biotic disturbance, recolonization, and early succession of bacterial assemblages in intertidal...

Biotic Disturbance, Recolonization, and Early Successionof Bacterial Assemblages in Intertidal Sediments

C.J. Plante and S.B. Wilde

Department of Biology, Grice Marine Laboratory, College of Charleston, Charleston, SC 29412, USA

Received: 1 April 2003 / Accepted: 24 June 2003 / Online publication: 3 May 2004

Abstract

The role of disturbance in structuring natural microbialcommunities has been largely unexplored. Disturbanceassociated with invertebrate ingestion can reduce bacte-rial biomass and alter metabolic activities and composi-tions of bacterial assemblages in marine sediments. Theprimary objectives of the research presented here were totest whether ingestion by a taxonomically diverse groupof deposit feeders constituted a disturbance, and todetermine the mechanisms by which bacterial assem-blages recover following deposit-feeder ingestion. To testthe question of disturbance, we compared fresh egesta vssurficial sediments with respect to bacterial assemblagestructure. In emersed intertidal sediments, microbialrecovery could be due to regrowth of bacterial popula-tions surviving gut passage or to immigration fromadjacent sediments. To differentiate between these modesof recolonization we used field manipulative experimentsto exclude migration by isolating freshly extruded fecalcoils of three deposit-feeding species from surroundingsediments. We then followed the quantitative and qual-itative recovery in egesta and sediments through timeusing epifluorescence microscopy and PCR-DGGE anal-ysis of 16S rDNA. Our findings indicate that (1) thedegree and nature of the disturbance to bacterialassemblages from deposit feeding varies among inverte-brate taxa, (2) recovery was significant but incompleteover 3 h, and (3) recolonization of biotically disturbedsediments is dominated by immigration.

Introduction

Bacteria in sediments are vital to ecosystem function inthat they play key roles in benthic food webs and are the

primary remineralizers of organic matter. Macrofaunalorganisms frequently ingest bacteria associated withsediments and can dramatically influence biomass [15,35], growth [8, 39] and activity [2, 8, 23] and altercommunity composition [9, 10, 54].

Although deposit-feeder injestion is usually thoughtof as a predator–prey (+, )) interaction, we argue herethat treatment as a disturbance is more appropriate.When compared to the typical relationship of a predatorand prey, the effect of a deposit feeder on ingested mi-crobes is less selective and a single encounter influencesmicrobial communities as opposed to individuals. Theimmense size disparity between deposit-feeding macro-fauna and bacteria is also uncharacteristic of classicpredator–prey interactions. In fact, it is not clear thatdigestion of bacteria provides a net gain to depositfeeders [24, 26, 40]. Furthermore, disturbances disruptthe physical environment as well as biota. Deposit feedingalters geochemical gradients within ingested sediments[1, 38], dramatically affects the quantity and architectureof microbial biofilms [6], and can change grain size,sorting, water content, and compaction of sediments [20,45].

Classic mechanisms of population regulation havebeen fairly well studied for sedimentary bacteria, yet al-most no empirical support exists for the importance ofeither competition [7, 51] or predation (by macrofauna[3, 24, 26], meiofauna [13, 26], or protozoa [12, 13, 25]).High levels of diversity [37, 50] and low stability [14, 15,48] of bacterial assemblages in sediments suggests animportant role for nonequilibrium determinants ofcommunity structure. Nonequilibrium models empha-size environmental disturbance and spatial heterogeneityin habitats in which a state of equilibrium is unusual.Community structure in this case is shaped by theinteractions of the heterogeneity of the physical–chemicalenvironment, disturbance, and recolonization.

The three mechanisms by which a species canrecolonize newly available space after disturbance areCorrespondence to: C.J. Plante; E-mail: [email protected]

154 DOI: 10.1007/s00248-003-1031-x d Volume 48, 154–166 (2004) d � Springer Science+Business Media, Inc. 2004

regrowth, immigration, and recruitment [44]. The rela-tive importance of these three mechanisms varies widelyamong systems and taxa. In the case of bacteria ininvertebrate egesta, regrowth refers to repopulation bysurvivors of gut passage, immigration to those bacteriacolonizing egesta from adjacent sediment, and recruit-ment to colonization from air (emersed egesta) oroverlying water (immersed egesta). By these definitions,‘‘recruitment’’ can be ignored in the present study inintertidal sediments, as bacterial densities in air are manyorders of magnitude less than in marine sediments.

The rate of recovery and the character of ensuingsuccessional changes will be dependent upon the mech-anism of initial recolonization. The significance of dis-turbances by deposit feeders to the structure of microbialcommunities in bulk sediments will be determined by thefrequency of disturbance relative to the rate of recovery.

Here we report on comparative studies used todetermine whether deposit-feeding activities by threediverse invertebrate taxa constitute significant distur-bances to sedimentary bacterial assemblages, and on fieldexperiments designed to ascertain mechanisms and rela-tive rates of recolonization. We employed epifluorescencemicroscopy, metabolic profiles, and molecular techniquesto characterize both quantitative and qualitative aspectsof the microbial community structure.

