L
Journal of Experimental Marine Biology and Ecology 245 (2000) 127–147
www.elsevier.nl / locate / jembe
Bacteria–flagellate coupling in microcosm experiments in the
Central Atlantic Ocean
a ,* b b
¨ ´
Klaus Jurgens , Josep M. Gasol , Dolors Vaque a
¨ ¨
Max-Planck-Institut f ur Limnologie, P.O. Box 165, D-24302 Plon, Germany
b
` ´
Institut de Ciencies del Mar, Pg. Joan de Borbo s /n, E-08039 Barcelona, Spain Received 18 May 1999; received in revised form 23 July 1999; accepted 20 October 1999
Abstract
The coupling between planktonic bacteria and bacterivorous protozoans was examined in microcosm experiments at several oligotrophic and ultra-oligotrophic sites in the subtropical and
5
tropical Atlantic Ocean. Bacterial concentrations at these stations were in the range 2.2–8.1310
21 21
cells ml , heterotrophic nanoflagellates (HNF) in the range 100–800 cells ml , bacterial doubling times (estimated from leucine incorporation) in the range 1–100 days, and chlorophyll a
21
levels in the range 0.03–0.36 mg l . The experimental uncoupling of the microbial loop by differential filtrations did not result in an increased growth and grazing by nanoflagellates despite a stimulation and increase of bacterial abundance and mean cell volume due to the bottle incubations. A strong response of the grazer population occurred after increasing bacterial numbers about 10-fold by the addition of a complex substrate source (yeast extract). Bacteria responded immediately to the substrate enrichment with an increase in mean cell size and abundance, and reached stationary phase already after about 24 h. In contrast, HNF development showed a pronounced lag phase, and it needed between 3 and 7 days until grazers reduced bacterial numbers to about the initial values. The grazing impact on the bacterial assemblage in the bottles resulted in feed-back effects that resembled those known from other, more productive systems: protozoan size-selective grazing removed preferentially larger sized bacteria and shifted the size-distribution towards the initial, natural situation with a dominance of small cocci. Grazing-resistant morphotypes consisted of bacterial aggregates embedded in a polysaccharide matrix whereas filamentous forms did not develop. These experiments provide evidence that bacterial assemblages have the capacity to respond to enhanced substrate availability (for example in micropatches) and to utilise these substrates without significant grazer control. 2000 Elsevier Science B.V. All rights reserved.
Keywords: Aggregates; Atlantic Ocean; Bacteria; Grazing; Nanoflagellates
*Corresponding author. Tel.: 149-4522-763-244; fax: 149-4522-763-310. ¨
E-mail address: [email protected] (K. Jurgens)
1. Introduction
About 30% of the surface of the Earth is covered by oligotrophic oceanic waters in which prokaryotes are the dominant primary and secondary producers of organic matter (Whitman et al., 1998). Determining the controlling factors of planktonic bacteria and their ecological interactions within the microbial food web is essential for an understand-ing of the biogeochemical oceanic fluxes. Most studies examinunderstand-ing these issues are from coastal and estuarine waters and comparatively few data exist for the open ocean. However, some consistent features of the most oligotrophic systems emerged from studies of recent years: Heterotrophic bacteria constitute the major carbon pool in the euphotic zone and this biomass can be several fold greater than phytoplankton biomass (Cho and Azam, 1988; Fuhrman et al., 1989), leading to a different biomass food web structure in oligotrophic compared to eutrophic systems (Dortch and Packard, 1989; Gasol et al., 1997). The implications are also that bacterioplankton contribute sig-nificantly to the particulate pools of inorganic nutrients (N, P) and affect light scattering and absorption phenomena in the open ocean (Morel and Ahn, 1990). In most of the oligotrophic subtropical and tropical oceanic areas also the primary producers are dominated by prokaryotic autotrophic picoplankton belonging to the taxa
Prochlor-ococcus (Chisholm et al., 1988; Olson et al., 1990) and SynechProchlor-ococcus (Waterbury et
al., 1979; Burkill et al., 1993). Grazing by heterotrophic protists is assumed to be the main loss factor of both heterotrophic and autotrophic picoplankton (Wikner and
¨
Hagstrom, 1988; Landry et al., 1995; Reckermann and Veldhuis, 1997).
