Hydrobiologia (2007) 583:223–230 DOI 10.1007/s10750-006-0532-2
PRIMARY RESEARCH PAPER
Glucose uptake and extracellular polysaccharides (EPS) produced by bacterioplankton from an eutrophic tropical reservoir (Barra Bonita, SP–Brazil) Rafael S. Panhota Æ Irineu Bianchini Jr Æ Armando A. H. Vieira
Received: 26 May 2005 / Revised: 22 November 2006 / Accepted: 5 December 2006 / Published online: 3 March 2007 Springer Science+Business Media B.V. 2007
Abstract We have studied the production of polysaccharides by bacterioplankton in an eutrophic tropical reservoir (Barra Bonita, SP–Brazil) through a decay experiment using glucose as carbon source. The temporal evolution was monitored by measuring the total organic carbon and the contents of monosaccharides. The glucose added to the reservoir sample water was consumed at higher rates within the first hours of incubation, and after 30 days 94.4% of the carbon contents were mineralized; 4.2% remained as particulate organic carbon (POC) form and 1.5% as dissolved organic carbon (DOC) form. The production of polysaccharides occurred in two stages: within the first 48 h, there was intense glucose consumption with small POC increment (ca. 16%) and release of small quantities of dissolved polysaccharide. In the second, more intense stage production was accelerated after the 9th day of incubation, with the highest polysaccharide concentration measured on the 20th day. Such formation of polysaccharides was related to the excretion of capsules and sheaths by bacte-
Handling editor: S. M. Thomaz R. S. Panhota I. Bianchini Jr (&) A. A. H. Vieira Departmento de Hidrobiologia, Universidade Federal de Sa˜o Carlos, Via Washington Luiz, Km 235, Sao Carlos, SP 13565-905, Brazil e-mail:
[email protected]
rioplankton, mainly in the senescence of heterotrophic populations, with release of reserve and structural intracellular materials. Keywords Extracellular polysaccharides EPS Monosaccharides Bacterioplankton Decomposition
Introduction In aquatic environments bacterioplankton release considerable quantities of cell material (Brophy & Carlson, 1989; Stoderegger & Herndl, 1998), usually referred to as extracellular polysaccharides or exopolysaccharides (EPS). The structure and function of these polysaccharides are still poorly understood, in spite of their importance for the limnology of tropical areas in which the organic carbon released may contribute significantly to the pool of organic compounds, in their dissolved (DOC) or particulate forms (POC). The substances released by bacterioplankton depend on their growth stage. When metabolically active, bacterioplankton release mainly polysaccharides derived from renewal of cell wall capsules (Stoderegger & Herndl, 1998), while during the senescent stage intracellular structures are preferentially released (Shibata et al., 1997). The compositions of such extracellular polymers vary with type of microorganisms (Vaningelgem et al.,
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2004), nutrient availability (Degeest & De Vuyst, 1999; Ricciardi et al., 2002), phase of microorganisms growth and environmental conditions (Fisher et al., 2003; Bahat-Samet et al., 2004). In general, such compounds consist primarily of high-molecular weight, hydrated polysaccharides (Heissenberger et al., 1996). The EPS (capsule or slime layer) are involved in cell protection against phagocytosis or dehydration (Shimada, 1997), surface fixation and formation of chelates (Welch et al., 1999; Flemming & Wingender, 2001a; Bahat-Samet et al., 2004), retention of exoenzymes, cellular debris, genetic material and nutrients sequestration from the water phase (Flemming & Wingender, 2001a). Usually, polysaccharides are complexed with metallic ions (Guibaud et al., 2005) and provide the structural unities of humic substances. In fact, one possible route for formation of humic compounds is polysaccharides condensation with amines that originate the melanoidins (Stevenson, 1994). According to Koivula & Ha¨nninen (2001), the humus arises not only from degradation of refractory compounds but also from microbial synthesis. Bacterioplankton utilize a considerable fraction of products generated by phytoplankton primary production as carbon source (FreireNordi & Vieira, 1996; 1998; Passow et al., 2001; Giroldo et al., 2003; Colombo et al., 2004; Pacobahyba et al., 2004). Jørgensen & Volleinweider (2000) estimated that 40% of phytoplankton primary production is consumed by microbial metabolism. In addition to being important as DOC mineralization agents, bacterioplankton perform an important role as organic carbon source to higher trophic levels of the food web, in the microbial loop process (Azam et al., 1983). DOC normally appears in aquatic environments as high-molecular weight compounds, but bacterioplankton may use enzymatic systems to transform such compounds in low-molecular weight substances for further consumption (Chro`st, 1991). In this context, the present study is aimed at identifying and quantifying monosaccharides generated in the release of polysaccharides by bacterioplankton in the aerobic decomposition of a simple monosaccharide (glucose). Use is made of high-performance liquid chromatography
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(HPLC) with pulse amperometric detection (PAD) (Clark et al., 1991; Gremm & Kaplan, 1997; Panagiotopoulos et al., 2001; Balance et al., 2005).
