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Applied and Environmental Microbiology, October 2007, p. 6112-6124, Vol. 73, No. 19
0099-2240/07/$08.00+0     doi:10.1128/AEM.00551-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Diatom-Derived Carbohydrates as Factors Affecting Bacterial Community Composition in Estuarine Sediments{triangledown}

Kelly Haynes,1 Tanja A. Hofmann,1 Cindy J. Smith,1 Andrew S. Ball,1,{dagger} Graham J. C. Underwood,1 and A. Mark Osborn1,2*

Department of Biological Sciences, University of Essex, Wivenhoe Park, Colchester CO4 3SQ, United Kingdom,1 Department of Animal and Plant Sciences, The University of Sheffield, Western Bank, Sheffield S10 2TN, United Kingdom2

Received 9 March 2007/ Accepted 26 July 2007


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ABSTRACT
 
Microphytobenthic biofilms in estuaries, dominated by epipelic diatoms, are sites of high primary productivity. These diatoms exude large quantities of extracellular polymeric substances (EPS) comprising polysaccharides and glycoproteins, providing a substantial pool of organic carbon available to heterotrophs within the sediment. In this study, sediment slurry microcosms were enriched with either colloidal carbohydrates or colloidal EPS (cEPS) or left unamended. Over 10 days, the fate of these carbohydrates and changes in ß-glucosidase activity were monitored. Terminal restriction fragment length polymorphism (T-RFLP), DNA sequencing, and quantitative PCR (Q-PCR) analysis of 16S rRNA sequences were used to determine whether sediment bacterial communities exhibited compositional shifts in response to the different available carbon sources. Initial heterotrophic activity led to reductions in carbohydrate concentrations in all three microcosms from day 0 to day 2, with some increases in ß-glucosidase activity. During this period, treatment-specific shifts in bacterial community composition were not observed. However, by days 4 and 10, the bacterial community in the cEPS-enriched sediment diverged from those in colloid-enriched and unamended sediments, with Q-PCR analysis showing elevated bacterial numbers in the cEPS-enriched sediment at day 4. Community shifts were attributed to changes in cEPS concentrations and increased ß-glucosidase activity. T-RFLP and sequencing analyses suggested that this shift was not due to a total community response but rather to large increases in the relative abundance of members of the {gamma}-proteobacteria, particularly Acinetobacter-related bacteria. These experiments suggest that taxon- and substrate-specific responses within the bacterial community are involved in the degradation of diatom-derived extracellular carbohydrates.


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INTRODUCTION
 
Intertidal estuarine mudflats are highly productive systems (24, 69) that can support abundant microphytobenthic (MPB) biofilms on the sediment surface. In sediments mainly comprised of fine silts and clays (<63 µm in size), these biofilms tend to be dominated by populations of motile, epipelic pennate benthic diatoms (2, 69). These photoautotrophs migrate vertically through the sediment, in response to tidal and diel cycles (14) and form conspicuous biofilms on the sediment surface during low tide exposure. During cycles of vertical migration, epipelic diatoms exude extracellular polymeric substance (EPS), primarily composed of a complex mixture of polysaccharides and glycoproteins (10) which, together with other extracellular carbohydrates excreted during photosynthesis, contributes to the mucilage component of intertidal biofilms (5, 16, 30). The pathway of EPS production, from photosynthetic carbon incorporation to EPS excretion, takes between 0.5 and 2 h (55, 66, 74) and is subject to variation in both rates of production and biochemical composition, in response to a variety of environmental factors including irradiance, temperature (75), and nutrient limitation (66, 67).

Diatom EPS exists as a continuum of material from closely attached "capsular" EPS surrounding cells, to loose "colloidal" carbohydrates and EPS secreted into the surrounding environment (67). Established extraction procedures, using a sequence of increasingly astringent methods can be used to fractionate these EPS. This results in the generation of a saline-soluble (colloidal) fraction containing both polymeric material, i.e., colloidal EPS (cEPS), and lower-molecular-weight carbohydrates; a hot water-extracted carbohydrate fraction (predominantly intracellular diatom storage carbohydrate, glucan, or chrysolaminarin); and a hot bicarbonate-extracted carbohydrate fraction (tightly bound and capsular EPS). These fractions differ in their chemical and physical properties (1, 5, 10, 68). Concentrations of these different EPS fractions can be as high as 8.7 mg of glucose equivalents per g (dry weight) of sediment (5). Consequently, there is a substantial pool of organic carbon being produced within these estuarine biofilms. Although the production patterns for these different EPS are relatively well described, little is known about the fate of MPB-derived EPS within sediments.