Whereas solid data exist meanwhile for the biomass distribution, much less is known about the population dynamics of the respective organism groups and how trophic coupling between dissolved organic matter (DOM), phytoplankton, bacteria, viruses and protozoa controls the carbon flux and biomass production of bacteria (Azam et al., 1994). It is important to understand these ecological interactions as they form the bases of the biogeochemical fluxes in the ocean. One research goal is to reveal the importance of inorganic and organic substrates (bottom-up control) or predation (top-down control) as limiting factors for bacterial biomass (e.g. Wright and Coffin, 1984; Ducklow et al., 1992; Shiah and Ducklow, 1995). Strong evidence for bottom-up control of hetero-trophic bacteria has accumulated from open ocean studies. One line of evidence comes from the fact that the addition of DOM (often in combination with inorganic nutrients) usually results in a stimulation of bacterial productivity in short-term microcosm experiments (Kirchman, 1990, Kirchman and Rich, 1997). Another line of evidence is the significant positive correlation between phytoplankton and bacterial biomass (Cole et al., 1988; Ducklow and Carlson, 1992; Dufour and Torreton, 1996), and between bacterial production and bacterial biomass (Ducklow, 1992). Both correlations point towards resource limitation as a mechanism of bacterial biomass regulation and imply that mean levels of bacterial biomass reflect the substrate supply or richness of the system (Billen et al., 1990).
main bacterivorous grazers (Fenchel, 1982b; Fuhrman and McManus, 1984; Rassoul-¨
zadegan and Sheldon, 1986; Wikner and Hagstrom, 1988). The relevance of predation control of bacteria in oligotrophic systems remains, however, controversial and the coupling between bacteria and bacterivores is not fully understood yet. This is partly due to the difficulty in obtaining precise measurements of bacterivory and biomass of heterotrophic nanoplankton as these rates and standing stocks are extremely low. Some comparative analysis from different aquatic systems suggest that top-down regulation of bacteria is more important in eutrophic and bottom-up control in oligotrophic environ-ments, although grazing and bacterial production are generally balanced (Sanders et al., 1992).
After the development of the ‘Microbial Loop’ concept, the prevailing picture was that a highly active and efficient protist grazer community controls bacterial abundance and consumes new bacterial production, which eventually results in the rather constant and homogenous low bacterial abundance observed in the open ocean (e.g. Ducklow, 1983). Efficient grazing control was also supported by experimental microcosm studies which revealed a close coupling between bacteria and bacterivores (Wikner and
¨
Hagstrom, 1988; Weisse, 1989) and by substrate addition experiments in which bacterial abundance remained relatively constant despite strong increases in bacterial production (Kirchman, 1990; Kirchman and Rich, 1997). Even less is known about the importance of grazing as a shaping factor for the phenotypic and genotypic bacterial community composition in oligotrophic systems. Studies in more productive coastal or freshwater environments have demonstrated that bacterial grazing has an important impact in this
¨ ¨
respect (Jurgens and Gude, 1994).
The purpose of the present study was to examine the bacteria–protozoan coupling in warm oligotrophic to ultra-oligotrophic ocean sites in microcosm experiments. Ex-perimental manipulations with substrate additions and size-fractionations were aimed at stimulating bacterial production and increasing predation pressure by small bacterivores. We were specially interested in (1) revealing the response time of bacterivores to increases in bacterial production and bacterial biomass, and (2) analysing the impact of grazers on bacterial size distribution and the appearance of grazing-resistant mor-photypes in response to increased predation.
2. Material and methods
Bottle incubation experiments were carried out during the cruise Latitud II across the Central Atlantic Ocean (from the Canary Islands (Spain) to Mar del Plata (Argentina),
´
Fig. 1. Latitude II transect across the Central Atlantic with the sampling stations at which microcosm experiments 1–4 were performed (arrows).
