Release of available nitrogen from river-discharged dissolved organic matter by heterotrophic bacteria associated with the cyanobacterium Microcystis aeruginosa/Mineraalsete lammastikuuhendite eraldamine joevee lahustunud orgaanilisest ainest tsuanobakteriga Microcystis aeruginosa assotsieerunud heterotroofsete bakterite poolt.
The cyanobacterium Microcystis aeruginosa is known to thrive mostly in eutrophic fresh waters. However, it is also frequently observed in brackish coastal waters and embayments (Kononen & Sellner, 1995). High biomass concentrations of M. aeruginosa occur also in estuarine areas transported there via rivers. In the coastal areas of the Gulf of Riga maximum growth of M. aeruginosa continues from late June till September. Although generally the development of M. aeruginosa proceeds sporadically, it often initiates blooms in coastal zones (Balode & Purina, 1996; Seppala & Balode, 1999), including the Gulf of Riga where the culture of the cyanobacterium Microcystis aeruginosa (MAGR-2) was isolated. High abundances of M. aeruginosa are related to high nutrient concentrations. Studies on this species have been intensified in the last decades mainly due to the discovery of potent hepatotoxic heptapeptides, called microcystins, shown to be detrimental to wild and domestic animals and humans (Lambert et al., 1994; Van Dolah, 2000; Ojaveer et al., 2003).
Wetland-derived dissolved organic matter (DOM) may be an important nitrogen source for coastal ecosystems in summertime when concentrations of inorganic nitrogen are low and the bioavailability of dissolved organic nitrogen (DON) is high (Stepanauskas et al., 1999). For instance, the cyanobacterium Nodularia spumigena reaches the highest cell density when cultured in the presence of DOM > 1000 D originating from a river, and it was concluded that DOM could substitute dissolved inorganic nitrogen (DIN) as a nutrient source (Panosso & Graneli, 2000). Although it is verified that the compounds of DON sustain the growth of cyanobacteria, including Microcystis aeruginosa (Purvina et al., 2008), heterotrophic bacteria are the primary contributors in the mineralization of organic matter. Therefore probably one of the main pathways for DOM utilization by phytoplankton in aquatic ecosystems is bacterially mediated enzymatic breakdown resulting in release of inorganic compounds (Berman et al., 1999; Grossart, 1999). According to investigations of Worm & Sondergaard (1998), Microcystis colonies are bacterial incubators for the surrounding water as well as the 'hotspot' for bacterial activity. Because of the particulate nature of Microcystis and the influence exerted on the physico-chemical environment of colonies by products of its metabolism at the micro-scale, bacteria associated with Microcystis colonies live in a microhabitat distinct from the surrounding water. This indicates close interaction between cyanobacteria and bacteria.
A number of literature sources contain information regarding the consumption of allochtonic organic matter by marine bacteria (Sondergaard & Milddelboe, 1995; Young et al., 2005), the usage of organic matter produced or excreted by phytoplankton (Lignell, 1990; Baines & Pace, 1991; Obernosterer & Herndl, 1995), as well as the degradation of toxins produced by M. aeruginosa and other cyanobacteria (Jones et al., 1994; Christoffersen et al., 2002; Bourne et al., 2006). Less is known on how associated heterotrophic bacteria contribute to the biodegradation of allochtonic DOM brought by rivers and whether they supply cyanobacteria with these compounds. Therefore, we performed an experiment to verify the hypothesis that bacteria associated with M. aeruginosa are involved in the process of allochtonic DOM biodegradation and production of bioavailable inorganic N compounds. In this study we report the metabolic pathway of DON, ammonia, nitrate, and nitrite in experimental treatments with the cyanobacterium Microcystis aeruginosa, associated bacteria, and land-derived DOM as the only source of nitrogen and phosphorus.
