Grazing of intertidal benthic foraminifera on bacteria: Assessment using pulse-chase radiotracing
M. Mojtahid ⁎
,1, M.V. Zubkov, M. Hartmann, A.J. Gooday
National Oceanography Centre, Southampton, University of Southampton Waterfront Campus, European Way, Southampton, SO14 3ZH, UK
a b s t r a c t a r t i c l e i n f o
Article history:
Received 24 September 2010
Received in revised form 7 December 2010 Accepted 14 January 2011
Available online xxxx Keywords:
Bacteria
Benthic foraminifera Radiotracing
Although foraminifera are a dominant component of many marine benthic communities, quantification of their predation on prokaryotes remains an experimental challenge. We have developed an approach that allows us to study grazing by adult specimens of the calcareous speciesHaynesina germanicaandAmmonia beccarii, and the single-chambered agglutinated speciesPsammophagasp., on bacteria (Halomonassp.), pulse- chase-labelled with3H- and14C-Leucine. The bacterivorous ciliateUronemasp. andflagellatePteridomonassp.
were used as positive controls. The rate of release of3H when protozoa were incubated with the labelled bacteria indicated the predator's grazing rate; the proportion of14C found in the foraminiferal biomass and shell indicated the prey assimilation rate. All three foraminiferal species grazed bacteria at a rate of 3.2–
5.7 ng C ind−1h−1depending on bacterial concentrations. About 23% of the biomass of the14C-labelled prey was most likely assimilated into foraminiferal pseudopodia, 12% was expelled in dissolved waste material, about 62% was respired and only 0.1% was incorporated into the carbonate shell. Extracellular digestion associated with pseudopodia could explain the very low proportion of the labelled food assimilated in the cell body and the significant proportion located in pseudopodial networks. These experiments also suggest that very little of the carbon ingested by adult calcareous foraminifera is incorporated into the shell. However, we cannot conclude that diet has no influence on the stable isotope composition of the shell since none of our calcareous specimens grew new chambers during the experiments.
© 2011 Elsevier B.V. All rights reserved.
1. Introduction
Benthic foraminifera are heterotrophic protists that are present in a wide range of marine environments, from shallow brackish waters to the deepest ocean depths. They constitute a large part of the biomass of the benthic communities (Snider et al., 1984; Alongi, 1992; Gooday et al., 1992; Moodley et al., 1998a, 2000) and may play a major role in the short to long term consumption of organic carbon in the surface sediments (Altenbach, 1992; Linke et al., 1995; Moodley et al., 2002).
Although foraminifera are a well-studied benthic taxon, little is known about their general ecology or their trophic strategies and preferences.
However, food and oxygen availability are believed to strongly affect foraminiferal growth, reproduction, abundance and diversity as well as their regional distributions (Lutze and Coulbourn, 1984; Altenbach et al., 2003; Gooday, 2003). The reticulate pseudopodia of foraminifera are highly efficient food-gathering organelles (Bernhard and Bowser,
1992; Bowser and Travis, 2002) that give them a central role in decomposing and recycling organic carbon (Mackensen et al., 1993;
Bowser and Travis, 2002). Algae and algal-derived detritus are often consumed by foraminifera and evidence for bacterial consumption is growing (Langer and Gehring, 1993; Goldstein and Corliss, 1994; Ernst et al., 2005). Several littoral benthic foraminifera require bacteria to reproduce (Muller and Lee, 1969) and have been shown to selectively ingest bacteria according to strain (Lee et al., 1966; Lee and Muller, 1973). Some epiphytic foraminifera have adopted a farming strategy, producing nutrient-rich substrata and then ingesting cultured bacteria (Langer and Gehring, 1993). Foraminifera are also able to feed actively on bacterial biofilms (Bernhard and Bowser, 1992), which can form on any solid surface immersed in water (Zubkov and Sleigh, 1999).
Prokaryotes are possibly a main food source for foraminifers in oligotrophic (Gooday et al., 1997) and dysoxic (Schmiedl et al., 2003) settings and on disturbed substrates such as ash falls (Hess and Kuhnt 1996) and turbidites (Hess et al., 2005).
Foraminiferal diets are an important topic in geology as well as biology. It has been proposed that the stable carbon isotope ratios of foraminiferal shells are affected by diet (Mackensen et al., 1993).
According to this idea, the stable carbon isotopic composition depends, among other things, on that of its food sources and on isotopic fractionation during the feeding process. However, this proposal has never been verified experimentally. Growth in
⁎Corresponding author at: Laboratoire des Bio-Indicateurs Actuels et Fossiles, UFR Sciences (Université d'Angers), 2 Bd Lavoisier, 49170 Angers, France. Tel.: + 33 2 41 73 50 02; fax: + 33 2 41 73 53 52.
E-mail address:meryem.mojtahid@univ-angers.fr(M. Mojtahid).
1Present address: Laboratory of Recent and Fossil Bio-Indicators (BIAF), UPRES EA 2644, University of Angers 2, Boulevard Lavoisier, 49045 Angers Cedex, France, and Laboratoire d'Etude des Bio-Indicateurs Marins (LEBIM), Ker Chalon, 85350 Ile D'Yeu, France.