Guinea dome, which is centered at 208W, 108N (Stramma and Schott, 1999). The Guinea dome appears south of Cape Verde, and creates a cyclonic structure and an upward displacement of the isolines (Siedler et al., 1992).
All experimental incubations were conducted in duplicate acid-washed 1.5-l poly-carbonate bottles. Experiments started within 2 h after sampling and lasted for 96–160 h. The bottles with surface water (5 m) were incubated in water tanks in order to simulate in situ conditions with approximate ambient light and temperature values. Water temperature in the tanks varied at the most 0.58 from in situ temperature. Light was reduced with a Nylon net covering the tanks. Light measurements during the cruise showed that we could simulate the light at roughly 5-m depth. The samples from the DCM were incubated in a climate chamber, at 17–198C, with light of roughly 1% of surface noon light.
The experiments 1, 2, and 4 consisted of differentially filtered treatments in order to enhance or eliminate predation on bacteria by bacterivores. One fraction, filtered through a ,5 mm filter (42 mm polycarbonate membrane, Millipore), was designed at eliminating micro- and mesozooplankton, thereby removing predators on HNF and increasing predation pressure on bacteria. One fraction filtered through a ,0.8mm filter should contain only bacteria and no eukaryotic bacterivores. In experiment 2 we had an additional fraction, for 5 m and DCM, filtered through a 50-mm mesh net in order to remove mesozooplankton. We always had two replicate treatments without nutrients and two replicates which received substrate additions to stimulate bacterial growth. We used yeast extract as a complex substrate source because in previous experiments this resulted in an immediate growth of bacteria (data not shown). The concentration of yeast extract
21
which was added, between 1.5 and 4.5 mg l , was sufficient to increase bacterial abundance by approximately one order of magnitude. The chemical analysis of the yeast extract we used (Difco) revealed a molar C:N:P ratio of 93:21:1, resulting in an addition of 50–150 mM organic C in the experiments. In experiment 3 only unfiltered water incubations from 5 m depth were used to which different substrate sources were added:
21
yeast extract (1.5 mg l ), glucose (7.4 mM), mixture of amino acids (5 mM), or inorganic nitrogen (2 mM NH NO ).4 3
Subsamples for bacteria and HNF were taken daily, fixed with cold glutaraldehyde (1% final concentration) and enumerated using epifluorescence microscopy and DAPI staining (Porter and Feig, 1980). Between 1 and 10 ml (depending on enrichment and time of incubation) were filtered on black 0.2 mm-polycarbonate filter (Millipore) for counting bacteria. The filters were frozen at 2208C until processing of the samples in an epifluorescence microscope (Zeiss Axiophot). For measuring bacterial cell size distribution and calculation of mean cell volume we used an automated image analysis
¨
system (SIS GmbH, Munster, Germany). Briefly, the procedure consisted of recording images of DAPI-stained cells (500–1500 cells per size analysis) with a CCD camera and measuring the cell dimensions (pixel area, perimeter) after edge detection with a second derivative filter, manual thresholding and binarization (e.g. Massana et al., 1997).
3
21
C (mol leucine) . This factor is an average conversion factor determined in several experiments during the cruise from parallel measurements of biomass increase and
3
H-leucine incorporation in 0.8-mm filtered bottle incubations (Gasol et al. submitted). For some of the substrate-enriched fractions we quantified bacterial abundance in aggregates at the end of the experiments. Aggregated bacteria were directly counted in normal DAPI preparations when samples contained only smaller aggregates. When large aggregates with more than several hundred bacteria appeared, these could not be properly quantified and samples were sonicated (Branson Sonifier 250, ten bursts of 3 s) to disperse aggregated bacteria. From a first DAPI preparation only the number of freely dispersed single cells was obtained. The total number of bacteria was counted after a second DAPI preparation of the sonicated sample. The difference of the two counts gives an estimate of the bacterial abundance in aggregates. In addition we counted and sized (maximal dimension) bacterial aggregates at the end of the experiments from the DAPI preparations. To visualize whether bacterial aggregates were embedded in a matrix of extracellular polysaccharides, selected samples were stained with alcian blue according to the procedure described by Logan et al. (1994).
