Mar Biol (2012) 159:2389–2398 DOI 10.1007/s00227-012-1927-4
ORIGINAL PAPER
Microbial interactions with the cyanobacterium Microcystis aeruginosa and their dependence on temperature Claudia Dziallas · Hans-Peter Grossart
Received: 2 October 2011 / Accepted: 13 March 2012 / Published online: 5 April 2012 © Springer-Verlag 2012
Abstract Associated heterotrophic bacteria alter the microenvironment of cyanobacteria and potentially inXuence cyanobacterial development. Therefore, we studied interactions of the unicellular freshwater cyanobacterium Microcystis aeruginosa with heterotrophic bacteria. The associated bacterial community was greatly driven by temperature as seen by DNA Wngerprinting. However, the associated microbes also closely interacted with the cyanobacteria indicating changing ecological consequence of the associated bacterial community with temperature. Whereas concentration of dissolved organic carbon in cyanobacterial cultures changed in a temperature-dependent manner, its quality greatly varied under the same environmental conditions, but with diVerent associated bacterial communities. Furthermore, temperature aVected quantity and quality of cell-bound microcystins, whereby interactions between M. aeruginosa and their associated commu-
Communicated by U. Sommer. Electronic supplementary material The online version of this article (doi:10.1007/s00227-012-1927-4) contains supplementary material, which is available to authorized users. C. Dziallas · H.-P. Grossart (&) Department of Limnology of StratiWed Lakes, Leibniz-Institute for Freshwater Ecology and Inland Fisheries, Alte Fischerhuette 2, 16775 Stechlin, Germany e-mail:
[email protected] Present Address: C. Dziallas Marine Biological Section, Department of Biology, University of Copenhagen, Strandpromenaden 5, 3000 Helsingør, Denmark H.-P. Grossart Institute for Biochemistry and Biology, Potsdam University, Am Neuen Palais 10, 14469 Potsdam, Germany
nity often masked this temperature eVect. Both macro- and microenvironment of active cyanobacterial strains were characterized by high pH and oxygen values creating a unique habitat that potentially aVects microbial diversity and function. For example, archaea including ‘anaerobic’ methanogens contributed to the associated microbial community. This implies so far uncharacterized interactions between Microcystis aeruginosa and its associated prokaryotic community, which has unknown ecological consequences in a climatically changing world.
Introduction In many regions, cyanobacterial abundance increased in fresh and brackish waters over the last decades (Hudnell and Dortch 2008), often leading to mass developments with sometimes detrimental eVects on animals and potentially on human health. Massive cyanobacterial blooms impair the use of water bodies for recreation, for watering animals and for abstracting drinking water. The latter is particularly relevant in regions where treatment technology is not available or eVective. Therefore, cyanobacterial blooms may be a hazard to human health and cause kills of livestock or wildlife. It is of great socio-economical interest to understand the underlying mechanisms for such cyanobacterial mass developments. Paerl and Huisman (2008) concluded that climate change leading to higher surface water temperatures as well as eutrophication favour growth and toxin production of cyanobacteria, in particular of Microcystis aeruginosa (Rouco et al. 2011). However, the ecological role of cyanobacterial toxins such as microcystins is still not fully understood. A proposed function is allelopathy against other photoautotrophs (Sukenik et al. 2002), but this suggestion was rejected due to the unexplained diversity of
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produced toxin variants that require additional energy from the cells (Welker et al. 2007). Previously, structural variations have been regarded as a response in the course of coevolution towards resistance of consumers to certain molecules (Futuyama 1983), which argues for the possibility of allelopathic implications. As genes for microcystin synthetase seem to have evolved long before the presence of metazoans on earth (Rantala et al., 2004), protection from grazing (e.g. Ghadouani et al. 2004; Czarnecki et al. 2006) is unlikely the primary function of microcystins and other cyanobacterial toxins. Recently proposed functions of microcystins include protein modulation (Zilliges et al. 2011), scavenging of oxygen radicals (Dziallas and Grossart 2011b) and involvement in photosynthesis (Jähnichen et al. 2001, 2011). Although it is known that associated bacteria can enhance or decrease the growth of cyanobacteria and algae in culture (Paerl and Millie 1996; Salomon et al. 2003), they have been largely overlooked as a driver for cyanobacterial growth and toxicity. Hence, we