0022-0981/$–see front matter © 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.jembe.2011.01.011
Contents lists available atScienceDirect
Journal of Experimental Marine Biology and Ecology
j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j e m b e
foraminifera is usually accomplished by intermittently adding new chambers.Angell (1967)provided thefirst ultrastructural account of chamber formation in a calcareous perforate foraminifer. First, with the aid of pseudopodia, the protist delineates a space that partially isolates the organism from the environment. It then forms an organic template in the shape of the new chamber and precipitates CaCO3on both sides of the thin organic layer (Goldstein, 1999; Erez, 2003; Erez et al., 2008). Consequently, the uptake of dissolved carbonate from porewater or the water column is energetically much easier (Ziveri et al., 2003). In coccolithophorids, the carbon incorporated into biomass and into the coccoliths originates from two distinct metabolic pathways (e.g.Anning et al., 1996). Whether the same dichotomy exists in foraminifera is an important question that is addressed in this laboratory culturing study.
Most of what we know about the biology of benthic foraminifera originates from studies on shallow-water species that can be easily collected and maintained in culture. Early studies of diets, feeding and nutrition have included both controlled laboratory protocols (e.g.Lee et al., 1966; Delaca, 1982; Bowser et al., 1985; Bernhard and Bowser, 1992) and observations on freshly-collected specimens (Alexander and Delaca, 1987). Recentin situand laboratory-based experiments have explored the effects of food and oxygen on foraminiferal assemblages (Alve and Bernhard, 1995; Moodley et al., 1998b;
Heinz et al., 2002; Geslin et al., 2004; Hess et al., 2005; Nomaki et al., 2005). The food uptake experiments have traced the uptake of algae and bacteria labelled with13C (e.g.Moodley et al., 2000; Nomaki et al., 2005, 2006),32P and14C (Lee et al., 1966) or15N pre-enriched bacteria (Pascal et al., 2008). In addition, direct observations of food vacuoles (Goldstein and Corliss, 1994) and bacteria labelled with fluorescent dyes (Langezaal et al., 2005) have revealed the ingestion of bacteria byAmmoniasp. Nevertheless, except forNomaki et al.
(2006), these studies do not yield quantitative data on the fate of ingested carbon, and the importance of bacteria in foraminiferal nutrition remains poorly constrained. In order to better understand the role that foraminifera play in benthic food webs, we need to have more information about their grazing rate on bacteria.
The central objective of this paper is to present an innovative radiotracer method to determine the metabolism of foraminifera and the fate of biomass that they ingest. This method, which involves depositing dual-radioactive-labelled bacteria (Halomonas sp.) on
filters, has been already used successfully with other protozoa (Zubkov and Sleigh, 1999). Its advantages over previous tracer methods are: (i) the amount of bacteria consumed can be quantified, (ii) assimilation rather than just uptake can be determined, (iii) the amount of food the foraminiferan recycles can be measured, (iv) the respiration can be assessed as well as (v) the contribution of labelled food to shell formation. Using this approach, we address the following two questions. 1) What are the grazing rates of foraminifera on bacteria? 2) How is the ingested biomass partitioned between the foraminiferal cytoplasm, shell, respiration and excretion?
2. Material and methods 2.1. Cultures
Foraminifera were collected from the intertidal zone of the Hamble estuary at Warsash, Hampshire, England (Alve and Murray, 1994, Fig. 1, W5–W8) on the 18th of September and 9th of November 2009.
Fresh-water inflow is small so the estuary has a strong marine influence and is not greatly disturbed by human activity (Alve and Murray, 1994). Surficial sediment was carefully transferred to a plastic tub and covered with water from the same area. In the laboratory, the sediment was stored in a microcosm system (Topping et al., 2006) in which the sediment was surrounded by aerated water to prevent the formation of anoxia. The oxygen, salinity, temperature and light exposure of the seawater in the microcosm tank (Hamble estuary, salinity 34) were monitored weekly. The oxygen was kept constant by the aeration of the seawater using an air gas diffuser. A lid to slow evaporation and the frequent addition of deionised water ensured that salinity remained constant. The tank was maintained at a temperature of 9–19 °C, depending on the time of the day and on the season, simulating the natural changes in the Hamble estuary (Alve and Murray, 2001). The tank was illuminated for 12 h during each 24 h period. Foraminifera for experiments were picked out under a dissecting stereomicroscope (Stemi SV11, Zeiss), using a Pasteur pipette, from sieved (125μm) fractions of stock cultures, washed with sterile artificial seawater in 3–5 serial baths, and starved for a week before the beginning of the grazing experiments. For each experiment, care was taken to select mature individuals of the same size (300– 400μm). Those from a single stock culture served as the sole source
Fig. 1.Images showing live intertidal species with orange/brown coloration of their endoplasm and pseudopodial activity.
for a single experiment. The calcareous speciesHaynesina germanica, Ammonia beccarii, and the monothalamous speciesPsammophagasp.