Materials and methods The Barra Bonita Reservoir is located within the Middle Tieteˆ River basin at 430 m of altitude; in the central area of Sa˜o Paulo State (2229† to 2244† S and 4810† W). The reservoir is one of the most populous and developed areas of Brazil; in a geographical transition between tropical and subtropical climates, where seasons are basically divided in rainy (Spring and Summer) and dry (Autumn and Winter) seasons. The Barra Bonita Reservoir is considered mainly polymictic (with rare periods of thermal stratification) and eutrophic tropical ecosystem, in which the seasonal cycles of limnological events seem to be dominated by precipitation, wind, outflow and residence time, which may vary from 1 to 6 months (Tundisi & Matsumura-Tundisi, 1990). It is also known that a large input of nutrients occurs during the rainy season (Henry et al., 1985; Calijuri & Dos Santos, 2001). Water samples were collected in March/2002 in a pelagic area at 3.0 km upside the dam (2232¢34.5† S and 4829¢26.4† W), at different depths (1, 10 and 20 m), in order to guarantee the vertical homogeneity of the sample. The equivalent water samples were mixed and filtered through glass wool (to remove the coarse detritus). In laboratory, the water samples were filtered in a fiberglass membrane (1.2 lm, Whatman) and amended with glucose (solution of 50 mg l–1 ” 20 mg l–1 of DOC). Two replicate solutions were incubated in flasks of 1 l for 30 days in the dark, under aerobic conditions and controlled temperature (20.0 ± 1.6C). Control solutions (n = 2) were also prepared with filtered (1.2 lm, Whatman) samples without glucose. To keep the aerobic condition, the solutions were maintained with dissolved oxygen (DO) in concentrations higher than 2.0 mg l–1, by bubbling clean air periodically. The concentrations of DO and temperature in the solutions were monitored daily (DO meter, YSI–model 58).
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TOC; model 5000A). After TOC analyses, the samples were filtered through an ester cellulose membrane (0.45 lm; Millipore) to separate the particulate and dissolved fractions, which had also their carbon contents measured. The concentrations of particulate organic carbon (POC) were estimated from the difference between the total (TOC) and dissolved organic carbon (DOC) concentrations. We assumed that POC concentrations (>0.45 lm) were associated with the biomass of microorganisms.
Results Figure 1 shows the temporal evolution of organic carbon. After 30 days, 94.4% of the glucose carbon were mineralized, i.e. converted into inorganic carbon by respiration in the indigenous microorganisms community, 4.2% were present in particulate carbon forms, which probably included bacterioplankton cells, and 1.5% was in dissolved forms. The particulate carbon fraction presented two well-defined concentration peaks, at 48 h (16%) and 13 days (29.8%). The formation of monomers from dissolved (DPM) and particulate (PPM) polysaccharides due to glucose consumption by bacterioplankton is shown in Fig. 2. The concentrations of monosaccharides derived from polysaccharides varied
30 25 20 -1
C (mg l )
During the incubation period (30 days), three aliquots (ca. 10 ml) of each flask were periodically sampled; two of them were filtered in ester cellulose membrane (0.45 lm; Millipore) to determine the free (dissolved) monosaccharide (FM); one of these filtered samples was hydrolyzed to determine the monosaccharides (FM) derived from dissolved polysaccharides (DPM). In this case, the DPM concentrations were calculated by the difference between the total monosaccharide and FM concentrations. No-filtered and hydrolyzed samples were used to quantify total monosaccharides, which includes: FM, DPM and those derived from particulate polysaccharides (PPM). The concentrations values of PPM were estimated from the difference between the total monosaccharides concentration (obtained in the non-filtered and hydrolyzed samples) and FM and DPM concentrations. Desalted samples without hydrolysis were directly analyzed on PAD-HPLC in order to detect and identify free dissolved monosaccharides. The hydrolysis was performed according to Gremm & Kaplan (1997). PAD-HPLC analysis was performed on Dionex DX500, consisting of a GP40 gradient pump module, ED40 electrochemical detector, and a LC5 manual injector with a Rheodyne 9125 valve and a 25-ll sample loop. ED40 was equipped with an amperometric flow cell, a gold working electrode, and a Ag/ AgCl reference electrode. PA-10 (Dionex) anion-exchange analytical column (250 · 4 mm), fitted with a corresponding guard-column (50 · 4 mm), was used to separate the monosaccharides. The mobile phase was NaOH 18 mM and NaOH 200 mM for the column recovering, at a flow rate of 1 ml min–1 (Gremm & Kaplan, 1997). All samples (polymeric and free monosaccharides) were desalted on a Bio-Rad ionic exchange resin (AG2X8-anion exchange and AG50W-cation exchange). In order to quantify the monosaccharides the following standards were used: glucose, fucose, arabinose, galactose, rhamnose and ribose (Sigma Chemical Co). Throughout the incubation time, one aliquot (ca. 10 ml) of each solution was periodically sampled for quantitative determination of carbon. The concentrations of organic carbon (TOC) were obtained with a carbon analyzer (Shimadzu