Materials and Methods

Study Site and Species. Samples were collected from,and experiments were performed at, Breach Inlet(32�46¢23¢¢ N, 79�48¢53¢¢ W), a semi-sheltered beach onthe northeast end of Sullivans Island, South Carolina,USA (see [42] for detailed description). Fecal materialswere collected from the three dominant deposit feeders:Nereis succinea (Polychaeta: Neriedae), Balanoglossusaurantiacus (Enteropneusta: Ptychoderidae), and Lepto-synapta tenuis (Holothuroidea: Synaptidae). Althoughthe distributions of all three species overlapped on thebeach, the mean location of N. succinea was slightlyhigher than that of B. aurantiacus, which was againslightly higher than the average location of L. tenuis.

Design of Studies. Here we report on studiesconducted both in November 1998 and in April 2002.The intent of the 1998 study was to test whether feedingby these three deposit feeders significantly altered struc-ture of bacterial assemblages relative to sediments, and tomake comparisons among the three deposit-feedingspecies. One goal of the 2002 study was to test the gen-erality of the trends observed in 1998. However, genome-based techniques were employed to compare qualitativeaspects of disturbance in place of the physiological pro-files (Biolog plates) used in the earlier study. In addition,

the 2002 study also included field experiments designedto distinguish mechanisms of recolonization.

Sample Collection and Preparation. To collectfresh egesta during the 1998 study, existing fecal moundsor coils were flagged and numbered to mark their loca-tions, then brushed away. After the next egestion (t = 0h), fresh fecal materials were collected from each speciesusing either a sterile 1-cc syringe (with Luer end re-moved) or clean spatula. Surficial sediments (top �3mm) were similarly sampled at this time. Initially, 20-mLnine-salt solution (NSS) [53] was added to each sample.Bacteria were dislodged from the sediment using a shortburst (20 s) of sonication with a 3-mm sonic probe(Branson Sonifier 250) at setting 4 on the output control(amplitude 306 lm, power output 65 W). Sonicationoptimization studies demonstrated that the highestaverage well color development (AWCD; index of cellrespiration, see below for details) in the Biolog platesoccurred with this duration and intensity of sonication.Serial dilutions (10)1, 10)2, 10)3) from these sampleswere used for Biolog plates, CTC assays, and total counts.Samples were then dried so as to normalize to dry gsediment)1. Values for blanks (tubes of 20 mL NSS) weresubtracted for each measurement.

For the comparative component of the 2002 studies,methods were similar but varied in that samples wereplaced into 10-mL sterile, filtered (0.2 lm) seawater, andthat sediment samples were taken separately, fromslightly different beach elevations, with each invertebratespecies (i.e., fecal and sediment samples were paired). Inaddition, samples for molecular analysis were also takenfrom ‡3 replicates of each type of egesta and associatedsediments. Approximately 0.5 g of each was collected viaspatula and placed into 2-mL conical microtubes con-taining �2.5 g 0.1-mm silica/zirconia beads (BiospecProducts, Battlesville, OH [34]). These samples werefrozen at )20�C until DNA extractions were performed.To follow bacterial recolonization of the egesta of thesedeposit feeders, fecal and ambient, surficial sedimentsamples were collected over a 2- to 3-h period. After theinitial egestion (t = 0 h), 100-mm diam. round polysty-rene plates (cut from petri dishes) were inserted at a 45�angle into the sediment beneath the egesta to block fur-ther addition of fecal material (cf [15]). To test whetherrecolonization was due to in situ growth vs migrationfrom underlying sediments, we placed ‘‘latrines’’ (see Fig.1 in [42]) over burrow exits to capture and isolate egestafrom sediments. Latrines, fecal materials on the sedimentsurface, and control areas for sediment samples werecovered with 5.5-cm high · 11.5-cm diam. cylindricalglass culture dishes to inhibit evaporation [42]. Allsamples, sediment (SED), naturally incubated fecal casts(FC), and fecal casts on latrines (LAT), were sampled asdescribed above for enumeration and molecular work.

C.J. PLANTE, S.B. WILDE: BACTERIAL RECOLONIZATION AND SUCCESSION IN SEDIMENTS 155

Enumeration. We enumerated metabolically ac-tive bacteria through time using the fluorogenic redoxdye 5-cyano-2,3-ditolyltetrazolium chloride (CTC; Poly-sciences, Inc., Warrington, PA) [47]. This tetrazolium saltis used as an artificial electron acceptor in the electrontransport system, indicating oxidative cell metabolism.Aliquots (2 mL) were taken from the diluted, sonicatedsample and transferred to 15-mL centrifuge tubes for theCTC reduction assay. CTC (200 lL of 25 mM) was addedto each tube and incubated (3 h) at 25�C in the darkwhile shaking (200 rpm). Samples were fixed in formalin(4% final concentration) for subsequent direct counts ofactive and total bacteria.

DAPI counts were made using a modified version ofthe protocol of Hymel and Plante [22]. Briefly, fixedsamples were centrifuged at 4000 g for 15 min, thenresuspended in Trizma buffer (0.05 M, pH 8.10) and adispersing agent (0.5% Triton X-100) and sonicated for20 s with a 3-mm sonic probe at 65 W. Samples werethen stained with DAPI (5 lL mL)1) for 20 min, recen-trifuged to remove stain, and concentrated onto 0.2-lmblack polycarbonate membranes (Poretics, Livermore,CA). For each sample and filter set, 20 grids (or more)were counted to include >200 total cells per slide.