(Larkin and Gooday, 2004), were selected. Only live foraminifera with orange/brown coloration and pseudopodial activity (seeFig. 1) were used. The condition of the foraminifera was checked daily during and at the end of the experiments. We ensured that they were alive by observing pseudopodia and the fact that each day they had moved to different positions on thefilters.
As a positive control for the grazing experiments, we used the scuticociliateUronemasp., isolated from the Solent and theflagellate Pteridomonas sp., isolated from Southampton Water. These two protozoan species were grown in the dark at a temperature of 10 °C.
For the tracing experiments, we used cultures that were between one and four weeks old.
2.2. Grazing experiments
The principal prey used to feed all protozoans wasHalomonassp.
The bacteria were maintained on marine agar plates (Difco) and harvested at the stationary phase. They were then suspended in freshlyfiltered (0.2μm cellulose acetate membranefilter, Minisart) artificial seawater (ASW) and the suspension subsequentlyfiltered through a 0.8μm polycarbonate filter to retain or break clumps, yielding a bacterial concentration of about ~ 2·108Halomonascells/ml in thefiltrate. ASW was prepared using the protocol described in Wilson et al. (1996). A preliminary estimate of the bacterial concentration was made from the absorbance at 280 nm using a spectrophotometer (Nanodrop) and an empirical calibration of
absorbance against the concentration of these suspended bacteria measured more precisely usingflow cytometry (Zubkov and Sleigh, 2005). To do this, a subsample of the initial bacterial suspension, was diluted 1:10 withfiltered artificial seawater,fixed with 1% parafor- maldehyde (PFA) final concentration, stained with SYBR Green (Sigma, Poole, UK) in presence of tripotassium citrate. Stained bacterial cells were enumerated by flow cytometry (FACSCalibur, BD Biosciences, Oxford, UK) using 0.5μm multifluorescent beads (Polysciences, Germany) as an internal standard.
2.3. Pulse-chase labelling of bacterial prey
The initial stock of suspendedHalomonascells was diluted with 0.2μm-filtered ASW to reach a final bacterial concentration of
~2 · 107cells/ml. Twenty millilitre of that bacterial suspension was simultaneously spiked with3H-leucine (37 MBq/ml; 7.03 · 10−6mol/l) to a final concentration of 0.38 nmol/l, and with 14C-leucine (1.85 MBq/ml; 1.56 · 10−4mol/l) to afinal concentration of 94 nmol/l.
The suspension was incubated for 30 min (Fig. 2a). The incubation time was monitored by taking subsamples of 200μlfixed with 10μl of 20%
PFA stock solution, 2, 5, 10, 15 and 30 min after the spiking andfiltered onto 0.2μm polycarbonatefilters. The specific activities of the3H- and
14C-labelled leucine were then reduced 750-fold compared to the original activities by addition of non-labelled leucine stock solution (3.2 · 10−3mol/l) (Fig. 2a). After a further 1-h incubation to saturate intracellular pools, label levels stabilised in these dual-radioactively labelledHalomonascells (Zubkov and Sleigh, 1995, 1999). This phase was monitored as described earlier by taking subsamples of 200μlfixed
Fig. 2.Schematic of the experimental set-up; seeMaterial and methodsfor details.
with 10μl of 20% PFA stock solution at 15 min intervals andfiltered onto 0.2μm polycarbonate filters. Previous experiments showed that live bacteria deposited onfilters metabolized the macromolecules with which they had been pulse-chase labelled. The rate of metabolism was sufficient to obscure the results of grazing by slowly growing protozoa (Zubkov and Sleigh, 1995, 1999). We therefore killed the labelled bacterial cells for experimental use with 6–10 bursts of 10 s each in a microwave oven (Zubkov and Sleigh, 1999) (Fig. 2a). Microwave irradiation was chosen because of its rapid action on bacterial cells and its maximal preservation of the label in the cells. Because microwave radiation can damage bacterial cells, this phase was also monitored usingflow cytometry. This is reflected in the cell size obtained by the side scatter of the laser (broken pieces are smaller than the actual cell size). Hence, by comparingflow cytometry data of bacterial cells before and after the treatment we can establish if any detrimental morpho- logical changes happened to the bacteria. The good retention of the label in the cell, especially in thefirst days of the experiment, indicated that lysis of the bacteria was not a major problem. Allfilters were washed with 5 ml of 0.2μm-filtered ASW, placed in scintillation vials,filled with 6 ml of scintillation cocktail, left for solubilisation of the biomass for at least 24 h, and the amounts of the two radioisotopes assayed simultaneously using a liquid scintillation counter (1220 Quantulus, Wallac, Finland). Two and four millilitres of the killed (microwaved) labelled bacterial suspension, representing ~43 · 106 and ~86 · 106 bacterial cells respectively, were filtered onto transparent, 0.4μm, collagen-coated membranefilters (4.2 cm2area) moulded in inserts (Corning) designed tofit into 6-well cell culturing plates, and washed with 5 ml of 0.2μm-filtered ASW. The inserts with deposited bacteria were placed in the wells of sterile plates containing 2 ml offiltered ASW, according to the manufacturer's instructions. An additional 3 ml were poured carefully down the inside wall of the inserts (Fig. 2a).
