Shifts in composition of microbial communities ... - Wiley Online Library

RESEARCH ARTICLE

Shifts in composition of microbial communities of subtidal sandy sediments maximise retention of nutrients Hugh Forehead1,2, Peter Thomson1 & Gary A. Kendrick2 1

CSIRO Division of Marine and Atmospheric Research, Hobart, Tas., Australia; and 2The UWA Oceans Institute and School of Plant Biology, University of Western Australia, Crawley, WA, Australia

Correspondence: Hugh Forehead, Institute for Marine and Antarctic Studies, University of Tasmania, Hobart, Tasmania 7001, Australia; Tel.: +61 3 6226 6379; fax: +61 3 6226 6391; e-mail: [email protected] Received 4 July 2012; revised 8 August 2012; accepted 9 August 2012. Final version published online 4 September 2012. DOI: 10.1111/j.1574-6941.2012.01472.x

MICROBIOLOGY ECOLOGY

Editor: Riks Laanbroek Keywords trophic index; autotrophy; biogeochemistry; microphytobenthos; sediment carbon.

Abstract The density and composition of microbial communities of subtidal sandy sediments determines their role in the cycling of nutrients in coastal waters. It has previously been found that sediments disturbed by waves and currents have reduced biomass, greater productivity to respiration (P/R) ratios and a tendency to take up nutrients. Conversely, with shelter and greater biomass, P/R ratios were smaller and nutrients released. This study, in warm temperate waters, examined the consequences of high and low levels of hydrodynamic energy on the microbial community structure and biogeochemistry at two locations at different times of year. Measurements included biomarkers, sediment properties and exchanges of gases and nutrients. Microbial communities were dominated by diatoms and bacteria. Exposed sites, relative to paired sheltered sites, had smaller ratios of bacteria to benthic microalgae (BMA), larger C/N ratios, smaller indices of diagenetic activity, but smaller P/R ratios. The bacteria in exposed sediments exhibited biomass-normalised rates of respiration almost double those in sheltered sediments. This increased activity was most likely fuelled by elevated concentrations of photosynthates, secreted by BMA attached to sand grains. Changes in community composition owing to different levels of disturbance led to shifts in functioning that resulted in consistently small exchanges of nutrients.

Introduction The microbial community in the surface millimetres of shallow subtidal sandy sediments is generally dominated by benthic microalgae (BMA) and bacteria. This community of microorganisms performs a number of roles in modulating the exchanges of nutrients across the sediment–water interface (MacIntyre et al., 1996). The BMA assimilate nutrients from pore waters and the water column (Rizzo, 1990; Sundba¨ck et al., 1991; Suzumura et al., 2002), and diazotrophs can bring otherwise inert N2 into the ecosystem (Carpenter et al., 1991). BMA secrete carbohydrates for motility or as a strategy for releasing excess fixed carbon in conditions of strong irradiance (Staats et al., 2000). These exudates are a highly labile substrate that is rapidly utilised by bacteria (Middelburg et al., 2000; Cook et al., 2004). Thus, the photosynthetic activity of BMA can modulate the activity of sediment bacteria (Murray et al., 1986) and the resulting FEMS Microbiol Ecol 83 (2013) 279–298

rate of breakdown of organic matter (Haack & McFeters, 1982; Alongi, 1994). Changes in the composition of the microbial community can have significant effects on the rates and pathways of nutrient cycling in sediments. The disturbance of subtidal sediments by waves and currents can affect the concentration, composition, chemistry and metabolism of the associated microbial communities. The bacterial fraction may be preferentially depleted; in an experiment in a flume, bacteria were more easily advected into the water column than BMA (Shimeta et al., 2002). These disturbances also change the composition of the BMA community, increasing the dominance by species attached to the substrate (Colijn & Dijkema, 1981; Busse & Snoeijs, 2003) and reducing biomass by advecting cells into the water column (Gabrielson & Lukatelich, 1985; Schofield et al., 2004), and may cause mechanical damage to cells (de Jonge & van den Bergs, 1987). Decrease in biomass of BMA has been found to result in a reduced capacity for the uptake and ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

280

retention of dissolved inorganic nitrogen (DIN) in sediments at temperatures < 21 °C (Sundba¨ck & Graneli, 1988; Sloth et al., 1996). The mixing of sediments into the water column by resuspension causes a release of nutrients from porewaters and the surfaces of sediment grains (Oviatt et al., 1981; Fanning et al., 1982). This depletes reserves in the sediments and can fuel the growth of phytoplankton (Davies, 1975; Fanning et al., 1982), bacteria (Wainright, 1987) and microheterotrophs (Lawrence et al., 2004) in the water column. At least in the short term, physical disturbance increases the depth of oxygen penetration into the sediments, allowing for more rapid mineralisation of organic matter (Hansen & Blackburn, 1991). These conditions of reduced biomass of BMA, reduced concentrations of nutrients in porewaters and increased oxygenation are quite different to those in sediments that are not resuspended or disturbed by waves and currents. Shelter from ambient wave energy has been shown to result in an increase in the concentration of BMA and bacteria in the sediments of coasts and estuaries (PlanteCuny & Bodoy, 1987; Alongi et al., 1996; Galois et al., 2000; Incera et al., 2003b; Nozaki et al., 2003). Shelter has also been found to result in increased concentrations of nutrients in coastal waters (Cerco & Seitzinger, 1997). For example, an experimental mesh enclosure of the water over sandy sediments resulted in an increase in concentrations of DIN in the overlying water column by two orders of magnitude (Forehead et al., 2011). These conditions can lead to greater phytoplankton biomass and reduction in the supply of light to BMA (Sundba¨ck et al., 2004). After extended periods of stable conditions in oligotrophic waters, inorganic nutrient limitation may lead to dominance by cyanophytes (Barranguet et al., 1997) or dinoflagellates (GA Kendrick, unpublished data), particularly when the waters are warm (Watermann et al., 1999). The composition of microbial communities in sediments has been found to influence nutrient exchanges in different ways. Western Australian sediments that were experimentally sheltered in mesocosms exhibited taxonomic changes to their microbial communities; the sheltered sediments exhibited a greater uptake of nutrients from the water column than the sediments than sediments that had not been sheltered (Kendrick et al., 1998). It has also been shown that the degree of dominance of the sediment community by autotrophs can be proportional to the tendency for the sediments to take up nutrients (Manini et al., 2003). The oxygen-based productivity to respiration ratio has been used as a means of predicting the nutrient cycling characteristics of the sediment microbial community. In these studies, autotrophically dominated (oxygen producing) communities take up nutrients and heterotrophic (oxygen consuming) ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

communities release them (Rizzo et al., 1996; Viaroli & Christian, 2004). Increases in temperature can raise the rates of photosynthesis, respiration (Masini & McComb, 2001; Hancke & Glud, 2004) and increase the loss of nutrients from the sediments to the water column because of increased rates of diagenesis of organic matter (Klump & Martens, 1989). This study combined a broad range of measurements to examine previously unstudied relationships between disturbance by waves and currents, sediment microbial community composition, community metabolism and the resulting nutrient exchanges. The study was located in warm temperate waters; the results are likely to be applicable to other warm oligotrophic coastal areas with sandy sediments. The three hypotheses tested in this study were that: (1) increased disturbance of the sediment by waves and currents would reduce the accumulation of organic matter in these sediments, (2) exposed sites would have a decreased ratio of bacteria and grazers to BMA relative to sheltered sites, (3) sediments with smaller ratios of bacteria to BMA would have reduced rates of release of nutrients into the water column relative to sediments with larger ratios. Paired sheltered and exposed sites were sampled at four locations. At two locations sampling extended across three seasons. The conditions during this study ranged from mild (16 °C) temperatures and frequent disturbance by large waves in winter (August), to warmer temperatures (22 °C) and decreased disturbance by waves in early summer (December) with these conditions persisting and warming further through to early autumn (March).

Materials and methods Study sites

The coastal waters of southern Western Australia are oligotrophic (Johannes et al., 1994); the sediments are composed of mainly carbonate sand. The area is microtidal; the tidal range is from 0.1 to 0.9 m at nearby Fremantle (Simpson et al., 1996); the water temperature is warm temperate (16–24 °C). The area experiences strong wave energy conditions in winter and calmer conditions in summer (Pattiaratchi et al., 1991). The waves consist of oceanic swell, mostly from the south-west, and as the swell approaches the coast, it is refracted to west–southwest and strengthened by wind-generated sea waves (Simpson et al., 1996). Swell heights at the shores of embayments north and south of Cockburn Sound, Owen Anchorage and Warnbro Sound, is reduced by 90% of the size in the adjacent ocean and by 95% at the southern end of Cockburn Sound. The two locations used in this study were in Cockburn Sound, on the south-west coast of Western Australia, FEMS Microbiol Ecol 83 (2013) 279–298

281

Sandy sediment microbial communities and nutrient exchanges

near Perth. Cockburn Sound is a sheltered embayment, 16 km long (north–south) and 9 km wide. The sediments are mainly carbonate sand, with areal seagrass coverage ranging from 0.6% in the east of the Sound, to 50.1% in the south (Kendrick et al., 2002). Samples were collected from a depth of 7 m, below the influence of wind-generated waves. Jervoise Bay is in the north of the sound (32° 13.971′S, 115°76.218′E); it contains a 1.5- by 0.5-km man-made embayment enclosed by rock walls. Mangles Bay is in the south (32°25.66′S, 115°69.686′E) and is bisected by a causeway that shelters the waters on its eastern side from ocean swells and currents. Abbreviations used in the following text are JB for Jervoise Bay and MB for Mangles Bay. There were paired sites at each of the two locations: inside (sheltered) and outside (exposed) the seawalls. Both locations were sampled on two occasions, in August (winter) and December (summer); JB was also sampled in March (early autumn). Site measurements

Photosynthetically active radiation (PAR) was measured with a Li-Cor LI-192SA sensor attached to a LI-250A light meter. Salinity, dissolved oxygen (DO) and temperature were measured with either a Seabird SB19+ (Sea-Bird Electronics Inc, Bellevue) or a Yeo-Cal model 611 Intelligent Water Quality Analyser (Yeo-Kal electronics Pty. Ltd., Brookvale, NSW, Australia). Collection of samples