Statistical analysis including t-test and analysis of variance (ANOVA) were performed using the software package STATISTICA.
3. Results
Yeast extract proved to be an ideal substrate for stimulating bacterial growth and increasing bacterial abundance, probably because besides organic carbon also inorganic macro- and micronutrients are supplied. Experiment 3 compares the effects of the relatively high substrate pulse of yeast extract addition (¯50 mM organic C) to the addition of glucose, inorganic nitrogen, or a mixture of amino acids, all supplied at low concentrations similar to previously published substrate addition experiments (Kirch-man, 1990; Kirchman and Rich, 1997). The strong effect of yeast extract on bacterial productivity and abundance is shown in Fig. 2. The addition of glucose or inorganic nitrogen had no effect on the development of bacterial abundance or leucine uptake compared to the unamended controls (one-way repeated-measures ANOVA of the effects of nutrient addition on bacterial abundance or leucine uptake, both p.0.05). The addition of amino acids resulted in a moderate increase in bacterial abundance ( p,
0.05) but not of leucine uptake ( p.0.05) compared to the controls.
For the coupling between bacteria and protozoans after nutrient enrichment we examined altogether seven size-fractionation and nutrient addition experiments: at three stations (Exp. 1, 2, and 4) incubations with water from two depths (5 m, DCM), and at one station nutrient enrichment of unfiltered water from 5 m depth. Although the stations were from different oceanic areas, the first two experiments from slightly more productive Northern subtropical areas and the second two experiments from the very unproductive Southern subtropical Atlantic gyre, they all represented warm, oligotrophic
21
situations. The chlorophyll a concentrations were in the range 0.03–0.23 mg l at the 21
surface and in the range 0.17–0.36mg l in the depth of the DCM (Table 1). Bacterial
5 21
Fig. 2. Bacterial response to the addition of different nutrients in unfiltered incubations of experiment 3 (5 m
21
depth). CONTR: Control bottles without nutrient additions, YE: yeast extract (1.5 mg l ), GLC: glucose (7.4 mM), AA: mixture of ten amino acids, (100mM); N: NH Cl (3.34 mM). (A) Leucine incorporation, mean and range of two replicate treatments, each measured with triplicate subsamples (not for AA because of the dilution with cold leucine); (B) development of bacterial cell numbers, mean and range of two replicate treatments.
5 21
Table 1
a
Physical and biological parameters for the stations at which microcosm experiments were performed Exp. Depth Temp. Bacteria HNF T Bactd Prochlorophytes Synechococcus Chl a
5 21 21 5 21 3 21 21
(m) (8C) (10 ml ) (ml ) (days) (10 ml ) (10 ml ) (mg l )
1 5 24.4 8.14 351 77.4 1.24 6.18 0.09
85 19.6 5.13 192 81.9 0.46 0.68 0.25
2 5 28.9 6.92 805 1.1 2.33 9.29 0.23
50 16.8 2.26 166 5.6 0.38 0.45 0.36
3 5 25.4 4.42 446 4.1 0.84 4.67 0.03
4 5 25.0 3.84 532 5.0 1.58 9.83 0.04
180 22.5 2.19 101 9.7 0.36 0.26 0.17
a
The second depth for experiments 1, 2 and 4 is the depth of the deep chlorophyll maximum (DCM). Bacterial doubling time (T ) is based on leucine incorporation data and the empirically determined conversiond
factor of 0.62 kg C per mol of leucine. Concentrations of prochlorophytes and Synechococcus were determined by flow-cytometry (as described in Gasol et al., submitted).