Ten specimens each ofHaynesina germanica,Ammonia beccarii, and Psammophagasp., and 20–200μl of a culture of protozoa containing a few tens of ciliate cells (Uronemasp.) and few hundreds offlagellate cells (Pteridomonassp.), were used for the experiments. The different species were inoculated in separate inserts. The inserts with deposited bacteria but without added protozoa were used as controls (Fig. 2b).
The numbers of ciliates andflagellates inoculated at the beginning of the experiments were measured more precisely usingflow cytometry (see previous discussion). The grazing of all protozoa was monitored by taking subsamples of 50μl of ASW from outside the insert every day for 15–20 days. The subsamples were mixed with 6 ml of scintillation cocktail and radioassayed (Fig. 2b). We found previously that protozoa grazing on deposited bacteria rapidly released most of the3H label upon ingestion of the labelled bacteria,14C being mostly respired (Zubkov and Sleigh, 1995, 1999). Therefore, the dissolved digested fraction in the sampled seawater corresponds to the difference between the amount of3H and14C labels in the dissolved fraction in the control series without protozoa and in the experimen- tal series with protozoa, as a proportion of the amount of3H and14C labels in the former. Since most of14C is respired by the protozoa and very little remained in the seawater, the3H-dissolved fraction gives an indirect estimation of the respiration rate. We assumed that the protozoa had consumed 100% of the deposited labelled bacteria when the amounts of labels in the dissolved fractions stabilised with time.
At the end of the experiment, foraminifera were picked out individually from the inserts, before the inserts were removed from the wells. The water within each insert wasfiltered through its insert filter, and 5 ml of 0.2-μm-filtered ASW was passed through to wash both insert andfilter. Eachfilter was then cut out and placed in a scintillation vial,filled with 6 ml of scintillation cocktail and radio- assayed (Fig. 2c). Assuming that all the bacteria were eaten at the end of each experiment, the remaining radioactivity on the filters originated from the particulate digested fraction and the foraminiferal pseudopodial cytoplasm retained on thefilters, or assimilated in the case of ciliates andflagellates.
To measure radioactivity separately in the shell and in the biomass of the calcareous species, we used the following technique (Fig. 2c). In a 20 ml gas tight glass vial, we placed two containers, one a 2 ml plastic Eppendorf vial containing the calcareous foraminifera and the other a 2 ml glass vial containing a GF/Bfilter wetted with 200μl of NaOH (5 M). Once the 20 ml gas-tight vial was sealed, we injected into the vial containing foraminifera 500μl of HCL (10%) using a syringefixed on a 0.8 mm-needle. After removing the syringe and the needle, the vials were left to equilibrate for 24 h. In thefirst container, the HCL dissolved the foraminiferal shell carbonate, leaving the biomass (cell). The CO2produced from this reaction was trapped by the alkalinefilter. Then 1.8 ml of scintillation cocktail was added to each 2 ml container. The 2 ml glass vial was sealed with a gas-tight cap. The 2 ml vials were placed inside scintillation vials and the amounts of the two radioisotopes assayed (Fig. 2c). This technique was verified and confirmed with several experiments using 14C- bicarbonates (simulating foraminiferal shells) with a known radioac- tivity and applying the protocol described earlier for foraminifera. In each experiment, the14CO2produced from this reaction was entirely trapped by the alkalinefilter.
2.4. Biomass determination
Because the biomass of foraminifera could not be accurately estimated from biovolumes (e.g. Hannah et al., 1994), the protein content of the foraminiferal cells was measured by the bicinchoninic acid (BCA, Sigma, Poole, UK) method (Smith et al., 1985), using bovine serum albumin as a standard. This yielded estimates of the average protein content per foraminiferal cell. The mean protein content of bacteria (Halomonas) used in grazing experiments is about 0.2 pg cell−1 (Zubkov and Sleigh, 1995). Since the carbon and protein content have a consistent 1:1 relationship with each another in unicellular organisms (Smith et al., 1985; Zubkov and Sleigh, 1995), we used the protein content to monitor the carbon content and hence the biomass of foraminifera. We measured the size of foraminifera under the stereomicroscope before protein determination. Following Pelegri et al. (1999), the biomass of Pteridomonas sp. and Uronema sp. is about 5.2 pg C cell−1and 121 pg C cell−1respectively.
3. Results
3.1. Labelling of bacteria
The bacteria started incorporating the labelled leucine immedi- ately after the addition of the spike. There was a linear increase in the amount of3H and14C incorporated throughout the 30 min of active labelling (Fig. 3). The uptake of labels was ~ 18% for3H and ~30% for
14C. When the specific activities of the radioactive precursors were sharply reduced with unlabeled analogues, the uptake rate levelled off to a more or less stable plateau, corresponding to the amount of the labels incorporated into macromolecules. After microwaving labelled Halomonas, the incorporated labels remained in bacterial biomass, showing that this method resulted in adequate, predictable and reproducible retention of both labels within bacterial cells, especially at the beginning of the grazing experiments (Figs. 3 and 4e, j).