Both total and active counts were made from eachslide using a Nikon epifluorescence scope at 1250·. Be-cause of interference of the CTC with DAPI, total bacteriawere taken as the sum of active + DAPI-staining cells [4].DAPI-staining bacteria were counted using a UV filter set(Omega XFO2, 330WB80 exciter, 400EFLP emitter).Active bacteria (CTC staining) were counted using arhodamine filter set (Omega 605DF55, center wavelength605 nm, discriminating filter, full band width at halfmaximum transmission 55 nm).

Metabolic Profiles. Biolog GN microtiter plates(Biolog Inc., Hayward, CA) were used to provide com-munity-level physiological profiles. Each plate is loadedwith 95 different carbon sources and a redox dye (tet-razolium violet). Bacterial respiration reduces the tetra-zolium dye to formazan in active cells, so that the patternof colored wells (different carbon sources) represents ametabolic fingerprint. To achieve the optimal dilution forthe Biolog inoculations, 20 mL more NSS was added toeach sample and was vortexed for 30 s at high speed.Biolog plates were then inoculated with 150 lL well)1 ofthe supernate using a multipipettor and incubated aer-obically at RT. Color formation in microplate wells wasanalyzed at t = 0, 24, 48, and 72 h (and 96 h for B.aurantiacus samples) on a Titertek Multiskan Plus(Titertek, Huntsville, AL) microplate reader at 570 nm.

DNA Extraction, PCR, and DGGE. Bacterial cellsin sediments were physically lysed using bead-millhomogenization. After frozen samples were thawed, 250

lL of 10% SDS (SDS–Tris–NaCl: 100 mM NaCl–500 mMTris pH 8–10% SDS) was added to sample in conicalmicrotubes and brought to volume with sodium phos-phate buffer (dibasic, 100 mM, pH 8). Each tube wasshaken at 1700 rpm for two cycles of 2.5 min using aMini Bead Beater-8 (Biospec Products, Bartlesville, OK).Following centrifugation (5 min, 16,000 g), supernateswere placed in sterile 1.5-mL microcentrifuge tubes. Twoextractions with equal volumes of phenol:chloro-form:isoamyl alcohol (25:24:1 vol) were performed, fol-lowed by one extraction with equal volume ofchloroform:isoamyl alcohol (24:1 vol). Sodium acetate (3M, pH 5.4) was added to the aqueous supernatant to givea final concentration of �0.3 M sodium acetate. DNAwas pelleted (5 min, 16,000 g) in an equal volume of ice-cold isopropanol (>1 h), then washed with an equalvolume of cold 70% ethanol (pelleted by centrifugation at16,000 g for 5 min). After evaporation of EtOH at roomtemperature, the pellet was resuspended in 50 lL TEbuffer. The Wizard PCR DNA purification system (Pro-mega, Madison, WI, USA) was used for final purificationusing manufacturer’s instructions.

The primer set for amplification of the V3 region (5¢-ATTACCGCGGCTGCTGG-3¢ and 5¢-CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCAGGGGGGCTACGGGAGGCAGCAG-3¢) of the 16S rDNA was usedfor PCR. PCR reactions (100 lL) consisted of PCR buffer(10 mM HCl, pH 9, 50 mM KCl, 0.1% Triton X-100;Promega), 2.25 mM MgCl2, 0.8 mM dNTPs, 0.5 lM ofeach primer, 1.2 units of polymerase (Taq DNA poly-merase; Promega), and 2 lL of template. PCR consistedof an initial step for 5 min at 94�C, and 30 cycles of 94�Cfor 1 min (denaturation), 1 min for annealing, and 72�Cfor 3 min (extension). Annealing temperature was 65�Cfor the first cycle, then decreased 1�C every other cycleuntil reaching 55�C, where temperature was unchangedover the last 10 cycles. The 30 cycles were followed by 7min incubation at 72�C [46], then samples were held at4�C until removal from thermal cycler.

PCR products were purified and concentrated usingthe Wizard PCR DNA purification system. Final elutionof DNA from each 100 lL reaction from the Wizardminicolumns with 35 lL TE resulted in a �3-fold con-centration. DNA was then quantified fluorometricallyusing the PicoGreen dsDNA quantitation kit (MolecularProbes, Eugene, OR, USA) according to manufacturer’sinstructions.

Denaturing gradient gel electrophoresis (DGGE) wasperformed with the D-Code Mutation Detection System(Bio-Rad Laboratories, Hercules, CA, USA). An 8%polyacrylamide gel with a gradient of DNA denaturantagent was cast by mixing solutions of 40% and 60%denaturant agent (100% denaturant is 7 M urea and 40%deionized formamide). Sample volumes were varied sothat 300 ng of PCR product was loaded for each sample

156 C.J. PLANTE, S.B. WILDE: BACTERIAL RECOLONIZATION AND SUCCESSION IN SEDIMENTS

and the gel was run at 70 V for 16 h at 60�C in 1· TAEbuffer (40 mM Tris [pH 7.4], 20 mM sodium acetate, 1mM EDTA). Gels were stained with 1· Sybr Gold(Molecular Probes) for 30 min and visualized with UVlight in a Model 1000 VersaDoc imaging system (Bio-RadLaboratories).