MATERIALS AND METHODS
Cultures and experimental design
The experiment was carried out in the subsequent series: (1) Microcystis aeruginosa culture (Control), (2) M. aeruginosa culture + DOM (AB + DOM), and (3) bacterial filtrate + DOM (B + DOM). Cultures were grown in filter-sterilized (0.22 [micro]m) seawater distributed (4 L) into 5-litre polycarbonate pre-autoclaved bottles. The experiment lasted for 15 days. The numbers of bacteria and cyanobacteria, bacterial production, and dissolved and particulate nutrient concentrations were monitored according to the growth curve of M. aeruginosa on days 0, 6, 7, 9, 12, and 15. Bacteria samples were additionally taken on Day 3. The in vivo fluorescence and M. aeruginosa cell number concentration were monitored daily till Day 15. The sampling volumes were calculated so that at least 1 L of experimental solution should remain for the last day of sampling. Daily DOM additions to treatments AB + DOM and B + DOM lasted till Day 6 with DON totalling 20 [micro]M and 22 mL concentrate per bottle (the normal in situ concentration of the summer waters from where M. aeruginosa MAGR-2 was isolated). Each treatment had three replicates.
The clonal culture of Microcystis aeruginosa Kutzing, MAGR-2, was isolated from the Gulf of Riga, Baltic Sea. The stock culture was grown in a medium based on the F medium of Guillard & Ryther (1962). Two-week-old M. aeruginosa inocula cultures were centrifuged in sterile conic flasks. The supernatant was removed; packed cells were re-suspended in nutrient-free sterile sea water and left for 24 h in the culture room. Centrifugation and re-suspension were repeated, providing partial nutrient depletion of M. aeruginosa cells and removal of dissolved nutrients from the mother suspension. Cell suspension of M. aeruginosa (13 mL) was inoculated into two treatment series: Control and AB + DOM, providing an initial concentration of 18.7 x [10.sup.6] cells [L.sup.-1]. Enrichments of Fe-EDTA, metal, and vitamin mixtures were added.
For the preparation of bacterial inoculum, a fraction of M. aeruginosa mother culture was centrifuged in sterile conic flasks; then the supernatant was filtered through a Nucleopore membrane filter with 2 [micro]m pore size to remove algae cells. As the bacterial mother suspension 300 mL of filtrate, containing 1.4 x [10.sup.9] bacteria [L.sup.-1], was used. It was inoculated in filter sterilized sea water. The inoculum cell counting was done under epifluorescence illumination. Neither bacterial grazers nor autofluorescent algae were detected in the filtrate.
The Control and AB + DOM treatments were incubated at 18 [+ or -] 1[degrees]C with the photoperiod of 16 h light and 8 h dark at 53 [micro]M [m.sup.-2] [s.sup.-1] irradiance from cool white fluorescent lamps. The B + DOM treatment was incubated in the dark.
Bacteria and algae counting
Samples (5 mL) for M. aeruginosa cell counting (Neubaeur cell) were fixed with acid Lugol solution and counted using inverted Leitz microscope. For counting bacteria 15 mL of samples were preserved by using 0.2 [micro]m filtered formaldehyde (final concentration 2.5%). Aliquots of 5 mL were taken from each preserved sample and filtered onto black polycarbonate filters (pore size 0.2 [micro]m) and stained with DAPI (4',6-diamidino-2-phenylindole) to get a final concentration of 0.1 mg [mL.sup.-1]. Bacteria were counted using epifluorescence microscope Leitz Dialux fitted with a 100 x 1.25 objective and UV excitation light (Hobbie et al., 1977; Porter & Freig, 1980). Bacterial production was measured by uptake of tritiated thymidine (Fuhrman & Azam, 1980, modified by Smith & Azam, 1992).
Dissolved organic matter
Water for DOM extraction was collected in the Parnu River, upstream from the town of Parnu. A tangential flow ultrafilter was used for the extraction process (Guo et al., 1995; Benner et al., 1997). Water was pre-filtered applying 1.2 [micro]m and 0.2 [micro]m Opticap filter units (Millipore). Then DOM was concentrated with a tangential device, the Prep/scale[TM] TFF 6 [ft.sup.2] cartridge (Millipore). Total dissolved nitrogen (DN) and phosphorus concentrations were measured after the persulphate digestion procedure at 120[degrees]C (Pujo-Pay & Raimbault, 1994). The concentration of DON was calculated as the difference between total DN and DIN (N[O.sub.3.sup.-] + N[O.sub.2.sup.-] + N[H.sub.4.sup.+]), and dissolved organic phosphorus (DOP) as the difference between total dissolved phosphorus (DP) and phosphate-P concentrations. The concentration of DON in the DOM extract was 639 [micro]M, and the concentration of DOP was 5.14 [micro]M. The DOM extract also contained 49 [micro]M N[O.sub.3.sup.-] and 3.37 [micro]M P[O.sub.4.sup.3-].