3.2. The release of radioactive labels by grazing protozoa
The dynamics of the3H and14C labels in the presence of protozoa were corrected for bacterial release of both labels in the control series (without protozoan grazers). The resulting dynamics clearly differ from those in the controls (Fig. 4). Generally, we observed a ~50–75%
release of3H label into the water for up to thirteen days, with a lag phase for ciliates andflagellates before the start of the release. Later, the radioactivity in the water either remained at the same high level or somewhat decreased. We assumed that when the release into the
water of3H label stabilised, the protozoa had ingested all the available labelled bacteria deposited on thefilters. Therefore, the dynamics of
3H label release indicates protozoan grazing rates. In contrast, the concentration of the14C label in the water never exceeded 20% of the total initial radioactivity of deposited bacteria. In other respects, the dynamics of14C generally resembled that of the dissolved3H label (Fig. 4). The ciliate Uronema sp. reached its maximum3H-release earlier than theflagellatePteridomonassp. and the foraminifera. The lag phase was also shorter forUronemasp. than forPteridomonassp.
(Fig. 4).
3.3. Grazing rates of protozoa and growth in biomass
Two initial concentrations of bacteria (~43 · 106 and 86 · 106 bacterial cells) deposited on the filters were used in the grazing experiments. The release of3H into the water represents the grazing of protozoa. Therefore, the maximum amount released by each species is equivalent to 100% of the bacterial cells ingested. Thus, the numbers of ingested bacteria is equivalent to the percentages of3H released by each foraminiferal species (Fig. 5). Then, we used a linear polynomial model to simulate 3H label release for each species excluding thefinal stabilisation and decrease of the concentration of dissolved 3H label (Fig. 5). This model is adequate (r2N0.98) to describe the observed dynamics. Ten specimens of H. germanica ingested ~43 · 106bacterial cells in 12 days and ~ 86 · 106bacterial cells in 13 days. Ten specimens of A. beccarii ingested ~43 · 106 bacterial cells in 11 days and ~86 · 106 bacterial cells in 13 days.
Finally, ten specimens ofPsammophagasp. ingested ~43 · 106bacterial cells in 11 days (Fig. 5). The quantity of bacteria on thefilters affected the3H release by protozoa. The average grazing rates for low bacterial concentration are ~14,900 bacterial cells ind−1h−1forH. germanica and ~16,300 cells ind−1h−1 for A. beccarii and Psammophaga sp.
When grazing higher bacterial concentrations, the grazing rates are higher: ~ 27,600 bacterial cells ind−1h−1for bothH. germanicaand A. beccarii. Converting these values into bacterial biomass (~0.2 pg per Halomonascell) gives an average grazing rate for one foraminiferal specimen of ~3200 pg C ind−1 h−1 at low bacterial concentration (~43 · 106cells) and ~5700 pg C ind−1h−1 at higher bacterial con- centration (~83 · 106cells) (Fig. 6a). The ingestion of bacteria by adult foraminifera was not followed by reproduction or an increase in biomass, as shown by their stable protein content measured during the grazing experiments (Fig. 6b).
Ciliates andflagellates grazed more quickly, and released more3H label in the water, when higher bacterial concentrations were used.
When grazing on lower bacterial densities (~ 43 · 106cells), the number ofUronema cells increased by ~ 500 times in 8 days (from
~20 to ~ 10,100 cells; seeFig. 6c) compared to ~ 620 times (from ~20 to 12,500 cells; seeFig. 6c) in 7 days when grazing higher bacterial concentration (~83 · 106cells). At low bacterial concentration, the number ofPteridomonascells increased by ~800 times (from ~100 to 79,500 cells; seeFig. 6c) in 10 days and by ~ 1000 times in 7 days (from ~100 to 100,000 cells; seeFig. 6c) at high bacterial concentra- tion. The average grazing rate of Uronema sp. was ~22 bacteria cells ind−1h−1(~ 4.5 pg C ind−1h−1) at low bacterial concentrations and ~ 41 bacteria cells ind−1h−1 (~ 8.4 pg C ind−1h−1) at high concentrations (Fig. 6a). The average grazing rate of Pteridomonas sp. was ~ 2 bacteria cells ind−1h−1 (~0.4 pg C ind−1h−1) when feeding at low concentrations (Fig. 6a) and ~5 bacteria cells ind−1h−1 (~1 pg C ind−1h−1) at high concentrations (Fig. 6a).
3.4. The assimilation of the radioactive labels in foraminiferal biomass and shell
A very small proportion of the label was present in the cytoplasm of all species (~0.14% of3H and ~ 0.31% of14C;Table 1). The proportion of14C in the shell of the two calcareous species (~ 0.07% of14C) was even lower, (Table 1).