Two possible pathways of EPS loss from sediments are the removal of EPS to the overlying estuarine water during high tide and the mineralization of EPS by heterotrophic bacteria in situ. In pelagic systems bacterial and algal production are closely correlated (13), but in muddy sediments that contain significant quantities of detrital material the bacterial heterotrophy is high and is primarily driven by allochthonous inputs of organic matter (31). Despite high background levels of organic matter in muddy sediments, correlative studies of algal, bacterial, and enzymatic activity suggest that some degree of algal-bacterial coupling does occur (45, 50, 70). 13C-tracer studies have demonstrated transfer of photosynthetically fixed carbon from MPB to bacteria within hours, indicating rapid utilization of labile carbon sources, possibly including EPS (41). Diatom EPS may be sequentially degraded; with particular monosaccharides being selectively utilized by heterotrophs, as has been seen with the degradation of EPS from the freshwater planktonic diatom Thalassiosira (25). However, structurally complex EPS molecules may not be accessible to generalist heterotrophs, rendering them relatively refractory (17). Recent field data from intertidal sediments in situ, suggest that tightly bound EPS may be converted into colloidal EPS prior to breakdown into small carbohydrate moieties, a process probably driven by extracellular enzyme activity (30). A suite of extracellular enzymes are produced by bacteria for the breakdown of carbohydrates, including, for example, the exo- and endoglucanases (61). The enzyme ß-glucosidase hydrolyzes terminal nonreducing ß-D-glucose and has been extensively used as a general indicator of extracellular enzymes (32, 52). Although coupling between bacterial ß-glucosidase activity and biofilm EPS dynamics has been observed in situ (70), it is not known to what extent coupling exists between EPS production and subsequent utilization by specific EPS-degrading bacteria and also the identity of the bacteria involved. In estuarine sediments, the dominant bacterial taxa have been identified as members of the subphyla Flavobacteria and Sphingobacteria (previously Cytophaga-Flexibacter-Bacteroides) and {alpha}- and {gamma}-proteobacteria (35, 51, 58). Bacteria from these taxa have also been found to be associated with (marine) diatoms (29, 50). In addition, members of the {alpha}-proteobacteria and Flavobacteria and Sphingobacteria have also been found to be associated with particular cultured diatoms, e.g., Dytilum brightwellii, Thalassiosira weissflogii, Asterionella glacialis, Chaetoceros socialis, Leptocylindrus danicus, and Coscinodiscus spp. (50). In our earlier field study of the response of estuarine sediment bacterial communities to diatom inputs (30), only 5 of 59 operational taxonomic units within the total bacterial community, as determined by terminal restriction fragment length polymorphism (T-RFLP) analysis, exhibited a significant correlation with the production of cEPS and colloidal carbohydrates by diatoms. This suggests that algal-bacterial coupling in estuarine sediments is likely to involve particular bacterial taxa, rather than a response from the entire bacterial community.

Consequently, we have used here microcosm experiments to investigate changes in bacterial community composition, carbohydrate fraction composition, and extracellular enzyme activity in response to additions of different MPB (diatom)-derived carbohydrates. Slurries of estuarine intertidal sediments were enriched with two different biofilm-EPS fractions: colloidal carbohydrate (labile material extracted in saline and including both low-molecular-weight [LMW] and polymeric components) and cEPS (EPS isolated from colloidal extracts by precipitation in ethanol). These experiments were used to determine whether particular bacterial taxa showed substrate-dependent shifts in presence and/or abundance in response to the different available carbon sources.


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MATERIALS AND METHODS
 
Field sampling.
Sediment samples for both the extraction of colloidal carbohydrates and cEPS and the subsequent establishment of slurry microcosm experiments were taken from Alresford Creek (51°50.2'N, 0°59.5'E) in the Colne Estuary, Essex, United Kingdom. Alresford Creek is a typical mid-estuary intertidal site, with mudflats supporting abundant biofilms, consisting predominantly of diatoms (30). Surface sediments, from which colloidal carbohydrates and cEPS were to be extracted for subsequent sediment enrichments, were collected at low tide on 7 February 2005 in an area of high MPB biomass (chlorophyll a [Chl a], 110.64 µg g–1). Sediment samples for the slurry microcosms were taken on 14 February 2005 from similar biofilms (Chl a, 97 ± 20 µg g–1) at the same site. On each occasion, 120 cores (6.5 cm [inner diameter] by 10 cm) were transported back to the laboratory within an hour of sampling. Sediment slices (top 2 mm from each core) were then pooled. Sediments to be used for colloidal carbohydrate and cEPS extractions were immediately frozen at –80°C for 1 h and freeze-dried overnight.

Extraction of colloidal carbohydrate and cEPS fractions.
Colloidal carbohydrates were extracted from freeze-dried sediments. Aliquots (4 g) of sediment were mixed in 20 ml of saline (25{per thousand} [wt/vol] NaCl), incubated for 30 min at 25°C, and then centrifuged at 3,000 x g for 15 min at room temperature (68). Then 10 ml of the resulting supernatant (colloidal carbohydrate) from each centrifuge tube was pooled to give a total volume of 180 ml of colloidal extract. The remaining 10 ml of supernatant from each sample was dispensed into centrifuge tubes, each containing 30 ml of ethanol, and left at 4°C for 24 h in order to precipitate the cEPS fraction. Samples were then centrifuged at 3,000 x g for 15 min at room temperature, the supernatant was discarded, and the resultant cEPS pellets was pooled and resuspended in a final volume of 100 ml of distilled water. Concentrations of both colloidal carbohydrates and cEPS were determined by using the phenol-sulfuric acid assay (19) as described by Hanlon et al. (30) with carbohydrate concentrations quantified (µg g–1) as glucose equivalents. The final concentrations of colloidal carbohydrate and cEPS were 235 and 105 µg ml–1, respectively.

Experimental microcosms.
Two separate slurry microcosm experiments were established. In the first experiment (experiment 1), 20 g (wet weight) of fresh sediment (from the top 2 mm) was weighed into each of 15 autoclaved 250-ml conical flasks to establish microcosms representing two carbohydrate enrichments (colloidal or cEPS) and one control (no enrichment) (n = 5 for each). For the colloidal enrichment, 18 ml of colloidal carbohydrate extract was added to each flask, and the total volume adjusted to 100 ml by the addition of M9 minimal medium (42), giving a final colloidal carbohydrate concentration of 42.3 µg ml–1. Similarly, cEPS enrichments (100-ml final volume) were established after the addition of 10 ml of cEPS to give a final cEPS concentration of 10.5 µg ml–1. Control microcosms were established containing 20 g of sediment and made up to a final volume of 100 ml with M9 minimal media. All microcosms had a final salinity of 20{per thousand} and were incubated aerobically in the dark at 12°C at 130 rpm for 10 days. Subsamples were taken from each flask at day 0, 2, 4, and 10, to analyze the carbohydrate composition (2 ml) and bacterial community structure (1 ml) and to determine ß-glucosidase activity (1 ml).

