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FEMS Microbiology Ecology 35 (2001) 1^9 www.fems-microbiology.org MiniReview Ecological and molecular investigations of cyanotoxin production Melanie Kaebernick, Brett A. Neilan * School of Microbiology and Immunology, University of New South Wales, Sydney, N.S.W. 2052, Australia Received 5 June 2000 ; received in revised form 11 September 2000; accepted 21 September 2000 Keywords : Cyanobacteria; Microcystin ; Saxitoxin ; Cylindrospermopsin; Non-ribosomal peptide; Function; Regulation 1. Introduction Cyanobacteria, broadly classi¢ed as oxygenic phototrophs containing chlorophyll-a and accessory pigments, are among the oldest life forms on earth. They may be unicellular, colonial or ¢lamentous, with cell sizes varying from less than 2 Wm to 40 Wm in diameter. They may live as symbionts with plants and fungi, in the benthos or in the water column. Cyanobacteria have a cosmopolitan distribution and are common in all kinds of habitats, including Antarctic lakes, thermal springs, arid deserts and tropical acidic soils. Most commonly they are known for their existence as planktonic members of the water column in marine and freshwater environments. Cyanobacteria have special adaptations such as; nitrogen ¢xation, ability to regulate buoyancy, light harvesting pigments, and di¡erentiated cell types for reproduction or resting, allowing them an advantage over many competitors. Anthropogenic factors such as increased nutrient loading in freshwater and coastal environments, have lead to the occurrence of cyanobacterial blooms world wide. While blooms are unpleasant both visually and due to released odour and taste factors, certain metabolites produced by some cyanobacterial species pose a more serious problem [1]. Due to their adverse e¡ects on higher organisms, these compounds have been labelled as `toxins' and from here on are referred to as such. The chemical structures of many cyanotoxins and their adverse e¡ect on animals have been elucidated and recently reviewed [2]. However, the actual physiological function and ecological regulation of cyanobacterial toxins remain largely a mystery. Studies on regulation and function of cyanotoxins are interrelated and have focused on their e¡ect on the local ecosystem, as well as the e¡ect of environmental parame- * Corresponding author. Tel. : +61 (2) 93853235; Fax: +61 (2) 93851591; E-mail: [email protected] ters on toxin production. Hypotheses regarding the ecophysiology of the cyanotoxins, stemming from such studies, are discussed in this review. With advances in molecular biology, it has more recently been possible to elucidate the genetics behind the biosynthetic pathways of some of these toxins [3] and begin to investigate the regulation of toxin synthesis on a molecular level [4]. While such studies on cyanotoxin gene regulation are limited, similar genetic systems have been discovered and studied in other bacteria [5]. An understanding of these systems will be helpful in guiding work and ideas in cyanotoxin research. This combination of genetic and ecological investigations, is part of a new era of research, in which ecosystem interactions are understood in terms of their molecular determinants. 2. Cyanobacterial toxins Cyanobacterial toxins can be classi¢ed according to their chemical structures as cyclic peptides (microcystin and nodularin), alkaloids (anatoxin-a, anatoxin-a(s), saxitoxin, cylindrospermopsin, aplysiatoxins, lyngbyatoxin-a) and lipopolysaccharides but are more commonly discussed in terms of their toxicity to animals. While there are several dermatotoxins (lyngbyatoxin and aplysiatoxins) produced primarily by benthic marine cyanobacteria (Table 1), most cyanotoxins are classi¢ed as either hepatotoxins or neurotoxins. 2.1. Hepatotoxins Hepatotoxins are produced by many genera of cyanobacteria (Table 1). These toxins, which target the liver due to speci¢c binding of the organic anion transport system in hepatocyte cell membranes, have been implicated in the deaths of birds, wild animals, agricultural livestock and ¢sh [6], and have been responsible for human illness and death, reported from India, China, Australia and Brazil. 0168-6496 / 01 / $20.00 ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII: S 0 1 6 8 - 6 4 9 6 ( 0 0 ) 0 0 0 9 3 - 3 FEMSEC 1186 27-2-01 Cyaan Magenta Geel Zwart 2 M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 Table 1 Cyanotoxins and toxin producing cyanobacterial genera (see text for references to the relevant literature) Toxin Hepatotoxic Microcystins Nodularin Cylindrospermopsin Neurotoxic Anatoxin-a Anatoxin-a(s) Saxitoxins Dermatotoxic Aplysiatoxins Lyngbyatoxin-a Cyanobacterial genera Microcystis, Oscillatoria, Nostoc, Anabaena, Anabaenopsis Nodularia Cylindrospermosis, Aphanizomenon, Umezakia Anabaena, Oscillatoria, Aphanizomenon Anabaena Anabaena, Aphanizomenon, Lyngbya, Cylindrospermosis, Planktothrix Lyngbya, Schizothrix, Oscillatoria Lyngbya Microcystins