propose that community composition of associated bacteria alter cyanobacterial growth as well as quality and quantity of produced cyanobacterial toxins. Associated bacteria interact with phototrophic cyanobacteria in manifold ways including exudate remineralization (Kirkwood et al. 2006), toxin degradation or production (Maruyama et al. 2003; Dziallas and Grossart 2011a), respiration increasing carbon dioxide availability (Kühl et al. 1996) and production of growth factors such as vitamins (Paerl and Pinckney 1996). On the other hand, cyanobacteria provide a unique microhabitat for associated microorganisms with increased concentrations of organic substrates including toxins (Cole 1982; Worm and Sondergaard 1998; Nicolaus et al. 1999). Additionally, cyanobacteria due to their gas vacuoles are able to stay in distinct water depths with relatively stable environmental variables and are rather resistant against predation due to mucilage and colony size (Casamatta and Wickstrom 2000). So far, it is largely unclear whether cyanobacteria can also provide protection of bacteria from phages. Microbial interactions with cyanobacteria, however, are poorly understood due to their complex nature, but it has been shown that they have the potential to control bloom development of potentially toxic cyanobacteria (Dziallas and Grossart 2011b). The ongoing climate change may aVect the availability of nutrients and hence cyanobacterial growth and toxin production in various ways, for example, due to changes in lake stratiWcation patterns (De Stasio et al. 1996). Both changes in temperature and nutrient availability have important consequences for microorganisms associated to cyanobacteria (Dziallas and Grossart 2011a). Accordingly, we propose that associated bacterial community composition depends on temperature even in well established and long-lasting interactions as in cultured cyanobacteria.
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Additionally, not only bacteria but also fungi (O⬘Brien et al. 2005) and archaea (Dziallas and Grossart 2011a) are associated to cyanobacteria. In particular, archaea belong to a phylogenetic group that has been assigned to anaerobic or extreme environments (DeLong 1992) and, thus, have been relatively little studied in oxic environments. Recently, methanogenic archaea have been also reported from the oxygenated water column (Grossart et al. 2011), where they can be of biogeochemical importance. In the present study, we frequently found archaea in cyanobacterial cultures indicating that they are a common constituent of microbial communities associated to cyanobacteria. All associated microorganisms are situated in or around the so-called phycosphere (Cole 1982), a microhabitat characterized by steep physicochemical gradients, for example, of organic matter concentrations. Hence, to better characterize the cyanobacterial phycosphere, we have measured gradients of important environmental parameters such as oxygen concentration and pH with microelectrodes. Additionally, exudates from cyanobacteria were analysed for dissolved organic carbon (DOC), toxins and amino acids. In particular, amino acids are of interest since they contain high amounts of nitrogen and hence represent high nutritious substrates for associated microbes. Therefore, we propose that cyanobacteria provide unique microenvironments for interactions with associated microorganisms and that these have potential eVects not only for cyanobacterial bloom development and toxicity but also for biogeochemical processes. Since cyanobacteria strongly respond to changes in temperature, the cyanobacterial microenvironments for associated bacteria will be altered by increasing temperatures. Thus, cyanobacterial growth and physiology will be aVected via global climate change by direct and indirect changes in dynamics of associated microorganisms. Therefore, the following four hypotheses were tested (1) incubation temperature alters bacterial communities associated to cyanobacteria, (2) amount and composition of cyanobacterial exudates depend on incubation temperature, (3) associated microbes impact quantity and quality of cell-bound toxins and (4) cyanobacterial phycospheres vary in a temperature- and light-dependent manner.
Materials and methods Experimental design Temperature eVects To test whether bacterial communities in long-established cyanobacterial cultures change with incubation temperatures, we tested three xenic cultures at three diVerent
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of the genus Pimelobacter (Actinobacteria) and (3) natural bacteria from Lake Stechlin, Northeast Germany. Five millilitre of the cyanobacterial culture was lyophilized, extracted with 75 % methanol and measured with a RP-HPLC (Agilent) after the protocol of Lawton et al. (1994) and Fastner et al. (1998).