A second microcosm experiment (experiment 2) was established in order to differentiate between abiotic- and biotic-carbohydrate turnover. This experiment was run over a 4-day period and consisted of control slurry microcosms with no enrichments (as described above) and microcosms poisoned with HgCl2 (to a final concentration of 2%) (n = 5 for each). Subsamples were taken from these microcosms on days 0, 1, 2, and 4 to analyze carbohydrate concentrations and to determine ß-glucosidase activities.

Analysis of water-soluble, hot-water- and hot-bicarbonate-extractable carbohydrates.
The 2-ml slurry subsamples taken for carbohydrate analysis were immediately frozen at –80°C for 1 h and freeze-dried overnight. Extraction of colloidal, cEPS, hot water-extracted (HW) and hot bicarbonate-extracted (HB) fractions was carried out as described previously (30), with concentrations subsequently determined as glucose equivalents by using the phenol sulfuric acid method (as described above). Concentrations of LMW carbohydrates were calculated by subtraction of the concentration of the cEPS fraction from that of the colloidal carbohydrate fraction.

Measurement of ß-glucosidase activity.
Maximum potential rates of extracellular ß-glucosidase activity were determined by using a fluorescently labeled methylumbelliferyl (MUF) substrate (32). Sodium azide (0.5 ml) was added to give a final concentration of 0.2% (to prevent de novo production of extracellular ß-glucosidases), and 8 ml of a 320-µg ml–1 (in 70% ethanol) solution of MUF-ß-D-glucopyranoside (Sigma-Aldrich, Gillingham, United Kingdom) was added to 1 ml of each fresh microcosm slurry subsample to give a final saturating substrate concentration of 1 mM. Five additional subsamples taken from control slurries were boiled to denature all enzymes prior to substrate addition to differentiate substrate release unrelated to ß-glucosidase activity. Samples were then incubated at 20°C for 4 h. Subsamples (2.5 ml) were taken at 0, 2, and 4 h and centrifuged at 3,000 x g for 15 min at room temperature. The resulting supernatants were transferred into cuvettes containing 0.2 ml of Tris-borate-EDTA buffer (pH 10), and fluorescence was measured by using a fluorimeter (Perkin-Elmer, Beaconsfield, Buckinghamshire, United Kingdom) at wavelengths of 365 nm (excitation) and 445 nm (emission). Enzyme activity rates were calculated based on MUF release from the linear part of the time-dependent activity curves.

Total nucleic acid extraction.
Total nucleic acids (DNA and RNA) were extracted from 1 ml of microcosm slurry samples (equivalent to 0.2 g [wet weight] of sediment) taken at days 0, 2, 4, and 10 using a Bio 101 lysing matrix and adapting the protocol used by Manefield et al. (39) via the addition of 500 µl of 0.1 M sodium phosphate (pH 8.0) and 500 µl of phenol-chloroform-isoamyl alcohol (25:24:1), followed by vortexing for 2 min and centrifugation at 16,100 x g for 1 min at 4°C, prior to an additional chloroform-isoamyl alcohol (24:1) extraction. Total nucleic acids in the resultant upper aqueous layer were then precipitated with absolute ethanol and resuspended in 100 µl of sterile diethyl pyrocarbonate (DEPC)-treated water, before being visualized after electrophoresis in a 0.7% (wt/vol) agarose Tris-acetate-EDTA gel.

PCR and reverse transcription-PCR (RT-PCR) amplification of 16S rRNA genes and 16SrRNA.
Total RNA was prepared by digestion of total nucleic acids using TURBO DNA-free (Ambion, Austin, TX). An aliquot (25 µl) of the total nucleic acid extract was diluted by adding an equal volume of DEPC-treated sterile water, and the digestion was carried out in accordance with the manufacturer's protocol. cDNA from 16S rRNA was generated by using the primer 518R (38) with Superscript III (Invitrogen, Paisley, United Kingdom). PCR amplification of 16S rRNA genes from DNA and from 16S rRNA (cDNA) was carried out as described in Hanlon et al. (30) with the primers FAM63F (5'-CAG GCC TAA CAC ATG GCA AGT C-3') and HEX518R (5'-CGT ATT ACC GCG GCT GCT CG-3') (26).

T-RFLP analysis.
Amplified 16S rRNA genes and RT-PCR-amplified 16S rRNA were purified by using a SureClean PCR purification kit (Bioline, London, United Kingdom) according to the manufacturer's protocol, as described for PCR product sizes greater than 100 bp. T-RFLP analysis was carried out as described by Hanlon et al. (30). Duplicate AluI and CfoI T-RFLP profiles from each DNA- or cDNA-derived PCR product were generated.

Clone library analysis.
RT-PCR-derived 16S rRNA amplicons were cloned by using a TOPO TA cloning kit (Invitrogen, Paisley, United Kingdom), using 3 µl of purified PCR product and according to the manufacturer's protocol. Cloned inserts were PCR amplified from liquid cultures of white transformants using the vector primers T3 (5'-ATT AAC CCT CAC TAA AGG GA-3') and T7 (5'-TAA TAC GAC TCA CTA TAG GG-3') from 25 clones per clone library using an initial denaturation at 95°C for 5 min, followed by 30 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 2 min, with a final extension step of 72°C for 10 min. PCR products were subsequently purified as described above, and 4 µl of each purified product was used as a template in sequencing reactions with vector primers using an initial denaturation at 95°C for 5 min, followed by 25 cycles of 96°C for 15 s, 55°C for 15 s, and 60°C for 4 min, and then desalted by ethanol precipitation, resuspended in 10 µl of deionized formamide, and denatured prior to sequencing using an ABI 3100 (Applied Biosystems). DNA sequences (ca. 400 to 500 bp) were edited by using Chromas v2.11 (Technelysium.com), and sequences were compared against the EMBL database using FASTA (44). Sequences were aligned by using CLUSTALX, (63), and distance matrices were calculated by using the DNADIST program in PHYLIP (22). A phylogenetic tree was created from the distance matrix by using the neighbor-joining method (49) and the Kimura substitution algorithm (34). A consensus tree was calculated after bootstrapping (1,000 replicate trees). The expected sizes of terminal restriction fragments (T-RFs) from cloned sequences were calculated in silico by using Webcutter 2.0 (http://rna.lundberg.gu.se/cutter2/). Clones corresponding to dominant T-RFs in community profiles, as identified by in silico analysis, were analyzed by T-RFLP as described above.