Samples were collected by SCUBA divers. At each site multiple samples were collected from within an area of about 16 m2. The sampling areas at each site appeared to be representative of each site in terms of topography and sediment type. Areas were selected haphazardly and marked with a Global Positioning System instrument (Garmin GPS III). Porewater for nutrient analysis was collected in situ from two depths in the sediment: 0–5 cm and 5–10 cm, using syringes attached to sippers (Kendrick et al., 1998). Water column nutrient samples were collected from about 0.5 m above the sediment surface. All water samples for nutrient analyses were filtered through precombusted GF/F filters (nominal pore size 0.7 lm) into 10-mL polypropylene tubes, cooled on ice and frozen within 3 h. Nutrient analysis was carried out within 8 months of collection. For flux measurements, sediment cores were carefully collected in clear polycarbonate tubes (internal diameter 47 mm), without visibly disturbing the sediment surface, and transported to the laboratory in cooled insulated containers. Replication was at n = 4 for oxygen measurements, N2 fixation and inorganic nutrients. In the laboratory, cores were FEMS Microbiol Ecol 83 (2013) 279–298

kept submerged in recirculated, filtered (2 lm) site water. Incubations were conducted at an in-situ temperature and PAR of 500 lmol photons m
2 s
1, the light intensity measured at 7 m depth on a clear day. Cores were stirred with individual Teflon-coated magnetic stirrer bars (25 mm long), driven by an external magnet rotating at 60 r.p.m. This speed was sufficient to circulate the water without visibly disturbing the sediment surface. Cores were allowed to equilibrate in the laboratory for at least 6 h before flux experiments were started. During the flux experiments, DO levels in capped cores were allowed to deviate by no more than 20% from initial values (after Blackburn, 1986). It is important to limit the changes in concentration during the incubation because the sediment and a volume of water are isolated in the cores and the concentrations of gases and solutes in the two phases change with time. These changes alter the diffusional gradient between the sediment and the water column (Miller-Way et al., 1994). Measurements of illuminated cores were taken in daytime and darkened cores at night, to capture any endogenous diurnal patterns. Material for pigment analysis was collected from the same cores that were used for oxygen, nutrient and N2 fixation fluxes after the measurements were completed (n = 8). The top 5 mm of the sediment surface was removed, gently homogenised, and the subsamples were placed in cryogenic vials and stored in liquid N2 at
176 °C within 30 min of harvesting. Because of the amount of sample material needed for bulk sediment parameters (granulometry, lipids and organic C and total N content), additional samples were simultaneously collected from the top 5 mm of sediment extruded from 10 cm diameter cores. Sample sizes were the following: August n = 1 (combined samples from eight cores); December and March n = 4, December MB exposed site organic carbon (OC) n = 1 datum as a result of equipment problems. The samples used for granulometry, lipids, % C and % N content were collected into solvent-rinsed glass jars on site, returned to the laboratory in the dark and on ice and then frozen at
20 °C. Bright-field microscopy (at 2009 magnification) was performed on sediment from the cores used for flux measurements. Qualitative observations were made to identify the most abundant taxa of BMA, for comparison with biomarker data. Analysis

Grain sizing Sediments from each site were dried at 50 °C and weighed before wet-sieving through an 11 sieve stack (2360–45 lm). The sieves containing the sediments were then dried at room temperature for 48 h, and the dried sediment fractions weighed. The degree of sorting was ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

282

calculated using the inclusive graphic quartile deviation index (QDI) (Giere et al., 1988): QDI ¼

u84
u16 u95
u5 þ ; 4 6:6

where φ =
log2 (grain size in lm), φx is the φ value of the xth percentile of the cumulative weight plot. The results for sediment grain size at different depths were classified into one of the seven categories based on their degree of sorting.

OC

Sediment samples were freeze-dried, ground to a powder with a mortar and pestle and weighed into aluminium cups (Elemental Microanalysis) for OC analysis. Before analysis for OC, samples were sequentially acidified with sulphurous acid to remove carbonate (Nieuwenhuize et al., 1994) and dried in an oven at 50 °C. Sediment samples for nitrogen analysis were weighed into tin cups. Samples were analysed for nitrogen and carbon contents, using a Carlo Erba NA1500 CNS analyser. Combustion and oxidation were achieved at 1090 °C and reduction at 650 °C. Where necessary (because of high carbon contents), the carbon signal was quantitatively diluted with helium. Samples were analysed at least in duplicate. The standard for carbon is Vienna PeeDee Belemnite, and the standard for nitrogen is atmospheric N2.

Lipids

Sediment samples were extracted three times by a one-phase dichloromethane–methanol–water mixture (3 : 6 : 1 v/v/v), according to a modified version of the Bligh and Dyer method (Bligh & Dyer, 1959). An aliquot of the total extract was saponified with 3 mL of 5% KOH in methanol/water (80 : 20) and heated at 80 °C for 2 h. The neutral lipid fraction was extracted into hexane/chloroform (4 : 1, the remaining extractions used this mixture), and then, the remaining material was acidified and the fatty acid fraction extracted. Fatty acid methyl esters were formed by treating the fatty acid fraction with MeOH/ HCl/CHCl3 (10 : 1 : 1) at 80 °C for 2 h and then extracting. The neutral fraction was treated with bis(trimethylsilyl) trifluoroacetamide, 100 lL at 60 °C for 2 h to convert hydroxylated compounds such as sterols and alcohols to their trimethylsilyl ethers. Initial gas chromatography (GC) was performed using a Varian CP 3800 with hydrogen as the carrier gas. Analysing the neutral fraction, the GC was equipped with a 50 m 9 0.32 mm i.d. cross-linked 5% phenyl-methyl silicone (HP5; Hewlett ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

Packard), fused-silica capillary column. The total fatty acid fraction (as methyl esters) was analysed on the same instrument, except that an septum programmable injector was used with the capillary column (Hewlett Packard HP1: fused-silica, 50 m long, 0.32 mm i.d. cross-linked 1% phenyl-methyl silicone). Sterol and fatty acid fractions were analysed using a flame ionisation detector (FID), with 5b(H)-cholestan-24-ol as the internal standard for sterols and the methyl ester of tricosanoic acid as the internal standard for fatty acids. Peak identifications were based on retention times relative to authentic and laboratory standards and subsequent GC-MS analysis. The detection limit for individual sterols and fatty acids was approximately 0.2 mg m
2 of 0.5-cm-deep sediment. Individual sterols and fatty acids were identified by GC-MS analyses performed on a Thermoquest/Finnigan GCQ-Plus bench-top mass spectrometer fitted with a direct capillary inlet and an automated on-column injector. Data were acquired in scan acquisition or selective ion monitoring. The nonpolar column (HP5) and operating conditions were the same as that described above for the GC-FID analyses, except that helium was used as the carrier gas. For the analysis of sediment microbial communities, fatty acids were ascribed to the sources listed in Supporting Information, Appendix S1 and neutral lipid sources in Appendix S2 using a range of references (Budge & Parrish, 1998; Pancost & Sinninghe Damste´, 2003; Volkman, 2003). Because of its large concentrations and potentially ambiguous origins, the 15:0 fatty acid was considered separately from the other bacterial fatty acids, and not included in the estimates of bacterial biomass. Fatty acids used to calculate total algal fatty acids were the following: 14:0, 16:4, 16:1x9, 16:1x7, 16:0, 18:2x6, 18:4x3, 18:1x9, 20:5x3, 22:6x3; diatoms: 14:0, 16:1x7, 16:0, 20:5x3. Quantities of other BMA fatty acids were probably from diatoms too, but proportions were not known. The chlorophyte fatty acids could also be found in some other classes, so assignations to that class were made in conjunction with pigments (chl b, lutein). Sterols are referred to by their trivial names for the ease of reading, that is, cholesta-5,22E-dien-3b-ol as 22-dehydrocholesterol, cholest-5-en-3b-ol as cholesterol, 24-methylcholesta-5,22E-dien-3b-ol as brassicasterol, 24-methylcholesta-5,24(28)-dien-3b-ol as 24-methylenecholesterol, 24-methylcholest-5-en-3b-ol as campesterol, 24-ethylcholesta-5,22E-dien-3b-ol as stigmasterol, 24ethylcholest-5-en-3b-ol as sitosterol, and 4a,23,24trimethyl-5a-cholest-22E-en-3b-ol as dinosterol. The conversion of stenols to stanols (hydrogenation of the delta 5 bond) is an early step in the process of diagenesis of stenols, and the stanol/stenol ratio is used here as an index of bacterial reprocessing and grazing by FEMS Microbiol Ecol 83 (2013) 279–298

283

Sandy sediment microbial communities and nutrient exchanges

metazoans (Gagosian et al., 1980; Canuel & Martens, 1993; Jaffe et al., 2001). Pigments

Sediments were extracted twice in 100% acetone at 4 °C, firstly for 18 h and then for 4 h. Following extraction, water was added to give a ratio of 9 : 1 acetone/water by volume and filtered (0.2-lm membrane filter; Whatman Anatop, Whatman plc, Brentford, UK) before analysis by HPLC. The analysis was by a Waters high-performance liquid chromatography, comprising a model 600 controller, 717 plus refrigerated auto-sampler, and a 996 photodiode array detector. Pigments were separated as described by (Wright et al., 1991), detected at 436 nm, and identified against standard spectra. Concentrations of chlorophyll a (chl a), chlorophyll b (chl b), b,b-carotene, and b,e-carotene in sample chromatograms were determined from standards, and all other pigment concentrations were determined from standards of purified pigments isolated from algal cultures. The pigments measured in this study were the following: chlorophyllide a, chlorophylls c1, c2 and c3, fucoxanthin, 19′-hexanoyloxyfucoxanthin (abbreviated to 19′-hex), diadinoxanthin, diatoxanthin, lutein, zeaxanthin, chlorophyll b, chlorophyll a allomer, chlorophyll a, chlorophyll a epimer, phaeophytin a, phaeophytin b, b,b-carotene, b,ecarotene, pyrophaeophytin a. Chlorophyll a concentration was calculated as the sum of chlorophyll a allomer, chlorophyll a, chlorophyll a epimer. Information for the assignment of microalgal marker pigments was drawn from Jeffrey et al. (1997). While fucoxanthin is widely regarded as a marker pigment for diatoms, it also occurs in prymnesiophytes and dinoflagellates, but the absence of chl c3, 19′-hex and peridinin makes these classes unlikely sources. Raphidophytes cannot be ruled out by pigment or lipid data, but the only reference that could be found to their occurrence in sediments was as resting stages (Stahl-Delbanco & Hansson, 2002). In this study, fucoxanthin was used as a diatom marker. Chlorophyll b is found in three classes of green microalgae, Prasinophyceae, Euglenophyceae and Chlorophyceae; however, no prasinoxanthin or violaxanthin was detected, making prasinophytes an unlikely source. The sediments contained other pigments found in euglenophytes, b,b-carotene and diadinoxanthin, but the presence of lutein (not found in euglenophytes) made chlorophytes the most likely source of the chl b. Correlation between lutein and chl b was strong and consistent (0.914, P < 0.001) across studies carried out in Cockburn Sound. The ratio of chl b to chl a decreases with light intensity (Falkowski & Owens, 1980), so was treated with caution as an indicator of the importance of FEMS Microbiol Ecol 83 (2013) 279–298

chlorophytes. In this study, zeaxanthin is used as a marker for cyanophytes, to estimate biomass, a ratio of 0.56 for zeaxanthin/chl a was used (Gibb et al., 2001). The ratio of phaeophytin a to pyrophaeophytin a and chl a were used as an indicator of the degradation of autotrophs (Bidigare et al., 1986; Millie et al., 1993; Sun et al., 1994). Great care was taken with sample preparation, and no macroalgal contamination was observed, but the possibility cannot be excluded. Nutrient fluxes