and nodularin inhibit eukaryotic protein phosphatases type 1 and type 2A resulting in excessive phosphorylation of cytoskeletal ¢laments, ultimately leading to liver failure [6]. They are cyclic hepta- and pentapeptides, respectively, both containing the same unusual C20 amino acid ; 3-amino-9-methoxy-2,6,8-trimethyl-10phenyl-4,6-decadienoic acid, abbreviated Adda [7]. Microcystins, cyclo (-D-Ala-X-D-MeAsp-Z-Adda-D-GluMdha-), with a molecular mass (Mw) between 900 and 1100 Da are the largest and most diverse group of the cyanotoxins (Fig. 1). Over 60 di¡erent chemical forms, mostly variable in L-amino acids, have been reported for microcystin and are summarised elsewhere [2]. Although similar in structure, only four nodularins have been identi¢ed from Nodularia spumigena [7] and a ¢fth (trivially named motuporin) isolated from a sponge in Papua New Guinea, which is probably produced by a microbial symbiont [8] (Fig. 1). Microcystin, and tentatively nodularin, is the only cyanotoxin for which the biosynthetic pathway and gene cluster has been identi¢ed [3]. A third hepatotoxin, cylindrospermopsin (C15 H21 N5 O7 S), belongs to a new class of alkaloids possessing a tricyclic guanidine moiety combined with hydroxymethyl uracil (Fig. 1) [9]. This toxin suppresses glutathione and protein synthesis [10,11]. 2.2. Neurotoxins Neurotoxins, produced in association with Anabaena spp. and Aphanizomenon £os-aquae blooms, have been responsible for several animal poisonings around the world. Anatoxin-a (C10 H15 NO), a secondary amine 2-acetyl-9azabicyclo[4.2.1]non-2-ene (Mw = 165), and anatoxin-a(s) (C7 H17 N4 O4 P), a unique N-hydroxyguanidine methyl phosphate ester (Mw = 252), inhibit transmissions at the neuromuscular junction, by molecular mimicry of the neurotransmitter acetylcholine and inhibition of acetylcholinesterase activity, respectively [1]. Similar to cylindrospermopsin, both are believed to be synthesised via arginine FEMSEC 1186 27-2-01 derivatives involving a retro-Claisen condensation [12] (Fig. 1). The cosmopolitan distribution of anatoxins cannot be extended to Australian waters, where these toxins have not been isolated, and Anabaena spp. produce saxitoxins instead (Fig. 1) [13]. Saxitoxins and neo-saxitoxins, also produced by A. £os-aquae, Lyngbya, Cylindrospermopsis and Planktothrix, are part of a greater group of neurotoxins, including the C-toxins and gonyautoxins, involved in paralytic shell¢sh poisoning (PSP) and consequently classi¢ed as PSPs [2,14]. These toxins, which inhibit nerve conduction by blocking neuronal sodium channels, are also produced by dino£agellate species (Alexandrium spp., Gymnodinium catenatum, Pyrodinium bahamense var. Compressum) manifesting as toxic `red tide' events [1]. 3. Environmental e¡ects on toxin production E¡ects of the environment on toxin production are widely disputed. Blooms in the same waterbody can be variably toxic or non-toxic from one year to the next. A di¡erent strain composition, i.e. toxic versus non-toxic, which cannot be distinguished microscopically if belonging to the same species, is a common explanation for this occurrence. However, some species are known to exhibit high or low toxicity under di¡erent laboratory conditions. The stimuli for toxin production in such species is currently unknown. Environmental parameters such as light intensity, temperature, nutrients and trace metals have been mimicked under laboratory conditions and investigated with respect to their e¡ect on cyanotoxin production. A comprehensive list of studies has recently been compiled [2]. Studies on light intensity and toxin production are highly variable, partly due to the di¡erent intensities and strains tested. However, lowest toxin concentrations have generally been documented at low light intensities (2^20 Wmol photons m32 s31 ), with highest levels between 20 and 142 Wmol photons m32 s31 depending on the study. Also important is the relationship between iron uptake and light intensity. High light intensities increase cellular iron uptake which may ultimately be responsible for higher toxin production [15]. In contrast, low concentrations of iron, implicated in slower cell growth, have led to higher microcystin concentrations [16]. Nutrients, such as nitrogen and phosphorus are essential for cyanobacterial growth. Phosphorus is usually the limiting factor in lakes, and hence small changes in this nutrient may in£uence toxin production merely as a result of in£uencing growth. Generally, decreased amounts of microcystin (produced by Anabaena, Microcystis and Oscillatoria), anatoxin-a (produced by Aphanizomenon) and nodularin (produced by Nodularia) have been reported under the lowest phosphorus concentrations tested [2]. An exception to this being increased microcystin, with Cyaan Magenta Geel Zwart M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 3 Fig. 1. Chemical structures of the most common cyanobacterial hepatotoxins (a,b,c) and neurotoxins (d,e,f). respect