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DOC POC MC
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Fig. 1 Changes in DOC (dissolved organic carbon), POC (particulate organic carbon) and MC (mineralized carbon) during decomposition of glucose under laboratory conditions
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Fig. 2 Monomers from dissolved (DPM) (B1) and particulate (PPM) (C1) polysaccharides derived from glucose decomposition (A1), performed by bacterioplankton of Barra Bonita Reservoir; A2, B2, C2 show nine initial days in detail
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The temporal changes in concentrations glucose, monomers from dissolved (DPM) and particulate (PPM) polysaccharides and other Glucose
Others
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in time, with polysaccharide formation occurring mainly after 9 days of incubation. After the 15th day, the concentrations of galactose (DPM and PPM) decreased and from the 20th day, the concentrations of rhamnose and arabinose (PPM) also decreased. In relation to the monosaccharides from dissolved polymers (DPM), the concentrations of fucose and rhamnose increased from the 5th until the 30th day. The concentrations of galactose increased from the 10th to the 15th day and decreased until the last day. The concentrations of arabinose increased from 1st to 5th day and decreased until the 15th; after the concentrations increased again until the 20th day and decreased until the end (Fig. 2). The concentrations of arabinose and fucose from particulate polymers (PPM) presented continuous increments up to the end of the experiments. The concentrations of galactose and rhamnose decreased until the last day since the 15th and 20th days, respectively.
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Fig. 3 Temporal changes of glucose, dissolved (DPM) and particulate (PPM) monomers of polysaccharides and others compounds (CO2 and humic compounds) derived from glucose decay
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products, due to glucose decomposition, are illustrated in Fig. 3. Mineralization was the main process during glucose consumption, as the production of polysaccharides corresponded to only 2% of the decomposition products.
Discussion When the data for DOC and glucose concentration decays were treated by linear regression, one could note that on average the DOC decay corresponded to 61% of the glucose decay. Considering that 40% of glucose weight is supported by carbon, this result means that the glucose concentrations might have been underestimated, probably due to the analytical procedures applied to the samples before submitting them to PAD-HPLC analysis. Because only a few monosaccharide standards were used, some monomers (e.g. mannose, xylose, fructose, N-acetylglicosamine, glucuronic acid and galacturonic acid) may not have been distinguished in the determination of polysaccharides composition (Characklis & Marshall, 1990). This would cause the polysaccharide quantification (DPM, PPM e FM) to be underestimated. Glucose was intensely consumed since the early stages of decomposition, e.g. after only 12 h, ca. 11% of the glucose contents had already been consumed (Fig. 2A). According to Antonio & Bianchini Jr. (2003) and Panhota & Bianchini Jr. (2003), glucose consumption by microorganisms includes basically the following competing processes: (i) biological assimilation (anabolic pathways); (ii) mineralization (catabolic routes) and (iii) reaction with other compounds, which results in formation of humic substances. By fitting the glucose decay data to a first-order kinetics model, we estimated a half-time of 6.9 days for glucose decomposition. Results obtained by Antonio (2004) from glucose incubated in water samples from Barra Bonita reservoir were fitted to a sigmoid curve model, showing a half-time of 3.2 days. In this context, our value is approximately 9 times lower than that obtained for incubations with water from the Barra Bonita reservoir collected in February of the same year. According to the empirical models adopted by Antonio (2004), glucose consumption was associ-
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ated with changes in DOC concentrations and physical aspects of the water (e.g. Secchi disk depth), which are mostly related to input of matter during the rainy season. Since the population of bacterioplankton varies with environmental changes (Christensen & Characklis, 1990), the changes in monosaccharide concentration could arise from variation in the heterotrophic metabolic activity. Furthermore, it is also possible that the concentration of monosaccharides changed due to consumption and chemical rearrangement in their molecular structures during the formation of humic substances (Thurman, 1985; Cunha-Santino & Bianchini Jr., 2004). For instance, Giroldo (2003) reported a selective microbial utilization for carbohydrates excreted by algae with specific consumption rates for monosaccharides. This study (Giroldo, 2003) showed that during degradation of algal polysaccharides, rhamnose and fucose were consumed slower than other monosaccharides. As the polysaccharides retained rhamnose and fucose, they became more hydrophobic, which led to formation of aggregates. In this