Inorganic nutrients were analysed on a Lachat 8000QC

flow injection system. NO
2 , NO3 and PO4 were analysed using Lachat standard methods. NH4 was analysed using o-phthalaldialdehyde derivatisation and fluorescence detection as described in Watson et al. (2004) except the oven temperature was set at 50 °C. Flux rates were calculated as nutrient flux ¼

D½nut  v ; A  Dt

where [nut] is the molar concentration of inorganic nutrient; v, the volume of the water column in the core; A, the cross sectional area of the sediment surface; and t, time. For calculating dissolved organic nitrogen (DON), total nitrogen was determined from autoclave digests with potassium persulphate, modified from Valderrama (1981). Net rates of inorganic nutrient fluxes over 24 h were calculated from hourly rates measured in the light and in the dark, multiplied respectively by the lengths of day and night at the time of sampling. Oxygen fluxes

Water above the sediment cores was measured for DO with a calibrated WTW Oxi 440 probe (precision ± 5%). The same cores were used for oxygen and inorganic nutrient fluxes. The tops of the core tubes were raised above the surrounding water level, capped with watch glasses and any bubbles were carefully removed. Measurements were taken at the beginning and end of the incubations, and the rate was calculated as oxygen flux ¼

D½O2   v ; A  Dt

symbols are the same as for nutrient flux calculation. Net primary production (NPP) was the rate of oxygen evolution during the day; community respiration (CR) was the rate of oxygen uptake in the dark. Gross primary production (GPP) was calculated as the sum of NPP and CR ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

284

and the productivity/respiration (P/R) ratio as GPP divided by CR. DO concentrations in capped cores were allowed to deviate by no more than 20% from initial values. To test for linearity, four measurements were taken through the course of incubations for different sediments and the fit to a straight line was consistently good (R2  0.991). N2 fixation

N2 fixation was measured by the acetylene reduction assay (Capone, 1993). Core tubes were raised until the tops were clear of the water surface and water was gently removed until a depth of 5 cm remained, enough to cover the stirrer bars. The cores were capped with clear lids and acetylene injected via a small septum to give a concentration of > 20% ethylene. For each incubation, a time series of three gas samples was collected via the septa into 5-mL evacuated glass tubes (Vacutainer, BD, North Ryde, Australia). Incubations were performed illuminated during the day and in the dark overnight, to capture diurnal patterns of N2 fixation (Bautista & Paerl, 1985). Two cores were incubated for each plot, one in the light and another in the dark. The concentration of ethylene was measured within a week using a Hewlett Packard 5890 gas chromatograph equipped with an Alltech AT Alumina column (30 m, 0.53 mm i.d.) and a FID. Plotted 24-h rates are calculated by multiplying daytime and night-time rates by in-situ periods of light and dark. The molar ratio of C2H2 reduced to N2 fixed varies from 3 to 4 (Charpy-Roubaud et al., 2001), and a ratio of 5 was used for this study. Attempts to experimentally determine the ratio for this study, using 15N2 (Capone & Montoya, 2001), failed because of an absence of N2 fixation activity. A ratio of 4 : 1 was used for conversion from ethylene produced to N2 fixed (Capone, 1993). Statistics

Statistical analysis included the generalised linear modelling (GLM) routine, owing to the continuous nature of the variables. Data were examined for normality and natural log transformation applied where necessary to stabilise variances. GLM was applied to comparisons between sites across locations and times of year. Student’s t-test was used for simple analysis for difference in a parameter between sites at one location; Pearson’s correlation statistics was used for testing for associations between biomarker compounds. Analyses were carried out in SPSS (SPSS Inc., Chicago) and R (R Development Core Team, 2008). ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

Results Site parameters

The differences in the degree of exposure between the sheltered and exposed sites were clearly observed by divers during sample collection. During the August (winter) sampling, water movement was strong enough that divers had difficulty working at the exposed sites, but the sheltered sites presented no such problems. An easily resuspended dark brown floc layer was often present on the surface of the sediments at sheltered sites, particularly in JB, but not at the exposed sites. Sediments were sorted by waves and currents to a greater degree in exposed than in sheltered sites. The QDI indicated that sediments were poorly sorted in sheltered sites (QDI at JB = 1.23, MB = 1.46), but moderately sorted at JB (QDI = 0.91) and moderately well sorted at MB (QDI = 0.69). Water temperatures remained warm on all visits to all sites (Table 1), the coolest being 15.6 °C in August in Cockburn Sound. There was a seasonal pattern, with summer temperatures reaching 24.2 °C in March at JB. Concentrations of dissolved inorganic nutrients in the water column were generally in the oligotrophic range
(Table 2). Concentrations of both NHþ 4 and NOx in the water column were greater at sheltered than at exposed sites at JB (t = 2.6, P = 0.02 and t = 2.9, P = 0.01 respectively). NH4 was the primary form of inorganic nitrogen in all samples. Concentrations of NHþ 4 were greater in the porewaters than in the water column at MB (t = 2.98, P = 0.02) and at JB (t = 2.04, P = 0.05). NO
x concentrations were greater in the water column at JB (t = 2.45, concentrations were not consistently P = 0.03). PO3
4 different between sheltered and exposed sites, but were greater in the porewaters than the water column at JB (t = 3.54, P < 0.01) less distinctly at MB (t = 2.19, P = 0.06). Water column concentrations of DIN were elevated in March at JB, N/P ratios were 151 and 269 at exposed and sheltered sites, respectively. There were no consistent trends in the concentrations of DON between sheltered and exposed sites or between the porewaters and the water column. Concentrations of SiOþ 4 in the Table 1. Day lengths and site temperatures at three sampling periods and two locations Site

August 2002

December 2002

March 2003

Day length (h) Mean daily max PAR (lmol phot m
2 s
1) JB (°C) MB (°C)

10.5 1017

15.1 2014

13.0 1641

15.6 15.6

20.9 21.9

24.2 n/s

n/s, Not sampled.

FEMS Microbiol Ecol 83 (2013) 279–298

285

Sandy sediment microbial communities and nutrient exchanges

Table 2. Water chemistry (lM) at exposed (exp) and sheltered (shelt) sites over three sampling periods August 2002

December 2002

JB exp Water column NO
0.62 x 0.20 PO3
4 NHþ 0.13 4 SiOþ – 4 DON 1.68 Porewater 0–5 cm NO
0.12 x 0.21 PO3
4 NHþ 10.49 4 – SiOþ 4 DON 0.00 Porewater 5–10 cm NO
0.05 x PO3
4.47 4 NHþ 28.85 4 SiOþ – 4 DON 2.51

MB shelt

exp

March 2003

JB shelt

MB

exp

shelt

exp

JB shelt

exp

shelt

2.23 0.20 0.30 – 3.48

0.48 0.28 0.37 – 3.24

0.08 0.04 0.14 – 0.74

0.02 0.21 0.24 2.29 9.80

1.30 0.09 1.16 4.75 2.16

0.29 0.41 0.64 2.01 8.90

0.06 0.13 0.16 – 2.99

0.26 0.08 1.42 1.11 –

1.34 0.02 4.04 2.20 –

0.71 2.05 13.39 – 1.13

0.54 2.58 3.26 – 0.86

0.12 4.17 2.39 – 1.08

0.38 5.93 6.33 101.00 0.00

0.39 5.26 64.18 5.14 0.00

0.29 9.09 35.23 – 0.00

0.52 2.42 37.64 112.91 0.00

0.26 2.31 14.62 18.77 –

0.40 3.93 106.16 72.83 –

0.12 4.46 22.57 – 0.99

0.36 3.31 1.87 – 2.33

0.12 3.35 46.26 – 2.23

0.39 8.03 6.16 89.61 0.56

0.39 0.39 56.13 125.21 0.00

0.46 1.33 54.54 – 0.00

0.34 5.77 52.12 104.62 0.00

0.23 2.36 15.41 24.93 –

0.39 4.89 76.18 65.80 –

‘–’indicates missing measurements of SiOþ 4 and DON owing to equipment problems.

porewaters were always greater than in the water column, sometimes by an order of magnitude. There were no consistent differences between the shallow and deep porewater samples in the concentrations of any nutrients (Table 2). Microscopy

Diatom species identified from the sediments in this area included the following: Cocconeis sp., Diploneis sp., Nitzschia closterium, Pleurosigma sp., a range of sizes of naviculoid cells, Trigonium sp., Gyrosigma sp., Amphora sp., and Chaetoceros sp., although centric diatoms were rare. Nondiatom microalgae were very rarely seen. Organic C and N

Carbon/nitrogen (C/N) ratios were significantly greater (F = 14.3, P < 0.01) at exposed than at sheltered sites (Fig. 1e and f) by an average of 3.5 times. The C/N ratios were consistently above the Redfield value of 6.6; the smallest value was 9.62 (n = 1) at MB in August. Concentrations of OC ranged from 1.1 to 11.5 mol m
2 (0.2–1.6% dry weight) and of nitrogen from 0.08 to 0.99 mol m
2 (0.004–0.096% dry weight). Concentrations of OC were greater (F = 98.73, P < 0.01) by an average of 64% at sheltered than at exposed sites (Fig. 1a–d). OC concentrations varied between cool (August) and warm (December) months increasing by 50–60%, except for the MB exposed site, which showed decreases of 62%. FEMS Microbiol Ecol 83 (2013) 279–298

Pigments

The concentrations of chl a were on average 67% greater (F = 41.24, P < 0.01) at sheltered than at exposed sites (Fig. 2a and b). Differences in chl a concentrations between sheltered and exposed sites ranged from a mean value of 72.3% [standard error (SE) = 2.6%, n = 8] to 14% (SE = 10%, n = 8). The BMA community in these sediments was dominated by diatoms; the ratios of fucoxanthin to chl a were around an order of magnitude greater than either zeaxanthin or chl b (Fig. 2b–f), and community composition was similar across all sites based on the pigment biomarker data. There were significant increases in the proportion of chlorophytes in August 2002 at the MB exposed site and cyanophytes in March 2003 at the JB sheltered site. There was a small increase in the proportion of cyanophytes (zeaxanthin) in March 2003 at JB sheltered site, to a maximum of 12% of chl a [using a ratio of 0.59 for zeaxanthin/chl a (Gibb et al., 2001)]. The maximum biomass of chlorophytes (chl b) was [using a ratio of chl b to chl a of 0.42 (Gibb et al., 2001)] about 17% of chl a in August at MB. There were significantly greater (F = 38.7, P < 0.001) proportions of phaeopigments at the sheltered than at the exposed sites (Fig. 2g and h). When analysed for each location and time of year, ratios were significantly different between exposed and sheltered sites on all occasions except JB in August (F = 0.15, P = 0.7). The ratios for all ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

286

Jervoise Bay 14

mol m–2

12

Organic carbon

(b)