Data Analysis. One-way ANOVA was used totest for effects of sample type (sediment or fecalmaterial associated with each invertebrate species) inthe 1998 comparative studies. For the 2002 enumera-tion data we used two-way ANOVA to test for effectsof time and sample type (sediment, isolated, or natu-rally incubated fecal casts) on total bacterial abundanceand numbers of active bacteria. Prior to analysis, data

were tested for normality (Lillefor’s test) and hetero-scedasticity (Levene’s test) and natural log transformedas required (and retested). We also tested for outlyingvalues and removed from analysis only those valuesdetermined to be ‘‘extreme’’ outliers (according to[49]). If main effects were significant, planned pairwisemultiple comparison tests employed Fisher’s LSD cor-rection; if main effects were not significant, the moreconservative Bonferroni’s adjustment was used [33].

Community level physiological comparisons wereconducted looking at two components: overall meta-bolic rate and resource utilization patterns [16]. Toanalyze the overall intensity of color development,average well color development (AWCD) after 48 h

Figure 2. Mean ± SEM of (a) active bacterial densities, and (b)total bacterial densities in 2002 Breach Inlet samples. SED: surfacesediment; Ba: B. aurantiacus; Ns: N. succinea; Lt: L. tenuis; FC: fecalmatter. Asterisks indicate significant difference between fecal andpaired sediment sample at the a < 0.10 (*) and 0.05 (**) levels.

Figure 1. Mean and SEM of (a) active bacterial densities, and (b)total bacterial densities in 1998 Breach Inlet samples. SED: surfacesediment; Ba: B. aurantiacus; Ns: N. succinea; Lt: L. tenuis; FC: fecalmatter. Different letters indicate significant (a = 0.05) differenceamong sample types.


incubation was contrasted among sample types usingANOVA, as described above for bacterial count data.The AWCD for each plate was calculated as the averageabsorbance of the 95 test wells after subtracting theabsorbance of the control well (blank) and setting anynegative values equal to 0. Similarities in carbon sourceutilization patterns of microbial communities wereinvestigated using multivariate analysis. To correct forunequal inoculum size and rates of plate development,utilization patterns at different incubation times (24 hfor L. tenuis egesta and sediment, 72 h for B. auran-tiacus and N. succinea egesta) were compared, whenAWCDs were roughly equal (�0.4). In order to visu-alize differences in community respiration patterns, weused principal component analysis (PCA) with theBiolog data [17]. Additionally, with the first fiveprincipal components as variables, we used MANOVAto test for differences among the sample types [19].

The Quantity One software (Bio-Rad) used forDGGE gel image acquisition was also employed foranalysis. Following removal of background intensity fromlanes using the rolling disk method, the software per-forms a density profile through lanes, detects individualbands, and matches bands occupying the same positionin the different lanes. Similarity indices were then cal-culated using band position and intensity (weighted) orband position only (unweighted) via the method of theDice Coefficient. Replicates (typically three) of eachsample type were run numerous times on multiple gels so

that quantitative (i.e., profile similarity values) andqualitative (e.g., bands appearing to be unique to asample type) aspects could be checked for consistency.

Results

Comparative Study. Numerical trends of bacteriapassing through the gut varied among the three deposit-feeding species but generally were consistent for a givenspecies on different dates. In 1998, densities of activebacteria varied significantly by sample type (P = 0.002).L. tenuis fecal mounds appeared to contain highestdensities (Fig. 1a), significantly greater than seen in B.aurantiacus (P < 0.001) or N. succinea (P < 0.001)mounds, but not significantly different from those inadjacent sediments (P = 0.133). Both B. aurantiacus andN. succinea mounds contained lower numbers of activebacteria as compared to sediments (P = 0.001 and<0.001, respectively), but did not differ from one another(P = 0.987). Total numbers showed similar, but notidentical, patterns (Fig. 1b). The effect of sample type wassignificant (P = 0.002), with densities in L. tenuis fecessignificantly higher than adjacent sediment (P = 0.026),B. aurantiacus (P = 0.007), and N. succinea feces (P <0.001). N. succinea mounds contained lower densities oftotal bacteria as compared to sediments (P = 0.046), butnumbers in B. aurantiacus mounds did not differ fromeither N. succinea mounds (P = 0.679) or sediment(P = 0.223).

Figure 3. Comparison of catabolic profilesfrom 1998 Breach Inlet sediment (SED) andfecal (FC) samples from B. aurantiacus (Ba),N. succinea (Ns), and L. tenuis (Lt); plot offirst (PC 1) and second (PC 2) principalcomponents derived from PCA for 48-hBiolog readings.


In the 2002 samples, active bacteria were also fewerin number in N. succinea egesta as compared to pairedsurface sediments (P = 0.071; Fig. 2a). In contrast, B.aurantiacus patterns differed between 1998 and 2002samples in that higher numbers were found in egesta in2002 (P = 0.036; Fig. 2a). Fecal samples and sedimentsdid not differ significantly for L. tenuis (P = 0.732; Fig.2a). The reduction in numbers of total bacteria from N.succinea ingesta to egesta (P = 0.012; Fig. 2b) also agreedwith 1998 results. Although densities of total bacteriaappeared to be greater in L. tenuis egesta compared topaired sediment samples, as in the 1998 results, thisdifference was not significant (P = 0.207; Fig. 2b), norwas there a significant difference between B. aurantiacusfecal samples and sediments (P = 0.712; Fig. 2b).