A general pattern in bacterial development, similar to all experiments, was observed. The bacterial population development is illustrated here only for the size-fractionation experiment 1, surface and DCM (Fig. 3), but the experiments 2 and 4 did not deviate essentially from this pattern. The addition of yeast extract always resulted in a rapid 21
increase of bacterial abundance, with growth rates in the range 0.07–0.13 h . Maximum bacterial numbers were proportional to the amount of added yeast extract, which differed slightly between the experiments, and reached values between 3.4 and
6 21
9.6310 ml (Table 2). This level was approximately one order of magnitude higher
Fig. 3. Example of a fractionation and enrichment experiment (exp. 1). Development of bacteria and HNF in
21
Table 2
a
Summary of the enrichment experiments
Exp. Enrichment Depth Maximum HNF
(yeast extract) (m) bacterial numbers response time
21 6 21
Maximum bacterial numbers (mean6S.D. of the fractions ,0.8 and ,5 mm) after substrate addition. HNF response time is defined as the time period after which bacteria in the,5-mm fractions were reduced to about the initial levels due to HNF grazing.
than in the natural situation at the beginning. Within 24 h bacteria seemed already to have reached stationary phase after which bacterial numbers stayed more or less constant until the end of the experiments (in the ,0.8-mm fractions) or until nanoflagellates developed (in the ,5-mm fractions). HNF were comprised mainly by naked, colourless forms, 3–5 mm in diameter, which increased within 1 day from nearly undetectable to
4 21
levels .5310 ml . For treatments in which we had fairly reliable estimates on the initial HNF abundance before the peak, we estimated the population doubling time to be in the range 5–8 h. The HNF population peaks coincided with a rapid decline of the
5 21
bacteria to slightly above the initial levels (5–9310 ml ). Peaks of HNF were short-lasting and they decreased again to low levels after the collapse of the bacterial community. We defined the HNF response time to the bacterial enrichment as the time period which it took for the grazers to reduce bacteria to about the initial levels. This corresponded generally to the peak in HNF abundance. The values were in the range of 3–6 days (Table 2) with lower values in the warmer surface waters than in the DCM. The variability in response time (Table 2) and in HNF peak abundance (Fig. 3) is due to the fact that HNF did not always increase exactly simultaneously in replicate treatments. Bacterial abundance in the unenriched fractions did not remain constant although this is not visible from Fig. 3 in which bacteria of enriched and unenriched treatments are plotted at the same scale. A 2–3-fold increase in bacterial abundance occurred in the unenriched treatments in all size fractions during the course of the experiments. Together
3
with an increase in mean cell volume from 0.05 to 0.07–0.09 mm this resulted in a considerable increase of bacterial biomass. For experiment 2, in which this stimulation of bacterial growth was especially pronounced, this pattern is illustrated in Fig. 4. The bacterial growth was also expressed in an immediate and drastic increase in leucine
21
incorporation. The leucine incorporation reached about 2–4 nM h after 24 h of incubation and remained at this high level until the end of the experiments. HNF
21
3
Fig. 4. Development of bacterial biomass (closed symbols) and H-Leucine incorporation (open symbols) in different size fractions of experiment 2 (water from 5 m depth) without nutrient addition. Means and ranges of two replicate treatments.
Fig. 5. Bacterial size distribution and mean cell volume at times 0, 72, and 115 h during the incubation of ,0.8 and ,5-mm filtered and substrate-enriched water (example from exp. 2, water from 5 m depth).
the size distribution shifted again to small coccoid cell volumes in the ,5-mm fractions after the peak in HNF development and the bacterial decline.
Fig. 6. Average size distribution (longest dimension) of bacterial aggregates, pooled data from different experiments (n57).
aggregates with about 5–20 mm in size and harboring approximately 20–100 bacteria / aggregate dominated (about 70% of total aggregates). Additionally, large aggregates in the range 50–100 mm occurred, with several hundred bacteria per aggregate. The aggregates could be stained with the dye Alcian blue, indicating that the bacteria are embedded in a polysaccharide matrix (Fig. 7) and thus looked similar to transparent exopolymer particles (TEP) described before (Alldredge et al., 1993).
