Effects of Current Velocity on the Nascent Architecture of Stream Microbial Biofilms

Current velocity affected the architecture and dynamics of natural,multiphyla, and cross-trophic level biofilms from a forestedpiedmont stream. We monitored the development and activity ofbiofilms in streamside flumes operated under two flow regimes(slow [0.065 m s^-1] and fast [0.23 m s^-1]) by combined confocallaser scanning microscopy with cryosectioning to observe biofilmstructure and composition. Biofilm growth started as bacterialmicrocolonies embedded in extracellular polymeric substancesand transformed into ripple-like structures and ultimately conspicuousquasihexagonal networks. These structures were particularlypronounced in biofilms grown under slow current velocities andwere characterized by the prominence of pennate diatoms orientedalong their long axes to form the hexagons. Microstructuralheterogeneity was dynamic, and biofilms that developed underslower velocities were thicker and had larger surface sinuosityand higher areal densities than their counterparts exposed tohigher velocities. Surface sinuosity and biofilm fragmentationincreased with thickness, and these changes likely reduced resistanceto the mass transfer of solutes from the water column into thebiofilms. Nevertheless, estimates of dissolved organic carbonuptake and microbial growth suggested that internal cyclingof carbon was more important in thick biofilms grown in slowflow conditions. High-pressure liquid chromatography-pulsedamperometric detection analyses of exopolysaccharides documenteda temporal shift in monosaccharide composition as the glucoselevels decreased and the levels of rhamnose, galactose, mannose,xylose, and arabinose increased. We attribute this change inchemical composition to the accumulation of diatoms and increasedincorporation of detrital particles in mature biofilms.

INTRODUCTION

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Introduction

An understanding of the structure-function relationships inmicrobial biofilms is central to interpret and predict biofilmimpacts on the habitat where they form (46). Our current perceptionof biofilm structure results from the interplay of mathematicalmodeling and experimental studies. At least three conceptualmodels have been derived from this research (57): (i) planar,homogeneous biofilms are typified in dental plaque exposed toextremely high substrate concentrations in the saliva; (ii)heterogeneous, mosaic biofilms consisting of stacks of bacterialcolonies embedded in extracellular polymeric substances (EPS)and appearing as columns surrounded by the bulk liquid havebeen described from drinking water distribution systems (21);and (iii) biofilms with complex three-dimensional structureswith channel networks through which liquid flow occurs havebeen described from studies by using confocal laser-scanningmicroscopy (CLSM), microelectrodes, and fluorescent particles(10, 23, 45). A unifying thread of these conceptual models isthat they attempt to explain biofilm structure as a functionof substrate concentration and hydrodynamics (47, 50, 57). Additionally,the biofilms described in these models were derived from nuisancegrowths that colonize medical and industrial systems, wheresunlight is absent and communities are largely bacterial monocultures.In contrast, biofilms from natural ecosystems exposed to sunlightinclude greater trophic (phototrophs and heterotrophs) and phylogenetic(eucaryotes and procaryotes) diversity and complexity.

Although some of the earliest biofilm research was related torock surfaces in stream ecosystems (14, 27, 28, 43), our currentknowledge of lotic biofilms lags far behind their medical andindustrial counterparts. However, biofilms represent the dominantmode of microbial life in streams (12, 14), where they can driveecosystem metabolism, nutrient cycling, and biodegradation.Stream biofilms are functionally (42) and phylogenetically (2,29) diverse communities and possess highly complex EPS chemistry(33, 34, 58). Flow is an inherent feature of stream ecosystemsthat influences function, as well as structure, and yet studiesof flow effects in streams primarily have focused on the compositionand shape of diatom communities (3, 4). Most of our currentunderstanding of the relationships between flow and structure-functioncoupling of biofilms is largely derived from bacterial biofilmsmanipulated in laboratory-scale reactors (see, for example,references 30, 38, and 56). Thus, there is a need to relatehydrodynamics to the structure and function of stream biofilmsand the resultant impacts on biological and chemical processesin stream ecosystems.