PCA (Fig. 3) and MANOVA results illustrate thesignificant differences between the carbon source utili-zation patterns of the microbial communities associatedwith two of the deposit feeders, B. aurantiacus and N.succinea, versus those of L. tenuis and ambient sediment.The metabolic fingerprint of the bacterial assemblage inL. tenuis egesta was not significantly different from that inambient sediment (P = 0.412), but was different fromthose in N. succinea (P = 0.002) and B. aurantiacus (P <0.001) egesta. N. succinea and B. aurantiacus egesta wereindistinguishable (P = 0.866), whereas both were distinctfrom sediment (P < 0.001 for both). Analysis of AWCDresults were similar in that L. tenuis fecal samples andsediment were indistinguishable (P = 0.535; 1-wayANOVA), while color development in L. tenuis sampleswas significantly greater than in egesta of both B. au-rantiacus (P < 0.001) and N. succinea (P < 0.001). Sedi-ment AWCD likewise was higher than both B.aurantiacus (P < 0.001) and N. succinea (P < 0.001; datanot shown).

DGGE banding patterns revealed subtle differencesamong compositions of bacterial assemblages in fecal andsediment samples (Fig. 4). In an attempt to add a mea-sure of rigor to our DGGE analyses we ran replicate DNAsamples (N = 2 or 3) for each sample type numeroustimes on multiple gels. Although gel-to-gel variation wasobserved, in general, within-sample (i.e., between-repli-cates) similarity was greater than that for between-sampletype comparisons. For instance, between-replicate meansimilarities for the gels shown here were 96.4, 94.1, and93.3% for Figs. 4, 7, and 8, respectively (all bands of gelsnot shown). Similarity values much below these levelslikely indicate real differences. Unweighted comparisonsamong the sediment samples taken from slightly differentelevations on the beach (associated with the differentdeposit feeders) were similar (similarities of 91.5, 93.5,and 91.2%; Table 1), but showed somewhat less similaritythan did replicate samples. Differences between animalfecal samples and their paired sediment samples (pre-sumed to be ingesta) showed greater differences (89.3,

90.6, and 71.7%; Table 1), with the greatest apparentcompositional disturbance seen with B. aurantiacusfeeding (Table 1). Comparisons among fecal samples alsoshowed relatively lower similarities (84.0, 72.7, and82.5%; Table 1), again indicating that sediments weredisturbed with gut passage and that each deposit-feedingspecies had a different effect on bacterial compositions.When weighted comparisons were performed, similaritieswere invariably lower (usually by 20–30%), but all trendsremained the same as for unweighted comparisons.

Figure 4. Representative DGGE gel profiles (unretouched) for2002 Breach Inlet fresh fecal and sediment samples. SED: surfacesediment; Ba: B. aurantiacus; Ns: N. succinea; Lt: L. tenuis; FC: fecalmatter. Indicated bands are unique or are more intense in par-ticular sample types and are discussed in text.


A more qualitative examination of gels indicated thata few bands (i.e., bacterial phylotypes) were unique to, orat least relatively more important in, particular sampletypes. For instance, bands 16 and 19 were most intense inL. tenuis samples (both ingesta and egesta), and band 9appeared to represent a phylotype especially important inN. succinea samples. In the additional gels on which thereplicates were run, absolute similarity index values var-ied, but the same general patterns were observed (i.e.,SED–SED comparisons showed greater similarity thandid FC–FC or FC–SED comparisons, and B. aurantiacusFC samples were most different from their paired sedi-ment samples and from the other FC samples). The morequalitative observations regarding specific bands (i.e.,bands 9, 16, and 19) were also consistent in these gels.Band 17, which appears to represent a phylotype stronglyassociated with sediment samples as compared to fecalsamples (Fig. 4), did not show this pattern on the othergels.

Recolonization Study. Bacterial recolonization ofthe egesta of N. succinea was evident in that total bacterialdensities increased through time (P = 0.011); althoughactive bacterial numbers in fecal matter were higher at 3 hthan at 0 h, this increase was not significant (P = 0.224;Fig. 5a). Significant temporal change was not observedfor latrine-isolated egesta (P = 0.330 and 0.844 for activeand total bacteria, respectively; Fig. 5). A decrease in totalbacterial densities in sediment samples through timetrended toward significance (P = 0.077; Fig. 5b), but thischange was not significant for the CTC–staining portionof bacteria (P = 0.180; Fig. 5a).

Results were more complex in the B. aurantiacusrecolonization experiment. As for N. succinea experi-ments, no significant temporal changes in active or totalbacterial densities were observed in latrine samples (Fig.6). However, the only real change in naturally incu-bated egesta was a decline in active bacteria between 0and 1 h incubation (P = 0.006; Fig. 6a). Surprisingly,active bacterial densities varied significantly betweenfresh FC and LAT samples (P < 0.001; Fig. 6a), which

at t = 0 h is essentially the same sample type. In sedi-ments, density of active bacteria declined through time(P = 0.003 for 0 vs 1 h; Fig. 6a), whereas total bacterial

Table 1. Percent similarity of DGGE banding patterns among fresh fecal (FC) and sediment (SED) samples

Similarity matrix

Lane Ba-FC Ns-FC Lt-FC SED(Ba) SED(Ns) SED(Lt)

Ba-FC 100 84.0a 72.7a 71.7b 75.0 70.6Ns-FC 100 82.5a 85.2 89.3b 81.4Lt-FC 100 90.9 85.2 90.6b

SED(Ba) 100 91.5c 93.5c

SED(Ns) 100 91.2c

SED(Lt) 100

The Dice Coefficient was used to compute similarities, and comparisons were unweighted (i.e., used only band position).SED: sediment; Ba: B. aurantiacus; Ns: N. succinea; Lt: L. tenuis; FC: fecal matter.aFC vs FC; bFC vs SED; cSED vs SED.