Bacterial aggregates were most obvious in the ,5-mm fractions after HNF had developed but that was due to the fact that nearly all freely suspended bacteria were eliminated and not to the fact that aggregates developed only in these treatments. Bacterial aggregates were also present in the substrate-enriched ,0.8-mm fractions but here their relative importance was much less due to the high number of freely suspended single cells. At the end of the experiments the ,0.8-mm fractions were dominated to 80–95% by freely suspended bacteria whereas in the ,5-mm fractions bacteria in aggregates made up 22–72% (mean 51%) of the total bacterial abundance. However, when looking at the total numbers it is obvious that aggregated bacteria appeared in about the same amounts in ,5-mm and ,0.8-mm fractions but that freely suspended bacteria were strongly reduced in the ,5-mm fractions (Fig. 8).
4. Discussion
Fig. 7. Microphotographs of a bacterial aggregate in the ,5-mm fraction which appeared after protozoan grazing in exp. 1. (A) Epifluorescence, DAPI staining; (B) transmission, Alcian Blue staining. Bar in panel B corresponds to 10mm.
Fig. 8. Proportion of free and aggregated bacteria at the end of the experiments in the substrate-enriched fractions ,0.8 and ,5mm.
adequate to mimic natural processes (Carpenter, 1996). This should be especially true for microbial processes as bacterial communities are known to be sensitive to bottle incubations during which signs of an enrichment effect often become evident (Ferguson et al., 1984). Plankton communities in ultra-oligotrophic waters are probably especially sensible to sample handling and confinement as small changes in DOC supply and availability might have strong alterations for the limitation conditions and could result in enrichment effects on the bacterial community (Gasol and Moran, 1999).
leucine after 24 h in our bottles could be due to a shift in growth rate in the bottle incubations and a resulting uncoupling of protein synthesis and cell division (Chin-Leo and Kirchman, 1990; Ducklow et al., 1992).
This presently unexplained bottle effect was, however, not a major restriction to our experimental goals as we were not intending to measure in situ rates of production and grazing losses at near steady state conditions in the system. Instead we wanted to examine the response of bacterial grazers to a transient state after a substrate pulse and an increase in bacterial production and biomass. The stimulation and increase of bacteria in the unenriched fractions was obviously not sufficient for a significant development of a flagellate grazer population within the incubation time (5–6 days). A slight increase in HNF concentration might have remained undetected with our methods but as no decrease in bacterial numbers nor a change in size distribution occurred, we judged the grazing impact not to be very important. This is further confirmed by the fact that there were no differences between fractions in which grazers were eliminated (,0.8 mm) compared to fractions in which grazers potentially remained or developed (,5, ,50
mm). We found it difficult to give a precise enumeration of heterotrophic nanoflagellates in the natural samples of most stations and in the unenriched fractions of the
21
experiments. Concentrations were often below 200 ml , cells were generally small (,3mm), did not contain peculiar features and were mostly without preserved flagella. The storage of frozen DAPI stained filters might have further decreased the quality of the preparations compared to fresh samples. However, nanoflagellates looked different, were clearly visible and easy to count when they developed in the substrate enriched incubations. This can indicate that the enrichment might have resulted in a shift of HNF community composition (Lim et al., 1999) or in a change of the physiological (and morphological) state of the nanoflagellates. Not much is known about the starvation– survival mode of heterotrophic flagellates which would enable their existence during prolonged periods of very low bacterial concentrations until a bacterial peak develops or until they encounter a micropatch with elevated bacterial concentrations. Estimates of HNF growth in the oligotrophic Sargasso Sea revealed very low rates, with doubling times in the range of days to weeks (Caron et al., 1999), and indicated severe food limitation. In short-term starvation experiments with HNF isolates, a rapid decrease in respiration rate and cell volume was found (Fenchel, 1982a) but it is not known whether, besides cyst formation, mechanisms for long-term survival exist.