temperatures (20, 26 and 32 °C—the range of present and expected summer temperatures in temperate lakes). The strains HUB P323 (isolated from Lake Pehlitz, Germany, in 1995), AB2002/21 (isolated from Nakuru Wnal sewage point, Kenya, in 2002) and AB2005/47 (isolated from Lake Taihu, China, in 2005) were grown until their exponential growth phase. Then cells were counted and transferred to 50 ml of Z medium (Zehnder and Gorham 1960) in Erlenmeyer Xasks. The inoculum contained ca. 1 £ 105 cyanobacterial cells per ml. These cultures were grown in triplicates at the three diVerent temperatures for 23 days with a 16:8 h light/dark cycle and a light intensity of 72 E. On day 17, in late exponential growth phase, 5 ml of culture was removed and Wltered Wrst on a polycarbonate Wlter with a pore size of 3.0 m for attached microbes and subsequently on a Wlter with 0.2 m pore size for free-living microbes and stored at ¡20°C until DNA extraction. The pH was measured each day with a pH electrode (WTW). Cyanobacterial cells were counted each third day with a FACSAria II Xow cytometer.
As cyanobacterial aggregates and scums can diVer greatly on a small scale, we measured oxygen and pH gradients in the aggregates and scums with microelectrodes. This should also indicate the presence of anaerobic bacteria and archaea. The same cultures as used for the growth experiment were incubated at 24 °C for 15 days. Afterwards, Xoating scums and aggregates obtained in a roller tank were measured at diVerent light intensities (ranging from 2.1 to 129 E) with microelectrodes (PreSens) for oxygen and pH. Measurements started at the surface of the scum or aggregate and proceeded in 100 m steps.
Exudate quantity and quality
Studies on bacterial community composition (BCC)
Quality and amount of exudates can have strong eVects on associated bacterial communities. Therefore, we investigated the exudates of two strains of M. aeruginosa incubated at three diVerent incubation temperatures. Cyanobacterial strains were checked for being axenic by epiXuorescence microscopy after staining with DAPI and SybrGold, showing no further microbes than cyanobacteria. Axenic strains of the toxic strain HUB 5-2-4 (isolated from Lake Pehlitz, Germany, in 1977) and the non-toxic strain HUB P461 (isolated from Lake Pehlitz, Germany, in 1995) were incubated in Wve replicates for 4 weeks at 20, 26 and 32 °C. After these 4 weeks, all treatments were in stationary growth phase with 2.5 £ 107 cells per ml culture. The incubation medium with exudates was Wltered over 3.0 m to get rid of cyanobacterial cells, and thereafter, the Wltrate was tested for DOC and transparent exopolymeric particles (TEPs). DOC was measured with a Shimadzu DOC analyzer, and TEPs were microscopically counted and measured after Logan et al. (1994).
Fluorescence in situ hybridization (FISH)
Characterization of the cyanobacterial microenvironment
To quantify the dominating groups of associated microorganisms, we conducted FISH or catabolized reporter deposition (CARD-) FISH. FISH with FITC-labelled probes was performed after Manz et al. (1992) and CARD-FISH after Sekar et al. (2003). DNA extraction, PCR, DGGE DNA was extracted after Zhou et al. (1996) with zirconium beads and phenol–chloroform. PCR for bacteria was done as described by Dziallas and Grossart (2011a) with primers 341f-GC and 907r (Muyzer et al. 1993, Teske et al. 1996). Afterwards, ca. 500 ng of ampliWed product was loaded on a lane of a 40–70 % DGGE gel, which was run for 20 h with 100 V. The DGGE gels were stained with SybrGold, shortly destained and photographed with an AlphaImager (BioRad).
Microcystin experiment Cluster analysis and statistics As speciWc strains of M. aeruginosa are often able to produce more than one speciWc variant of the cyanobacterial toxin microcystin, we wanted to investigate whether the associated bacterial community can potentially aVect toxin quality. Therefore, the axenic and toxic strain HUB W333 (isolated from Lake Wannsee, Germany, in 1995) was incubated at 20, 26 and 32 °C for 19 days in triplicates. In addition, this strain was inoculated with (1) an isolate of the genus Sphingomonas (Alphaproteobacteria), (2) an isolate
Pictures of DGGE gels were analysed with GelCompar II using Dice coeYcient. The similarity matrices were loaded into Primer6 and tested with two-way cross-analysis of similarities (ANOSIM) and non-metric multidimensional scaling (MDS). MDS is a method to show similarities between samples in a two- or three-dimensional manner. Further two-way ANOVA was done (Holliday 2011) with a signiWcance level of 0.05.