*

10 8 6

2

2 0 Dec

Mar

Aug

(d) 1.2

Total N

Total N

1.0

*

0.8

**

0.6

mol m–2

mol m–2

1.0

Dec

**

0.8 0.6

0.4

0.4

0.2

0.2

0.0

0.0 Aug

Dec

Mar

45

Aug

(f)

C:N

**

40 35

16

30

14

25

Dec

20 18

Ratio

Ratio

6 4

Aug

(e)

8

4

1.2

Organic carbon

10

0

(c)

14 12

** mol m–2

(a)

Mangles Bay

C:N

12 10

20

8

15

6 10 Aug

Dec

Mar

Aug

sites were different at the three times of year (F = 11.9, P < 0.001), averaging 0.30 (SE = 0.07, n = 4) in August, 0.34 (SE = 0.05, n = 4) in December and 0.43 (SE = 0.08, n = 2) in March. Lipids

BMA markers dominated the fatty acids in the sediments, fractions ranged from 64.2% (SE = 0.6, n = 4) of total fatty acids in December, to 71.8% (SE = 0.1, n = 4) in March. Bacterial fatty acids also formed a large component of the community, 28.1% SE = 0.54 in December to 23.3% SE = 0.2 in March. Total bacterial fatty acids correlated strongly with algal fatty acids across all sites and times (0.99, P < 0.001). ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

Dec

Fig. 1. Plots of sediment OC (a, b), total nitrogen concentrations (c, d) and C/N ratios (e, f) at exposed and sheltered sites at JB and MB in August, December 2002 (JB and MB) and March 2003 (JB). Sheltered sites are represented by filled symbols and exposed sites by empty symbols. Error bars are standard error; *significant difference between sites at 0.01 < P < 0.05 at each sampling time; **P < 0.01. August 2002 n = 1, pooled samples each from eight cores; December and March n = 4, except in A December MB exposed site OC n = 1 owing to equipment problems.

The ratio of bacterial to BMA fatty acids was greater at sheltered than at exposed sites (F = 5.117, P = 0.034) except at MB in December; there was no consistent effect of time of year (F = 0.717, P = 0.500). Sheltered sites had greater concentrations of fatty acids from algae, that is, 16:1x7 (F = 8.53, P = 0.008), 18:0 (F = 7.55, P = 0.012), total BMA (F = 5.426, P = 0.029), and from bacteria (F = 4.505, P = 0.045) than exposed sites. A similar suite of lipids was found in the sediments at all sites, only the proportions and concentrations of the compounds differed. Concentrations of the phytosterols (brassicasterol, campesterol, sitosterol and stigmasterol) correlated strongly (Table 3), suggesting that most of the sterol (eukaryote) content of the sediments had a common source in BMA. FEMS Microbiol Ecol 83 (2013) 279–298

287

Sandy sediment microbial communities and nutrient exchanges

Jervoise Bay (a)

Mangles Bay (b)

100

*

60 40 20

(d)

0.30 0.25

*

0.20 0.15

Dec

0.20 0.15

0.14 0.10

0.08

**

0.06 0.04 0.02

Pigment : chl a

0.12

0.10

0.08 0.06 0.04 0.02

0.00

0.00

–0.02

–0.02

Dec

1.2

Mar

**

1.0

*

0.8

Aug

(h)

0.6 0.4

Dec

0.30 0.25

Phaeo : chl a

Pigment : chl a

Aug

0.25

Mar

(f)

Aug

Phaeo : chl a

Dec

0.30

0.12

** **

0.20 0.15 0.10 0.05

0.2

0.00

0.0

Aug

Brassicasterol

Sitosterol

Stigmasterol

0.783 (< 0.001) 0.931 (< 0.001) 0.924 (< 0.001)

0.924 (< 0.001) 0.953 (< 0.001)

0.856 (< 0.001)

The total concentration of neutral lipids showed greater biomass at sheltered than at exposed sites, at both JB and MB (t = 4.39, d.f. = 21, P < 0.001). Concentrations of FEMS Microbiol Ecol 83 (2013) 279–298

Dec

0.14

Table 3. Correlations between concentrations of brassicasterol and other phytosterols (Pearson’s correlation coefficient R), n = 22

Campesterol Stigmasterol Sitosterol

Aug

0.10

Aug

(g)

40

Mar

Fucoxanthin : chl a

Fucoxanthin : chl a

Dec

0.10

Fig. 2. Plots of sediment data from exposed and sheltered sites; chl a concentrations (a, b), ratios of fucoxanthin to chl a (c, d), zeaxanthin (circle symbols) and chlorophyll b (triangle symbols) to chl a (e, f), phaeopigments to chl a (g, h) at JB and MB in August, December 2002 (JB and MB) and March 2003 (JB). Sheltered sites are represented by filled symbols and exposed sites by empty symbols. Error bars are standard error, n = 8, bars marked with * indicate significant difference between sites at 0.05 > P > 0.01, **P < 0.01.

60

0

Aug

(e)

**

20

0

(c)

**

80

mg chl a m–2

mg chl a m–2

80

100

Dec

Mar

Aug

Dec

phytol, a part of the chlorophyll a molecule, were also greater at sheltered than at exposed sites at both JB and MB (t = 3.31, d.f. = 21, P = 0.003). The concentration of 24-methylenecholesterol, a marker for centric diatoms, was greater at sheltered than at exposed sites (F = 14.782, P = 0.001): as was the ratio of 24-methylenecholesterol to brassicasterol (diatoms) (F = 5.399, P = 0.030). Cholesterol was the second most plentiful sterol found in these sediments. Compared to exposed sites, sheltered sites had greater concentration of cholesterol (F = 9.35, P = 0.006) and a larger ratio of cholesterol to brassicasterol ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

288

(F = 6.70, P= 0.003). The stanol/stenol ratios were greater at sheltered than at exposed sites, except at MB in August (Fig. 3); the difference was highly significant (F = 27.05, P < 0.001). The ratio of bacterial to BMA fatty acids (Fig. 4a and b) was greater at sheltered than at exposed sites (F = 7.04, P = 0.015). Different components of the sediment community made greater contributions at different times of year. Multiple regression of phytol and summed bacterial plus microheterotroph fatty acids against OC concentrations: OC ¼ a ðphytolÞ þ b ðbacteria & heterotrophsÞ: There was a dominance of autotrophic biomass in August and December [R = 0.726, standardised coefficients in the regression: phytol (a = 0.74) t = 3.50, P = 0.005, bacterial and heterotroph fatty acids (b =
0.15) t =
0.73, P = 0.48], with bacterial and heterotrophic dominance in March (JB) [R = 0.883, standardised coefficients in the regression: phytol (a =
0.32) t =
1.40, P = 0.22, bacterial fatty acids (b = 0.96) t = 4.19, P = 0.009]. Dinosterol was found in concentrations of 7.9% (0.09 mg m
2) to 65.7% (0.6 mg m
2) of brassicasterol concentrations, except in March 2003 (JB), where the concentration (11.7 mg m
2) reached 150% that of brassicasterol. Dinosterol also increased in concentration relative to phytol between winter and the later two sampling times; the ratios were 0.020 (SE = 0.012, n = 4) in August, 0.100 (SE = 0.016, n = 16) in December and 0.24 (SE = 0.068, n = 16) in March. The differences were significant between August and December (P = 0.013), August and March (JB, P < 0.001), but not between December and March (P = 0.16) at JB. The large increases in the dinosterol/total sterol ratio over time suggested that heterotrophic dinoflagellates increased as a proportion of biomass.

Jervoise Bay

(a)

O2 fluxes

P/R ratios (GPP/CR) were significantly greater at sheltered than at exposed sites (F = 6.3, P = 0.019); average differences were 25% (SE = 8, n = 24) (Fig. 5c and f), except for MB in August, where 3 of the 4 replicates at the exposed site were net heterotrophic, but the other had a P/R ratio of 4.04. Mean rates of NPP at different sites ranged from
0.39, SE = 0.27 (n = 3) to 2.44, SE = 0.25 (n = 3) mmol O2 m
2 h
1 (Fig. 5a and b) and CR from 0.15, SE = 0.04 (n = 3) to 2.12, SE = 0.21 (n = 3) mmol O2 m
2 h
1 (Fig. 5c and d). Over all seasons, rates of NPP were consistently greater (F = 26.2, P < 0.001) at sheltered than at exposed sites by an average of 55% (SE = 18, n = 30). Considered individually, all sheltered sites were significantly different on all dates except the December sampling of JB. There was no consistent relationship between CR with the degree of shelter between the two sites at JB in August or March (Fig. 5c). At MB, CR was always significantly greater (F = 31.4, P < 0.001) at the sheltered than at the exposed site (Fig. 5d). When CR was normalised to OC, rates were around double at exposed (28.4%, SE = 0.69%, n = 5) than at paired sheltered (15.0%, SE = 1.2%, n = 5) sites (t =
2.48, P = 0.0685). The ratio was similar between sites at MB in August; if these data were removed from the analysis, the difference became highly significant (t =
6.57, P = 0.0072) and remained a factor of 2.1. There was an increase in CR between August and December at all sites (F = 12.3, P < 0.001). N2 fixation

Rates of N2 fixation were small, generally at least an order of magnitude less than DIN fluxes (Fig. 6a and b); the

0.4

0.2

0.0

**

**

**

Aug

Dec

Mar

Stanol/Sterol ratio

Stanol/Sterol ratio

0.3

0.1

Mangles Bay

(b)

0.4

0.3 0.2

**

**

0.1 0.0

Aug

Dec

Fig. 3. Plots of averaged stanol/sterol ratios of most abundant sterols as follows: cholesterol, brassicasterol, campesterol, stigmasterol and sitosterol for sediments at sheltered and exposed sites at JB and MB in August, December 2002 (JB and MB) and March 2003 (JB). Sheltered sites are represented by filled symbols, exposed sites by empty symbols. For August, n = 1 (combined samples from eight cores), December, March n = 4; error bars are standard error; **significant difference at P < 0.01 between sheltered and exposed sites.

ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

FEMS Microbiol Ecol 83 (2013) 279–298

289

Sandy sediment microbial communities and nutrient exchanges

Jervoise Bay

Mangles Bay (b)

0.7

**

0.6

*

0.5 0.4 0.3

0.7

Bacterial : BMA f.a. ratio

Bacterial : BMA f.a. ratio

(a)

Aug

Dec

0.4 0.3 0.2

Mar (d)

**

14 12 10

**

8 6 4 2 0

Dinosterol : sterol ratio

Dinosterol : sterol ratio

(c) 18 16

Aug

Dec

Aug

Dec

10 8 6 4 2 0

Aug

maximum rate measured was 31.3 lmol N2 m
2 day
1 or 1.3 lmol N2 m
2 h
1 fixed (see Materials and methods for ratio). In August 2002 there was no detectable acetylene reduction activity at either JB or MB. In December 2002, JB sheltered water column concentration of NHþ 4 was an order of magnitude greater than at JB exposed; N2 fixation was greater at the exposed site. At MB exposed site NHþ 4 concentrations were greater than MB sheltered site; again, N2 fixation rates were greater at the sheltered site. A similar pattern occurred in March 2003 at JB, with smaller water column concentrations of NHþ 4 and greater rates of N2 fixation occurring at sheltered than at exposed sites. Inorganic nutrients

Night-time (dark) and daytime (light) hourly rates of fluxes of DIN were significantly different (t = 2.88, P < 0.007) over all sites and sampling occasions. There was an average increase from 8.9 lmol m
2 h
1 (SE = 3.3, n = 30) to 20.0 lmol m
2 h
1 (SE = 5.6, n = 30), presumably the effect of photosynthesis on fluxes. NH4 was the major DIN flux, often up to two orders of magnitude larger than NOx. While fluxes often differed between the exposed and sheltered sites (Fig. 7), the differences were usually not FEMS Microbiol Ecol 83 (2013) 279–298

0.5

0.1

0.2

Fig. 4. Plots of sediment lipid ratios from exposed and sheltered sites at JB and MB; bacterial/BMA fatty acids (a, b) and dinosterol/ total sterols (9100) (c, d), in August, December 2002 (JB and MB), and March 2003 (JB). Sheltered sites are represented by filled symbols and exposed sites by empty symbols. For August, n = 1 (combined samples from eight cores); December, March n = 4; error bars are standard error; *significant difference between sites at 0.01 < P < 0.05; **P < 0.01 between sheltered and exposed sites.

0.6

Dec

Mar

temporally consistent between the locations JB and MB. For example, in August (winter), DIN uptake at the MB exposed site was significantly greater than at the sheltered (Fig. 7b), while at JB fluxes at the exposed and sheltered sites were nearly the same (Fig. 7a). DIN fluxes at JB in August were two orders of magnitude less than in December and March; the sediments at MB had larger DIN fluxes in August than in December.

Discussion Exposure of the microbial communities in sediments to disturbance by waves and currents resulted in reduced biomass and changes in the composition and metabolism of the biota. The pattern of reduced biomass at exposed sites relative to paired sheltered sites at the two locations, MB and JB, was consistent across seasons sampled. The communities were always dominated by diatoms and bacteria, but the ratio of bacteria to BMA was 29% smaller in exposed than in sheltered sediments. The exposed sediments also had significantly greater C/N ratios, averaging 3.5 times those in sheltered sediments and significantly smaller P/R ratios. Metabolism at exposed sites was on average 36% more heterotrophic than the paired sheltered sites, most likely due to the bacterial respiration associated with increased production of labile carbohydrates by ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

290

Jervoise Bay 3

Mangles Bay (b)

NPP

*

**

2

3

mmol O2 m–2 h–1

mmol O2 m–2 h–1

(a)

1

0

–1

1

* 0

Mar

Aug

(d) 2.5

CR

mmol O2 m–2 h–1

mmol O2 m–2 h–1

Dec

2.0 1.5 1.0 0.5

Aug

P:R 3

**

1.5 1.0

**

0.5

Dec

Mar

Aug

Dec

P:R

(f)

*

3 2

**

Ratio

Ratio

CR

2.0

2 1

1

0

0

–1

–1

Aug

µmol m–2 d–1

Dec

0.0

0.0

(e)

2

–1

Aug

(c) 2.5

**

NPP

Dec

Mar

Aug

(a)

(b)

140 JB

140

120

120

100

100

80

80

60

60

*

40

MB

**

40 20

20

0

0 Aug

Dec

Mar

Aug

Dec

Fig. 6. Plots of nitrogen fixation, measured using acetylene reduction rates (a, b) at exposed and sheltered sites at JB and MB over 24 h in August, December 2002 (JB and MB) and March 2003 (JB). Sheltered sites are represented by filled symbols, exposed sites by empty symbols. n = 4, error bars are standard error; *significant difference between sites at P < 0.05, **significant difference at P < 0.01.

ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

Dec

Fig. 5. Plots of sediment–water fluxes of oxygen from exposed and sheltered sites at JB and MB; NPP (a, b), CR (c, d) and N/P ratio (e, f); August 2002, December 2002 (JB and MB), March 2003 (JB), n = 3. Sheltered sites are represented by filled symbols and exposed sites by empty symbols. Error bars are standard error; *significant difference between sites at 0.01 < P < 0.05; **P < 0.01.

BMA. Indices of the breakdown of organic matter were greater at sheltered sites. Exchanges of nutrients between the sediment and water column remained similar between paired sheltered and exposed sites. The changes in these microbial communities with disturbance provided a mechanism for the retention of nutrients in sediment porewaters. This conclusion was reinforced by the near absence of nitrogen fixation, a process typically associated with conditions of N limitation. Exchanges of N2 gas by diazatrophy were two orders of magnitude smaller than those of DIN in these sediments. The BMA in this system comprised the major fraction of the autotrophic biomass in these shallow coastal waters. Concentrations of chl a in the top 0.5 cm of the sediments ranged from 32.0 mg m
2, 2.4 times greater than the integrated phytoplankton chl a concentration (Wilson & Paling, 2003) at JB sheltered site in December, FEMS Microbiol Ecol 83 (2013) 279–298

291

Sandy sediment microbial communities and nutrient exchanges

Jervoise Bay

(a)

Mangles Bay

0.5

(b)

0.2

–0.5 –1.0

mmol m–2 d–1

mmol m–2 d–1

0.0

DIN

–1.5 –2.0

–3.0

–0.2

Dec

Mar

Aug

(d)

PO4

Dec

10

**

60 0

40

µmol m–2 d–1

µmol m–2 d–1

*

–0.6

Aug 80

0.0

–0.4

–2.5

(c)

0.4

20 0

–10

–20

–20

–40 –30

Aug

(e)

2.0

Dec

Mar

Aug

(f)

Si

1.0

**

0.5

**

1.0 0.5

ND

0.0

0.0

ND –0.5

–0.5

Aug

to 4.2 mg m
2, 14.5 times greater concentration at the exposed site (Wilson & Paling, 2003). With a mean chl a concentration of 48 mg m
2 (SE = 2.0, n = 30) and OC of 7080 lmol m
2 (SE = 756, n = 22), the sediments in Cockburn Sound were similar to those in oligotrophic coastal environments elsewhere, in terms of their relative contribution to autotrophic biomass (Nelson et al., 1999; Gillespie et al., 2000) and concentration of OC (Alongi et al., 1996; Grenz et al., 2003). An experiment to simulate wave resuspension of sediments in Cockburn Sound (Forehead, 2006) resulted in the removal of sufficient material to create a 170% increase in the concentration of OC in the 7-m-deep water column. These microbial communities comprised a substantial reserve of labile OC in Cockburn Sound. FEMS Microbiol Ecol 83 (2013) 279–298

mmol m–2 d–1

mmol m–2 d–1

Fig. 7. Plots of inorganic nutrient fluxes over 24 h at exposed and sheltered sites in August, December 2002 (JB and MB) and March 2003 (JB). Sheltered sites are represented by filled symbols, exposed sites by empty symbols. Note much smaller scale used in B and E; n = 4, error bars standard error; *significant difference between sites at P < 0.05; **P < 0.01; n/d, no data.

2.0 1.5

1.5

Dec

Dec

Mar

Aug

Dec

The elevated C/N ratios in these sediments were attributed to the presence of extracellular polymeric substances (EPS) secreted by BMA. The differences in C/N ratios of the sediment organic matter were then attributed to differences in concentrations of EPS generated by BMA between paired sheltered and exposed sites. Epipsammic diatoms are the dominant type in disturbed sediments generally (Round, 1971) and in the carbonate sediments of south-western Australia (Kendrick et al., 1998). As they are attached to sediment grains, they are unable to migrate away from excessive irradiance. These cells are obliged to channel surplus fixed carbon into the production of EPS as one mechanism to protect against damage to their photosynthetic apparatus when exposed to excessive PAR. Large C/N ratios can also indicate the presence ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

292

of refractory organic matter in sediments influenced by terrestrial, or seagrass organic matter, such as partially degraded material from higher plants (Jaffe et al., 2001). The biomarker evidence argued against the presence of any significant concentrations of such materials in these sediments. The principal sources of OC in this system were BMA, bacteria and the products of their metabolism. The activity of grazers and bacteria, as shown by ratios between pigment and lipid degradation products (Figs 2g, h and 3), was usually less at exposed sites than at sheltered sites. The ratio of bacterial to BMA fatty acids was also an average of 29% less at the exposed than at the sheltered sites. The ratios of cholesterol to brassicasterol and of dinosterol to total sterols were always less at exposed than at sheltered sites, suggesting a smaller proportion of grazers to BMA. Exposed sites also generally had 35% smaller ratios of phaeopigments to chlorophyll a and 10% lower ratios of stanols to their respective sterols, indicating that there was reduced grazing activity and breakdown of organic matter at exposed sites (Sun, 2000). The differences in organic matter processing with degree of shelter were consistent with similar observations worldwide (Plante-Cuny & Bodoy, 1987; Incera et al., 2003a, b), but the exchanges of oxygen and nutrients were not. The wave-exposed sediments had 25% (SE = 8%) smaller P/R ratios than equivalent sheltered sediments, even though they had proportionally less bacteria and smaller indices of grazing and diagenetic activity. At each location, the smaller P/R ratios consistently coincided with the greater C/N ratios (Fig. 8). Rates of NPP were always greater in sheltered than in exposed sites, consistent with the greater concentrations of BMA. However, the metabolism of the microbial communities in sheltered sediments showed a P/R ratio > 1 and was only < 1 (heterotrophic) once (MB August). The ratio in exposed sediments was always < 1, with the exception of JB in August. The cause of the increased rates of bacterial activity in exposed sediments relative to sheltered sediments was most likely the greater concentrations of EPS. The EPS secretions of BMA provide a highly labile source of OC for bacteria (Middelburg et al., 2000; Jensen et al., 2005; Evrard et al., 2008). Glucose can comprise 90% of the photosynthates or EPS secreted by BMA (de Brouwer & Stal, 2001), and it is a preferred substrate for bacteria that can be metabolised rapidly (Sawyer & King, 1993). The rates of activity of bacterial communities are not necessarily proportional to their biomass (Creach et al., 2003). In-situ rates may have been greater than those measured in this study, owing to the replenishment of oxygen in porewaters of permeable sediments by advective transport (Shum & Sundby, 1996). Although bacteria ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