Concentrations of inorganic nutrients were measured with a Skalar autoanalyser, following the standard procedures for seawater analyses (Valderrama, 1995). Samples for the analyses of particulate organic carbon (POC) and particulate organic nitrogen (PON) were collected using precombusted glass microfibre filters Whatman GF/F, decarbonated (HCl fumes, overnight), and analysed by applying a CHN analyser (model Carlo Erba 1500). Samples for particulate organic phosphorus (POP) analysis were collected using similar methods as for POC and PON. POP concentration was determined as P[O.sub.4.sup.3-] concentration using the persulphate digestion procedure at 120[degrees]C (Pujo-Pay & Raimbault, 1994). The initial nutrient concentrations in Control and AB + DOM were 8 [micro]M N[O.sub.3] + N[O.sub.2] and 0.4 [micro]M P[O.sub.4], in B + DOM treatment 10 [micro]M N[O.sub.3] + N[O.sub.2] and 0.4 [micro]M P[O.sub.4].
Analysis of variance (ANOVA) was used to test the significance of differences in M. aeruginosa and bacterial growth, biomass increase, and nutrient dynamics between the treatments under different experimental conditions.
The growth of M. aeruginosa started without a lag phase. A certain number of M. aeruginosa were in the exponential growth phase till Day 6 in both treatments with cyanobacteria added, reaching higher values in the treatment with DOM addition (AB + DOM, 0.74 x [10.sup.9] cells [L.sup.-1]) than in Control (0.54 x [10.sup.9] cells [L.sup.-1]). On Day 6 M. aeruginosa entered the stationary phase in Control, whereas in the AB + DOM treatment, DOM additions facilitated a continuous growth of cell concentration, which reached a maximum on Day 13 (1.02 x [10.sup.9] cells [L.sup.-1]) (Fig. 1). Contrary to the instantaneous growth of M. aeruginosa cell numbers, in vivo fluorescence started to increase after a 2 days long pause. In Control the in vivo fluorescence increased until Day 10, but in AB + DOM fluorescence continued a gradual increase until the end of the experiment. The cell numbers of M. aeruginosa and in vivo fluorescence had higher values in the AB + DOM treatment than in Control (p < 0.05) (Fig. 1).
The Heterotrophic bacterial community grew quickly in all treatments, reaching maximal values of bacterial production on days 3 and 5, whereas cell numbers reached the peak 2 to 3 days later, on days 5 and 7 (Fig.
1). After five incubation days similar values of total bacterial numbers (TBN) and bacterial biomass (BB) were registered in both treatments with DOM addition irrespective of the presence or absence of M. aeruginosa (2.37 x [10.sup.9] and 2.58 x [10.sup.9] bacteria cells [L.sup.-1]; 0.37 and 0.38 mgC [L.sup.-1] in AB + DOM and B + DOM, respectively). The heterotrophic bacterial community in the AB + DOM and B + DOM treatments remained in the stationary phase until Day 9, followed by a decline. The dynamics of heterotrophic bacteria was similar in the AB + DOM and B + DOM treatments (ANOVA, p = 0.98, n = 21). Similarity in the fluctuation of the bacterial number curves both in the presence and absence of cyanobacteria indicates that the growth of associated bacteria was not suppressed owing to M. aeruginosa absence in the B + DOM treatment. Bacterial production (BP) demonstrated similar shapes (r = 0.81, n = 63) (data not presented) to the fluctuation of TBN.
[FIGURE 1 OMITTED]
No or an insignificant amount of grazers of heterotrophic bacteria were found in all treatments until Day 12. Due to its suppressing influence on heterotrophic flagellates, M. aeruginosa manifested itself during the exponential and stationary growth phases. In the bacterial treatment, flagellates started their rapid development after Day 12.