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Results Temperature eVects on cyanobacterial growth and bacterial community composition (BCC) Incubation of three M. aeruginosa strains from Germany, China and Kenya at diVerent temperatures (20, 26 and 32 °C) revealed the same maximum cell numbers at all incubation temperatures (Suppl. Fig. 1). However, signiWcant temperature-dependent diVerences in BCC (20, 26 and 32 °C) were obvious on day 17 at the end of the exponential growth phase of the cyanobacterium (Table 1; Fig. 1). For each strain, diVerences in BCC of diVerent temperature regimes were usually more pronounced than between replicate experiments of a single temperature regime. However, the variability between replicates was surprisingly high. In all samples, attached and free-living bacteria signiWcantly diVered from each other. Each sample consisted of 20–25 DGGE bands, whereby each band should stand for one bacterial genotype representing at least 1% of the whole associated bacterial community (Muyzer et al. 1993). Sequencing of DGGE bands revealed that Bacteriodetes and Alphaproteobacteria were both free-living and attached. Whereas Gammaproteobacteria were always attached, Betaproteobacteria were mainly free-living. This holds true for all three investigated cyanobacterial strains at all temperatures. Changes in BCC are apparently driven by diVerences in environmental factors such as pH, oxygen concentration, nutrient availability or
cyanobacterial cell density and temperature itself. Indirect eVects of environmental factors on BCC include dissolved compounds from lysed cells and excreted polymers (e.g. peptides) from growing cyanobacterial cells. Temperature eVects on exudate quantity and quality Incubation of a toxic (HUB 5-2-4) and a non-toxic (HUB P461) M. aeruginosa strains at 20, 26 and 32 °C for 4 weeks (stationary growth phase) was performed to test whether they show the same temperature-dependent exudation pattern. Whereas the toxic strain did not show any temperature-dependent changes in concentrations of DOC, the non-toxic strain showed a DOC increase with temperature from 9.29 § 1.44 mg l¡1 at 20°C to 17.95 § 1.76 mg l¡1 at 32°C (Suppl. Fig. 2), whereby the number of cyanobacterial cells did not diVer with temperature at the end of the experiment. In contrast, amount and composition of amino acids showed no clear temperature pattern and were highly variable between replicates (data not shown). The production of transparent exopolymeric particles was also independent of temperature for both toxic and non-toxic M. aeruginosa strains (Suppl. Fig. 3). EVects of BCC on amount of measurable microcystins Microcystins were detected in exudates only when toxic cyanobacterial strains were incubated at 20 °C (HUB
Table 1 Results of A) two-way cross-ANOSIM analyses and B) two-way ANOVA and subsequent Post hoc test on DGGE banding pattern with bacteria-speciWc primers in late exponential growth phase Cyanobacteria strain
Free-living versus attached
Temperature
AB2005/47
R = 0.96, p = 0.001
R = 0.66, p = 0.001
20 versus 26°C: R = 0.73, p = 0.01 20 versus 32°C: R = 0.81, p = 0.01 26 versus 32°C: R = 0.62, p = 0.02
AB2002/21
R = 0.45, p = 0.12
R = 0.49, p = 0.002
20 versus 26°C: R = 0.49, p = 0.03 20 versus 32°C: R = 0.43, p = 0.04 26 versus 32°C: R = 0.19, p = 0.16
HUB P323
R = 0.98, p = 0.003
R = 0.75, p = 0.001
20 versus 26°C: R = 0.33, p = 0.05 20 versus 32°C: R = 0.96, p = 0.01 26 versus 32°C: R = 0.82, p = 0.01
(A)
Cyanobacteria strain
Free-living versus attached
Temperature
Combined
Post hoc (Tukey)
AB2005/47
F = 100.4, p < 0.001
F = 24.2, p < 0.001
F = 2.9, p = 0.085
20 versus 26°C: p < 0.001 20 versus 32°C: p = 0.002 26 versus 32°C: p = 0.032
AB2002/21
F = 2.8, p = 0.110
F = 11.2, p = 0.001
F = 1.7, p = 0.206