Real-time PCR quantification of 16S rRNA genes.
Real-time PCR was used to quantify 16S rRNA genes in sediments as a proxy for bacterial numbers using an ABI Prism 7000 detection system as described by Smith et al. (54) using the primers 1369F (5'-CGG TGA ATA CGT TCY CGG-3') and 1492R (5'-TAC GGY TAC CTT GTT ACG ACT T-3') and the TaqMan probe TM1389F (5'-CTT GTA CAC ACC GCC CGT A-3') (62).

Statistical analysis.
Changes in carbohydrate fractions and enzyme rates in each treatment over the time course were analyzed with one-way analysis of variance, followed by the post-hoc Tukey test (76). The data were checked for normality with Bartlett's test (4) and, where necessary, log transformed prior to analysis. Where overall changes in concentrations with time were not significant (by analysis of variance) but for which short-term increases or decreases were apparent between two specific times, these data were further analyzed by using the Student t test. The relationship between carbohydrate fractions and ß-glucosidase activity was further analyzed by using product-moment correlation.

Duplicate T-RFLP profiles from triplicate samples of each (RT-)PCR amplified 16S rRNA gene or 16S rRNA from each treatment and time point were grouped by restriction enzyme (AluI or CfoI) and by 5' and 3' T-RFs. Each group was then aligned on the basis of T-RF lengths, and the individual peak areas of the T-RFs identified by using T-Align (53) based on a 0.5-bp moving average, resulting in the generation of datasets of aligned T-RFs that gave individual relative peak areas as a proportion (%) of the overall profile. All T-RFs that contributed less than 1% of the total peak area for a profile were excluded from further analysis. Using Primer v6 (Primer-E, Plymouth, United Kingdom) (12), the aligned T-RFs were transformed by log(X+1) to remove any weighting from dominant peaks and analyzed by using a Bray-Curtis similarity matrix (7). Using the resultant resemblance matrix, the data were analyzed in two formats: a group average hierarchical cluster dendrogram and a two-dimensional multidimensional scaling (MDS) plot. The clusters from the dendrogram were overlaid onto the MDS plot based on a similarity cutoff point determined from SIMPER analysis (Primer v6) (11). The two-way crossed SIMPER analysis was carried out on all of the transformed aligned T-RF data to determine the percent contribution of each individual T-RF to the similarity matrix and the similarity or dissimilarity of subsequent clustering based on time and treatment.

The resultant resemblance matrices from the Bray-Curtis analysis of the total and active bacterial communities (from molecular data) were compared to the corresponding datasets describing carbohydrate composition and enzyme activities (biochemical data) by using Primer v6. These latter datasets were analyzed by using a Euclidean distance matrix (23) to produce resemblance matrices. The two resemblance matrices—one for the molecular data and the other for the corresponding biochemical data—were compared by using BIO-ENV in Primer-E (11). The BIO-ENV routine calculates the similarity between the two matrices by rank correlation, using a Spearman rank correlation coefficient ({rho}), and was used to identify the individual or combinatorial contribution of biochemical variables that best grouped the composition of both the total (DNA) and active (RNA) bacterial communities according to time and treatment. The biochemical data resemblance matrix was further analyzed by using principal components analysis for the days corresponding to the total bacterial samples and the active bacterial samples with the resultant vectors of the important biotic variables for each set, as determined by BIO-ENV, overlaid.

Nucleotide sequence accession numbers.
Nucleotide sequences were deposited in the EMBL database with the accession numbers AM411540 to AM411572.


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RESULTS
 
Changes in carbohydrate concentrations and enzyme activity.
In the 10-day sediment slurry microcosm experiment (experiment 1), the initial addition of colloidal carbohydrate resulted in an ~25% increase in cEPS and an ~300% increase in the LMW carbohydrate concentration compared to the unamended-control microcosm (Table 1). The initial addition of cEPS resulted in an ~105% increase in the cEPS concentration but also an ~600% increase in the LMW carbohydrate concentration (Table 1 and Fig. 1A to C).


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  		TABLE 1. Initial carbohydrate concentrations in sediment slurry microcosmsa


Figure 1
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  		FIG. 1. Changes in mean concentrations (± the standard error) of LMW carbohydrate and cEPS (colloidal EPS)-carbohydrate extracts in sediment slurry microcosm experiments. (A) Control slurries, (B) colloidal enrichments, and (C) cEPS enrichments over a 10-day time course; (D and E) control (D) and poisoned (E) slurries over a 4-day time course; (F) ß-glucosidase activity in control and poisoned slurries.

There were significant temporal changes in cEPS concentrations over the following 10 days (experiment 1; Fig. 1A to C) in the controls (F5,24 = 2.710, P < 0.05), colloidal enrichments (F5,24 = 10.438, P < 0.001), and cEPS enrichments (F5,24 = 40.056, P < 0.001). Changes in cEPS concentrations showed two distinct phases over the course of the experiment. In the first phase (0 to 2 days), cEPS concentrations decreased to 80 to 100 µg of glucose equivalents g (dry weight) of sediment–1 in all treatments, despite the different starting concentrations. The rate of loss was greatest in the cEPS-enriched slurries (–140 ± 25 µg g–1 day–1) (Fig. 1C). During the second phase (days 4 to 10), the cEPS concentrations gradually increased in all treatments, with final values (day 10) being significantly higher than the initial concentrations in the control slurries (70% increase, P < 0.05) and in the colloidal enrichments (40% increase, P < 0.05) and substantially higher in cEPS enrichments (300% increase, P < 0.001) (Fig. 1A to C).