were a smaller fraction of the communities of exposed sediments, their rates of activity were greater than those of the bacteria in sheltered sediments. Other studies in Cockburn Sound sediments have made the same conclusion that there was more ‘heterotrophic’ metabolism in microbial communities with greater autotrophic dominance and that this is associated with disturbance. Measurements of BMA with a pulse amplitude-modulated fluorometer showed that the rates of photosynthetic activity normalised to biomass were similar in sheltered and exposed sediments (Forehead & Thompson, 2010). The sediments also showed a pattern of bacteria/BMA ratio relative to wave disturbance (depth) as found in this study. The most exposed sediments had NPP of near zero, which required that the daytime respiratory activity was around 2.5 times that of CR (Forehead & Thompson, 2010). The close interactions between BMA and heterotrophic bacteria in sediments can be synergistic. In an experimental system (Murray et al., 1986), thymidine uptake in bacteria was accelerated in the presence of EPS from BMA, and in turn, the production of EPS has been shown to depend on a suite of chemical interactions with bacteria (Bruckner et al., 2011). Elevated rates of bacterial respiration consume both DO from the water column and the oxygen output of closely associated, rapidly photosynthesising BMA (Haack & McFeters, 1982; Glud et al., 1999). Much of the oxygen is exchanged within the biofilm, not released to the water column (Bo¨er et al., 2009), giving rise to the lower values of NPP sometimes found in these exposed sediments. The difference in P/R ratios between sheltered and exposed sites was apparent even when the biomass of BMA was not very different, such as at JB in August and in March (Figs 2a and 5e). The differences in rates of production of EPS (inferred from C/N ratios), in response to the different degrees of exposure, are most likely due to the prevailing functional types of diatoms (epipelic or epipsammic) in the communities of BMA. The exchanges of nutrients were small, although the composition of the microbial community was generally distinct between sheltered and exposed sites (Fig. 9). The ratios of the nutrients exchanged across the sediment– water interface were rarely close to Redfield and thereby suggest the effects of other processes in addition to their uptake or release by BMA. There was only one occasion when nutrient fluxes were in close to Redfield proportions of P, N and Si, at MB exposed site in August. More commonly, the activity of BMA on nutrient uptake was obscured by other processes, such as bacterial cycling of nutrients (Riaux-Gobin et al., 1987; Nelson et al., 1999; Qu et al., 2005) or exchange of P adsorbed to sediment surfaces (Mindl et al., 2005; Evrard et al., 2008). Twentyfour-hour rates of DIN exchange were all less than FEMS Microbiol Ecol 83 (2013) 279–298

293

Sandy sediment microbial communities and nutrient exchanges

5

August JB August MB December JB December MB March JB

** 4

Exposed

3 *

P : R ratio

Fig. 8. Scatter plot of P/R ratio vs. C/N ratio showing consistent pattern of differences between paired sheltered and exposed locations at different times of year (MB for August and December 2002 only). Error bars are standard error: for C/N, August n = 1; December n = 3 except MB n = 1, March n = 4; for P/R, n = 4 except August JB exp n = 3, August MB shelt n = 3, December JB shelt n = 2. For P/R ratio, *significant difference between paired sites at each location at each sampling time at P < 0.05; **P < 0.01.

Sheltered

2

1

0 5

10

15

20

25

30

35

40

–1

0.5

Exposed

Aug JB Aug MB Dec JB Dec MB Mar JB

Sheltered

Shelt. 0.0

24 h DIN exchange (mmol m–2 d–1)

Fig. 9. Scatter plot of 24-h DIN exchanges vs. bacteria/BMA fatty acids ratio, showing pattern of differences in community composition, but similarity of DIN exchanges between paired sheltered and exposed locations at different times of year (MB for August and December 2002 only). Ellipse around MB sheltered site shows exception to pattern of community composition related to degree of shelter. Error bars are standard error: for DIN, n = 4; for fatty acid ratio, August n = 1 (combined samples from eight cores), December and March n = 4. For fatty acid ratio, *significant difference between paired sites at each location at each sampling time at P < 0.0.

–0.5

–1.0

** *

–1.5

–2.0

–2.5

0.1

0.2

0.3

0.4

0.5

0.6

0.7

Bacteria : BMA ratio –3.0

1.5 mmol m
2 day
1, and the exchanges were assimilatory on all but one occasion. This, together with the small size of the PO4 fluxes, suggested that much of the nutrient demand of both the autotrophic and heterotrophic sediment community was being met from reserves in the porewaters and through tight recycling of organic matter. Diatoms are found to be the dominant group in BMA assemblages in many parts of the world (Barranguet et al., 1998; Seroˆdio et al., 2001; Saburova & Polikarpov, 2003; Consalvey et al., 2004). They have been reported dominant in the Perth region (Kendrick et al., 1998) and remained the dominant group of BMA reported herein, across sites, locations and times. The only consistent trends in community composition with time across all sites were an increase in the concentration of heterotrophic dinoflagellates (Fig. 4c and d) and a 5.8-fold increase FEMS Microbiol Ecol 83 (2013) 279–298

45

C : N ratio

in concentrations of the scarce cyanophytes (one to two orders of magnitude smaller concentrations than diatom pigments) from August to December to March. These increases coincided with increasing temperature and decreasing frequency of storms. The green pigmented fraction of the BMA community showed only nonsignificant decreases in chl b in response to shelter. The nutrient status of the water column can be important in determining community composition, for example, a low N/P ratio or low concentrations of Si favour a community shift from diatoms to cyanophytes (Risgaard-Petersen et al., 2005). However, nutrient limitation is probably rare in BMA (Watermann et al., 1999; Rocha et al., 2002). In Cockburn Sound, concentrations of inorganic nutrients were generally much greater in porewaters than in the water column. There was release of Si from the ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

294

sediment on all occasions. Si fluxes were measured, suggesting that concentrations in porewaters were in excess for BMA requirements. The various BMA communities in this study differed most in biomass, not in coarse taxonomy. This, together with an absence of a shift to alternate modes of nutrient uptake, such as diazatrophy, further suggested that these BMA communities were not limited by nutrients. The reduced hydrodynamic forces in sheltered sites allowed the accumulation of BMA, mostly autochthonously produced cells but also some settled from the water column. An in-situ experiment in the same region demonstrated that sheltered and enriched sediments showed a 40% increase in BMA biomass over 10 days because of autochthonous production (Forehead et al., 2011). On nearby Parmelia Bank (from 1.5 to 14 m depth) BMA concentration increased with depth and decreasing influence of waves and currents (Forehead & Thompson, 2010). In the current study, a brown, easily resuspended layer was often seen on top of sediments in sheltered sites. Layers of living algal biomass or detritus and associated bacteria on the sediment surface have been associated with a decrease in water movement in a number of other studies (MacIntyre & Cullen, 1995; Kendrick et al., 1998; Beaulieu, 2003; Gerbersdorf et al., 2004). The increased concentrations of 24-methylenecholesterol in sheltered relative to exposed sediments indicated that reduced disturbance led to a greater accumulation of normally planktonic centric diatoms. However, microscopy found that planktonic cells always remained a small proportion of the diatom community. The rates of N2 fixation were less than DIN fluxes by around two orders of magnitude. Physical disturbance reduced rates of N2 fixation in an experimental resuspension of nearby sediments (Forehead, 2006), and in winter, these sediments are frequently disturbed by storms. N2 fixing is most likely to occur when there is N limitation (Villbrandt et al., 1991; Capone, 2001; Marino et al., 2002) and is usually inhibited by concentrations of NHþ 4 > 10–100 lM (Howarth et al., 1988). Concentrations of NHþ 4 in the shallow (0–5 cm) porewaters of Cockburn Sound averaged 29.4 lM (ranged from 6.33 to 106.16 lM) during December and March. Denitrification also played a very minor role in N cycling in these sediments (Forehead, 2006; Forehead et al., 2011). This finding was consistent with an inhibition of the process by oxic conditions caused by BMA activity and low concentrations of NO
3 in the water column (Christensen et al., 1990; Sundba¨ck et al., 2000; Risgaard-Petersen et al., 2005). Anammox is a bacterial process that consumes þ NO
2 and NH4 from sediments and produces N2 (Dalsgaard et al., 2005). It has been found in stable sediments under conditions of anoxia or hypoxia; anammox rates ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

are proportional to concentrations of organic matter. It is unlikely to have been a significant path for N cycling in these sediments.

Conclusion The decrease in the ratio of bacteria to BMA in the microbial communities of these sediments was related to increased hydrodynamic energy. The changes in community composition were accompanied by shifts in metabolism which resulted in retention of nutrients and relatively small exchanges of inorganic nutrients regardless of the degree of exposure to disturbance. The mechanism proposed here for this phenomenon in sediments in oligotrophic waters is the high rates of production of EPS by the BMA that drives heterotrophy. The production of EPS by epipsammic diatoms is likely to be reduced whenever BMA have access to sufficient nutrients or under conditions of low PAR. Functionally similar sediment communities are likely to be found in sandy sediments in mild oligotrophic waters elsewhere. Sandy sediments cover about 70% of the world’s continental shelves (Boudreau et al., 2001), and warm, low-latitude oceans are mostly oligotrophic (Falkowski et al., 1992). Large areas of coastal sandy sediments may have greater capacity as sinks for nutrients and for the photosynthetic uptake of CO2 than oxygen flux measurements alone would suggest. A more robust evaluation of the trophic status of such sediments would include (less frequent) additional measurements, such as analysis for C and N content, or a determination of community composition.

Acknowledgements We would like to thank Dr A. Waite for her helpful advice, field support and laboratory resources. Thanks to L. Clementson, A. Revill and J. Volkman for their useful discussions, and to S. Grove, A. Brearley and K. Kilminster for their assistance with field work; P. Bonham for their assistance with pigment analyses and chromatography; D. Holdsworth and S. Armand for their assistance with lipid chromatography; and V. Latham and R. Watson for their assistance with nutrient analysis. Thanks to two anonymous reviewers whose input improved the manuscript. The authors wish to acknowledge financial support of a PhD scholarship for H.F. and research funding from the Strategic Research Fund for the Marine Environment (a joint initiative of the Western Australian state government and Commonwealth Scientific and Industrial Research Organisation Division of Marine and Atmospheric Research). FEMS Microbiol Ecol 83 (2013) 279–298