Bacteria responded quickly to the substrate additions as would be expected at these high water temperatures and has been shown in previous DOM enrichment experiments (Kirchman and Rich, 1997). Our enrichment with a high concentration of DOM resulted in an approximately 10-fold increase of bacterial abundance within 24 h. The consistent pattern in our enrichment experiments was that bacterial abundance rapidly reached a saturation level and stayed there without strong alterations for 3–6 days, until HNF developed in higher abundance. The delay of HNF development could simply be the result of a very low initial abundance but continuous exponential growth. For example,
21
an initial HNF abundance of 200 cells ml would achieve, with a mean growth rate of
21 4 21
changes, for several days and then increased with very high growth rates (4–6 h doubling time) within less than 1 day. Similar high growth rates of HNF were found by
¨
Weisse and Scheffel-Moser (1991) in prolonged bottle incubations, also following a bacterial peak with a lag-phase of about 2 days. This pattern suggests that HNF had to adapt to the enriched food conditions and could not immediately respond with increased growth rates.
Temporary lags in grazing pressure on increased bacterial abundance have also been reported from other incubation studies with oligotrophic seawater (Van Wambeke and Bianchi, 1985) and observed also in situ (Kirchman et al., 1989). Whereas bacteria seem to respond within hours to enhanced substrate concentrations, grazers cannot keep up with an increase in bacterial production and seem to need, at least in very oligotrophic systems, several days for adaptation and population increase. In more productive freshwater and marine systems the response time of bacterivores to increases in bacterial abundance or production are generally much shorter, in the range of a few hours to ¯1 day (e.g. Andersen and Fenchel, 1985; Hadas et al., 1990).
More close coupling between bacteria and grazers is indicated by microcosm experiments which found constant bacterial numbers despite increased production rates. For example, in experiments in the equatorial Pacific Ocean the addition of low amounts (1 mM) of glucose or amino acids resulted in a strong stimulation of thymidine incorporation but only in negligible changes in bacterial abundance (Kirchman and Rich, 1997). This was interpreted in a match of bacterial production by bacterial mortality. If we consider, however, that unbalanced growth occurs in these bottle incubations and that the incorporation of thymidine or leucine into non-growing cells has been demonstrated (Marden et al., 1987, Marden et al., 1988), we must be cautious with conclusions on grazing mortality from data on thymidine incorporation and bacterial abundance.
Imbalances between bacteria and their main grazers might occur on different scales. Probably most important is a microspatial scale at which bacteria benefit from increased nutrient supply (Azam et al., 1994). For example, abundant marine snow aggregates exist also in the oligotrophic ocean (Alldredge and Silver, 1988). In these organically enriched microenvironments much higher substrate (Shanks and Trent, 1979, Herndl, 1992), bacterial (Alldredge et al., 1986), and protozoan (Caron et al., 1982) con-centrations can be found. But also on temporal scales bacteria can benefit from transient events such as episodic phytoplankton blooms which occur even in the oligotrophic ocean (Lohrenz et al., 1988). On both scales protozoans would follow bacteria with a lag phase, exerting grazing pressure and top-down control after bacteria had multiplied already several times and released new cells into the environment.
Feed-back effects at the bacterial level (reduced vulnerability) in response to protozoan grazing are widespread at least in more productive systems (see review by
¨ ¨
Jurgens and Gude, 1994). In planktonic freshwater environments, filamentous bacteria, belonging to different phylogenetic groups, are often a dominant form of
grazing-¨
resistant bacterial morphotype (Jurgens et al., 1999), and filamentous bacteria were also observed in enclosure experiments with coastal marine water (Shiah and Ducklow, 1995; Havskum and Hansen, 1997). We expected that the combination of high grazing pressure and sufficient substrate supply should allow the development of potentially resistant
¨ ¨
bacterial strains (Jurgens and Gude, 1994). In the substrate enriched size-fractions of our experiments we never observed the appearance of filamentous bacteria. It is presently not known whether the bacterioplankton in the open, oligotrophic ocean consist of a very distinct bacterial community composition compared to coastal areas.