We report here on the nascent architecture and dynamics of natural,microbial biofilms in a forested piedmont stream. Biofilms werecultivated under two different current velocities in streamsideflumes to incorporate the natural dynamics of stream water chemistry,particle behavior, and hydrodynamics into our experimental design.We describe biofilm development from bulk biomass measurements,we used CLSM, along with cryosectioning, to explore the spatialorganization of bacterial and algal cells, bulk exopolysaccharides,and particles within biofilms, and we analyzed the monosaccharidecomposition of the EPS.

MATERIALS AND METHODS

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Materials and Methods

Cultivation of biofilms.
Biofilms were grown in four streamside flumes (length, 30 m;width, 0.30 m; depth, 0.30 m) adjacent to a forested reach ofWhite Clay Creek (WCC; Stroud Water Research Center, Pa.). Flumeswere continuously fed in a once-through mode with streamwaterfrom a header tank to ensure constant flow. WCC nutrient concentrationstypically range from 3 to 4 mg of NO₃-N liter^-1 and 10 to 60µg of PO₄-P liter^-1 (35). All flumes were packed withthe same clean, commercially available cobble and pea gravel(median grain size, 20.9 ± 0.8 mm) and unglazed ceramiccoupons (1 by 1.5 by 0.3 cm) (6). Flume slopes were adjustedto yield duplicate open channel flow velocities of 0.065 ±0.003 and 0.23 ± 0.005 m s^-1. The average water temperaturewas 18.4°C and was not affected by water residence timein the flumes.

Microbial biomass and exopolysaccharides.
Bacterial abundance was estimated by epifluorescence directmicroscopic counts (17). Briefly, preserved biofilms (2.5% formaldehyde)were treated with 0.1 mM sterile tetrasodium pyrophosphate (54)for 10 min, and cells were removed by sonication (75 W, 60 s).Aliquot samples (1 ml) were transferred into sterile vials containing1 ml of tetrasodium pyrophosphate, thoroughly vortexed, andthen sonicated (75 W, 30 s) again. Samples were then mixed withsterile glycerol (30% final concentration), vortexed, and spunto pellet particles that likely cause background fluorescence(2). After staining with propidium iodide, bacterial cells inthe supernatant were enumerated on a black 0.2-µm-pore-sizeGTBP Millipore filter in 10 to 30 randomly selected fields toaccount for 300 to 500 cells. Two filters were counted per ceramiccoupon. Chlorophyll a was assayed to provide an estimate ofalgal biomass (44). Bulk organic and inorganic biofilm accumulationswere derived from dry and ash masses (450°C, 5 h).

Carbohydrate analyses were performed on bulk EPS harvested frombiofilms by extraction in 50 mM EDTA (53) on a rotary shaker.The extract was filtered (0.2-µm pore size) to removeparticles and cells, and EPS in the filtrate was precipitatedwith ice-cold ethanol (99% [vol/vol]) and held at -20°Cfor 24 h. The polymeric fraction was pelleted by centrifugation(1,500 x g, 30 min), and the supernatant was discarded. Thepellet was redissolved in 2 ml of deionized water, and EPS waswashed a second time in ice-cold ethanol. The resulting pelletwas lyophilized and subsampled. One aliquot was assayed fortotal carbohydrates by the sulfuric acid-phenol method (11),and another aliquot (0.5 ml) was digested (100°C, 5 h) with4 M trifluoroacetic acid in precombusted glass ampoules thatwere flushed with nitrogen and analyzed for monosaccharidesby high-pressure liquid chromatography (HPLC)-pulsed amperometricdetection (PAD) (7). Bulk EPS carbohydrates, bacterial abundance,and chlorophyll a were measured on each sampling date, whereasmonosaccharides were determined on days 3 and 5 (young biofilms)and days 18 and 21 (mature biofilms) of the biofilm development.