context, the POC concentration increased in the first hours of incubation, accompanied by a small release of dissolved polysaccharides (0.30%), which suggests that DPM formation could be related to polysaccharides excretion along with formation of capsules and sheaths by bacterioplankton (anabolic processes). The low efficiency of glucose conversion into exopolysaccharides was also observed in another study performed with batch cultures (Williams & Wimpenny, 1977). However, the higher concentrations of polysaccharides produced by bacterioplankton occurred after the 9th day, with maximum concentration being measured on the 20th day (3.35%). As the highest POC concentration was measured on the 13th day of incubation, the major release of polysaccharides was related to the senescence of microorganisms, when intracellular, structural and maintenance materials are usually soluble and available to the catabolic processes. The formation of dissolved and particulate polysaccharides (ca. 1%) depended on the type of resource. In this context, glucose has been considered as an important substrate for metabolic pathways that comprise energy gain (Lehninger,
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1990) (Fig. 3). In addition to polysaccharides, other compounds such humic compounds and CO2 were included as products from the glucose consumption (Fig. 3). In earlier works (Antonio & Bianchini Jr., 2003; Panhota & Bianchini Jr., 2003), CO2 derived from mineralization was the predominating product of decomposition. In relation to the polymeric polysaccharides produced by bacterioplankton, the followings monosaccharides were detected: rhamnose, fucose, galactose and arabinose. Rhamnose showed higher concentrations, followed by fucose, galactose and arabinose (Fig. 2). In earlier experiments on polysaccharides microbial heterotrophy using substrates from the same reservoir, rhamnose was the predominant monosaccharide among others excreted by Anabaena spiroides (Colombo et al., 2004; Antonio, 2004). The adherence strength of EPS in aggregates depends on the relative concentration of monosaccharides such as rhamnose, fucose and arabinose in the polysaccharide, which defines the features of EPS, especially with regard to adherence and aggregate formation (Zhou et al., 1998; Bahat-Samet et al., 2004). Another aspect still poorly explored in biofilms is its role in locating extracellular enzymes near the cells, because the polysaccharides of the EPS matrix can ensure a protected microenvironment for bacterial extracellular enzymes. It is possible that the specific associations between enzymes and EPS matrix can prolong its activities and resilience during environmental changes (Decho, 2000). The physical state of EPS (i.e. gel, slime, dissolved state) can influence its ability of ion binding. For example, the gel state exhibits higher affinity for binding cations than a looser slime state of a similar EPS (Geesy & Jang, 1989). Another factor that might influence metal chelation is the EPS change owing to UV irradiation. Earlier studies have shown increased availability of carboxyl groups and other organic molecules after EPS was exposed to UV irradiation (Kieber et al., 1990). Owing to its being a potentially labile carbon source, EPS may also play the role of transferring contaminated chelated metals through the food web, thus increasing metal bioavailability significantly (Schlekat et al., 2000). In any case, the ability of EPS to complex
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metal depends on its physical states (e.g. capsules, gel, slime, DOC) and chemical composition (Decho & Lopez, 1993). Finally, many EPS aspects remain unknown and require ecological studies as much as analysis of biotechnological potential (Flemming & Wingender, 2001b).
Conclusion The aerobic glucose consumption by bacterioplankton followed mainly the energetic metabolic pathway resulting in CO2. In a smaller proportion, catabolic processes also produced monosaccharides; those derived from polysaccharides (dissolved and particulate) prevailed. The formation of polysaccharides was associated to a lower extent to bacterioplankton excretion (e.g. capsules, sheaths), but depended mainly on the senescence of bacterioplankton and liberation of intracellular, reserve and structural material. Polysaccharides were then utilized or changed by chemical reactions. Thus, the decomposition of low-molecular weight compounds may produce high-molecular weight compounds, contributing to the pool of refractory organic compounds in the Barra Bonita reservoir. Although humification is not the predominant route of glucose decay, the formation of melanoidins may constitute an additional factor to maintain DOC concentration in this environment. Acknowledgements The authors thank Coordenadoria de Aperfeic¸oamento de Pessoal de Nı´vel Superior (CAPES) and Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq) for the scholarship and to Fundac¸a˜o de Amparo a` Pesquisa do Estado de Sa˜o Paulo (FAPESP, process no. 99/07766-0) for financing this study. Dr. D. Giroldo for HPLC analysis, Dr. R. M. Antonio for suggestion in the translation and Dr. Osvaldo N. Oliveira Jr. (IFSC-USP) for his critical proofreading of the manuscript.
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