to dry weight, recorded under phosphorus limiting conditions [17]. Nitrogen a¡ects the production of cyanotoxins di¡erently in nitrogen-¢xing and non-nitrogen-¢xing cyanobacteria. Anabaena, Aphanizomenon, Nodularia and Cylindrospermopsis strains, all capable of nitrogen-¢xation show highest levels of microcystin, anatoxin-a, or nodularin when in a nitrogen-free medium [18^21]. In contrast, Oscillatoria and Microcystis strains (non-nitrogen ¢xing) show highest levels of toxin at high levels of nitrogen [2]. The e¡ect of temperature on toxin levels is comparable in most cyanobacteria. Anatoxin-a production by Anabaena spp. and Aphanizomenon is highest at 20³C compared to 30³C or lower temperatures [22]. Similarly, microcystin FEMSEC 1186 27-2-01 and nodularin concentrations, as studied in Anabaena, Microcystis and Nodularia, are highest between 18 and 25³C, with lower levels experienced at either higher or lower temperatures tested [2]. Temperature e¡ects on saxitoxin production in cyanobacteria have not been investigated, but their concentration in the dino£agellate Alexandrium sp. is increased at low temperatures and phosphorus limitation [23]. Di¡erent temperatures can also be correlated with di¡erent chemical forms of toxin produced [19]. At temperatures below 25³C, Anabaena spp. produce microcystin-LR, instead of microcystin-RR which is preferentially synthesised at higher temperatures [19]. Similarly, more microcystin-LR compared to microcystin-RR is synthesised by Microcystis sp. under phosphorus limiting conditions [17]. Microcystin-LR also correlates to higher lev- Cyaan Magenta Geel Zwart 4 M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 els of light compared to [D-Asp3] microcystin-LR which is more common at lower light intensities [19]. In many cases, outlined above, highest toxin concentrations are reported under conditions which are optimal for cell growth. This may re£ect a direct relationship between toxin production rates and growth, as recently investigated by Orr and Jones [24]. A linear correlation was established between microcystin production rates and cell division when examined under both nitrogen and phosphorus limiting conditions [17,24]. Such correlations were not observed for the production of cylindrospermopsin by Cylindrospermopsis raciborskii [21]. 3.1. Considerations in the measurement of cyanotoxin production rates Unfortunately, no conclusive results regarding factors which in£uence toxin production, have been gained from the studies stated above. Individual studies are not readily comparable due to the di¡erent bacterial strains, culturing methods and toxin analyses employed by the various laboratories. Toxins (or toxicity) have commonly been measured via mouse bioassays, activity inhibition assays (for microcystin and nodularin) or HPLC. These methods cannot distinguish between all the di¡erent chemical isomers of the toxins. Structural di¡erences alter bioactivity, and hence may in£uence overall toxicity as a result of changes in the ratio of variably toxic isomers. Toxin data have been correlated to a variety of parameters including dry weight, total protein and cell numbers, which led to widely varied results. Dry weight, for example, is expected to increase under high light conditions due to the production of high molecular mass fatty acids and carbohydrate accumulation. In addition, total protein is a¡ected by high light as a result of the breakdown of photosynthetic components such as the phycobilisomes [25]. Hence, cellular responses unrelated to toxin production need to be considered when comparing results between studies. 4. Putative extracellular toxin functions A cyanobacterial bloom event has various implications, direct and indirect, upon the local ecosystem. An understanding of the e¡ects of cyanotoxins on such a system may guide our knowledge towards putative toxin functions. Higher animals incapable of selective feeding, such as wild birds and ¢sh are directly a¡ected by both hepatoand neurotoxins from cyanobacteria. Fish death during cyanobacterial blooms, frequently linked to the depletion of oxygen as a result of increased algal productivity, has also been attributed to the total inhibition of PP1 and 2A by microcystin, resulting in liver necrosis and death [26]. Interesting are the responses of lower eukaryotic organisms (zooplankton and phytoplankton) and bacteria to cyanotoxins. These may engage in selective feeding, be FEMSEC 1186 27-2-01 positively attracted to cyanobacterial cells, or be killed by the toxins. 