Figure 5. Mean ± SEM of (a) active bacterial densities, and (b)total bacterial densities through time in N. succinea recolonizationexperiments. SED: surface sediment; LAT: latrine-isolated fecalmatter; FC: naturally incubated fecal matter. Asterisks indicatesignificant temporal change at the a < 0.10 (*) and 0.05 (**) levels.


numbers increased over this same time span (P = 0.002;Fig. 6b).

DGGE similarity index values suggest that N. succi-nea–associated sediment samples changed little, if any,through time (94.1% similarity; Fig. 7, Table 2). Natu-rally incubated fecal samples (FC) showed the mostchange over 3 h (87.5% similarity; Table 2), whereas la-trine-isolated fecal samples (LAT) showed the leastchange (98.5% similarity between t = 0 and 3 h; Table 2).Although this suggests that migration is responsible forrecovery, this does not appear to be a simple case ofindiscriminate recolonization by the sedimentaryassemblage, as the fecal sample community at 3 h is nomore similar to the sedimentary community than is the 0

h sample (Table 2). It is more likely that a highly motilesubset of the assemblage is responsible for numericalincreases and compositional changes. Although the twointense bands that appear to be unique to FC-3 samples(bands 16 and 31 of Fig. 7) would seem to support thisidea, only band 16 was found consistently on other gelswith replicate samples. These additional gels did, how-ever, support the patterns seen in the gel in Fig. 7, i.e.,that aged fecal sample did not come to more closelyresemble sediment samples, and that the greatest changeswere in naturally incubated fecal samples with the leastchange in latrine samples (data not shown).

Likewise for B. aurantiacus time series data illustratedin Fig. 8, DGGE similarity index values suggest that la-trine-incubated fecal samples changed least through time(96.1% similarity between 0 and 3 h; Table 3). Naturallyincubated fecal samples (FC) again showed the mostchange over 3 h (81.8% similarity; Table 3), whereassediment samples were 87.3% similar between 0 and 3 h(Table 3). Again the fecal sample community at 3 h wasno more similar to the sedimentary community than wasthe 0 h sample (Fig. 8, Table 3). Two bands of specialnote in Fig. 8 appear to represent a phylotype that isrelatively more abundant in surficial sediments than inegesta (and so may be especially susceptible to digestive

Figure 6. Mean ± SEM of (a) active bacterial densities, and (b)total bacterial densities through time in B. aurantiacus recolon-ization experiments. SED: surface sediment; LAT: latrine-isolatedfecal matter; FC: naturally-incubated fecal matter. Asterisks indi-cate significant temporal change at the a < 0.10 (*) and 0.05 (**)levels.

Figure 7. Representative DGGE gel profiles (unretouched) of N.succinea fecal and sediment samples at 0 and 3 h incubation. SED:surface sediment; LAT: latrine-isolated fecal matter; FC: naturallyincubated fecal matter. Indicated bands are unique or are moreintense in particular sample types and are discussed in text.


lysis) and another that is much stronger in fecal samplesand presumably the gut (bands 16 and 21, respectively).In the two additional gels run with replicates of thesesample types (data not shown), patterns for these notablebands are consistent, as was the pattern that the fecalsample community at 3 h was no more similar to thesedimentary community than was the 0 h fecal sample.However, the observation that greatest changes were innaturally incubated fecal samples and least change in

latrine samples was less clear in the additional gels. Withthese replicate samples there were smaller differences inthe percent change over 3 h among the LAT, SED, or FCsample types.

Discussion

In general, numerical results from the comparativestudies of 1998 and 2002 were in agreement. In bothstudies, higher numbers of active and total bacteria werefound in the egesta of L. tenuis as compared to nearbysurface sediments, whereas bacteria were removed fromsediment with passage through the gut of N. succinea.Evidence for this type of disturbance was less clear in B.aurantiacus and may show temporal variation. Priorstudies [42] also noted significantly fewer bacteria in N.succinea egesta vs sediments, with no significant removalobserved for B. aurantiacus. Thus the disturbance trendsnoted in this study would appear to be generalizable toother times and sites.

Community structure and function are also depen-dent upon the composition of microbial assemblages.Our assessments of these more qualitative effects of de-posit feeding on bacterial assemblages were generallyinternally consistent and agreed with previous studies. Interms of both metabolic potential and DGGE bandingpattern, ingestion by L. tenuis appeared to have minorimpacts on the bacterial assemblages of sediments. Incontrast, both N. succinea and B. aurantiacus ingestionaltered microbial community metabolic potential andappeared to alter assemblage composition, although thelatter effect was less clear. One reason for the moreobvious disturbance effect in Biolog plates is that DGGEgels and similarity matrices shown here were from un-weighted rather than weighted comparisons. This may beespecially important as it has often been observed thatdisturbances normally have little effect on species rich-ness—more likely, major shifts in relative dominance andspecies evenness result [44]. Also important is the highlikelihood that bacteria could be inactivated or killed bydigestive processes within the animal gut, yet DNA would

Figure 8. Representative DGGE gel profiles (unretouched) of B.aurantiacus fecal and sediment samples at 0, 1, and 3 h incubation.SED: surface sediment; LAT: latrine-isolated fecal matter; FC:naturally incubated fecal matter. Indicated bands are unique or aremore intense in particular sample types and are discussed in text.