A few cells of M. aeruginosa were observed in the bacterial treatment, but their numbers as well as fluorescence did not increase because of the lack of light. We considered that there was no influence of algae and grazers in the B + DOM treatment until Day 12. Therefore the consumption or release of nutrients was calculated from the first day after the interruption of DOM addition, between Day 7 and Day 12, when grazers of bacteria started to develop.
Nitrates and nitrites were used fast in both treatments where M. aeruginosa was present (Fig. 2). In Control and AB + DOM the concentration of N[O.sub.3.sup.-] was very low (0.7 [micro]M) on Day 7 and fell below detection limits at the end of the experiment. In the bacterial treatment B + DOM the consumption of nitrates was observed during the exponential growth phase. At the end of the stationary phase nitrites and nitrates were released (Fig. 2). At first, the concentration of nitrites increased reaching a maximum on Day 9 (0.35 [micro]M), a peak of nitrates followed on Day 12 (4.45 [micro]M). Altogether 2.46 [micro]M of nitrates was released until Day 12 (B + DOM) (Table 1).
Till the end of the experiment a slow and gradual accumulation of ammonia took place in Control, indicating a gradual decay of organisms without additional nutrient supply (Table 1, Fig. 2). In the AB + DOM treatment a slight increase in the ammonia concentration occurred on Day 12. In the bacterial treatment (B + DOM) both the release and consumption of ammonia occurred, with the concentration of ammonia doubling on Day 9, followed by partial depletion on Day 12.
[FIGURE 2 OMITTED]
As DOM additions ended on Day 6, the cultures evolved as batch cultures. Dissolved organic nitrogen (DON) was depleted in Control, but in the AB + DOM treatment its concentration was enriched by cyanobacteria by 13%, whereas bacteria alone managed to reduce the bulk of DON by 53% (Table 1, Fig. 2). A similar pattern was observed for DN: in Control 60% of DN was used up, but in the AB + DOM treatment a 34% increase was observed; heterotrophic bacteria alone used 10% of DN (Table 1).
The addition of DOM increased biomass indices, such as particulate organic carbon (POC), particulate organic nitrogen (PON), and particulate organic phosphate (POP) concentrations in AB + DOM versus un-enriched Control (p < 0.05). In the AB + DOM treatment the increase of POC content was 36%, that of PON 18%, but of POP over 100%. The added DOM prompted a decrease of the PON : POP ratio of M. aeruginosa by 24%, changing the content of cyanobacteria cells from P-limited (PON : POP = 21.1) to N : P-balanced (PON : POP = 15.6). The POC : PON ratio of M. aeruginosa cells rose from 11.9 on Day 6 to 20.1 on Day 12 (Table 2). We therefore conclude that M. aeruginosa took up proportionally more C and P than N, and the surplus N was released as DON.
Heterotrophic bacteria used in our experiment live freely in water or are attached to aggregates of M. aeruginosa, utilizing allochthonous organic matter brought by rivers and/or autochthonous organic matter released by cyanobacteria. The fact that these heterotrophic bacteria reduced the riverine DON to half of its initial concentration indicates that they are adapted to this kind of nutrient substrate. Our results showed that heterotrophic bacteria associated with M. aeruginosa in the presence of riverine DOM in the treatments conducted with and without autotrophic cyanobacteria had similar growth with similar numbers and production curves. This is an evidence of poor advantages of the presence of photosynthetic extracellular products from algae. Hence, bacteria living in eutrophic brackish waters with cyanobacterial aggregates have a high capability to mineralize riverine organic matter.