20 versus 26°C: p = 0.001 20 versus 32°C: p = 0.118 26 versus 32°C: p = 0.039
HUB P323
F = 9.1, p = 0.007
F = 3.0, p = 0.072
F = 0.5, p = 0.588
20 versus 26°C: p = 0.071 20 versus 32°C: p = 0.623 26 versus 32°C: p = 0.294
(B)
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Fig. 1 MDS of the associated bacterial DGGE banding pattern of three strains of M. aeruginosa incubated at 20, 26 and 32 °C for 17 days. a Strain AB2005/47 isolated from Lake Taihu, China; b Strain AB2002/21 isolated from Nakuru Wnal sewage point, Kenya; c HUB P323 isolated from Lake Pehlitz, Germany
5-2-4). Amount and quality of cell-bound microcystin in cultures of the axenic, toxic strain (HUB W333), however, increased with temperature (Fig. 2a). Comparison of the same strain incubated with diVerent bacterial communities revealed that the presence of a single bacterial isolate or of a natural bacterial community from a lake result in changing amount and quality and hence toxicity of the produced microcystin (Fig. 2b–d, Suppl. Tab. 1). At 32 °C, presence of a single bacterial isolate or a whole microbial community from a lake led to a reduction in the cell-speciWc amount of the less toxic microcystin variant MC-YR (Suppl. Fig. 4A). This reduction was less pronounced for the more toxic variant MC-LR (Suppl. Fig. 4B). When microcystin concentrations were expressed per active transcribing microcystin synthetase gene D, the pattern was similar for the less toxic variant MC-YR (Suppl. Fig. 4C), but amounts of the more toxic variant MC-LR in cultures with the Pimelobacter isolate were as high as in the axenic culture (Suppl. Fig. 4D). Changes in the cyanobacterial phycosphere in dependence on temperature and light Temperature-dependent changes in pH during cyanobacterial growth were found in cultures of three strains of M. aeruginosa (AB2005/47, AB2002/21, HUB P323) incubated at 20, 26 and 32 °C (Fig. 3). These diVerences were most distinct during the exponential growth phase and equal at the beginning as well as in the stationary growth phase. Our pH measurements correlate well with cyanobacterial cell counts (Suppl. Fig. 1), indicating a relationship between cyanobacterial cell density and pH. However, the
dip in the pH curves after the Wrst maximum is not visible for the cell counts. To elucidate potential impacts of the cyanobacterial phycosphere on bacterial dynamics, we measured gradients of oxygen and pH in the scums of the M. aeruginosa strain AB2005/47 from Lake Taihu, China, during the end of its exponential growth phase (Fig. 4). Gradients of both parameters showed a strong light-dependent pattern, whereby maximal oxygen as well as pH concentrations was observed at highest light intensities in the centre of the scum—the most distant point from the edge of the scum. This strain has been kept in culture for 2 years, but still showed colony formation as well as intact gas vacuoles. Scum thickness varied between 3 and 4.5 mm in this xenic strain consisting mostly of large colonies (>300 m as deWned by Ibelings and Mur 1992). Further observations The results of the microelectrode measurements disagree with our Wndings of potentially anaerobic bacteria (sulphate reducers) and methanogenic archaea by speciWc PCR in cultures of M. aeruginosa (Table 2). In addition to these functional groups, we could also detect bacteria and archaea of other functional groups that also or exclusively occur in oxic environments. Interestingly, we could Wnd Crenarcheota that even represented over 50 % of all associated microorganisms in one M. aeruginosa culture during late exponential growth. However, in most cultures, the majority of associated microbes consisted of Alpha- and Betaproteobacteria. There were some exceptions where also Actinobacteria dominated.