Table 2. Percent similarity of DGGE banding patterns among N. succinea naturally incubated fecal (FC), latrine-incubated fecal (LAT),and sediment (SED) samples at 0 and 3 h

Similarity matrix

Lane FC-0 FC-3 LAT-0 LAT-3 SED-0 SED-3

FC-0 100 87.5a 94.1 95.5 95.7 92.5FC-3 100 87.5 88.9 89.2 85.7LAT-0 100 98.5b 95.7 95.5LAT-3 100 94.1 93.9SED-0 100 94.1c

SED-3 100

The Dice Coefficient was used to compute similarities, and comparisons were unweighted (i.e., used only band position).aFC0 vs FC3; bLAT0 vs LAT3; cSED0 vs SED3.


persist long enough to be extracted, and subsequentlyproduce DGGE bands.

Taken together, the numerical, Biolog, and DGGE dataindicate that L. tenuis feeding represents a negligible dis-turbance, N. succinea most clearly removes bacteria andalters community structure, and B. aurantiacus egesta andsediments vary little in bacterial number but differ quali-tatively at least as much as do bacterial assemblages in N.succinea egesta. That bacterial numbers are actually higherin the egesta of L. tenuis is most likely due to particle sizeselection by the animal. L. tenuis ingests particles with biasagainst both small and large grain sizes as compared tosurrounding bulk sediments [36, 43]; thus it ‘‘samples’’ thesediment in a way that we could not duplicate. Although itis impossible to conclude from the numbers alone thatdigestive removal did not occur, certainly any lysis of bac-teria did not exceed the effects of selection. Furthermore,previous studies have shown that the digestive fluids of thisanimal have relatively weak bacteriolytic capacity [41]. Thelack of apparent compositional changes (Biolog and DGGEresults) support the notion that this animal does not rep-resent a significant source of removal, or disturbance, tosedimentary bacteria.

N. succinea, on the other hand, most clearly removesbacterial biomass and also qualitatively alters bacterialassemblages. In general, polychaetes show high relativerates of digestive potency (i.e., enzymatic activity andsurfactancy) relative to other taxonomic groups of de-posit feeders [30], and, more specifically, higher bacte-riolytic capacity [41].

Bacterial numbers in B. aurantiacus egesta showedlittle difference from sediments, yet compositional andmetabolic changes were relatively large. This suggests thatour assumption that surficial sediments represent in-gested materials may be incorrect. It is likely that B.aurantiacus also ingests deeper, anoxic sediments. At ourstudy site, at least a fraction (�20%, [54]) of fresh B.aurantiacus coils are gray in color, and almost certainlywere reducing when ingested. Studies of Duncan [11]

suggest that surface sediments are rapidly subducted todepths of 3–10 cm to be ingested by B. aurantiacus.However, he also demonstrated that these deposit feedersshift the positions of their burrows and feeding funnelson a daily basis. Thus, anoxic sediments likely are in-gested both when the animal relocates and, to a lesserdegree, in normal feeding modes. If individuals of the B.aurantiacus population ingest vastly different assemblagesof bacteria from a range of sediment depths, this mighthelp explain the high variability observed in active bac-terial densities in fresh egesta (Fig. 6a), the disparitybetween fresh fecal (naturally incubated) and latrinesamples in our recolonization experiments (Fig. 6a), andthe lack of consistency with respect to numerical changeswith gut passage for this species [42, 54]. The ‘‘disturbed’’patches of sediment, containing bacterial assemblagesmetabolically and compositionally different from adja-cent surficial sediments, result then from animal trans-location of deep sediments to the surface, not digestion.That hemichordates also typically exhibit weak bacteri-olytic activity [41] supports this idea.

Given that numerical recovery was absent when N.succinea egesta were isolated on latrines, yet apparent infecal materials incubated on sediments, we can concludethat recolonization is primarily by immigration fromadjacent sediments. The modest change in DGGEbanding patterns of latrine-isolated samples relative tonaturally incubated fecal materials for both deposit-feeding species further supports this hypothesis. Theseresults corroborate earlier findings [42] that employedmetabolic profiles (i.e., Biolog plates) to show thatrecolonization of exposed intertidal sediments is pri-marily via immigration. In that study, metabolic fin-gerprints in latrine-isolated egesta remained unchangedand statistically distinct from those of sediments overtime, whereas fingerprints of naturally incubated fecesdiverged from those on the latrines and came to moreclosely resemble profiles of sedimentary bacterialassemblages.

Table 3. Percent similarity of DGGE banding patterns among B. aurantiacus naturally incubated fecal (FC), latrine-incubated fecal(LAT), and sediment (SED) samples at 0, 1, and 3 h

Similarity matrix

Lane FC-0 FC-1 FC-3 LAT-0 LAT-1 LAT-3 SED-0 SED-1 SED-3

FC-0 100 93.3a 81.8a 93.3 87.3 91.9 80.0 82.2 79.5FC-1 100 81.2a 94.9 91.9 93.5 82.2 86.8 81.6FC-3 100 84.1 80.0 79.4 78.1 77.6 77.6LAT-0 100 91.9b 96.1b 82.2 86.8 81.6LAT-1 100 93.2b 84.1 86.1 80.6LAT-3 100 80.6 85.3 82.7SED-0 100 95.8c 87.3c

SED-1 100 91.9c

SED-3 100

The Dice Coefficient was used to compute similarities, and comparisons were unweighted (i.e., used only band position).aFC0 vs FC1 vs FC3; bLAT0 vs LAT1 vs LAT3; cSED0 vs SED1 vs SED3.