4.1. Zooplankton ^ cyanobacterial interactions There is compelling evidence, that cyanobacteria and their toxins (both neuro and hepato) have an e¡ect on zooplankton (cladoceans and rotifers) population structure, and that this in turn may guide ecological processes responsible for cyanobacterial success. Cyanobacterial cells are generally a poor food source for zooplankton and are often selectively avoided [27]. As a result, zooplankton feed on phytoplankton otherwise in competition with cyanobacteria. In the process they release essential nutrients, further fertilising cyanobacterial growth. During cyanobacterial blooms, when alternative food sources for zooplankton have been exhausted, Daphnia populations are known to decline [28]. Some zooplankton (Daphnia pulicaria, Daphnia pulex) have evolved physiological and behavioural adaptations to survive in the presence of certain toxic cells [27]. Di¡erential adaptations of these species, create a change in zooplankton population dynamics. Finally, feeding pressure by adapted zooplankton on cyanobacteria is reduced by ¢sh predation, which again releases nutrients such as phosphorus, subsequently taken up by cyanobacteria [28]. It would be premature to postulate that cyanobacterial dominance is planned and guided by cyanotoxin production. However, feeding deterrence has been one of the earliest roles suggested for these metabolites. Whether the compounds causing toxicity and deterrence are one and the same has recently been questioned [29]. While daphnids are killed when feeding on toxic Microcystis cells, they show no selection between ingesting toxic or nontoxic cells, indicating that microcystins are not responsible for feeding inhibition [29]. 4.2. Bacterial^ and phytoplankton^cyanobacterial interactions Heterotrophic eubacteria, fungi, phyto£agellates and protozoans commonly associated with cyanobacterial cells may also be a¡ected by cyanotoxin production. Many bloom-forming cyanobacteria exhibit optimal growth in the presence of contaminant heterotrophic bacteria. Pseudomonas aeruginosa (Schroeter) Migula is chemotactically attracted to heterocysts of Anabaena, subsequently setting up a mutualistic relationship between the two bacteria, sharing the ¢xed N2 available. Similarly, Microcystis aeruginosa cells exhibit greater cell-speci¢c rates of CO2 ¢xation when in association with bacteria and protozoan grazers [30]. It has been suggested that the production and excretion of extracellular metabolites, such as cyanotoxins, may play a role in attracting these bene¢cial hosts while at the same time repelling antagonistic microbes and higher order grazers [30]. An allelopathic function for mi- Cyaan Magenta Geel Zwart M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 crocystin has also been suggested as a result of the toxin's growth inhibition of various algal species (Chlamydomonas, Haematococcus, Navicula and Cryptomonas) [31]. 4.3. Cyanobacterial^cyanobacterial interactions Studies of the interactions between di¡erent populations of cyanobacteria have noted the occurrence of Anabaena sp. prior to Microcystis during a bloom succession. Such a succession may be the result of numerous environmental conditions, such as changes in nutrient availability, light and temperature, favouring individual strains. However, with the recent identi¢cation of genes encoding eukaryotic type Ser/Thr protein kinases [32] and protein phosphatases (PPs) from Anabaena PCC7120 [33] and Nostoc commune UTEX584 [34], the possibility of microcystin functioning as an inhibitor of PPs against other cyanobacteria may be postulated. The activity of protein phosphatases found in Microcystis sp. are not inhibited by microcystin [35]. A factor of uncertainty is the intra- or extracellular movement of the toxins. Putative extracellular functions require export of these metabolites from the cell. No cyanotoxin transport system has been identi¢ed so far but some microcystin has been detected in the cell wall and sheath area of the cell, indicating possible release of this toxin from intact cells [36]. However, most of the toxin is located in the thylakoid and nucleoid areas [36] and toxin found in the media of laboratory cultures has generally been attributed to cell lysis as a result of some detrimental growth factor, or culture age [19]. While the same data may also suggest an export of the toxin from the cell under certain conditions this has not been investigated. 5. Putative intracellular toxin functions Most putative intracellular roles described are derived from observations of environmental e¡ects on toxin production. As most research in this ¢eld has focused on microcystin the discussion of putative intracellular functions is limited to this toxin. Toxin production as a mechanism for shunting excess metabolites during periods of environmental stress has been considered [30], yet appears unrealistic when considering the complex biosynthetic pathway (described later) and high energy costs required for production. Microcystin's a¤nity for iron and other cations, such as copper and zinc, provides the molecules siderophoric properties [37] and a putative role as an iron scavenging molecule [15]. Extracellular siderophores bind Fe2 from the environment, for use by the cell, under conditions of low iron availability (i.e. low iron = high siderophore/toxin [16]). Intracellularly, however, such siderophoric properties may