DOC uptake.
To estimate biofilm functional dynamics, we monitored the flume-scaleuptake of dissolved organic carbon (DOC) during biofilm growth.DOC concentrations in the flume influent and effluent waterswere measured with an OI-700 organic carbon analyzer (20). DOCuptake was estimated from mass balance calculations and relatedto the flume bed surface area.

Cryosections and CLSM.
Developmental patterns were derived from the CLSM investigationof the biofilm xy plane from ceramic coupons that were randomlysampled for structural analyses, carefully removed from theflumes, and transferred into microchambers (Fisher Scientific)containing streamwater (0.2-µm pore size filtered). Within30 min of sampling, bacterial cells were stained with the fluorescentnucleic acid stain SYTO 13 (Molecular Probes, Inc., Eugene,Oreg.) at a concentration of 20 µg ml^-1 (10 min, roomtemperature). Microbial EPS was stained with the lectin probeConcanavalin A (ConA)-Texas red (Molecular Probes), which purportedlybinds to terminal ${alpha}$ -D-glucose and ${alpha}$ -D-mannose, although ConA mightbind to non-EPS targets or nonspecifically to the biofilm matrix(19). We stained EPS with a lectin concentration of 5 mg ml^-1(10 min at room temperature). Excess staining solutions wereremoved by gently rinsing the samples with filtered (0.2-µmpore size) streamwater. These samples were used either for directCLSM analysis or cryoembedding. Cryoembedding was performedby embedding samples in Tissue-Tek O.C.T. compound (Miles, Inc.,Elkhart, Ind.) for 15 min and then freezing them on dry ice(18, 60). Once completely frozen, the biofilm was removed fromthe ceramic coupon, turned over, placed back on dry ice, andreembedded in O.C.T. compound. Embedded biofilms were cut into6-µm sections (yz plane) with a Leica (Germany) cryostat.Cryosections were transferred onto poly-L-lysine-coated slides,allowed to air dry, and subsequently mounted in Citifluor medium(Citifluor, Canterbury, United Kingdom). Sections were immediatelyanalyzed with a Zeiss LSM 510 scanning confocal microscope (CarlZeiss, Jena, Germany) equipped with an ArKr laser (488 and 568nm) and an HeNe laser (633 nm). For routine imaging, a x10 PlanNeo-Fluar lens was used to cover cryosections over a lengthof 922 µm. For high magnifications, we used x40 Plan Neo-Fluar(NA 1.3, oil immersion) and a x63 C-Apochromat (NA 1.2, waterimmersion) objective lenses. The algal autofluorescence wasrecorded in the far-red channel, and its overlay with Conavalin-Texasred resulted in a purple appearance.

Image acquisition and analysis.
Digitized cryosections (16 bits pseudocolor TIFF files, 1,024by 1,024 pixels) were imported from the LSM 510 into the GISand mapping software Arc-View 2. 2 (ESRI, Inc.) for image analysis.Each imported image layer (red-green-blue [RGB]) was convertedto a unique grid in a Cartesian coordinate system. We applieda fixed threshold to all images (15). Thresholding results ina matrix with "zero values" in positions where the numbers ofRGB pixels are <6 and "one values" where the numbers of RGBpixels are between 6 and 255. One values represent positionscontaining microbial cells, ConA-EPS, or particles; zero valuesrepresent positions with voids. After thresholding, we removedbackground noise by using the Spatial Analyst (version 1.1;ESRI). Biomass pixels that were not connected to adjacent biomassclusters and that contained <50 RGB pixels were set to thezero value. These images were used for the quantification ofmicrostructural heterogeneity in the xz plane. Cryosectionsproved appropriate for quantitative analyses of biofilm structure(31).