As a result of DON mineralization by heterotrophic bacteria, inorganic N compounds were released. It is noteworthy that during the elimination of reduced nitrogen (ammonia) oxidized compounds of N were produced. The total concentration of nitrites increased by 186% and that of nitrates by 82% (Table 1). During the stationary phase ammonia was produced by bacteria, with a decline following. Goldman & Dennett (1991) concluded that elimination of N[H.sub.4.sup.+] in the marine environment occurs when the nitrogen source is an organic compound (e.g. aminoacids, proteins, etc.). Tupas & Koike (1991) showed that bacteria can mineralize DON to N[H.sub.4.sup.+] even when the net N[H.sub.4.sup.+] uptake by bacterial assemblages is high and when C : N models predict no net production of ammonium. Our results indicated the capabilities of heterotrophic bacteria separated from M. aeruginosa to eliminate ammonia, nitrates, and nitrites when maximal biomass of bacteria cells was reached and the convenient substrate was depleted. Ammonium release by bacteria occurs commonly in marine and freshwater environments while the simultaneous release of nitrites or nitrates is poorly studied. Berman & Chava (1999) expressed an opinion that bacterial degradation of organic nitrogen will lead to the release of N[H.sub.4.sup.+] and/or urea and perhaps even N[O.sub.3.sup.-], which would be subsequently utilized by phytoplankton. Our findings supported this hypothesis.
Microcystis aeruginosa does not belong to the N-fixing cyanobacteria. It is estimated that the best nitrogen sources for M. aeruginosa are nitrate and ammonium; besides, Berman & Chava (1999) reported its good development sustained by urea. Maestrini et al. (1999) found that addition of urea, glycine, hypoxantine, putrescine, and spermine as the single source of nitrogen significantly provokes the growth of the M. aeruginosa culture, and dissolved organic matter from the Daugava River sustains the growth of cyanobacterial biomass at a level of 38% of that accrued from nitrate. Since the cultures of autotrophic cyanobacteria used in the foregoing experiments were not axenic, the contribution of heterotrophic bacteria in the mineralization of DON substances is not known. In our experiment the cyanobacteria could obviously benefit from nitrogen mineralization by heterotrophic bacteria or could utilize DON themselves, as utilization of organic nitrogen by cyanobacteria becomes significant during the summer season in brackish waters when DIN resources are depleted and DON bioavailability increases (Stepanauskas et al., 1999). However, 70% of DN transported by rivers worldwide, as well as to the Gulf of Riga, is DON (Meybeck, 1982; Laznik et al., 1999; Stalnacke et al., 1999). The primary production in marine and fresh waters is often limited because of low bioavailability of nitrogen. The estimated range of bioavailable DON in the Baltic Sea is broad, between 8% and 72% (Stepanauskas et al., 2002). In our case, after the bacterial growth bioassay, 53% of DON from the Parnu River could be characterized as biologically labile, and bacterially mediated final products were the main nutrients determining the growth of M. aeruginosa.
Besides nitrogen uptake by M. aeruginosa, the excretion of DON occurred, amounting to 0.54 [micro]M (between Day 7 and Day 12). In some studies the excretion of organic matter by phytoplankton is considered as a continuous process (Larsson & Hagstrom, 1979), but others suggest that significant excretion is a response to environmental stress, such as changes in the quantity and quality of light (Zlotnik & Dubinsky, 1989) or nutrient limitation (Myklestad & Haug, 1972; Lancelot, 1983; Obernosterer & Herndl, 1995). Obernosterer & Herndl (1995) hypothesized that large amounts of photosynthetic extracellular release (PER) from phytoplankton occur under severely (N : P = 100) P-limited primary production. In our case the release of DON by cyanobacteria can be explained by changing cell PON and POP content from P-limited (PON : POP = 21.5) at the beginning of the experiment to N : P balanced (PON : POP = 15.6) (AB + DOM treatment, Table 2). Altogether the PON : POP ratio of M. aeruginosa cells decreased by 27%. In total, DOM sustained the relative increase of the PON concentration by 18%, while POP increased by 103%. Therefore we can conclude that M. aeruginosa took up more P in relation to N from DOM with a subsequent release of surplus N.
Our findings support the hypothesis that bacteria associated with the cyanobacteria species Microcystis aeruginosa isolated from brackish waters can release ammonia, nitrite, and nitrate as a result of DON mineralization providing M. aeruginosa with simple and ready to use nitrogen compounds.
