2.5 mMeach), 50 pmol of each primer, 200 mg of bovineserum albumin, 10 mL of 53 Eppendorf TaqMas-ter™ PCR-enhancer (Brinkmann Instruments,Inc., Westbury, NY), and 15 ng of DNA extract.Thermocycling was performed using a Bio-RadiCycler™ thermal cycler (Bio-Rad Laboratories,Hercules, CA). After an initial denaturation at94°C for 5 min (hot start), 2.5 units of Takara ExTaq DNA polymerase was added to the reactionat 80°C. Thirty-five cycles of 1 min each at 94°C(denaturation), 60°C (annealing), and 72°C (ex-tension) were performed, and the reaction fin-ished with a final extension at 72°C for 9 min.Gels were imaged using a Bio-Rad Fluor-S™ Mul-tiImager system. For DGGE band sequencing,each band was excised using a sterile scalpel, andDNA was allowed to diffuse out for $3 days at4°C in 50 mL of 10 mM Tris buffer. One micro-liter of the solution was PCR-amplified using thesame primers, reaction mixture, and thermocy-cling conditions. A kit was used to purify PCRproduct (Qiagen, Inc., Valencia, CA) and 150 ngwas commercially sequenced in two separate re-actions (59 to 39 and 39 to 59). Complementary se-quences were matched, aligned, and edited usingSequence Navigator (Applied Biosystems, Foster
BEBOUT ET AL.388
City, CA) and submitted to the BLAST search en-gine (National Center for Biotechnology Infor-mation; www.ncbi.nlm.nih.gov) for phylogeneticmatching.
Lipid biomarker analysis. Gas chromatographicprofiles of fatty acids (FA), as fatty acid methylesters (FAME), were used to monitor changes ingroup-specific biomarker compounds in the sur-face layers of the greenhouse and field mat com-munities. A section (,25 cm2 and 1 cm in depth)was removed from a mat in each of the six flowboxes after 4 months, dissected, frozen, andfreeze-dried. A similar-sized section was re-moved from freshly collected mats at the field sitein Guerrero Negro in both December 1998 andJune 2001. The sections from the field-collectedmats were dissected shortly after collection,frozen, and returned to NASA Ames ResearchCenter where they were freeze-dried andprocessed.
Field and greenhouse mat communities weredissected and processed as follows. The floccu-lent orange material on the surface of the mat wasremoved by scraping the surface with a spatula.An exposed leathery layer of the mat, containingextremely high densities of the filamentouscyanobacterium M. chthonoplastes (hereaftercalled the Microcoleus layer), ,1–1.5 mm thick,was then separated using a razor blade from thedarker-colored gelatinous material that composesthe undermat. The freeze-dried mat layers wereground using a glass mortar and pestle. Totallipid extracts were prepared by a Bligh and Dyersingle-phase modification (Jahnke et al., 1992).Multiple extractions of the cell material recoveredby centrifugation were done until no further pig-ment coloration was apparent (normally four orfive times). A polar fraction (phospholipids andsome glycolipids) was prepared by precipitationin cold acetone. The remaining acetone-solublelipids were separated by thin-layer chromatogra-phy using a methylene chloride solvent system(Jahnke et al., 1992). The remaining glycolipidspresent in the origin zone (0–1 cm) of the thin-layer chromatography plate and the sterols(,2.5–3.5 cm) were recovered from the silica gelby Bligh and Dyer extraction. The polar lipid frac-tions were pooled, and FAME were prepared bya mild alkaline methanolysis procedure (Jahnkeet al., 2001). Gas chromatography–mass spec-trometry analyses of FAME or sterols as tri-methylsilyl esters were performed using an HP
5890 gas chromatograph equipped with a J&WDB-5ms (30 m 3 0.25 mm, 0.25-mm film) capillarycolumn and an HP 5971 mass-selective detector.Identification of individual compounds was by acombination of standard compounds, publishedspectra, and relative retention times. FAME werequantified using an Agilent 6890 gas chromato-graph equipped with a flame ionization detectorand HP-5 (30 m 3 0.32 mm, 0.25-mm film) capil-lary column. Methyltricosanoate (FAME) wasused as an internal standard. Chromatographswere programmed to operate for FAME from60°C to 120°C at 10°C/min, then at 2°C/min from120°C to 280°C. Total organic carbon (TOC) andstable isotope composition (d13C and d15N) forbiomass was determined using a Carlo Erba CHNEA1108 elemental analyzer interfaced to a Finni-gan Delta Plus XL isotope ratio mass spectrome-ter (Jahnke et al., 2001).