Microstructural parameters.
Microstructural parameters estimated from these images includedthe average thickness and roughness of the biofilm. Roughnesswas determined as the deviation from 200 to 250 individual thicknessmeasurements per section (cf. reference 31). Biofilm porositywas determined as the ratio of the void surface area (zero valueRGB pixel) to the total cross-sectional surface area (sum ofzero and one value RGB pixels). The sinuosity of the biofilmsurface front was determined as the ratio between the numberof connected RGB pixels of the biofilm surface between two givenpoints and the corresponding linear distance between these twopoints. This parameter is analogous to surface enlargement,which is thought to affect mass transport (40). Biofilm fragmentationwas determined as the ratio of the perimeter and the surfacearea of biofilm clusters. Biofilm fragmentation is analogousto the average diffusion distance suggested as a measure ofthe distance over which a substrate molecule must diffuse fromthe void to bacterial cells within a cluster (59). The biofilmareal density was calculated from average thickness and totalmass estimates of biofilms (39).

Statistical analyses.
Hydraulic parameters and DOC uptake are presented as the meanof duplicate flume-scale measurements performed at each samplingdate. Biofilm biomass parameters were determined from two tothree ceramic coupons per flume per sample date. Biofilm microarchitectureparameters were determined from duplicate coupons collectedfrom a single flume representing each flow treatment on eightoccasions. Because low sample numbers contribute to uneven variation,we used the nonparametric Mann-Whitney Wilcoxon procedure totest for flow effects on biofilm structures (13). Correlationsbetween biofilm microarchitecture parameters were explored byusing the Pearson r coefficient. All values are given as mean± the standard deviation (SD).

RESULTS

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Results

Hydraulic characteristics of the flumes.
The two flow regimes resulted in a 13-fold difference in shearstress and a 4-fold difference in Reynolds numbers (Table 1).The Reynolds numbers for open channels were indicative of transitionaland fully turbulent flow.

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TABLE 1. Hydraulic characteristics of the experimental flumes

Biomass development and flume-scale dynamics.
Flow significantly affected bulk microbial biomass, with 3.4-fold-higherbacterial abundance and 2.5-fold-higher chlorophyll a concentrationsin the slow flow treatments from approximately day 10 onward(Fig. 1). After an initial 3- to 5-day lag phase, chlorophylla and bacterial abundance steadily increased and then decreaseddue to massive biofilm detachment and grazing by invertebrates,as revealed by visual inspection. The development of long filamentousstreamers (up to 6 cm long) contributed to large within-flumevariations of mature biofilms.

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FIG. 1. Developmental dynamics of microbial biomass as indicated by chlorophyll a (a) and bacterial abundance (b). Closed symbols denote slow-flow treatments, and open symbols denote fast-flow treatments. Horizontal arrows indicate periods of increased biofilm detachment and grazing. The results are averages ± the SDs of two to three coupons for each experimental flume.

The trajectory of flume-scale uptake of DOC followed the broadpattern of biomass development (Fig. 2). DOC uptake, measuredas a decline in DOC concentrations between the header tank inflowand the flume outflow, occurred consistently when the influentwaters were pumped from the stream at baseflow and had a DOCconcentration of $<=$ 2.4 mg of C liter^-1. Outflow concentrationsfrom the treatments were very similar (average difference of0.028 mg of C liter^-1 over a range of values from 1.49 to 2.29mg of C liter^-1) between treatments. However, when the uptakevalues were multiplied by the discharge to obtain estimatesof the uptake flux, average rates were 3.9 times higher in thefast-flow treatment. The percentages of influent DOC removedwithin the flumes gradually increased with biofilm developmentfrom initial values of 2 to 6% from days 3 to 7, to 7 to 11%from days 9 to 17, and finally to 32% when last measured onday 27. The 32% DOC removal represented a difference of 0.479mg of C liter^-1 from a starting concentration of 1.49 mg ofC liter^-1.

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FIG. 2. Temporal dynamics of DOC uptake from the streamwater. The results are averages ± the SDs from duplicate flow treatments.