Previous studies have indicated that the majority ofsedimentary bacteria are physically attached to sedimentgrains (e.g., [5, 7]). Immigration then could occur withfree-living porewater bacteria, or attachment must bereversible over short (minutes to hours) time scales.Considering the first option, most studies suggest thatfree-living bacteria in aquatic sediments number less than10% of attached bacteria (e.g., <1% [31], <10% [18]).Given the levels of recovery noted in this study (Fig. 5),we conclude that porewater bacteria are an unlikelysource of colonists. There is evidence, however, thatparticle-attached bacteria could contribute to recovery, asLeff et al. [27] have demonstrated rapid attachment anddetachment of bacteria in freshwater sediments. In eithercase, bacterial swimming speeds (>50 lm s)1; [52])would not preclude this mechanism of recolonization.

Although rapid bacterial growth in the hindgutand/or feces of deposit feeders has previously beenrecorded [8, 39], this potential mode of recolonizationwas not important in this study (i.e., no recovery inlatrine samples). Given the short bacterial doublingtimes that have previously been reported for bacteria inguts (i.e., 1–3 h; [8, 39]), numerical recoveries reportedin this study certainly could have resulted from re-growth. However, these rapid rates of bacterial growthin deposit-feeder guts and egesta have not always beenobserved (e.g., [21, 29]), and growth rates have notbeen examined in relation to the three species studiedhere. In fact, in some of the studies in which rapidgrowth of bacteria in egesta had been concluded basedon numerical increases [21, 28, 32], other mechanismsof recolonization cannot be excluded.

Interestingly, as disturbed sediments ‘‘recovered,’’they did not become more similar to surficial sedi-ments in their DGGE banding patterns. This wasunexpected as metabolic profiles of naturally incubatedfeces did become significantly more similar to sedi-mentary profiles over time [42]. It is likely then thatthese disturbed patches are first recolonized by a highlymotile, opportunistic subset of the bacterial assemblageof adjacent sediments. The presence of a few uniquebands in aged egesta supports this idea, although itcannot be ruled out that these few bands representphylotypes introduced from the animal itself. To dis-tinguish the source of these particular bands, samplingfrom within the gut would be required.

Comparisons of the mid- and later successionalstages should provide more insight into the nature ofcommunity structuring in these disturbed patches ofsediment. It is unknown whether these communities aredominance-controlled, with predictable species sequencesfrom superior colonizers to competitors, or founder-controlled, in which all species are roughly equivalentcompetitors, species diversity is unpredictable, andcomposition is determined largely by chance. Ideally, B.

aurantiacus samples at 1 and 3 h allow such comparisons.Based on the observations of modest temporal changes inDGGE patterns and few new bands arising at 3 h, wewould conclude little competition-driven turnover ofspecies (i.e., a founder-controlled community). However,whether a significant portion of the full successional se-quence was observed in this study is in question. Al-though significant numerical and qualitative recovery ofbacterial assemblages in disturbed sediments was notedover the short (£ 3 h) periods of observation during lowtide, it was clear especially from DGGE results that thesepatches had not completely recovered or come toresemble adjacent surficial sediments. Similar workemploying subtidal deposit feeders would allow study oflater stages of succession.

Nonequilibrium models emphasize spatial heteroge-neity of the physical–chemical environment, disturbance,and recolonization as key elements in biological com-munity structure. Nonequilibrium theories attribute thecoexistence of similar species and high species diversity toprocesses of disturbance and recovery. In this study,however, because recolonization was primarily throughimmigration from adjacent sediments, we did not ob-serve greater species richness in egesta relative to sedi-ments—the source for colonists was restricted to thebacterial assemblage of those sediments. At the largerlandscape scale, however, we would predict greaterdiversity in areas with deposit-feeder disturbances rela-tive to undisturbed areas. Future work will attempt to testthis hypothesis with deposit-feeder removal and additionexperiments.

Temporal aspects of processes such as disturbanceand recolonization must also be understood for fullappreciation of the significance in structuring thesemicrobial communities. For instance, depending ondisturbance regime (disturbance frequency or magnitude,and rate of recovery), steady-state or constantly fluctu-ating mosaics can result. When disturbances are smallrelative to the landscape and/or infrequent relative to therecovery period of the biota, steady-state mosaics wouldbe predicted. We observed fairly subtle disturbances withsignificant recovery in short spans of time. The appar-ently ephemeral nature of these disturbances must beconsidered in light of the return intervals and lifespans ofthe sedimentary bacteria in question. Spatial and tem-poral aspects must then be integrated in patch dynamicsmodels to provide predictive capacity regarding com-munity structure and stability.

Acknowledgments

We thank Jeremiah Easley and Tom Busby for their helpwith field work and with epifluorescence microscopy.This work was supported by NSF grants OCE 95-04505,DEB 01-08615, and DBI 01-22336. Contribution #258 of


the Grice Marine Laboratory, College of Charleston,Charleston, SC.

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