Fluxes of oxygen estimated by microsensor measurements
Net fluxes of oxygen across the mat surface (J)were calculated according to Fick’s first law ofdiffusion (Berner, 1980):
J 5 2fDs (C/z)
where C/z is the change in oxygen concentra-tion with depth near the mat–water interface in(mM/mm), Ds is the sediment (mat in this case)diffusion coefficient (cm22 s21), and f is theporosity (volume of water/total volume of waterand sediment). fDs within the mat was estimatedas follows: For each salinity treatment, a high-res-olution (50-mm step) vertical profile through thediffusive boundary layer (DBL) and the mat wasmade. D0 (the free solution diffusion coefficient)in the DBL was calculated based on salinity andtemperature of the water column at the time ofthe profile using the equations of Li and Gregory(1974). Since f in water is 1 by definition, fD0 inthe DBL is known. fDs in the mat was then esti-mated as (C/z) fD0 (in DBL)/(C/z) (in mat).Mat fDs estimated in this way was similar topublished values (Glud et al., 1995; Wieland et al.,2001), ,1025.
Fluxes of oxygen, dissolved inorganic carbon(DIC), methane, and hydrogen
Fluxes of oxygen (O2) and DIC between themats and the overlying water were measured us-
A MICROBIAL MAT GREENHOUSE COLLABORATORY 389
ing glass benthic flux chambers that allowedunimpeded illumination of the surface, with noappreciable (,1°C) heating of the chamber. Thechamber design and operation follow those ofCanfield and Des Marais (1993). Briefly, eachglass chamber (covering an area of mat ,0.019 or0.012 m2) was fitted with a central stirring pad-dle and two sampling ports with septa. The glasspaddle rotated at a constant rate of 4.5 rpm.Chamber incubations were conducted for full 24-h daily cycles, taking samples by syringe every 6h. When deployed at sunrise, each chamber wasinjected with ,100 mL of pure nitrogen (N2) gasto create a headspace to sample at noon and be-fore sunset for acquisition of gas samples (O2,CH4, H2, etc.). At sunset, each chamber was re-deployed and injected with 70 mL of N2 and 30mL of O2 to accommodate O2 demand during thenight. Samples were taken at midnight and be-fore sunrise. Dissolved and headspace O2 valueswere measured using a Clark-style electrode withan internal reference mounted inside a 21-gaugesyringe needle (model 768, Diamond General De-velopment Corp.) and a gas chromatograph, re-spectively. The gas chromatograph (ShimadzuGC-14A) was fitted with a thermal conductivitydetector and a CTRI column (Alltech Associates,Deerfield, IL) held at 25°C. Helium, at a flow rateof 30 mL/min, was the carrier gas. DIC samples(5 mL) were filtered and analyzed in triplicate us-ing a “flow injection analyzer” (Hall and Aller,1992), where a sample was injected into an acid-ified aqueous solution that subsequently flowspast a gas-permeable membrane. Carbon dioxidefrom the acidified DIC sample traverses thismembrane and enters an alkaline solution thatsubsequently flows through a conductivity de-tector.
Hydrogen partial pressures
While actively photosynthesizing, mats gener-ate small (,1-mL) bubbles that are retained at themat surface with residence times of tens of min-utes. These bubbles were collected by means of a3-mL plastic syringe with a plastic pipette tiplodged in the luer fitting (flaring outward, to forma collecting funnel). When filled with water andsubmerged, this apparatus allows individualbubbles to be picked from the mat surface, with-out atmospheric contamination, and pooled to avolume of 10–25 mL. The pooled bubble gas is immediately subsampled with a gas-tight volu-
metric syringe and quantified by gas chromatog-raphy with HgO-reduction detection (modelRGA-3, Trace Analytical, Sparks, MD). The timefrom sample collection to analysis was generally,30 s. Partial pressures in bubble samples weredetermined by comparison with standards pre-pared by serial dilution of pure H2 into N2(Hoehler et al., 1998). Precision and accuracy aretypically about 65% for sample volumes in therange of 10–25 mL.
RESULTS
The greenhouse facility as a simulation of the natural environment
Microbial mats kept in the greenhouse facilityretained an overall appearance remarkably simi-lar to that of freshly collected mats. In particular,no evidence of the mat “greening,” in whichmotile cyanobacteria migrate to the surface of themat (Bebout and Garcia-Pichel, 1995), was ap-parent. During the first few weeks of greenhouseincubation, there was a notable increase in theabundance of loosely attached microbial “floc” atthe surface of the mats, as well as the develop-ment of small dark green spots containing largenumbers of cyanobacterial filaments in somemats. However, after the first 2 months, the loosefloc disappeared, and the mat surface was smoothand homogeneous in appearance once again. Ex-tensive, but nonquantitative, microscopic obser-vations revealed no major changes in communitycomposition. More specifically, the major popu-lations of cyanobacteria did not seem to change,and M. chthonoplastes remained the dominantphototroph in all of the sections of mat charac-terized microscopically.