Exopolysaccharides.
HPLC-PAD analyses revealed glucose as the dominant monomer inall biofilms, followed by galactose and mannose (Fig. 3). Youngbiofilms showed similar monomer compositions in both flow treatments.The exopolysaccharide fingerprints of biofilms under the slow-flowtreatment changed with biofilm development as follows: the molepercent value for glucose decreased 3.2 times; the mole percentvalues for rhamnose, galactose, mannose, and xylose increased2 to 2.8 times; and the mole percent value for arabinose increased17 times (Fig. 3). The average levels of exopolysaccharides(as glucose [cf. reference 11]) were significantly higher inthe fast-flow treatments and, when combined with the significantlylower bacterial abundance, yielded a nearly threefold-higherexopolysaccharide/cell ratio in the fast-flow treatments (Table2).

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FIG. 3. Monosaccharide composition of young and mature biofilms from slow- and fast-flow treatments. FUC, fucose; RHA, rhamnose, ARA, arabinose; GALN, galactosamine; GLUN, glucosamine; GALA, galactose; GLUC, glucose; MAN, mannose; XYL, xylose; FRUC, fructose; RIB, ribose. Molar ratio is the ratio of the number of moles of the monomer to the total number of moles of saccharides; data are averages ± the SDs from duplicate coupons.

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TABLE 2. Effect of flow velocity on bulk exopolysaccharides

Structural development of the xy plane.
The progression of biofilm growth began with bacterial colonization,which by day 4 consisted of isolated microcolonies and bacterialcells scattered over the entire substratum and the first appearanceof ripple-like structures with distinct ConA staining of theEPS (Fig. 4a and d). At this developmental stage, long filamentousbacteria were prominent in the fast flow treatment (Fig. 4d).By day 8, anastomoses developed between ripples, along witha first appearance of diatoms and amorphous material (Fig. 4band e). The overall orientation of these initial structureswas parallel to the main flow direction. This was particularlypronounced in the fast-flow treatments. By day 21, a prominentnetwork of quasihexagonal structures developed from the basalpattern, especially in those biofilms grown in slow flow (Fig.4c). Elongated pennate diatoms (Syndera ulna) formed the sidesof the hexagons around a central void and amorphous materialaccumulated around the diatoms. Pores or voids defined by thequasihexagonal configuration of biomass had an average diameterof 160 ± 36 µm (n = 14). At the same time, biofilmsfrom fast-flow treatments consisted of a dense network of conspicuousripples with a high degree of anastomosis, creating centralvoids that were similar to those in the slower flow but withoutthe same depth and layering (Fig. 4f).

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FIG. 4. CLSM images showing the structural dynamics and formation of quasihexagonal structures in stream biofilms (xy plane). (a, b, and c) Images of 4-, 12-, and 21-day-old biofilms, respectively, from slow-flow treatments; (d, e, and f) images of 6-, 12-, and 24-day-old biofilms, respectively, from fast-flow treatments.

Structural development of the xz plane.
Low-magnification CLSM analysis of cryosections generally supportedthe observed xy patterns of living biofilms, with loose butcopious clusters of ConA-stained EPS (Fig. 5a, e, and f). Virtuallyno algal cells were detectable in these 3- to 5-day-old biofilmspredominantly composed of bacteria. With continued growth, adistinct base layer and a canopy developed. The base layer wasgenerally recognized by low ConA-stained EPS and the presenceof accumulated particles and bacterial cells (Fig. 5c, d, g,and h) arrayed around arcade-like void spaces (Fig. 5c and h).Initially, the canopy consisted of finger-like protuberances(Fig. 5c) and, as biofilms developed, algal cells became thedominant canopy component (Fig. 5d, h, and i). A cross-sectionof a quasihexagonal space with a long algal cell bridging thegap shows this pattern (Fig. 5b). Depending on the developmentalstage of biofilms, valleys between the canopy protuberancescan extend into the biofilm base to form extensive channel networks(Fig. 5c and d). In some cases we observed a conspicuous stratificationof ConA-stained EPS layers and diatoms (Fig. 5g and h).