In the control treatments, the concentration of LMW carbohydrate showed an initial significant increase to maximum values of 790 µg g–1 (P < 0.05) on day 2, followed by a gradual decline to 250 µg g–1 by day 10 (Fig. 1A, overall changes of F5,24 = 2.667, P < 0.05). The colloidal and cEPS enrichments both showed an initial, significant decline in concentrations of LMW carbohydrate over the first day (colloidal enrichment, t = 2.818, P < 0.05; cEPS enrichment, t = 2.895, P < 0.05; Fig. 1B and C). No further significant patterns of change were seen in the colloid-enriched microcosms over the remainder of the experiment. However, the concentrations of LMW carbohydrate subsequently increased significantly in the cEPS enrichments, attaining maximum values on day 7 (1,800 µg g–1, P < 0.05; Fig. 1C, overall changes in EPS treatment, F5,24 = 3.216, P < 0.05).

Microcosm slurries poisoned with mercuric chloride (experiment 2) were used to investigate whether changes in slurry carbohydrate concentrations were due to abiotic factors or biological activity. In the control microcosms receiving no additional carbohydrate amendment (Fig. 1D), the concentrations of cEPS and LMW carbohydrate followed patterns over the 4-day time course similar to those observed in the controls of experiment 1 (Fig. 1A and D), with the concentration of colloidal carbohydrates significantly higher on days 2 and 4 compared to day 0 (P < 0.05) and with decreases in cEPS (t = 3.458, P < 0.05). In contrast, the sediment slurries poisoned with mercuric chloride (2%) showed no changes in cEPS and colloidal carbohydrate concentrations (Fig. 1E). The mean ß-glucosidase activity increased significantly in unpoisoned controls (F3,16 = 61.390, P < 0.001), reaching maximum values on day 1 (Fig. 1F) and declining thereafter. In contrast, in poisoned and boiled slurry microcosms, ß-glucosidase activities were reduced to values of between <0.1 and 0 nmol g (wet weight)–1 h–1.

During experiment 1, both ß-glucosidase activity and the concentrations of HB carbohydrate fractions increased significantly with time in colloidal (ß-glucosidase: F5,24 = 30.642, P < 0.001; HB concentration: F5,24 = 5.045 colloidal; P < 0.01) and cEPS (ß-glucosidase: F5,24 = 124.098, P < 0.001; HB concentration: F5,24 = 3.554 cEPS; P < 0.05) enrichments (Fig. 2A and B). Although no relationship existed between ß-glucosidase activity and concentrations of HB carbohydrates in control treatments (data not shown), the MUF substrate release rate was positively correlated with concentrations of HB carbohydrates in both of the carbohydrate-enriched microcosms (Fig. 2A and B). The extracellular enzyme activity was significantly positively correlated (P < 0.001) to the daily rate of increase in cEPS in all three treatments, but particularly in the colloidal and cEPS enrichments (Fig. 2C). During the first phase of the experiment (between days 0 and 2), when cEPS concentrations were decreasing, the MUF substrate release rate was low (<5 to 6 nmol g [wet weight]–1 h–1). Higher ß-glucosidase activity coincided with periods of increasing cEPS concentrations in the slurries toward the end of the experiment (days 4 and 10), especially in cEPS enrichments.


Figure 2
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  		FIG. 2. Relationship between concentration of HB carbohydrates and the MUF substrate release rate due to ß-glucosidase activity. (A to C) Colloidal enrichments (A), cEPS enrichments (B), and MUF release rate (n = 20) and cEPS gain/loss in all treatments (n = 20 per each treatment) (C). Microcosms were sampled at days 0, 2, 4, and 10 (day 0 [D0], D2, D4, and D10, respectively). The results of product-moment correlation analyses are shown (*, P < 0.05; ***, P < 0.001).

Changes in the composition of the bacterial communities within sediment microcosms.
Changes in the structure of the total bacterial community in the sediment slurry microcosms were assessed by T-RFLP analysis, which revealed clear temporal- and treatment-related shifts in community composition (Fig. 3). The total bacterial communities (16S rRNA genes) at day 0 from the three microcosms—no enrichment sediment control and the colloid- and cEPS-enriched sediments—showed 77.9% similarity (using SIMPER) to each other (Fig. 4A, cluster I). By day 2, the total bacterial communities from the sediment slurries were still more than 77% similar to each other (Fig. 4A, cluster II) but 41% similar to those at day 0 (cluster I). However, there was additional separation within cluster II of the total bacterial community from the cEPS-enriched sediment. By day 4, the total bacterial community from the cEPS-enriched sediment formed a separate cluster (Fig. 4A, cluster III) distinct from the communities in the colloid-enriched sediment and the no-enrichment control (cluster II). By day 10, an additional shift in the composition of the total bacterial community was observed in the cEPS-enriched sediment (Fig. 4A, cluster IV), and the community remained distinct from those in the colloid-enriched sediment and the no-enrichment control.


Figure 3
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  		FIG. 3. T-RFLP fingerprints of the total bacterial communities in sediment slurry microcosms. AluI digestion of PCR amplified 16S rRNA genes (5' T-RFs, black lines, 3' T-RFs, gray lines). Microcosms: S, no-enrichment sediment control; C, colloidal enrichment; E, cEPS enrichment sediments sampled at days 0 (D0) and 4. T-RFs that are important contributors toward the separation of the three communities at day 4 are indicated (see also Table 2).