For the three species of foraminifera, approximately 25% of the3H and14C labels remained on thefilter. We presume that these were located in the pseudopodial networks that we systematically observed to be left behind on thefilters when the tests were removed, and in particulate waste. For ciliates, the corresponding average proportions were 28% (3H) and 29% (14C) and forflagellates 25% (3H) and 21% (14C). In these treatments, the labels were located in protozoan biomass and particulate waste. The proportion of these labels was slightly higher when the protozoans fed on higher concentrations of bacteria, especially in the case of ciliates and flagellates, which grew in biomass (Table 1).
4. Discussion
Our experiments demonstrate thatHaynesina germanica,Ammonia beccariiandPsammophagasp. grazed avidly on bacteria. The presence of white areas (concentrated bacteria) around the foraminiferal tests, a few hours after they had been deposited on thefilters, provided Fig. 3.Incorporation of3H-leucine and14C-leucine into the macromolecules of bacteria, suspended in 20 ml of artificial seawater. Thirty minutes after the addition of labelled leucine, non-radioactive leucine was added, reducing specific radioactivity approximately 750 times. Two hours later, bacteria were killed by microwave irradiation.
direct visual evidence for bacterial consumption.Haynesina germanica andA. beccariiare the dominant species in UK estuaries (Alve and Murray, 2001; Murray and Alve, 2000);H. germanicais considered to
be infaunal (Murray, 1991), whileA. becariican be infaunal (review in Murray, 1991) or live at, or close to, the sediment-water interface (Debenay et al., 1998).Ammonia beccariiis believed to be a deposit Fig. 4.3H and14C label release into the water in various incubation experiments, expressed as a percentage of the amount initially present in the stock of microwave-treated bacteria deposited on insertfilters. In control experiments (e,j) bacteria were incubated without protozoan predators; in other experiments the bacteria were incubated with foraminifera (a,b,f,g), the ciliateUronema(c,h) or theflagellatePteridomonas(d,i). The continuous lines correspond to a bacterial concentration of 43 · 106cells and the dotted lines correspond to a bacterial concentration of 86 · 106cells. The values in a–d and f–i are corrected for the release of labels by bacteria in the control experiments.
feeder, mode of nutrition that involves the ingestion of numerous bacteria (Goldstein and Corliss, 1994). Previous studies of H. germanica (Alexander and Banner, 1984; Banner and Culver, 1978; Austin et al., 2005) suggest that it is an herbivore that uses its test ornamentation to break open food items, especially diatoms.
Psammophaga sp. is a soft-shelled monothalamous species (an
‘allogromiid’) that is common in the Warsash area (Larkin and Gooday, 2004). Nothing is known about its feeding biology. However, various lines of evidence suggest that, in general, allogromiids often feed on bacteria (Gooday, 2002).
4.1. Comparison with previous tracer studies
The indispensible role that bacteria play in foraminiferal nutrition (Muller and Lee, 1969; Lee, 1980) is consistent with the probable widespread occurrence of deposit feeding in these heterotrophic protists (Nomaki et al., 2006). The advantages of using radioisotopes as tracers over stable isotopes to assess the metabolism of these species arise chiefly from three properties: (1) analysis is rapid and little sample preparation is usually necessary, (2) radioisotopes are detectable in extremely minute concentrations, and (3) analysis for radioisotopic content often can be achieved without alteration of the sample. To our knowledge there have been no published radiotracer studies to monitor food uptake by foraminifera sinceLee et al. (1966) showed that different species of littoral foraminifera selected32P and
14C labelled food (diatoms, chlorophytes and bacteria). Recent techniques used to assess food uptake by foraminifera have involved in situ feeding experiments using13C-labelled unicellular algae and bacteria in order to understand the fate of lipid compounds in
phytodetritus at the deep-seafloor (Nomaki et al., 2005, 2006), the quantification of bacterivory by benthic organisms using15N-enriched bacteria (Pascal et al., 2008), and cell-permanentfluorescent staining of food for foraminifera (Langezaal et al., 2005). These techniques allow the estimation of food uptake by foraminifera, but not the partitioning of the ingested carbon into the shell, biomass, excretion and respiration. Of course, the fate of ingested carbon can be traced using13C labelling, but the amount of label required is much greater than in the case of the14C approach adopted in the present study.
4.2. Experimental procedure and limitations
Like all approaches to quantifying bacteriovory, the method used in the present study involves methodological shortcomings that complicate interpretation of the results. In the grazing experiments, about 10% of the original3H label, taking into account the amount the
3H in the dissolved fraction, was not recovered. This could reflect the Fig. 5.Averaged number of eaten bacteria estimated from the equivalent percentages of
3H released for each foraminiferal species excluding thefinal stabilisation of dissolved
3H label and modelled by polynomial approximation in the case of a) 43 · 106bacterial cells and b) 86 · 106bacterial cells.