This study was supported by the European Community Project 'DOMTOX' (MAS3-CT97-0149) and the European Social Fund. Particular thanks to colleagues from the Estonian Marine Institute at the University of Tartu for help in water collection from the Parnu River and M. Pfeifere (Latvian Institute of Aquatic Ecology) for the isolation of the M. aeruginosa culture.
Received 16 May 2008, revised 19 March 2010 and 15 June 2010
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Santa Purvina (a) [email], Christian Bechemin (b), Maija Balode (a), Celine Verite (b), Christophe Arnaud (b), and Serge Y. Maestrini (b)
(a) Latvian Institute of Aquatic Ecology, Daugavgrivas Str. 8, LV-1048 Riga, Latvia
(b) Centre de Recherche en Ecologie Marine et Aquaculture de L'Houmeau (CNRS-IFREMER), B.P. 5, F-17137 L'Houmeau, France
[email] Corresponding author, email@example.com
Table 1. Concentration of dissolved inorganic nitrogen (N[H.sub.4], N[O.sub.2], N[O.sub.3]), dissolved organic nitrogen (DON), and dissolved nitrogen (DN) on Day 7 and uptake (indicated by -) or release (+) of nutrients between Day 7 and Day 12 Treatment N[H.sub.4] Control Concentr., [micro]M 0.59 [+ or -] 0.15 Uptake/release, % +69 AB + DOM Concentr., [micro]M 0.72 [+ or -] 0.05 Uptake/release, % +145 B + DOM Concentr., [micro]M 0.91 [+ or -] 0.06 Uptake/release, % -17 Treatment N[O.sub.2] Control Concentr., [micro]M 0.07 [+ or -] 0.02 Uptake/release, % 0 AB + DOM Concentr., [micro]M 0.05 [+ or -] 0.00 Uptake/release, % +212 B + DOM Concentr., [micro]M 0.07 [+ or -] 0.00 Uptake/release, % +186 Treatment N[O.sub.3] Control Concentr., [micro]M 0.77 [+ or -] 0.3 Uptake/release, % -54 AB + DOM Concentr., [micro]M 0.73 [+ or -] 0.02 Uptake/release, % -62 B + DOM Concentr., [micro]M 2.44 [+ or -] 0.70 Uptake/release, % +82 Treatment DON Control Concentr., [micro]M 1.31 [+ or -] 0.28 Uptake/release, % -100 AB + DOM Concentr., [micro]M 4.20 [+ or -] 0.48 Uptake/release, % +13 B + DOM Concentr., [micro]M 5.47 [+ or -] 0.36 Uptake/release, % -53 Treatment DN Control Concentr., [micro]M 2.73 [+ or -] 0.22 Uptake/release, % -60 AB + DOM Concentr., [micro]M 5.70 [+ or -] 0.52 Uptake/release, % +34 B + DOM Concentr., [micro]M 8.88 [+ or -] 0.51 Uptake/release, % -10 Table 2. Mean particulate organic nitrogen (PON), particulate organic phosphorus (POP), and particulate organic carbon (POC) concentrations and POC : PON and PON : POP ratios (atom : atom) recorded on Day 12 in different treatments. Numbers within parentheses show the relative biomass increase due to the addition of DOM (%) Treatment PON, [micro]M POP, [micro]M Control 14.3 [+ or -] 1.3 0.53 [+ or -] 0.23 AB + DOM 16.9 [+ or -] 1.7 (18%) 1.08 [+ or -] 0.17 (103%) B + DOM 3.7 [+ or -] 0.7 0.35 [+ or -] 0.1 Treatment POC, [micro]M POC : PON Control 250 [+ or -] 23 17.5 [+ or -] 0.1 AB + DOM 340 [+ or -] 37 (36%) 20.1 [+ or -] 0.3 B + DOM 29.0 [+ or -] 4.6 7.8 [+ or -] 0.2 Treatment PON : POP Control 26.6 [+ or -] 8.3 AB + DOM 15.6 [+ or -] 1.7 B + DOM 10.6 [+ or -] 1.6
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|Author:||Purvina, Santa; Bechemin, Christian; Balode, Maija; Verite, Celine; Arnaud, Christophe; Maestrini, S|
|Publication:||Estonian Journal of Ecology|
|Date:||Sep 1, 2010|
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