Figure 4
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  		FIG. 4. MDS plots showing changes in the composition of the total and active bacterial communities in sediment slurry microcosms as determined from a Bray-Curtis similarity matrix with data from AluI and CfoI T-RFLP profiles. (A and B) Total community (PCR-amplified 16S rRNA genes) (A) and active community (RT-PCR-amplified 16S rRNA) (B). Microcosms as indicated, sampled at days 0, 2, 4, and 10; n = 3 for each sediment at each time point. Clusters are indicated with Roman numerals.

SIMPER analysis revealed that the T-RF that contributed most to the separation of the total bacterial community in the cEPS-enriched sediments was a 5' T-RF of 197.35 bp formed by AluI digestion (see also Fig. 3). Sequence analysis of nine 16S rRNA clones that also generated AluI 5' T-RFs of 197 bp after T-RFLP analysis revealed that these clones were most closely related to the marine {gamma}-proteobacterium Acinetobacter junii (AB101444, 95 to 99% identity).

Variation in the composition of the active bacterial communities (16S rRNA) in the sediment microcosms was investigated at days 0 and 4; the point at which the total bacterial communities diverged (Fig. 3). A similar pattern of divergence was observed in the active community, as revealed by T-RFLP analysis, as had been found in the total community (Fig. 4), wherein by day 4 the active community in the cEPS-enriched sediment formed a discrete cluster (Fig. 4B, cluster III) distinct from those in the colloid-enriched and the unamended sediments (Fig. 4B, cluster II). The active communities in all three slurries were 68.9% similar to each other at day 0 (Fig. 4B, cluster I, SIMPER analysis), but by day 4 these communities were only 33.1% similar overall to the original communities present at day 0 (Fig. 4B, clusters II and III). Two T-RFs contributed most to this shift in community structure: a 5' AluI T-RF of 32.83 bp for which the taxonomic lineage is unknown and a CfoI 3' T-RF of 156 bp that corresponded to a 16S rRNA clone most closely related to the marine {gamma}-proteobacterium Shewanella affinis (AF50080, 98.7% identity). At day 4, the active bacterial community in the cEPS-enriched sediment (Fig. 4B, cluster III) was 49% similar to that of the colloid-enriched sediment and 50% similar to the unamended sediment. A CfoI 5' 170.08-bp T-RF contributed most to the separation of the cEPS-enriched sediment community that corresponded to three cloned 16S rRNA sequences most closely related to the marine {gamma}-proteobacterium Acinetobacter johnsonii (AB099655, 98.7% identity).

The T-RFs that made the greatest contribution to the separate clustering of active bacterial communities from the three sediment slurry microcosms, as assessed by SIMPER analysis, are shown in Table 2. Comparison of the T-RFs generated from cloned 16S rRNA sequences suggested that these T-RFs corresponded to the presence of marine {gamma}-proteobacteria, namely, Acinetobacter sp. and Shewanella baltica, in all three treatments. Other T-RFs that contributed to the clustering seen in the MDS plot (Fig. 4B and Table 2), as determined by SIMPER analysis, were not represented in the clone libraries, and thus their taxonomic lineage is unknown.


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  		TABLE 2. 16S rRNA T-RFs important in cluster separation of the active bacterial communities as determined by SIMPER analysis

The 16S rRNA clone library data from both the colloid- and cEPS-enriched sediments at day 4 were dominated by sequences most closely related to Acinetobacter spp., with 13 of 20 clones in the colloid-enriched sediment clone libraries and 17 of 19 clones in the cEPS-enriched sediment clone libraries sharing >95% identity to Acinetobacter 16S rRNA sequences (Fig. 5 and data not shown). Other 16S rRNA clones were most closely related to other {gamma}-proteobacteria: Shewanella sp. (>97% identity; three clones from colloid-enriched sediment and one clone from cEPS-enriched sediment), Pseudomonas aeruginosa (>95% identity, two clones from colloid-enriched sediment), Pseudoalteromonas sp. (95% identity for one clone from colloid-enriched sediment), and Oceanisphaera sp. (>97% identity, one clone each from the colloid- and cEPS-enriched sediments).


Figure 5
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  		FIG. 5. Neighbor-joining tree of cloned RT-PCR-amplified 16S rRNA sequences from sediment microcosms at day 4. Clone designations: C, colloidal-; E, cEPS-enriched sediments. Scale bar indicates the genetic distance between sequences. Percent bootstrap values (1,000 replicates) are indicated.

Variation in 16S rRNA gene numbers in the sediment microcosms as assessed by Q-PCR analysis.
Quantitative PCR (Q-PCR) assays were performed to target 16S rRNA genes to investigate variation in the abundance of total bacteria within the sediment microcosms. Q-PCR amplification of 16S rRNA genes from the total bacterial community showed a log increase in the unamended control sediment from day 0 to day 2, followed by a subsequent decline to 6.44 x 109 16S rRNA genes g of sediment–1 by day 4, followed by an increase to 4.57 x 1010 by day 10 (Fig. 6). The number of 16S rRNA genes in the colloid-enriched sediment showed only a gradual increase (1.93 x 109 to 1.2 x 1010 16S rRNA genes g of sediment–1) over the 10-day experiment. In the cEPS-enriched sediment, there was a log increase in 16S rRNA gene numbers from day 0 to day 2, followed by a further log increase to 1.8 x 1011 genes g of sediment–1 by day 4, by which time 16S rRNA gene numbers were significantly higher than those found at day 0 (P < 0.05) and were also significantly higher than in the corresponding (day 4) no-enrichment sediment control microcosm (P < 0.02). 16S rRNA gene numbers in the cEPS-enriched microcosm then decreased to 5.31 x 1010 genes g of sediment–1 by day 10 (Fig. 6).