Sandy sediment microbial communities and nutrient exchanges

References Alongi DM (1994) The role of bacteria in nutrient recycling in tropical mangrove and other coastal benthic ecosystems. Hydrobiologia 285: 19–32. Alongi DM, Tirendi F & Goldrick A (1996) Organic matter oxidation and sediment chemistry in mixed terrigenouscarbonate sands of Ningaloo Reef, Western Australia. Mar Chem 54: 203–219. Barranguet C, Herman PMJ & Sinke JJ (1997) Microphytobenthos biomass and community composition studied by pigment biomarkers: importance and fate in the carbon cycle of a tidal flat. J Sea Res 38: 59–70. Barranguet C, Kromkamp J & Peene J (1998) Factors controlling primary production and photosynthetic characteristics of intertidal microphytobenthos. Mar Ecol Prog Ser 173: 117–126. Bautista MF & Paerl HW (1985) Diel N2 fixation in an intertidal marine cyanobacterial mat community. Mar Chem 16: 369–377. Beaulieu SE (2003) Resuspension of phytodetritus from the sea floor: a laboratory flume study. Limnol Oceanogr 48: 1235–1244. Bidigare RR, Frank TJ, Zastrow C & Brooks JM (1986) The distribution of algal chlorophylls and their degradation products in the Southern Ocean. Deep Sea Res A 33: 923–937. Blackburn TH (1986) Nitrogen-cycle in marine sediments. Ophelia 26: 65–76. Bligh EG & Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37: 911–917. Bo¨er SI, Arnosti C, Van Beusekom JEE & Boetius A (2009) Temporal variations in microbial activities and carbon turnover in subtidal sandy sediments. Biogeosciences 5: 4271–4313. Boudreau BP, Huettel M, Forster S et al. (2001) Permeable marine sediments: overturning an old paradigm. EOS Trans Am Geophys Union 82: 133–136. Bruckner CG, Rehm C, Grossart H-P & Kroth PG (2011) Growth and release of extracellular organic compounds by benthic diatoms depend on interactions with bacteria. Environ Microbiol 13: 1052–1063. Budge SM & Parrish CC (1998) Lipid biogeochemistry of plankton, settling matter and sediments in Trinity Bay, Newfoundland. II. Fatty acids. Org Geochem 29: 1547–1559. Busse S & Snoeijs P (2003) Gradient responses of diatom communities in the Bothnian Sea (northern Baltic Sea), with emphasis on responses to water movement. Phycologia 42: 451–464. Canuel EA & Martens CS (1993) Seasonal-variations in the sources and alteration of organic-matter associated with recently-deposited sediments. Org Geochem 20: 563–577. Capone DG (1993) Detemination of nitrogenase activity in aquatic samples using the acetylene reduction procedure. Handbook of Methods in Aquatic Microbial Ecology (Kemp PF, Sherr BF, Sherr EB & Cole JJ, eds), pp. 621–631. Lewis Publishers, Tokyo.

FEMS Microbiol Ecol 83 (2013) 279–298

295

Capone DG (2001) Marine nitrogen fixation: what’s the fuss? Curr Opin Microbiol 4: 341–348. Capone DG & Montoya JP (2001) Nitrogen fixation and denitrification. Methods Microbiol 30: 501–515. Carpenter RC, Hackney JM & Adey WH (1991) Measurements of primary productivity and nitrogenase activity of coralreef algae in a chamber incorporating oscillatory flow. Limnol Oceanogr 36: 40–49. Cerco CF & Seitzinger SP (1997) Measured and modeled effects of benthic algae on eutrophication in Indian River Rehoboth Bay, Delaware. Estuaries 20: 231–248. Charpy-Roubaud C, Charpy L & Larkum AWD (2001) Atmospheric dinitrogen fixation by benthic communities of Tikehau Lagoon (Tuamotu Archipelago, French Polynesia) and its contribution to benthic primary production. Mar Biol 139: 991–997. Christensen PB, Nielsen LP, Sorensen J & Revsbech NP (1990) Denitrification in nitrate-rich streams – diurnal and seasonal-variation related to benthic oxygen-metabolism. Limnol Oceanogr 35: 640–651. Colijn F & Dijkema KS (1981) Species composition of benthic diatoms and distribution of chlorophyll a on an intertidal flat in the Dutch Wadden sea. Mar Ecol Prog Ser 4: 9–21. Consalvey M, Paterson DM & Underwood GJC (2004) The ups and downs of life in a benthic biofilm: migration of benthic diatoms. Diatom Res 19: 181–202. Cook PLM, Revill AT, Clementson LA & Volkman JK (2004) Carbon and nitrogen cycling on intertidal mudflats of a temperate Australian estuary. III. Sources of organic matter. Mar Ecol Prog Ser 280: 55–72. Creach V, Baudoux AC, Bertru G & Le Rouzic B (2003) Direct estimate of active bacteria: CTC use and limitations. J Microbiol Methods 52: 19–28. Dalsgaard T, Thamdrup B & Canfield DE (2005) Anaerobic ammonium oxidation (anammox) in the marine environment. Res Microbiol 156: 457–464. Davies JM (1975) Energy flow through the benthos in a Scottish sea loch. Mar Biol 31: 353–362. de Brouwer JFC & Stal LJ (2001) Short-term dynamics in microphytobenthos distribution and associated extracellular carbohydrates in surface sediments of an intertidal mudflat. Mar Ecol Prog Ser 218: 33–44. de Jonge VN & van den Bergs J (1987) Experiments on the resuspension of estuarine sedments containing benthic diatoms. Estuar Coast Shelf Sci 24: 725–740. Evrard V, Cook PLM, Veuger B, Huettel M & Middelburg JJ (2008) Tracing carbon and nitrogen incorporation and pathways in the microbial community of a photic subtidal sand. Aquat Microb Ecol 53: 257–269. Falkowski PG & Owens TG (1980) Light-shade adaptation. Two strategies in marine phytoplankton. Plant Physiol 66: 592–595. Falkowski PG, Greene RM & Geider RJ (1992) Physiological limitations on phytoplankton productivity in the ocean. Oceanography 5: 84–91.

ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

296

Fanning KA, Carder KL & Betzer PR (1982) Sediment resuspension by coastal waters: a potential mechanism for nutrient re-cycling on the ocean’s margins. Deep-Sea Res 29: 953–965. Forehead HI (2006) The ecology and biogeochemistry of sandy sediments in the warm temperate coastal waters of Western Australia. PhD Thesis, University of Western Australia, Perth, Western Australia. Forehead HI & Thompson PA (2010) Microbial communities of subtidal shallow sandy sediments change with depth and wave disturbance, but nutrient exchanges remain similar. Mar Ecol Prog Ser 414: 11–26. Forehead HI, Thompson PA & Kendrick GA (2011) The effects of shelter and enrichment on the ecology and nutrient cycling of microbial communities of subtidal carbonate sediments. FEMS Microbiol Ecol 80: 64–76. Gabrielson JO & Lukatelich RJ (1985) Wind-related resuspension of sediments in the Peel-Harvey Estuarine System. Estuar Coast Shelf Sci 20: 135–145. Gagosian RB, Smith SO, Lee C, Farrington JW & Frew NM (1980) Steroid transformations in recent marine sediments. Advances in Organic Geochemistry (Douglas AG & Maxwell JR, eds), pp. 407. Pergamon, Oxford. Galois R, Blanchard G, Seguignes M, Huet V & Joassard L (2000) Spatial distribution of sediment particulate organic matter on two estuarine intertidal mudflats: a comparison between Marennes-Ole´ron Bay (France) and the Humber Estuary (UK). Cont Shelf Res 20: 1199–1217. Gerbersdorf SU, Meyercordt J & Meyer-Reil LA (2004) Microphytobenthic primary production within the flocculent layer, its fractions and aggregates, studied in two shallow Baltic estuaries of different eutrophic status. J Exp Mar Biol Ecol 307: 47–72. Gibb SW, Cummings DG, Irigoien X, Barlow RG, Fauzi R & Mantoura C (2001) Phytoplankton pigment chemotaxonomy of the northeastern Atlantic. Deep Sea Res Part 2 Top Stud Oceanogr 48: 795–823. Giere O, Eleftheriou A & Murison DJ (1988) Ch. 5: abiotic factors. Introduction to the Study of Meiofauna (Higgins RP & Thiel H, eds), pp. 61–78. Smithsonian Institution, Washington, DC. Gillespie PA, Maxwell PD & Rhodes LL (2000) Microphytobenthic communities of subtidal locations in New Zealand: taxonomy, biomass, production, and foodweb implications. NZ J Mar Freshwat Res 34: 41–53. Glud RN, Kuhl M, Kohls O & Ramsing NB (1999) Heterogeneity of oxygen production and consumption in a photosynthetic microbial mat as studied by planar optodes. J Phycol 35: 270–279. Grenz C, Denis L, Boucher G, Chauvaud L, Clavier J, Fichez R & Pringault O (2003) Spatial variability in sediment oxygen consumption under winter conditions in a lagoonal system in New Caledonia (South Pacific). J Exp Mar Biol Ecol 285: 33–47. Haack TK & McFeters GA (1982) Nutritional relationships among microorganisms in an epilithic biofilm community. Microb Ecol 8: 115–126.

ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

Hancke K & Glud RN (2004) Temperature effects on respiration and photosynthesis in three diatom-dominated benthic communities. Aquat Microb Ecol 37: 265–281. Hansen LS & Blackburn TH (1991) Aerobic and anaerobic mineralization of organic material in marine sediment microcosms. Mar Ecol Prog Ser 75: 283–291. Howarth RW, Marino R & Cole JJ (1988) Nitrogen-fixation in fresh-water, estuarine, and marine ecosystems. 2. Biogeochemical controls. Limnol Oceanogr 33: 688–701. Incera M, Cividanes SP, Lo´pez J & Costas R (2003a) Role of hydrodynamic conditions on quantity and biochemical composition of sediment organic matter in sandy intertidal sediments (NW Atlantic coast, Iberian Peninsula). Hydrobiologia 497: 39–51. Incera M, Cividanes SP, Lastra M & Lopez J (2003b) Temporal and spatial variability of sedimentary organic matter in sandy beaches on the northwest coast of the Iberian Peninsula. Estuar Coast Shelf Sci 58: 55–61. Jaffe R, Mead R, Hernandez ME, Peralba MC & DiGuida OA (2001) Origin and transport of sedimentary organic matter in two subtropical estuaries: a comparative, biomarker-based study. Org Geochem 32: 507–526. Jeffrey SW, Mantoura RFC & Wright SW (1997) Phytoplankton Pigments in Oceanography: Guidelines to Modern Methods. UNESCO, International Council of Scientific Unions, Scientific Committee on Oceanic Research, Paris. Jensen MM, Holmer M & Thamdrup B (2005) Composition and diagenesis of neutral carbohydrates in sediments of the Baltic-North Sea transition. Geochim Cosmochim Acta 69: 4085–4099. Johannes RE, Pearce AF, Wiebe WJ, Crossland CJ, Rimmer DW, Smith DF & Manning C (1994) Nutrient characteristics of well-mixed coastal waters off Perth, Western-Australia. Estuar Coast Shelf Sci 39: 273–285. Kendrick GA, Langtry S, Fitzpatrick J, Griffiths R & Jacoby CA (1998) Benthic microalgae and nutrient dynamics in wavedisturbed environments in Marmion Lagoon, Western Australia, compared with less disturbed mesocosms. J Exp Mar Biol Ecol 228: 83–105. Kendrick GA, Aylward MJ, Hegge BJ, Cambridge ML, Hillman K, Wyllie A & Lord DA (2002) Changes in seagrass coverage in Cockburn Sound, Western Australia between 1967 and 1999. Aquat Bot 73: 75–87. Klump VJ & Martens CS (1989) The seasonality of nutrient regeneration in an organic-rich coastal sediment: kinetic modeling of changing pore-water nutrient and sulfate distributions. Limnol Oceanogr 34: 559–577. Lawrence D, Dagg MJ, Liu HB, Cummings SR, Ortner PB & Kelble C (2004) Wind events and benthic-pelagic coupling in a shallow subtropical bay in Florida. Mar Ecol Prog Ser 266: 1–13. MacIntyre HL & Cullen JJ (1995) Fine-scale vertical resolution of chlorophyll and photosynthetic parameters in shallowwater benthos. Mar Ecol Prog Ser 122: 227–237. MacIntyre HL, Geider RJ & Miller DC (1996) Microphytobenthos: the ecological role of the “secret