Fig. 6.a) Grazing rates of protozoa; the undashed columns represent the lower bacterial concentrations (43 · 106cells) and the dashed columns represent the higher bacterial concentrations (86 · 106cells). b) The protein content measured in foraminiferal cells using BCA analyses. c) Graph showing the increase inUronemaandPteridomonascells during the grazing experiments.
retention of some bacteria in the wall of the inserts. At the same time, about 63% of14C-labeled macromolecules were completely metabo- lised and the major part of14C label converted to14CO2, allowing the respiration to be estimated (Table 1). The water-air equilibration of
14CO2would result in its release to the atmosphere, and this could explain the irreversible loss of14C label from the dissolved fraction. In contrast,3H-labelled metabolic products, mainly3H2O, accumulated in the water. Although particular care was taken to clean the foraminifera, we believe that other bacteria were introduced with them and could have taken part in the decomposition of the dead labelled bacteria. In order to control for this problem, we introduced foraminifera from the same cultures, cleaned them in the same manner, placed them on thefilters with the labelled bacteria and removed them after 1 h. These treatments generally showed a similar pattern of label release as the controls without protozoa. However, a minority (20–25%) exhibited anomalous peaks in the release of3H into the water, suggesting that some live bacteria had been introduced, although the level and consistency of release was never comparable to that observed in the experiments with foraminifera.
Over-estimation of the grazing rate could also arise because some food remained unconsumed when the release of the3H stabilised in the water. Also, in the experiments where bacteria were grazed, the final gradual decrease of the radioactivity in the water could reflect the reincorporation by introduced bacteria of some small labelled molecules released by the protozoa (Zubkov and Sleigh, 1999).
Finally, our experimental procedure cannot differentiate between the particulate (protein) fraction and the fraction in the pseudopodial network of foraminifera left on thefilters. To do this would involve the labelling of DNA using thymidine in addition to the labelling of proteins using leucine.
4.3. Grazing rates: comparison with other studies
The estimated grazing rates of the three intertidal species were comparable; ~3.2 ng C ind−1h−1(~16,000 bacterial cells ind−1h−1) for low bacterial concentration (~43·106cells) and ~5.7 ng C ind−1h−1 (~28,000 bacterial cells ind−1h−1) for higher bacterial concentration (~86· 106cells). According to Holling's prey-dependent type II func- tional response (Holling, 1959), food uptake by a protozoan grazer should increase with the abundance of food.Langezaal et al. (2005) found thatAmmonia beccariigrazed 90 bacteria during a 20 h period.
Converting this uptake rate into bacterial biomass (Norland et al., 1995) gives a grazing rate of 1.7 pg C ind−1h−1.Pascal et al. (2008)reported a rate of 78 pg C ind−1h−1for the same species. These rates are much lower than those found in the present study. This may be because they were based on suspended, rather than attached, bacteria. The very low uptake of bacteria in the study ofLangezaal et al. (2005)is probably partially the result of the low bacterial concentration in their microcosms (1.4· 103cells ml−1), substantially lower than benthic
bacterial abundance in natural environments (c.a. 109cells ml−1), and the use of dead DAPI stained bacteria that could have caused the death of foraminifera. The low grazing rate of foraminifera on bacteria in the study ofPascal et al. (2008)may reflect the presence of algae that depressed the ingestion of bacteria. Many studies suggest that foraminifera have a preference for algal food but some species will still ingest bacteria, even when other food resources are available (e.g.
Lee et al., 1966; Muller, 1975; van Oevelen et al., 2006; Pascal et al., 2008). In particular,Nomaki et al. (2006) showed a random (non- selective) uptake of bacteria, together with sediment particles and detritus, in species such as Chilostomella ovoidea and Cyclammina cancellata.
4.4. Partitioning of14C and3H in foraminifera
Foraminiferal pseudopodia form an extensive network that is used to trap food particles. Despite the importance of pseudopodial activity in foraminiferal feeding (see Lipps, 1983 for review), and the importance of bacteria in their nutrition (Muller and Lee, 1969; Lee, 1979), the mechanism by which foraminifera acquire bacteria has received little attention.Anderson and Lee (1991)showed that the primary lysosomes in foraminifera are dispersed throughout the cytoplasm of outer chambers and pseudopodia and provide a reserve of digestive enzymes to be utilized when food is engulfed. Small prey, including phytoplankton, zooplankton and bacteria, are caught in the peripheral pseudopodia and engulfed within food vacuoles by invagination of the surface of pseudopodial membrane, a process termed circumvallation. The food vacuoles fuse with acidosomes to form digestive vacuoles. In some benthic species there is evidence that much, if not all, of the digestion occurs in the extra-shell cytoplasm, especially for small prey such as bacteria (Anderson and Lee, 1991).
Meyer-Reil and Köster (1991)attributed high levels of hydrolytic enzymatic activity in marine surface sediments to the extracellular digestion of organic material by benthic foraminifera.Lee et al. (1991) suggest that pseudopodial digestion is widespread among benthic foraminifera.