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Fig. 2 Microcystins of the strain HUB W333 of M. aeruginosa incubated for 4 weeks at 20, 26 and 32 °C. a Axenic culture; b culture with a Sphingomonas sp. isolate; c culture with a Pimelobacter sp. isolate; d culture with a mixture of bacteria from Lake Stechlin. For more information on a and d, please see also Dziallas and Grossart (2011b)
Fig. 3 pH in cultures of three strains of M. aeruginosa incubated at 20, 26 and 32 °C. a Strain AB2005/47 isolated from Lake Taihu, China; b strain AB2002/21 isolated from Nakuru Wnal sewage point, Kenya; c HUB P323 isolated from Lake Pehlitz, Germany
Discussion Interestingly, all axenic M. aeruginosa strains kept their morphology. Two of them (HUB W333 and HUB P461) formed even larger colonies when growing without heterotrophic bacteria. Achieving axenic cultures is not trivial and can require months due to a delayed growth response. However, the same maximal cell numbers as for xenic
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strains can be reached in the stationary growth phase. In addition, gas vacuoles were still present and intact, which was true for axenic as well as xenic strains. This observation shows that cyanobacteria have a growth advantage when kept together with heterotrophic bacteria—at least in many cases—but do not existentially need them. The fact that we did not manage to axenize all our cyanobacterial strains suggests that cyanobacteria and heterotrophic bacteria
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2395 Table 2 Detected bacterial and archaeal groups and functions in cultures of M. aeruginosa by PCR and Xuorescence in situ hybridization (FISH)
Bacteria Alphaproteobacteria
Detected by speciWc PCR
Detected by FISH
+
+
+
+
Betaproteobacteria
+
Gammaproteobacteria
+
Planctomycetes
+
Actinobacteria
+
Cytophaga–Flavobacteria– Bacteroides Sphingomonades
+ + +
¡
Sulphate reducers
+
+
Ammonium-oxidizing bacteria
+
¡ and +
Denitrifying bacteria
¡
¡
Nitrogen-Wxing bacteria
¡
+
+
Euryarcheota
Temperature aVects bacterial communities associated to cyanobacteria Associated bacterial communities of three investigated M. aeruginosa strains clearly varied with incubation temperature. Thereby, diVerences between BCC of the tested cultures were highest between incubations at 20 °C and at 26 and 32 °C. Thus, temperature seems to have a severe impact on BCC, but also has an eVect on the tested cyanobacteria, for example, exudation. It has been shown earlier that bacterial communities associated with cyanobacteria change with incubation temperature when inoculating axenic strains with natural lake bacteria (Dziallas and Grossart 2011a). In addition, associated bacteria of the present study were adapted for a long time to culture conditions at 24 °C,
+ +
Crenarcheota
are closely linked to each other. In addition, cyanobacterial strains diVer in their sensitivity to lysozyme treatment, which is together with physical treatment the key to our axenization procedure and hence greatly aVect the axenization success rate.
+
Methane oxidizers
Archaea
Fig. 4 Oxygen saturation and pH in a Xoating scum of AB2005/47 isolated from Lake Taihu, China. a Oxygen saturation after 8 h of incubation at four light intensities; b pH after 8 h of incubation at three light intensities
Function is regarded as anoxic
+
Methanogens
+
+
Ammonium-oxidizing archaea
+
¡
No sign indicates that these groups or functions were not tested with the respective method. Detected functions that are regarded as anoxic processes are indicated in the last row
but still change in composition when incubated at diVerent temperatures under the same culture conditions. This result has important implications for cyanobacterial cultivation, as some bacteria favour and other hinder growth of cyanobacterial cells (Salomon et al. 2003). Taking the natural temperature regime into account, this implies that the associated bacterial community during cyanobacterial bloom events can signiWcantly change over time and also aVect cyanobacterial growth. The most signiWcant diVerence was observed in our experiment for strain AB2005/47 from Lake Taihu, China, which has been kept in culture for only 2 years before the experiments. Thus, it may host a more variable associated bacterial community. Consequently, diVerences in cyanobacterial growth can be related directly to temperature, but also indirectly to alterations of the associated BCC. Cyanobacterial exudates and cell-bound microcystins depend on temperature and BCC Whereas amount of DOC in one cyanobacterial culture depended on temperature, DOC quality was highly variable
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and did not show any relationship to temperature. In contrast, amount and composition of cell-bound microcystins were clearly temperature dependent in axenic M. aeruginosa cultures. DOC concentration and amount as well as quality of microcystins may have severe eVects on community composition of associated bacteria. Additionally, temperature itself may aVect BCC and thereby alters interactions with cyanobacteria, for example, due to increased growth of microcystin-degrading bacteria (Ho et al. 2007, Surono et al. 2008). On the other hand, our results also show that presence of associated bacteria can change quantity and quality of cell-bound toxins (see also Dziallas and Grossart 2011b). Depending on their chemical structure, microcystins greatly diVer in their toxicity (Gupta et al. 2003, Blom and Jüttner 2005), whereby