Molecular analyses based on 16S rRNA genesthrough DGGE fingerprinting were consistentwith the absence of significant shifts in the com-munity. The banding pattern of DNA extractedfrom the greenhouse mats was essentially identi-cal to that of freshly collected mats (Fig. 2). Twomajor bands were present in all of the mat sam-ples (Band b and Band c). In the greenhouse treat-ments (Fig. 2A), Band c represented 84% of the ob-served cyanobacterial diversity, gauged as apercentage of total PCR amplificate, while Band brepresented 10%. These results are consistent withthe structure of field samples (Fig. 2B), where Bandc represented 79% of the observed diversity with
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Band b accounting for 18%. Two other minorbands were found, but not in all treatments. Banda was present in three of the six greenhouse sam-ples and in the field samples, accounting for 3%and 2% of the observed diversity, respectively.Band d was found in all six greenhouse samplesat much lower levels (0.8%), and accounted for1.2% of the observed diversity in the field samples.BLAST similarity searches for the sequencedbands (,380 bp in length) were performed. Bandb had no close matches to any cultured cyanobac-terium (only 89% similar to Oscillatoria amphigran-ulata strain 23-2, a typical distance between bacte-rial genera), but did match an unidentified,uncultured cyanobacterium from a microbial matin Solar Lake, Sinai Peninsula, Egypt (Abed andGarcia-Pichel, 2001) with 94% similarity. TheBLAST results for the remaining two bandsyielded very high similarity (99%) to known cul-tured cyanobacteria. Band c matched the M.chthonoplastes cluster (Garcia-Pichel et al., 1996) andBand dmatched Euhalothece sp. strain MPI 95AH13(Garcia-Pichel et al., 1998). Though we were un-able to sequence it, Band a most likely representsa diatom plastid (according to previous researchand band placement in the denaturant gradient).
The compositions of the total esterified FA(TEFA) in natural and greenhouse mats were inmost respects similar for both the surface and un-derlying Microcoleus layer (Table 1). The amountsof TEFA recovered for the individual mat sam-ples were reasonably consistent. The majority ofthis TEFA consisted of n-16:0, n-16:1, and n-18:1,which together accounted for 60–77% of all FA.Smaller amounts of other straight-chain saturatedand unsaturated FA (i.e., n-14:0, n-17:0, n-18:0, n-17:1, n-20:1, n-20:2) together with various termi-nally branched FA (i-14:0, i-15:0, ai-15:0, i-16:0, i-17:1, i-17:0, ai-17:0) were present in relativelyuniform amounts in all samples. However, sub-tle differences for TEFA compositions in the sur-face layer and underlying Microcoleus layer of nat-ural mats, which were also apparent in thegreenhouse mats even after 4 months of experi-mental treatment (Fig. 3), were noted. The sur-face layer tended to have higher levels of iso-15:1and iso-16:1 than the Microcoleus layer. In naturalmat, these two iso-FA were composed primarily(90%) of D4 positional isomers, previously iden-tified in field-collected Nostoc spp. (Potts et al.,1987). All surface layers (in both natural andgreenhouse mats) also contained small amounts
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FIG. 2. DGGE fingerprints of PCR-amplified cyanobacterial 16S rRNA genes of the six flume treatments (A) andfield samples (B). Arrows indicate bands used for further analysis of the fingerprints. Bands b, c, and d were excised,re-amplified, and sequenced.
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TABLE1.
COM
POSITIO
NOFTW
ON
ATURALM
ATS
FROM
THEA
REA4 M
ATD
ISSE
CTED
ATG
UERRERON
EGRO
AND
OFM
ATSC
OLLECTED
FROM
THESA
MEA
REAA
FTERM
AIN
TENANCE
INTHEFL
UM
ESY
STEM
FOR
$4 M
ONTHS
12/98 P4n51
90‰ flume
120‰
flume
6/01 P4n51
Surface
Microco
leus
Surface
Microco
leus
Surface
Microco
leus
Surface
Microco
leus
Ester-linke
d FA (mg g2
1TOC)
17.760
0000
31.800
0000
25.670
0000
023
.380
0000
26.520
0000
027
.850
000
30.870
0000
022
.090
000
C/N R
atio
9.80
0000
ND
9.40
0000
09.20
0000
12.100
0000
09.60
000
ND
ND
d13 C
(‰)
210
.72
6 0.04
210
.23
6 0.28
29.01
6 0.05
210
.59
6 0.13
29.70
6 0.17
210
.50
6 0.1
28.02
6 0.07
210
.56
0.65
d15 N
(‰
)20.13
6 0.4
ND
21.27
6 0.06
20.92
6 0.38
20.23
6 0.35
20.26
6 0.18
ND
ND
Error estim
ates sho
wn indicate instrumen
tal precision. Sam
ples labe
led N
D w
ere too sm
all to permit a reliable an
alys
is.