The only bacteria which were obviously not heavily grazed by nanoflagellates at the end of the incubations, were those attached to aggregates, mostly embedded in a polysaccharide matrix (Fig. 7). These bacterial–detrital particles which developed in the experiments resemble those described as transparent exopolymer particles (TEP), gel-like polysaccharide particles produced by algae and bacteria (Alldredge et al., 1993). TEP are abundant in the ocean, provide the matrix of marine snow and are often colonized by substantial amounts of bacteria (Alldredge et al., 1993; Passow and Alldredge, 1994). The distribution of aggregated and free bacteria in our experimental treatments with and without grazers (Fig. 8) implies that the bacterial aggregates developed already before the HNF peak and remained there when freely suspended bacteria were eliminated. In a continuing succession these particles would probably become colonized by other groups of protozoans, specialized to feed on attached and aggregated bacteria (Caron, 1987; Artolozaga et al., 1997). The general shape of the size distribution of TEP resembles those found in the ocean, especially during situations in which diatoms were absent (Passow and Alldredge, 1994). However, we have to be aware that the fixation and filtration procedures probably modify the particulate material and the actual in situ dimensions might deviate from those measured microscopically.
Another well-known feed-back effect of flagellate grazing on bacterial assemblages, which occurred in our experiments, is the shift in bacterial size distribution
(Kuuppo-ˇ
Leinikki, 1990; Simek and Chrzanowski, 1992). This effect and the interplay between substrates and grazing as determinants of bacterial cell size distribution, was evident when comparing enriched fractions with and without grazers (Fig. 5). Bacterial rods
3
with a cell volume .0.08 mm were preferentially eliminated by flagellates whereas these remained in the grazer-free fractions (,0.8 mm). This mechanism can be responsible for the return of bacterial size distribution to the original state after a substrate pulse has shifted it to larger cell sizes.
The general picture of bacteria–protozoan coupling in the oligotrophic ocean is not easy to resolve and contradictory results exist. Bacterial growth rates, based on thymidine incorporation and estimated grazing mortality, are probably very low, with doubling times, in the range of one to several weeks (Cho and Azam, 1988; Fuhrman et al., 1989; Zohary and Robarts, 1992). This also suggests a low grazing pressure on bacteria. Low bacterial grazing was probably also the case in our unenriched incubations although we did not directly measure grazing rates.
a lower threshold where bacterial grazing becomes very inefficient (Wikner and ¨
Hagstrom, 1991). A numerical refuge of bacterial communities was estimated to exist
5 21
¨
below 7310 cells ml by Wikner and Hagstrom (1991), and Cho and Azam (1990)
5
found a lower threshold of bacterial concentrations in the euphotic zone of ¯3310 21
ml . If bacterial concentrations in the oligotrophic ocean are close to the lower threshold, active growth of bacterivores might be restricted to microniches such as marine snow. However, autotrophic picoplankton (Synechococcus, Prochlorococcus) can be an important additional food resource for protozoan grazers (Burkill et al., 1993; Landry et al., 1995; Reckermann and Veldhuis, 1997). It has been shown that bacterial growth and predation losses in oligotrophic oceanic systems are probably more or less balanced (Weisse, 1989; Landry et al., 1995; Goosen et al., 1997; Caron et al., 1999) although sophisticated methods might be necessary to obtain precise measurements in these systems (Zubkov and Sleigh, 1998; Zubkov et al., 1998). Bacteria should be able to quickly respond to DOM inputs, exploit the readily available substrate pool, and form ‘blooms’ on a temporal or microspatial scale without significant grazer control due to the time lag in HNF development and the absence of other efficient bacterivores.
Acknowledgements
´
We appreciate the help of Evaristo Vazquez during the experiments. The study was ´ financially supported by the Latitude grant CICYT [AMB94-0739 to Susana Agustı. [SS]
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