a single strain of M. aeruginosa can produce several variants of microcystin (e.g. Wiedner et al. 2003; Dziallas and Grossart 2011b). Thus, the associated bacterial community can alter actual toxicity of cyanobacterial bloom events and, therefore, has consequences for cyanobacterial toxicity on animal as well as human health and well being. Yet, our understanding on how associated bacteria may change the quality of cell-bound microcystins is marginal. Bacteria could directly alter the freshly produced toxins via exoenzyme activity. As most of the toxins remain inside the cells and thus are inaccessible for bacteria surrounding the cyanobacteria, this mechanism seems to be unlikely. Another possibility is that associated bacteria interact with cyanobacteria in a way that stimulates production of diVerent toxins depending on the mode of interaction. For example, associated bacteria may provide speciWc substrates or scavenge speciWc molecules interfering with toxin production and function. Clarifying the underlying mechanism requires a better mechanistic understanding on the role of associated bacteria aVecting toxin production. Our experiments show a complex response in measurable toxins of M. aeruginosa depending on temperature and on the presence of associated microbes. These Wndings imply that cyanobacterial toxin production in response to global warming may change in a so far unpredictable manner. Another aspect complicating this picture is that the presence of other cyanobacterial strains also aVects microcystin production by a given cyanobacterial strain in laboratory experiments (Dziallas and Grossart 2011b). Our observation (based on higher microcystin contents per active toxin-producing cyanobacterial cell at elevated temperatures) suggests an enhanced potential of toxin production at increasing water temperatures and thus requires understanding the mechanisms of bacteria–cyanobacteria interactions in a climatically changing world. Changing conditions in the cyanobacterial phycosphere aVect associated bacteria Besides temperature, cyanobacteria structure their phycosphere via their photosynthetic activity and the subsequent
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exudation of organic matter. As a result of photosynthesis, pH increased over time to values of up to 11.5, which is unsuitable for many other microorganisms including algae (Shapiro 1990). In our experiments, pH greatly varied with temperature and growth phase. The high pH as well as the cyanobacterial cell density can have a great impact on community composition and function of the associated microbial communities. Consequently, mass events of cyanobacteria alter major environmental variables and hence aVect growth of other phototrophs (e.g. Shapiro 1990; Dokulil and Teubner 2000) and heterotrophic microorganisms. For example, Ploug (2008) even found anoxic zones in decaying cyanobacterial aggregates in the Baltic Sea. Hence, cyanobacterial phycospheres represent unique microhabitats for the associated microbial community. Our microelectrode measurements revealed great diVerences depending on where the cells are situated: in a Xoating scum or an aggregate depending on the light conditions present. In contrast to Paerl (1996) and Ibelings and Mur (1992), we could not observe anoxic conditions in the middle of a cyanobacterial scum nor at the downward edge of the scum even after 8 h of darkness. This Wnding is remarkable as it questions the reason for the presence of bacteria and archaea with known anaerobic functions. Two possible alternatives may explain this obvious discrepancy: (1) anoxic zones are situated in patches that we did not touch with our microelectrodes or (2) microorganisms previously believed to be anaerobic can withstand oxic conditions and are active at least under micro-oxic conditions or they are just facultative anaerobes. However, the question whether functional genes related to anaerobic processes, for example, for sulphate reduction or methanogenesis, can be only used under oxygen depletion needs to be answered in the future. Since we only tested for the presence of these potential anaerobes, we are not able to state whether they were really active under oxic conditions. Nevertheless, as we also found these potential anaerobes in cultures established for over 30 years, it is very unlikely that they are not active. The main groups of obligate anaerobes we found are sulphate reducers and methanogens (Table 2), which have been mostly investigated in sediments and other anoxic environments. Recently, by using gas-tight bottles, it has been shown that an enrichment of methanogenic archaea lead to methane accumulation in a cyanobacterial culture even when oxygen is present (Grossart et al. 2011). Hence, it is likely that hydrogen- or acetate-producing cyanobacteria provide a preferential niche for methanogenic archaea, which—due to their capability to form methane—are of great importance for biogeochemical cycling in aquatic systems. Acknowledgments We thank Solvig Pinnow for technical assistance and Andreas Ballot and Manfred Henning for providing us a variety of M. aeruginosa strains. We further acknowledge Helle Ploug for scientiWc input and technical help with the microelectrode measurements. We thank three anonymous reviewers for their very helpful suggestions
Mar Biol (2012) 159:2389–2398 and comments. This study was funded by the German Science Foundation (DFG, GR 1540/11-1,2).
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