1 Collected
in D
ecem
ber 19
98 and
June
200
1 from
the
field site, Pon
d 4 nea
r Area 5.
FIG. 3. Relative distribution of various FA characteristic of the surface layer (A) and/or the underlying Microcoleus layer(B) for two natural mats collected from Pond 4 near Area 5 in June 2001 (black bars) and December 1998 (gray bars), andfor similar mats maintained for $4 months in flumes at salinities of 90‰ (hatched bars) or of 120‰ (dotted bars).
BEBOUT ET AL.394
of anteiso-17:1, which was not detected in the Mi-crocoleus layer. The most striking difference in thesurface layer, however, was the presence of rela-tively large amounts of several polyunsaturatedFA (PFA) (n-16:2, n-18:2, n-18:3, n-18:4, n-20:4,and n-20:5). The more highly unsaturated C20 FAare generally considered diatom biomarkers (Co-belas and Lechado, 1989). The lower Microcoleuslayer was also considerably enriched in severalFA. Most pronounced were a 10-methyl-16:0 as-sociated with some sulfate-reducing bacteria(Dowling et al., 1986; Kohring et al., 1994) and acy-19:0 that has been used as a biomarker forThiobacillus spp. (Kerger et al., 1986). In addition,microbial mats maintained in the greenhouse re-tained the carbon and nitrogen stable isotopiccomposition of recently collected mats (Table 1).
Measurements of rates of biogeochemical cy-cling in greenhouse mats were similar to ratesmeasured on freshly collected mats (Table 2). Pro-files of oxygen concentration within the green-house mats resembled those obtained in situ, inboth absolute concentration and distribution withdepth (data not shown). Oxygen fluxes to the wa-ter column, calculated from the concentrationgradients of these profiles, agreed with oxygenfluxes calculated from in situ profiles over a widerange of light intensities (Fig. 4). Similarly, netrates of O2 and DIC production and consumptionas determined using flux measurements weresimilar to those measured in freshly collectedmats (Table 2) after allowances were made for therelationship between rates of photosynthesis andseasonally-dependent levels of illumination inthe greenhouse. Fluxes of methane from thegreenhouse mats were reduced to 60% of thevalue measured in freshly collected mats andwere essentially equal day and night (Table 2). Ingas bubbles collected at the mat surface, H2 par-tial pressures in the low-salinity control mats were very similar to those observed in the field(Table 2).
The greenhouse as a facility for experimental manipulations
Over the course of the experiment, dramaticdifferences in the appearance of mats maintainedat in situ (NORMAL 5 90‰) and elevated(HIGH 5 120‰) salinities became apparent.Mats held at high salinities assumed a much moreorange color than those incubated at lower salin-ities. A change in color can result from physio-
logical (photopigment) changes in an unchangedcommunity or population shifts. It is known thatincreased salinity promotes increasingly oxida-tive conditions (Garcia-Pichel et al., 1999) and thatM. chthonoplastes can respond to increased salin-ity by increasing the cellular levels of carotenoid(López-Cortés, 1990). It is important to note, how-ever, that in natural field mats yellow-orangemats are typically associated with the dominanceof cyanobacteria from the Halothece cluster.
Nonquantitative microscopic observations re-vealed an increase in the abundance of unicellu-lar cyanobacteria that resembled the Halothecetype at the surface of the HIGH salinity mats rel-ative to those maintained at NORMAL salinity.Initial DGGE analyses supported this observa-tion, with a faint novel band attributable toHalothece, which appeared in samples collectedfrom all three HIGH salinity greenhouse matsamples (data not shown). However, these initialDGGE samples had been compromised by an ac-cidental thawing. The second set of samples,taken at the same time but not thawed beforeDNA extraction, did not show this band (Fig. 2).Rather, this second set of samples showed simi-lar fingerprints for all greenhouse samples, re-gardless of the treatment. Upon further investi-gation, we found that thawing and refreezing ofmat samples before DNA extraction caused thepreferential breakage of large-celled M. chthono-plastes, and the subsequent degradation of itsDNA. Because M. chthonoplastes provides the ma-jority of template for PCR amplification, DGGEanalyses of samples that had been thawed yieldeda high-resolution fingerprint of the communitymembers present at low numbers. It was only insuch analyses that we could detect communityshifts in the form of new bands appearing in allHIGH salinity greenhouses (data not shown). Se-quencing and phylogentic analyses of these novelbands revealed that the new community mem-bers belonged to the Halothece cluster of ex-tremely halotolerant, unicellular cyanobacteria(Garcia-Pichel et al., 1998). Thus we may havebeen witnessing an incipient community shift,but not a full replacement of the principalcyanobacterial populations.