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FIG. 5. CLSM micrographs of cryosections showing the structural development of the biofilm xz plane in slow flows (a to d) and fast flows (e to i). (a) Coalescing microcolonies of a 4-day-old largely bacterial biofilm; (b) a relatively compact biofilm (12 days) with finger-like protuberances and the first appearance of algal cells; (c) mature biofilm (17 days) with elongated algal cells bridging a possibly quasihexagonal void; (d) 24-day-old biofilm with distinct structural differentiation into a base layer and a canopy; (e and f) images showing initial microcolonies with filamentous bacterial cells in initial biofilms (at 3 and 6 days, respectively); (g and h) images showing the structural differentiation into a base layer and a diatom-rich canopy (g) and clear microstratification with a ConA-EPS-rich layer (h); (i) cross-section of ridge-like structure with void, possibly resulting from metazoan burrowing. Note the arcade-like void system within the base layer of mature biofilms (c and h).

With higher magnification, we were able to observe that mineralparticles coated with ConA-stained EPS were the major componentof biofilm microlayers (Fig. 6a). Diatoms constituted a centralbuilding block of the filamentous streamers that also includedabundant bacterial cells and noticeable peripheral ConA-stainedEPS (Fig. 6b). High magnification revealed large bacteria denselycolonizing a broken algal cell and its exudates, as visualizedby ConA-stained EPS (Fig. 6c), suggesting an autotrophic-heterotrophiccoupling.

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FIG. 6. CLSM images showing the incorporation of ConA-EPS-coated particles in a mature biofilm (a), a cross-section of the distal part of a streamer of a 14-day-old biofilm (b), and an example of the close coupling between an algal cell and bacteria (c).

Quantification of biofilm microstructures.
Biofilms grown in slow-flow treatments were thicker and hadhigher average sinuosity and areal density and yet had lessroughness than biofilms from fast-flow treatments (Table 3).Biofilm fragmentation and porosity were not affected by flow.Surface roughness decreased, whereas both sinuosity and fragmentationincreased, with biofilm thickness (Fig. 7). Fragmentation didnot correlate with biofilm thickness.

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TABLE 3. Microstructural biofilm parameters for slow (n = 18)- and fast (n = 18)-flow treatments

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FIG. 7. Relationships between biofilm thickness and roughness, sinuosity, porosity, and fragmentation. Error bars indicate the SDs from 200 to 250 single-thickness measurements. See the text for further explanation.

DISCUSSION

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Discussion

Structural development.
The development of ripple-like or ridge-like structures thatwe have documented are similar to biofilm features observedin other systems. For example, migratory ripples were describedfrom mixed-species bacterial biofilms cultivated in turbulentflow (48). Ridges aligned in the main flow direction that ultimatelydeveloped into dense parallel structures were observed in biofilmsgrown on polycarbonate slides within rotating annular reactorsfed by stream water (31).

Quasihexagonal structures similar to those we observed as anastomosesbetween parallel ridges evolved into a basal pattern have beendescribed for biofilms from wastewater (36) and marine sulfate-reducingbacteria (61). This mode of biofilm growth thus seems to bemore common than hitherto thought, and yet the mechanisms underlyingthis pattern remain largely speculative. This growth form maybe a response to the hydrodynamics of the flume microenvironmentsand facilitate the competition for a substratum during initialbiofilm formation as described by Picioreanu (41), for instance.Alternatively, bacterial cells that initiate biofilm growthare believed to skip over crevices to avoid zones of reducedmass transfer (32), and this could potentially lead to biomassgrowth clustered around pores in the substratum. However, closeCLSM inspection of the coupon texture revealed that the distributionof pores did not agree with biomass patterns.

The increased physical stability of quasihexagonal structuresis another possible cause of this biofilm architecture. Stoodleyet al. (49) described biofilm rheology as viscoelastic and demonstratedstructural deformation of bacterial biofilms grown in turbulentflow. The quasihexagonal patterns fortified by pennate diatomsas major building blocks could prevent biofilms from detrimentaldeformations and heavy erosion when faced with changes in currentvelocity in stream ecosystems. Furthermore, pores or voids withinthe interior of these quasihexagons may facilitate advectivesolute transport as envisioned by current biofilm models (see,for example, references 8 and 9).