In our study, extracellular digestion associated with pseudopodia could explain the very low proportion of the labelled food assimilated in the cell body and the significant proportion found on thefilter, which probably retained much of the pseudopodial network after removal of the tests. This idea is also corroborated by the unchanged protein content of the cell body during the grazing experiments, which implies that the food was not converted into biomass. The adult specimens (~360μm) that we used are more likely to be using food to produce free energy to maintain the integrity of the cell, for motility, and for various processes that enhance the survival of the cell, rather than converting it into biomass (Fenchel, 1987). Similarly, since foraminifera increase in biomass when they build new chambers, the even lower proportion of14C (0.1%) incorporated into the shells of the Table 1
Summary of the ingested food and the proportion of both3H and14C labels measured in different compartments (dissolved and particulate waste, cell body, shell, pseudopodia) and the estimated respiration (the remaining fraction from 100% of total carbon budget). The underlined values represent the lower bacterial concentration (43 · 106cells) and italic values represent the higher bacterial concentrations (86 · 106cells). DOW: dissolved organic waste; Ps: pseudopodia.
Dissolved waste Foraminiferal cell body Foraminiferal shell Filters Respiration
(estimated)
Budget3H
Species 3H2O +
3H-DOW
14C-DOW 3H 14C 3H 14C 3H biomass
(Ps.) +3H PW
14C biomass (Ps.) +14C PW
14CO2
H. germanica 70.0 59.6 7.7 11.3 0.14 0.11 0.26 0.32 -0.07 -0.02 0.05 0.1 20.2 21.9 22.4 23.7 69.59 64.58 90.27 81.59 A. beccarii 58.3 62.4 13.5 11.5 0.15 0.14 0.28 0.32 -0.05 0.02 0.05 0.09 25.6 28.3 25.4 27 60.77 61.09 84 90.86
Psammofagasp. 60.8 7.3 0.15 0.39 27 23.5 68.81 87.95
Uronemasp. 53.4 68.4 8.9 2.9 27.7 29.11 23.5 34.8 67.6 62.3 81.1 97.51
Pteridomonassp. 65.5 69.7 10.9 6.6 20.6 28.7 18 23.6 71.1 69.8 86.1 98.4
No grazers 10.1 11.7 3.7 3.2 68.2 78.7 49.1 45.6 47.2 51.2 78.3 90.4
two calcareous foraminifera could reflect the use of adult individuals rather than actively growing juveniles. Alternatively, it is possible that the carbon incorporated into the shell originates from two distinct metabolic pathways so that diet has no effect on the formation, and hence isotopic signature, of the foraminiferal test.
4.5. Flagellates and ciliates
We used theflagellatePteridomonassp. and the ciliateUronemasp. as positive controls because they are exclusively bacterivorous heterotro- phic protozoa. Both species create substantial water currents from which bacteria are intercepted by the tentacular arms ofPteridomonas (Fenchel, 1982) or by cilia located on the paroral membrane ofUronema (similar to Cyclidium, Fenchel, 1980). These two protozoans have different feeding adaptations;Pteridomonasfeeds primarily on bacteria suspended in the water andUronemafeeds on bacteria attached to surfaces (Zubkov and Sleigh, 2000). This could explain the delayed growth ofPteridomonascompared toUronemain our experiments and the plasticity of their feeding behaviour (Zubkov and Sleigh, 1999). Our experiments yielded growth and grazing rate comparable to those found in previous studies byZubkov and Sleigh (1995, 1999, 2000). In the present study, the grazing rates of single foraminiferal cells are much higher than singleflagellate and ciliate cells. However, one foraminiferal cell (~0.93μg C cell−1) is about 7750 times larger than one cell of Uronema(~120 pg C cell−1) and about 179,000 times larger than one cell ofPteridomonas(~5.2 pg C cell−1). Normalising to a standard cell size yields average grazing rates of 4030 pg C h−1for the three studied species of foraminifera, ~49,980 pg C h−1forUronemasp. and ~125,300 pg C h−1 forPteridomonassp.
5. Conclusion
The radioisotope labelling technique described here is a promising tool to study foraminifera–prey interactions and to understand the trophic pathways in foraminifera and the fate of ingested carbon. Under laboratory conditions, the intertidal calcareous species Haynesina germanica, Ammonia beccarii, and the single-chambered species Psammophagasp., consume and digest large quantities of bacteria. The very low proportion of14C incorporated into the cytoplasm within the shell, and the higher proportion located in thefilters and assumed to be located in detached pseudopodia, strongly suggests that the digestion of organic material occurred outside the shell and within the pseudopodia.
The very low proportion of 14C derived from the food that is incorporated into the shell provides no support for the existence of a distinct metabolic pathway for calcification. If dietary carbon is incorporated into foraminiferal carbonate, this is most likely to happen in actively growing, juvenile shells that are adding new chambers. There was no evidence of chamber formation in any of the calcareous specimens used in our experiments.
Acknowledgements
We thank Ross Holland for his help in flow cytometry and Professor Ralf Schiebel for his help in writing the proposal leading to this study. This work wasfinancially supported byMarie-Curie Action:
Intra-European Fellowships (IEF), by a research grant (NE/E016138/
1) of the Natural Environment Research Council (NERC), UK, and by the National Oceanography Centre Core Programme.[RH]
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