Some differences between the NORMAL andHIGH salinity mats, and between greenhouseand freshly collected mats, were apparent in theFA composition. An increase in the diatom pop-ulation in the HIGH salinity mats was readilydocumented by increased amounts of two diatom
A MICROBIAL MAT GREENHOUSE COLLABORATORY 395
TABLE2.
MEASU
REM
ENTS
OFBIO
GEOCHEM
ICALPROCESS
ES
INFR
ESH
LYC
OLLECTEDM
ATS
AND
MATSM
AIN
TAIN
ED
INTHEG
REENHOUSE
FACIL
ITY
ATNORMAL A
NDHIG
H S
ALIN
ITIE
S
Greenhouse-maintained mats
Freshly collected mat
May 19–20, 2000
NORMAL
HIGH
NORMAL
HIGH
NORMAL
HIGH
Tem
perature (°C
)Nighttime
16
19
19
16
16
16
16
Day
time
16
21
21
19
19
19
19
DIC
flux (m
mol m
22 h
21 )
Nighttime
2.60
(0.02
)4.86
(0.36
)—
3.6 (N
/A)
3.07
(0.05
)3.8 (N
/A)
3.3 (N
/A)
Day
time
23.29
(0.14
)26.86
(0.15
)—
25.6 (N
/A)
23.74
(0.08
)22.1 (N
/A)
20.6 (N
/A)
Oxy
gen flux
(mmol m
22 h
21 )
Nighttime
23.52
(0.01
)24.45
(0.02
)—
22.4 (N
/A)
22.41
(0.01
)24.00
(0.25
)23.34
(0.01
)Day
time
3.49
(0.62
)7.20
(0.31
)—
5.6 (N
/A)
3.98
(0.04
)4.66
(0.37
)4.43
(0.02
)Metha
ne flux (nmol m
22 h
21 )
Nighttime
381 (5.47)
179 (23.6)
—18
0 (43.3)
170 (25.8)
152 (3.14)
524 (32.1)
Day
time
328 (27.6)
214 (63.5)
—17
6 (68.3)
174 (11.6)
200 (7.38)
653 (21.1)
[H2] (pp
m)
Stea
dy-state
10.7 (3.3)
0.54
(0.16
)—
0.67
(0.18
)0.28
(0.08
)—
—Bubb
le4.6 (2.9)
4.3 (2.0)
2.9 (1.0)
4.0 (0.007
6)1.1 (0.12)
——
n5
26n
58
n5
6n
52
n5
2
Values in
paren
theses rep
resent the
SD of mea
n va
lues. F
or flux mea
suremen
ts, n
52. The
num
ber of rep
licates for hyd
roge
n bu
bble m
easu
remen
ts is
indicated.
An SD
value
of N/A in
dicates a flux measu
remen
t of a single sample ( n
51). T
emperatures at w
hich
the
mea
suremen
ts w
ere mad
e are also ind
icated
.
June 14–15, 2000
July 6–7, 2000
November 8–9, 2000
biomarkers: the PFA 20:4 and 20:5, and the sterolcomposition. Together 20:4 and 20:5 accountedfor 4.38 mg g21 TOC in the HIGH salinity matsand only 3.16 mg g21 TOC in the NORMAL salin-ity mats. Both of these values are considerablyhigher than the natural mat samples (,0.6 mgg21). Sterols, a direct indication of the presenceof microeukaryotes in these mat samples, werealso more abundant in the HIGH salinity treat-ment with 1.20 mg g21 versus 0.74 mg g21 forNORMAL salinity mats. The two sterols thatserve as diatom biomarkers, 24-methylcholesta-5-en-3b-ol and 24-methylcholesta-5,22-dien-3b-ol(Volkman, 1986), accounted for ,80% of the to-tal in both greenhouse samples.
There were no differences observed in oxygenprofiles in the NORMAL and HIGH salinity mats(Fig. 5). Fluxes of O2 were measured at the out-set (June 2000) of the parallel treatment of themats at two salinities, and 5 months later (No-vember 2000). In both cases, no differences in O2
fluxes were apparent between the two salinitytreatments with regard to the rate of export of O2to the overlying water column during the day, orthe uptake of O2 at night (Table 2). In contrast,rates of methane production from the NORMALand HIGH salinity mats, which showed no dif-ference for the majority of time the experimentwas running, differed at the last sampling (No-vember 8–9, 2000), with higher rates recorded inthe normal-salinity treatment. H2 partial pressurein photosynthetic surface bubbles from the HIGHsalinity mats was about half that in the NORMALsalinity mats after running the experiment for 5months.