FEMS Microbiol Ecol 83 (2013) 279–298

Sandy sediment microbial communities and nutrient exchanges

garden” of unvegetated, shallow-water marine habitats. 1. Distribution, abundance and primary production. Estuaries 19: 186–201. Manini E, Fiordelmondo C, Gambi C, Pusceddu A & Danovaro R (2003) Benthic microbial loop functioning in coastal lagoons: a comparative approach. Oceanol Acta 26: 27–38. Marino R, Chan F, Howarth RW, Pace M & Likens GE (2002) Ecological and biogeochemical interactions constrain planktonic nitrogen fixation in estuaries. Ecosystems 5: 719–725. Masini RJ & McComb AJ (2001) Production by microphytobenthos in the Swan-Canning Estuary. Hydrol Process 15: 2519–2535. Middelburg JJ, Barranguet C, Boschker HTS, Herman PMJ, Moens T & Heip CHR (2000) The fate of intertidal microphytobenthos carbon: an in situ C-13-labeling study. Limnol Oceanogr 45: 1224–1234. Miller-Way T, Boland GS, Rowe GT & Twilley RR (1994) Sedinment oxygen-consumption and benthic nutrient fluxes on the Louisiana continental-shelf – a methodological comparison. Estuaries 17: 809–815. Millie DF, Paerl HW & Hurley JP (1993) Microalgal pigment assessments using high-performance liquid-chromatography – a synopsis of organismal and ecological applications. Can J Fish Aquat Sci 50: 2513–2527. Mindl B, Sonntag B, Pernthaler J, Vrba J, Psenner R & Posch T (2005) Effects of phosphorus loading on interactions of algae and bacteria: reinvestigation of the ‘phytoplankton– bacteria paradox’ in a continuous cultivation system. Aquat Microb Ecol 38: 203–213. Murray RE, Cooksey KE & Priscu JC (1986) Stimulation of bacterial DNA synthesis by algal exudates in attached algalbacterial consortia. Appl Environ Microbiol 52: 1177–1182. Nelson JR, Eckman JE, Robertson CY, Marinelli RL & Jahnke RA (1999) Benthic microalgal biomass and irradiance at the sea floor on the continental shelf of the South Atlantic Bight: spatial and temporal variability and storm effects. Cont Shelf Res 19: 477–505. Nieuwenhuize J, Maas YEM & Middelburg JJ (1994) Rapid analysis of organic-carbon and nitrogen in particulate materials. Mar Chem 45: 217–224. Nozaki K, Darijav K, Akatsuka T, Goto N & Mitamura O (2003) Development of filamentous green algae in the benthic algal community in a littoral sand-beach zone of Lake Biwa. Limnology 4: 161–165. Oviatt CA, Hunt CD, Vargo GA & Kopchynski KW (1981) Simulation of a storm event in marine microcosms. J Mar Res 39: 605–626. Pancost RD & Sinninghe Damste´ JS (2003) Carbon isotopic compositions of prokaryotic lipids as tracers of carbon cycling in diverse settings. Chem Geol 195: 29–58. Pattiaratchi CB, Lavery PS, Wyllie A & Hick PT (1991) Remote Sensing of Water Quality in Cockburn Sound: Development of Multi-Temporal Algorithms. Centre for Water Research, Perth, pp. 48.

FEMS Microbiol Ecol 83 (2013) 279–298

297

Plante-Cuny MR & Bodoy A (1987) Biomass and primary production of phytoplankton and microphytobenthos in a sheltered and an exposed beach (Gulf of Fos, France). Oceanol Acta 10: 223–237. Qu W, Morrison RJ, West RJ & Su C (2005) Diagenetic stoichiometry and benthic nutrient fluxes at the sedimentwater interface of Lake Illawarra, Australia. Hydrobiologia 537: 249–264. R-Development-Core-Team (2008) R: A Language and Environment for Statistical Computing. R-DevelopmentCore-Team, Vienna, Austria. Riaux-Gobin C, Llewellyn CA & Klein B (1987) Microphytobenthos from 2 subtidal sediments from NorthBrittany. 2. Variations of pigment compositions and concentrations determined by Hplc and conventional techniques. Mar Ecol Prog Ser 40: 275–283. Risgaard-Petersen N, Meyer RL & Revsbech NP (2005) Denitrification and anaerobic ammonium oxidation in sediments: effects of microphytobenthos and NO3
. Aquat Microb Ecol 40: 67–76. Rizzo WM (1990) Nutrient exchanges between the water column and a subtidal benthic microalgal community. Estuaries 13: 219–226. Rizzo WM, Dailey SK, Lackey GJ, Christian RR, Berry BE & Wetzel RL (1996) A metabolism-based trophic index for comparing the ecological values of shallow-water sediment habitats. Estuaries 19: 247–256. Rocha C, Galvao H & Barbosa A (2002) Role of transient silicon limitation in the development of cyanobacteria blooms in the Guadiana estuary, south-western Iberia. Mar Ecol Prog Ser 228: 35–45. Round FE (1971) Benthic marine diatoms. Oceanogr Mar Biol Annu Rev 9: 83–139. Saburova MA & Polikarpov IG (2003) Diatom activity within soft sediments: behavioural and physiological processes. Mar Ecol Prog Ser 251: 115–126. Sawyer TE & King GM (1993) Glucose uptake and end product formation in an intertidal marine sediment. Appl Environ Microbiol 59: 120–128. Schofield KA, Pringle CM & Meyer JL (2004) Effects of increased bedload on algal- and detrital-based stream food webs: experimental manipulation of sediment and macroconsumers. Limnol Oceanogr 49: 900–909. Seroˆdio J, da Silva JM & Catarino F (2001) Use of in vivo chlorophyll a fluorescence to quantify short-term variations in the productive biomass of intertidal microphytobenthos. Mar Ecol Prog Ser 218: 45–61. Shimeta J, Amos CL, Beaulieu SE & Ashiru OM (2002) Sequential resuspension of protists by accelerating tidal flow: implications for community structure in the benthic boundary layer. Limnol Oceanogr 47: 1152–1164. Shum KT & Sundby B (1996) Organic matter processing in continental shelf sediments–the subtidal pump revisited. Mar Chem 53: 81–87. Simpson CJ, Burt JS, Cary JL, D’Adamo N, Masini RJ & Mills DA (1996) Southern Metropolitan Coastal Waters Study

ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

H. Forehead et al.

298

(1991–1994) Final Report. Department of Environmental Protection, Western Australia, Perth, Western Australia, pp. 288. Sloth NP, Riemann B, Nielsen LP & Blackburn TH (1996) Resilience of pelagic and benthic microbial communities to sediment resuspension in a coastal ecosystem, Knebel Vig, Denmark. Estuar Coast Shelf Sci 42: 405–415. Staats N, Stal LJ, de Winder B & Mur LR (2000) Oxygenic photosynthesis as driving process in exopolysaccharide production of benthic diatoms. Mar Ecol Prog Ser 193: 261–269. Stahl-Delbanco A & Hansson LA (2002) Effects of bioturbation on recruitment of algal cells from the “seed bank” of lake sediments. Limnol Oceanogr 47: 1836–1843. Sun MY (2000) Mass spectrometric characterization of 13Clabeled lipid tracers and their degradation products in microcosm sediments. Org Geochem 31: 199–209. Sun MY, Aller RC & Lee C (1994) Spatial and temporal distributions of sedimentary chloropigments as indicators of benthic processes in long-island sound. J Mar Res 52: 149–176. Sundba¨ck K & Graneli W (1988) Influence of Microphytobenthos on the nutrient flux between sediment and water – a laboratory study. Mar Ecol Prog Ser 43: 63–69. Sundba¨ck K, Enoksson V, Graneli W & Pettersson K (1991) Influence of sublittoral Microphytobenthos on the oxygen and nutrient flux between sediment and water – a laboratory continuous-flow study. Mar Ecol Prog Ser 74: 263–279. Sundba¨ck K, Miles A & Goransson E (2000) Nitrogen fluxes, denitrification and the role of microphytobenthos in microtidal shallow-water sediments: an annual study. Mar Ecol Prog Ser 200: 59–76. Sundba¨ck K, Linares F, Larson F, Wulff A & Engelsen A (2004) Benthic nitrogen fluxes along a depth gradient in a microtidal fjord: the role of denitrification and microphytobenthos. Limnol Oceanogr 49: 1095–1107. Suzumura M, Miyajima T, Hata H, Umezawa Y, Kayanne H & Koike I (2002) Cycling of phosphorus maintains the production of microphytobenthic communities in carbonate sediments of a coral reef. Limnol Oceanogr 47: 771–781. Valderrama JC (1981) The simultaneous analysis of total nitrogen and total phosphorus in natural waters. Mar Chem 10: 109–122.

ª 2012 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

Viaroli P & Christian RR (2004) Description of trophic status, hyperautotrophy and dystrophy of a coastal lagoon through a potential oxygen production and consumption indexTOSI: Trophic Oxygen Status Index. Ecol Ind 3: 237–250. Villbrandt M, Krumbein WE & Stal LJ (1991) Diurnal and seasonal-variations of nitrogen-fixation and photosynthesis in cyanobacterial mats. Plant Soil 137: 13–16. Volkman JK (2003) Sterols in microorganisms. Appl Microbiol Biotechnol 60: 495–506. Wainright SC (1987) Stimulation of heterotrophic microplankton production by resuspended marinesediments. Science 238: 1710–1712. Watermann F, Hillebrand H, Gerdes G, Krumbein WE & Sommer U (1999) Competition between benthic cyanobacteria and diatoms as influenced by different grain sizes and temperatures. Mar Ecol Prog Ser 187: 77–87. Watson RJ, Butler ECV, Clementson LA & Berry KM (2004) Flow-injection analysis with fluorescence detection for the determination of trace levels of ammonium in seawater. J Environ Monit 7: 37–42. Wilson C & Paling EI (2003) Water Quality of Cockburn and Warnboro Sounds. Institute for Environmental Science, Murdoch University, Perth, Western Australia, pp. 31. Wright SW, Jeffrey SW, Mantoura RFC, Llewellyn CA, Bjornland T, Repeta D & Welschmeyer N (1991) Improved HPLC method for the analysis of chlorophylls and carotenoids from marine phytoplankton. Mar Ecol Prog Ser 77: 183–196.

Supporting Information Additional Supporting Information may be found in the online version of this article: Appendix S1. Sources of fatty acids. Appendix S2. Sources of neutral lipids. Please note: Wiley-Blackwell is not responsible for the content or functionality of any supporting materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.

FEMS Microbiol Ecol 83 (2013) 279–298