Figure 6
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  		FIG. 6. Variation in 16S rRNA gene numbers g of sediment–1 in slurry microcosms as assessed by Q-PCR. Microcosms: S, no-enrichment sediment control; C, colloidal enrichment; E, cEPS enrichment sampled at days 0, 2, 4, and 10. Gene numbers were derived from a standard curve with r2 = 0.995, a y intercept of 39.63%, PCR efficiency (E) of 106%, and a no-template control CT value of 33.73 ± 1.26.

Multivariate analysis of molecular and biochemical datasets.
Relationships between the composition of the total bacterial communities (T-RFLP datasets) and biochemical datasets (ß-glucosidase activity and the concentrations of Chl a, colloid, cEPS, and HW and HB carbohydrate fractions) were investigated at days 0, 2, 4, and 10 by using BIO-ENV (Fig. 7A). The three variables shown by BIO-ENV to make the greatest contribution to the composition of the total bacterial communities were ß-glucosidase activity, and the concentration of the cEPS and HW carbohydrate fractions ({rho} = 0.466; Fig. 7A). In the cEPS-enriched sediments, in which the total bacterial communities at days 4 and 10 had diverged from those in the colloid-enriched and unamended sediments (Fig. 4A), the concentrations of cEPS and HW carbohydrates in the sediments were identified as most important in determining this separation (Fig. 7A).


Figure 7
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  		FIG. 7. Identification of biochemical variables that were most significant in determining the clustering of total (A) and active (B) bacterial community compositions in sediment microcosms, as determined by BIO-ENV analysis. Principle components analysis show clustering (indicated by Roman numerals) based on biochemical variables in microcosms at days 0, 2, 4, and 10. Vectors indicate biochemical variables important in clustering. Euclidean distances are shown.

Similarly, relationships between the composition of the active bacterial communities and the biochemical variables were also investigated at day 0 and day 4 (Fig. 7B). The ß-glucosidase activity and the concentrations of colloid, cEPS, and HW carbohydrates made the greatest contribution to the composition of the active communities ({rho} = 0.652; Fig. 7B). From the overlying vectors, the biochemical variables that contributed most to the separation of the active bacterial communities in the cEPS-enriched microcosms, from those in the colloid-enriched and the unamended sediments, was the concentration of the HW carbohydrate fractions, whereas the ß-glucosidase activity was most important in the temporal separation (day 0 versus day 4) between the active communities in the unamended control and colloid-enriched sediments (Fig. 7B).


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DISCUSSION
 
MPB biofilms are ubiquitous in almost all freshwater and marine intertidal and subtidal habitats exposed to light (8, 69). Biofilms provide potential sites for close coupling between the activity of primary producers and heterotrophic organisms, interactions that may play an important role in maintaining the resilience of biofilms to environmental variability. Although there is some evidence that a greater diversity of autotrophs helps to maintain the rates of primary production (24, 65, 73), the importance of taxonomic diversity of the heterotrophic community in determining biofilm properties is not clearly understood (59). We hypothesized that inputs of different organic carbon fractions would select for particular bacterial taxa, resulting in the identification of bacteria that preferentially utilize cEPS and/or colloidal carbohydrates. The data presented here indicate that particular constituents of the bacterial assemblage increased in their relative abundance within the community in response to the inputs of different, diatom-derived, organic carbon fractions, whereas other taxa showed a decline in their relative abundance. Coupled with data of changing sediment carbohydrate fractions and enzymatic activities, these data support the initial premise that there is specific algal-bacterial coupling within estuarine biofilms with respect to EPS cycling.

In the unamended sediment control slurries, the colloidal carbohydrate pool consisted of ~50% cEPS and ~50% LMW carbohydrate. Heterotrophic activity in these slurries over the first 2 days of the experiments resulted in declines in cEPS concentrations, increases in the standing stock of LMW carbohydrate (Fig. 1A), and increases in ß-glucosidase activity (Fig. 2). Although there was a shift in bacterial community composition in the unamended sediment in the first 2 days (Fig. 4), no further major shifts occurred between days 2 and 10, and the LMW carbohydrate initially produced (from the degradation of cEPS, but also from other refractory carbohydrate pools in the sediment) was utilized by the end of the experiment. LMW carbohydrate forms a highly bioreactive fraction, with concentrations fluctuating rapidly in situ (55, 64), and the lack of major shifts in the total bacterial community correlated with LMW carbohydrate dynamics indicates that LMW organic carbon can be utilized by a diverse range of bacteria (27, 50, 60).