DISCUSSION
For a period of at least 1 year, microbial matsmaintained in the greenhouse facility resembledfield-collected microbial mats with respect to
BEBOUT ET AL.396
FIG. 4. Microelectrodes were used to monitor oxygen fluxes across the Area 4 mat surface–water interface dur-ing greenhouse [NORMAL (squares) and HIGH (circles) salinity] and in situ (triangles) experiments.Positive fluxesindicate net flux of photosynthetically-generated oxygen out of the mat, while negative fluxes indicate net oxygenconsumption. Error bars are SE values. Mats maintained in the greenhouse retain the general diel pattern observedin situ, including a morning peak in photosynthesis. Note shading events caused by greenhouse window crossbarsresulting in temporary light limitation of photosynthesis in small areas of the greenhouse mats (e.g., at 13:30 on Day1 and at 10:30 on Day 2).
their overall appearance, cyanobacterial commu-nity composition, and rates of biogeochemical cy-cling. High rates of activity of microbial popula-tions in mats contributed to the adaptability ofthese ecosystems. However, this attribute alsomakes the preservation of mat systems in theiroriginal, natural states especially challenging.The phenomenon known as “greening,” in whichmotile cyanobacteria migrate to the surface of themats and remain there in response to lowered ir-radiance, is common in mats maintained underartificial illumination (Bebout and Garcia-Pichel,1995). For example, when removed from theirnatural environment and maintained in large out-door ponds, microbial mats collected from hy-persaline Solar Lake, Sinai, Egypt became “over-grown” with populations of cyanobacteria thatare not necessarily well-represented in situ (Abedand Garcia-Pichel, 2001).
Oxygen microprofiles, measured using micro-electrodes, and oxygen and carbon fluxes, mea-
sured using flux chambers, were found to be com-parable in greenhouse and freshly collected nat-ural mats. While it is reassuring to observe thissimilarity in mat net community metabolism, itis likely that these parameters are among the leastsensitive to change since measurements of netcarbon and oxygen flux integrate the rates ofmetabolic activity of most of the microorganismsin the community. Increases in the activity of onegroup of microorganisms may be compensatedfor by decreases in the activity of others. We alsomeasured a close similarity between greenhouseand in situ bubble H2 partial pressures. H2 par-tial pressures in bubbles are extremely sensitiveto changes in the environment of photosynthesisin the uppermost mat layer. For example, matsremoved from their native pond and placed un-der conditions of high solar irradiance and lowor no water flow frequently exhibited an increasein bubble H2 partial pressures of up to 2 ordersof magnitude. The combination of light and flow
A MICROBIAL MAT GREENHOUSE COLLABORATORY 397
FIG. 5. Oxygen microelectrode pro-files within microbial mat in the day-time (1000h, open symbols) and night-time (2200h, solid symbols). Littledifference can be seen in profiles takenin mats incubated at NORMAL (squares)and HIGH (circles) salinity.
regimens appears to exert strong and instanta-neous control on bubble H2 partial pressures bysetting the balance between diffusive DIC supplyand light intensity (Lambert and Smith, 1980;Houchins, 1984). The fact that little or no differ-ence in bubble H2 partial pressures was apparentbetween greenhouse and natural mats suggeststhat the basic flow and light regimens of the nat-ural environment are recreated to a very high de-gree in the greenhouse setting.
We believe that the greenhouse approach doc-umented here is among the most successful of theefforts to date to produce an environment ap-propriate for the long-term study of these im-portant microbial communities. When comparedwith previous efforts to maintain microbial mats,our results indicate that two factors—water flowand the light regime—are likely to be more im-portant than others in simulating the field envi-ronment. We discuss in more detail the impor-tance of these two factors in our greenhousefacility.
The importance of flow in microbial mat me-tabolism has been recognized for at least a decade(Jørgensen, 1994; Boudreau and Jørgensen, 2001).The DBL, a thin (,0.5 mm) layer of stagnant wa-ter, is situated at the interface between any mi-crobial mat community and the water column,even under conditions of high water flow. Thetransport of all solutes between the water columnand the mat occurs exclusively by molecular dif-fusion through the DBL. Therefore the thicknessof this layer sets the rates at which this exchangemay take place. Rates of activity, including ratesof primary production in microbial mat commu-nities, may be controlled by the rate at which car-bon dioxide and oxygen diffuse across the DBL(Garcia-Pichel et al., 1999).
In the greenhouse facility, water was con-stantly circulated over the mats. The flow veloc-ity was set to ,5 cm s21 by adjusting the outputof our water pumps with valves, and measuredand maintained by periodic observations of thetransit of small particles or bubbles (through ameasured distance in the flow box over a givenperiod of time). We chose a value of 5 cm s21 forthe greenhouse experiments as (1) it was a rea-sonable approximation of average in situ valuesand (2) that flow velocity is used in a large num-ber of published microelectrode measurements inmicrobial mats (Revsbech and Jørgensen, 1986),which facilitated the ability to compare our re-sults with those of others.