CLSM has become a powerful tool for studying the spatial organizationof biofilms. However, where limited penetration depth of theconfocal laser beam, along with photobleaching of fluorescentprobes, becomes an issue, sectioning of the specimen prior toCLSM can provide better images and information on the spatialorganization (26, 55). The combination of cryosections withCLSM yielded xz images of stream biofilms with high-qualityresolution that could not have been achieved with confocal opticalsections and subsequent reconstruction (cf. reference 33). OurCLSM analyses of cryosections conveyed excellent insight intothe vertical growth of biofilms and underscore the importanceof organic and inorganic particles in stream biofilms.

Upper layers of initial microcolonies coalesced through growth,which resulted in the development of voids within the base layerof the biofilm. Algal cells extended away from the basal layerto form the biofilm canopy, along with inorganic particles andorganic debris. In some instances we found a microstratificationof particles coated with EPS and diatoms, which probably reflectsparticle dynamics in the stream water. Particles can increasebiofilm surface area and transport attached bacteria to biofilms,but they may also affect mass transfer by increasing the diffusionalresistance within biofilms. Organic particles can furthermoreconstitute a significant carbon source for heterotrophs. Thecombination of these factors certainly contributes to the shell-likegrowth with elevated bacterial abundances and activities closeto the biofilm surface observed in various systems (see, forexample, reference 33) and derived from multidimensional modeling(40). On the other hand, bacterial growth in streams can alsoincrease particle deposition velocity through enhanced filtrationand adhesion [T. J. Battin, L. A. Kaplan, and J. D. Newbold,Eos. Trans. AGU, no. 82(47), Fall Meet. Suppl., abstr. H12G-02,2001]. Once accumulated and aggregated within biofilms, particlescan be exported from the ecosystem through detachment of entirebiomass clusters.

We found disparate developmental patterns of surface roughnessand sinuosity in the stream biofilms. Roughness decreased withbiofilm maturity in our stream systems, whereas it has beenshown to increase in Pseudomonas aeruginosa biofilms (39) andmultidimensional modeling (41). However, laboratory-grown biofilmsare not exposed to particles and do not include photoautotrophsor heterotrophic grazers. In streams, biofilm structural developmentis the net result of growth and detachment of microbial biomass,hydrodynamics, substrate availability, and grazing (cf. reference25). Among other things, enhanced particle filtration and accumulationwithin the heterogeneous canopies that were particularly welldeveloped in biofilms from our slow-flow treatments could alsoexplain the higher areal density of these communities.

The increase of surface sinuosity with biofilm thickness andhence with maturation agrees with modeling results by Picioreanuet al. (40, 41). These authors showed that the width of thevalley defined by two neighboring protuberances (i.e., sinuosity)determines advective transport to the biofilm rather than surfaceroughness. Thus, we postulate that sinuosity increases withbiofilm thickness to assure the mass transfer of substratesnecessary to satisfy microbial demand. Considering the shell-likegrowth of biofilms, enhanced transport of particles and substratesbetween neighboring protuberances would thus successively reducesurface roughness. Similarly, fragmentation increased with biofilmthickness, and this likely enhanced solute transport withinbiofilms by increasing the channel network and thus reducingthe average diffusive pathway for substrate molecules (59).

Exopolysaccharides.
The monosaccharide composition we described from the streambiofilms generally agrees with other gas chromatography- orHPLC-derived carbohydrate fingerprints from intertidal sediments(52) or terrestrial biofilms (51). Glucose is always the dominantmonomer, followed by mannose, rhamnose, galactose, or xylose.Noninvasive lectin-binding assays support the frequent occurrenceof these monomers and further illustrate their overwhelmingspatial heterogeneity (33, 34, 58). Patterns of ConA-stainedEPS in the present study also support the HPLC-PAD-derived patternsof glucose and mannose as dominant carbohydrates in the streambiofilms.