The addition of the two different carbohydrate (cEPS and colloidal) fractions resulted in significant increases in both cEPS and LMW carbohydrate concentrations (P < 0.05) in both treatments compared to the controls (Fig. 1). The enrichment levels that were used resulted in at least a doubling of the concentrations of potential substrates in the sediments (up to ~1,000 µg of colloidal carbohydrate g [dry weight] of sediment–1 and ~400 µg of cEPS g [dry weight] of sediment–1) compared to those in the control microcosms. These inputs nevertheless remained well within the range of carbohydrate concentrations encountered in estuarine biofilms in the Colne estuary (up to >4,000 µg of colloid g [dry weight] of sediment–1 and ~1,000 µg of cEPS g [dry weight] of sediment–1) (5), and in other mud-rich estuarine environments (67). During the first 2 days of the experiment there was a rapid reduction in concentrations of cEPS and LMW carbohydrate concentrations in all treatments, and an increase in bacterial numbers (16S rRNA gene numbers; Fig. 6) coupled with a change in the composition of the total bacterial community (Fig. 4), although treatment-specific effects were not observed. The decline in the concentrations of cEPS and LMW carbohydrates suggests rapid (within hours) utilization of the substrate amendments by the bacterial community, in particular, of the LMW components (see above); with timescales similar to those determined both in situ and experimentally using 13C and 14C labeling experiments (28, 41, 55, 70). The addition of colloidal carbohydrate did not result in further significant differences from the unamended sediment control microcosm, in particular, with respect to bacterial numbers (16S rRNA gene numbers; Fig. 6) or community structure (T-RFLP profiles; Fig. 3 and 4). However, in contrast, by day 4 of both experiments in the cEPS-enriched sediments, there were major shifts in the composition of both the total and the active bacterial communities (Fig. 4), accompanied by higher bacterial numbers (16S rRNA gene numbers) (Fig. 6) and increased intracellular enzyme activity (Fig. 2) for the remainder of the 10-day experiment. Associated with the initial cEPS input was a large pool of LMW carbohydrate (Table 1). These LMW molecules would have been trapped within the "gel" structure of the EPS in the internal pore spaces generated by complex polymers (17). This LMW carbohydrate was very rapidly utilized in the cEPS addition slurries (by day 2). Subsequently, in the cEPS-amended sediments there was an increase in the enzymatic hydrolytic activity, with increased liberation of HB-soluble EPS and accumulation of cEPS in the slurries over time. The precipitation step used to isolate cEPS has an approximate size cutoff of ~10 kDa (67). There is generally a positive relationship between the structural complexity of EPS and the difficulty of extraction (1, 17), with cEPS (and other more tightly bound sediment carbohydrate fractions, e.g., HB extracted) having increasingly complex structures and being more recalcitrant to microbial attack (1, 10, 17). Polysaccharides and large oligosaccharides require breakdown by extracellular enzymes prior to transport across membranes into bacterial cells (46, 71). Such dissolved organic matter can have complex tertiary structures that will affect substrate recognition and binding by hydrolytic enzymes (33), with enzymatic hydrolysis of high-molecular-weight material the rate-limiting step in the process of organic matter oxidation in sediments (40, 57).

Microbial ß-glucosidase production can be induced by increasing concentrations of structural polysaccharides, while ß-glucosidase production is reduced when other suitable substrates are available (6). This is a response similar to that shown in Fig. 2. Previous studies have similarly identified increased rates of ß-glucosidase activity in intertidal sediments enriched with organic matter (28, 36, 37). Extracellular enzymatic degradation of complex carbohydrates within the sediment would result in increased lability, allowing more EPS material to be extracted in less-astringent solvents. This may explain the observed gain in cEPS when concentrations of HB carbohydrates and enzyme activities were high (Fig. 2). cEPS would then be further hydrolyzed to generate LMW carbohydrate. Thus, the increases in concentrations of LMW carbohydrates in the cEPS-enriched sediments, when enzyme activities were high (toward the end of the experiment) would be the combined result of breakdown of the HB fraction by ß-glucosidase and hydrolysis of cEPS by other extracellular enzymes (30).

In both the colloid- and cEPS-enriched microcosms the dominant bacterial taxa identified were members of the subphylum {gamma}-proteobacteria (Fig. 5). The addition of cEPS led to the largest shifts in community composition (Fig. 4) and was coupled to increased bacterial numbers (Fig. 6) and enzyme activity (Fig. 2). Previous studies of the effects of substrate enrichment on pelagic marine bacteria (20, 21), organic matter degradation by bacterioplankton (45), and association with planktonic diatom blooms (48) and in estuarine waters (15) have found community shifts leading to increased numbers of the members of the subphyla Flavobacteria and Sphingobacteria and of {alpha}- and {gamma}-proteobacteria. Members of these taxa appear to be the dominant bacteria involved in algal organic matter degradation. Members of the {gamma}-proteobacteria have been identified previously in both estuarine and marine sediments (15) and, more specifically, have been linked to algal and diatom exudates (3, 29, 47, 51). {gamma}-Proteobacteria were also found to prevail in the surface sediment of a tidal flat in the German Wadden Sea (58), while in enrichment experiments, using riverine organic matter from estuarine waters, Kisand et al. (35) identified members of the {gamma}-proteobacteria and of the phylum Bacteroidetes to be the dominant taxa. In the present study, MPB-derived carbohydrate enrichment was associated with increases in the relative abundance of members of the {gamma}-proteobacteria, but especially Acinetobacter (Table 2 and Fig. 5) and in particular in the cEPS-enriched sediments. Acinetobacter spp. have previously been shown to be important in activated sludge sewage treatment (9, 56, 72), with some species able to degrade oil, naphthalene, phenol, and glucose (18, 43). To our knowledge, the present study is the first to identify Acinetobacter associated with either estuarine sediments or diatom exudates. These experiments have demonstrated a forced shift in the community driven by substrate enrichment (although still within the concentration ranges experienced by natural microbial assemblages in situ), whereby the {gamma}-proteobacteria outcompete other taxa, thus altering the composition of the sediment bacterial community structure. In conclusion, we have revealed here taxon-specific bacterial responses to the production of MPB (diatom)-derived EPS within estuarine sediments and highlight a hitherto-unknown role for the members of the genus Acinetobacter in carbohydrate degradation in estuarine sediments.


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ACKNOWLEDGMENTS
 
This study was funded by a grant to G.J.C.U., A.S.B., and A.M.O. from the United Kingdom Natural Environment Research Council (NER/A/S/2001/00536). K.H. was funded by a Natural Environment Research Council Ph.D. tied studentship (NER/S/J/2002/11016A).


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Animal and Plant Sciences, The University of Sheffield, Western Bank, Sheffield S10 2TN, United Kingdom. Phone: 44 (0114) 22 24626. Fax: 44 (0114) 22 20002. E-mail: A.M.Osborn{at}sheffield.ac.uk Back

{triangledown} Published ahead of print on 3 August 2007. Back

{dagger} Present address: School of Biological Sciences, Flinders University of South Australia, Adelaide, GPO Box 2100, Adelaide, SA 5001, Australia. Back


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Applied and Environmental Microbiology, October 2007, p. 6112-6124, Vol. 73, No. 19
0099-2240/07/$08.00+0     doi:10.1128/AEM.00551-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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