Previous work with these mats revealed that,compared with the dramatic change in DBL thick-ness that occurs as flow velocities increase from0 (stagnant) to 1–3 cm s21, the DBL thickness isrelatively constant once flow velocities exceed1–3 cm s21 (Jørgensen and Des Marais, 1990).Therefore, flow variations around the 5 cm s21
value will minimally affect the fluxes in and outof the mat. For the flux chamber measurementsreported here, motor speeds were adjusted todrive the paddles to produce similar (5 cm s21)flow velocities, which were also estimated by tim-ing the transit of small particles within the fluxchambers across the surface of the mats. Actualin situ flow velocities, also measured by observa-tions of naturally occurring particles underwaterby divers, range from 0 (in the morning beforethe wind comes up) to .20 cm s21 under windyafternoon conditions, though these values shouldbe regarded as rough estimates owing to the tur-bulent nature of the flow and the difficulty in-herent in making the observations. No attemptwas made in the greenhouse to simulate thesedaily fluctuations in flow velocities. However, thesimilarity of microelectrode-based measurementsof DBL thickness (not shown) and microelectrodeprofile-based oxygen fluxes to those measured insitu (Fig. 4) suggests that the greenhouse flumesystem closely reproduces the environmentalflow field and DBL control on mat–water soluteexchange.
In photosynthetic microbial mats, all of the en-ergy necessary for growth and maintenance of thecommunity is ultimately derived from the sun.Oxygen production and consumption by thesephototrophic organisms, as well as their sensitiv-ity to UV radiation, determine the physical struc-ture of the entire mat community. Furthermore,many mat microorganisms are motile, utilizinglight and/or UV radiation as a cue to adjust theirposition in the mats vertically (Castenholz, 1994;Bebout and Garcia-Pichel, 1995). Therefore, thesemats are highly sensitive to changes in PAR (lightwithin the wavelength range from 400 to 700 nm).The use of artificial illumination to drive photo-synthetic activity in microbial mats over longerperiods of time is not attractive from a logisticalperspective owing to the large amounts of heatthat is generated. In addition, sources of artificiallight rarely reproduce the high irradiance, and(even more rarely) the spectral composition ofnatural sunlight. For both of these reasons, the useof natural sunlight is preferred for the mainte-
BEBOUT ET AL.398
nance of these microbial communities. Use of agreenhouse facility for microbial mat maintenanceis also the most cost-effective way to deliver therequired visible and UV radiation to microbialmat communities, particularly when large areasof mat are required for destructive samplingand/or large numbers of replicate measurements.
Mats maintained in the greenhouse facilityhave access to natural solar radiation filtered onlythrough OP-4 acrylic and a few centimeters ofwater. Therefore, both the total PAR irradianceand the exact spectral composition of that PARare essentially unchanged from that of sunlight.The difference in latitude between our green-house and the field site, as well as the absence ofa 1-m water column in the greenhouse, con-tributed to making the irradiance at the surfaceof the greenhouse mats similar to that in situ. Forexample, the irradiance measured at the field site(on cloud-free days at high noon) 5 1,845 6 196.9mE m22 s21 (average reading from four field trips,in May 2000, June 2001, October 2001, and Sep-tember 2002). Attenuation of light by the watercolumn resulted in values measured at the matsurface (on those same cloud-free days) of1,130 6 301.9 mE m22 s21. In comparison, the av-erage reading at the mat surface in the green-house was 1,005 6 396.2 mE m22 s21 (mean 6 SD,n 5 548, values recorded at 10-min intervals from1200h to 1300h from June 2001 through Septem-ber 2001). The greenhouse irradiance record in-cludes cloudy days, but these were relatively fewin number owing to the relatively clear summerskies at NASA Ames Research Center.
Microbial mats in situ are likely to experiencesome changes in the spectral composition of lightincident upon their surface (relative to sunlight)due to particulates present in the water column.These changes in spectral composition of the in-coming PAR (relative to sunlight) vary (e.g., withtime of year due to changes in the communitycomposition of phytoplankton in the salterns, andwith time of day due to wind-driven resuspen-sion of detrital material). No attempt was madeto simulate these changes in spectral compositionin the greenhouse. Owing to the high diversity ofphototrophic organisms in mats, and the wide va-riety of pigments used by those organisms to cap-ture light for photosynthesis, the total quantity ofphotons incident upon a mat community is of fargreater importance for community metabolismthan the exact spectral composition of the incom-ing radiation (Prufert-Bebout et al., 1998). Addi-
tionally, cyanobacterial motility and photosyn-thetic state transitions provide for efficient use ofthe rapidly changing spectral compositions withinmicrobial mat communities over the 1,000-mmdepth range, within which visible irradiance is ex-tinguished (Ploug et al., 1998; Prufert-Bebout et al.,1998).