Our results are the first to show a relative shift of the monosaccharidecomposition during biofilm development. Whether this changeis attributable to a shift of the community structure remainsunknown. For instance, diatoms, which are well known to producecopious amounts of EPS (16), became particularly abundant inmature biofilms from slow-flow treatments. Alternatively, increasedincorporation of allochthonous EPS (e.g., via particles) couldalso shift the polysaccharide fingerprint. No lysis controlwas included in our EPS extraction, and so we cannot assessthe contribution of intracellular polysaccharides to our exopolysaccharideestimates.

We did find higher exopolysaccharide/cell ratios in biofilmsgrowing in a fast flow. This agrees with findings from Pseudomonasfluorescens biofilms grown in flow cell reactors (38) and frombiofilms collected from a high-gradient glacial stream (2).Elevated exopolysaccharide/cell ratios should improve adhesionunder elevated shear stress and thus reflect a functional responseto avoid detachment through erosion and sloughing.

Biofilm function.
Our results on DOC uptake indicate that biofilms grown in fastflow rely more on organic substrates from the water column,a finding that agrees with the uptake of experimentally injectedglucose (Battin et al., Eos. Trans. AGU abstr. H12G-02, 2001).We attribute this difference between flow treatments to theoverall lower diffusive resistance in the fast-flow treatments.In fact, the external boundary layer is thinner in a fast flow,and the internal diffusive resistance is lower due to reducedthickness and areal density. The converse effects isolate slow-flowbiofilms from external supply, and heterotrophs within thosebiofilms predominantly rely on internal energy sources. Algalexudates (see Fig. 6c) and DOC immobilized within the EPS matrixfigure among these internal sources (see, for example, reference1). Close coupling between heterotrophs and autotrophs is thusbelieved to be more important in thicker biofilms, a notionthat agrees with findings from microbial mats (see, for example,reference 22). This implies elevated internal cycling—orclose spatial coupling of carbon production and consumption—withinthese slow-flow communities, leading to reduced downstream cycling.

Conclusion.
Performing the present study in large-scale streamside flumeshelped place our biofilm research into an ecosystem context(sensu Palmer and White [37]) since the flumes received sunlight,natural concentrations of solutes and particles, and simulatedhydrodynamics that occur at the water-sediment interface. Theseconditions allowed us to observe the dynamics of biofilm structureand function at a level of complexity that could not be obtainedwith, for instance, more artificial, laboratory-scale rotatingannular bioreactors (24, 33). We found that stream microbialbiofilms grown under quasinatural conditions were highly dynamicand complex communities composed of organisms that cross trophiclevels and represent multiple phyla. An open system such asa stream provides a broad repertoire of structural (inorganicand organic particles) and functional (procaryotic and eucaryoticphototrophs and heterotrophs) building blocks or componentsfor biofilms. This expansive view of microbial biofilms complicatesour ability to probe the evolutionary, molecular, and geneticbases for biofilm formation, such as the description of genesinvolved in the maintenance of biofilm architecture in a bacterialmonoculture (5). Alternatively, it opens up possibilities forobserving interactions that are common in natural environments,at a scale that is relevant to organic matter dynamics and biogeochemicalprocesses in stream ecosystems.

ACKNOWLEDGMENTS

We thank Sherman Roberts and Mike Gentile for technical assistance,Kirk Czymmek for CLSM support, and Stefan Würtz and twoanonymous reviewers for constructive comments.

This research was supported by a FWF Erwin Schrödingergrant (J1879-BIO) to T.J.B., U.S. National Science Foundationgrant DEB-9904047 to L.A.K., and the Pennswood No. 2 Fund forEnvironmental Research.

FOOTNOTES

* Corresponding author. Mailing address: Department of Limnology, IECB, University of Vienna, Althanstr. 14, A-1090 Vienna, Austria. Phone: 43-1-4277-4350. Fax: 43-1-4277-9542. E-mail: tomba{at}pflaphy.pph.univie.ac.at.

REFERENCES

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