Future modifications in greenhouse flow andlight regimens may further improve our ability tosimulate the field environment. The one poten-tially significant difference in community com-position observed between greenhouse and nat-ural mats, namely, the apparent increase in theabundance of diatoms in surface layers as sug-gested by the FAME biomarker data, may, in fact,be explained by differences in the flow regimenexperienced by mats incubated in the greenhouserelative to in situ conditions. In the greenhouse,the flow of water over the mats, while reproduc-ing natural flow velocities and DBL thickness, oc-curs 24 h a day. In the field environment, waterflow over mats is primarily wind-driven and, assuch, is variable over the course of the day. Strongwinds are characteristic of the afternoon, as ris-ing warm air is replaced by strong onshorebreezes. At night, and into the morning, there isvery little wind and, therefore, a much lower flowof water over the mats. Recently, we have docu-mented a density stratification of the ponds atnight. This density stratification, in combinationwith high rates of oxygen consumption by themat, resulted in a layer of anoxic bottom waterseveral centimeters thick, which persisted untillate morning (S.R. Miller, unpublished data). Hy-drogen sulfide, produced in the mat by sulfate re-duction, reached levels in excess of 185 mM in thislayer (B. Thamdrup, personal communication).Additionally, we recently documented that tem-peratures at the mat surface are often several de-grees higher than those in the middle of the water column and those that we have been sim-ulating in the greenhouse. This is presumably be-cause of greater heat retention by the sedimentsunderlying the microbial mats in their natural en-vironment. Constant flow over the greenhousemats would tend to diffuse heat from the surfacelayers of the mat, resulting in temperatures lowerthan those recorded in situ.
The lack of exposure to anoxic, sulfidic waterat night, as well as greenhouse water tempera-tures lower than those experienced by the matsin situ, would tend to increase the abundance ofdiatoms relative to unicellular cyanobacteria in
A MICROBIAL MAT GREENHOUSE COLLABORATORY 399
the surface layers of the greenhouse mats. Thephotosynthetic performance of the diatoms maybe inhibited in situ as a result of the sulfide sen-sitivity of oxygenic photosynthesis (Oren et al.,1979), and unicellular cyanobacteria from micro-bial mats are known to grow optimally at rela-tively high temperatures (Dor and Paz, 1989; Garcia-Pichel et al., 1998). Future work in thegreenhouse will investigate the extent to whichperiodic and/or nighttime periods of low flow,as well as periodic higher temperature excur-sions, might reduce the tendency of the diatomsto dominate surface layers of the mat.
The microbial mats in our greenhouse simula-tion facility represent a resource from which wewould like to obtain diverse observations and asmuch data as necessary to address key questionsin ecology. To increase the accessibility of thegreenhouse to members of our research group, thefacility is being transformed into a “collaboratory.”The collaboratory will enable a geographically dis-persed group of scientists to plan experiments, op-erate scientific equipment, take experimental mea-surements, share results, and collaborate in realtime with remote colleagues. Within the collabo-ratory, intelligent software agents will assist in theexperimentation process controlling the hardware,troubleshooting, re-cording results, and reportingback to collaborating experimenters. The xyz posi-tioning table over the mats is capable of automat-ically positioning sophisticated instruments at anylocation over and within the mats. The instrumentpackage currently includes microelectrodes, a lightsensor, chlorophyll fluorometer, a surface detec-tion device, and a fiber optic spectrometer. The po-sitioning system and instrumentation package areviewable over the Internet (http://greencam.arc.nasa.gov) via a webcam connected to a com-puter located in the greenhouse. The advanced au-tomation capabilities in the greenhouse will enablelonger-term experimentation, 24 h a day, withoutthe costs associated with extensive human super-vision. The remote collaboration capabilities willextend this experimental resource to colleaguesnot physically present in the greenhouse.
In summary, in addition to reproducing ratesof in situ biogeochemical processes, greenhousemicrobial mats were responsive to manipulationsof environmental conditions. Increasing the salin-ity of water in the greenhouse facility reproducedsome of the community composition changes ob-served in natural mats growing at higher salini-ties. The differences observed in the salinity re-
sponses of natural and greenhouse mats may beconsistent with expected differences betweenfield and greenhouse environmental conditions(e.g., differences in temperature and nighttimeoxygen and hydrogen sulfide concentrations).Therefore, we are optimistic that this facility maybe used not only to maintain mats with naturallevels of activity and species composition, butalso to explore environmental conditions that arenot presently found in the natural environment(e.g., low sulfate and low oxygen environments).
ACKNOWLEDGMENTS
This study was supported by grants from theNASA Astrobiology Institute, the NASA Exobi-ology Program, and the NASA Ames Director’sDiscretionary Fund. The greenhouse xyz tablewas designed and constructed by Dan Andrewsand Brian Koss, Code FE, NASA Ames ResearchCenter, and funded by the NASA Intelligent Sys-tems Program. Comments by two anonymous re-viewers greatly improved the manuscript.
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