have a negative impact on cell functions, due to competition with primary metabolites for limited essential FEMSEC 1186 27-2-01 5 iron. Alternatively, under conditions of high cellular iron, an intracellular siderophore may have a protective function by chelating ferrous ions, forming iron^microcystin complexes and thus keeping the cellular level of free radicals low [15]. Such iron^microcystin complexes may be stored until low iron conditions allow the release of Fe2 for cellular processes. However, microcystin or complexes thereof have not been identi¢ed as part of the stored substances in cellular inclusions [36]. Instead, most microcystin is located in the thylakoid and nucleoid regions of the cell [36]. It is believed that the `Adda' moiety of the toxin may bind to the thylakoid (where microcystin is primarily located) leaving the polar peptide ring to bind metals from the cytoplasm [24]. This association with the photosynthetic apparatus of the cell, may indicate a putative function in the light harvesting and chromatic adaptation mechanisms exhibited by some cyanobacteria. Genetic regulation studies have shown increased transcription of genes involved in microcystin production at high light levels [4]. Whether this is indicative of microcystin function directly in cellular light mechanisms, or a role that is required for other cellular functions under these conditions, remains to be investigated. Intracellular regulation by microcystin in the producing organism, is highly speculative, but may be suggested by analogy to okadaic acid (OA) produced by the dino£agellate Prorocentrum lima. Similar to microcystin, OA is a speci¢c inhibitor of PPs. The upregulation of its production at increasing OA levels, has been speculated to regulate behavioural and physiological responses, which are controlled by reversible phosphorylation [38]. The e¡ect of microcystin on microcystin production or other cellular processes has not been tested. Putative roles for microcystin, discussed so far, in relation to the factors in£uencing toxin production, are in agreement with categorising the peptides as secondary metabolites. The de¢nition as `Tthose compounds that are not used by the organism for its primary metabolism' [6] is supported by the survival of microcystin-de¢cient strains and engineered mutants. Studies showing direct correlation between toxin production and growth rate have disputed this claim and suggest the production of non-toxic alternatives with the same function as the toxic peptides, for non-toxic strains [24]. While such a de¢nition dispute does not in£uence research results, it does change the focus of where to look for putative functions. 6. Molecular studies on microcystin biosynthesis and regulation In recent years, the advances of molecular biology, have furthered our understanding of secondary metabolite production and transcriptional regulation on a genetic basis. Unfortunately, most cyanotoxin genes, with the exception Cyaan Magenta Geel Zwart 6 M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 Fig. 2. Proposed biosynthetic model for microcystin-LR, showing the organisation of the gene cluster mcyA-J and the toxin, microcystin. Biosynthesis is via a multienzyme complex consisting of both peptide synthetase and polyketide synthase modules [3]. Numbered circles indicate the order of amino acids incorporated into the growing peptide chain by NRP genes (mcyA, B, C, EP , GP ). mcyA and mcyB contain two modules, A1/A2 and B1/B2, respectively. A1 also encodes an N-methyltransferase domain. Numbered rectangles show the order of polyketide synthesis in the formation of Adda (mcyGK , EK , D). Additional ORFs of putative microcystin tailoring function are indicated by `T'. mcyH shows high identity to ABC transporter genes [3]. The relative sizes of the mcy ORFs have been approximated, with the entire gene cluster comprising some 55 kb. of microcystin, have not been identi¢ed and can not be discussed in this section. Microcystin and nodularin peptides are produced, or putatively produced in the case of nodularin, non-ribosomally via a multifunctional enzyme (a peptide synthetase) that employs the thio-template mechanism [39] (for reviews on this topic see [40,41]). The synthetase has individual sites (modules) catalysing amino/hydroxy acid activation and thioester formation in the same order in which their residues are incorporated into the growing peptide chain. Chain elongation is catalysed by the enzyme-bound cofactor 4P-phosphopantetheine [40,41]. Genes encoding peptide synthetases are typically clustered into large operons with repetitive domains in which highly conserved core sequences have been identi¢ed [41]. Using these sequences it has been possible to locate peptide synthetase genes in M. aeruginosa [42]. Insertional mutagenesis of some of these genes (mcyA,B and D) pro- FEMSEC 1186 27-2-01 duced non-toxic mutants, providing evidence for their involvement in microcystin synthesis [3,42,43]. The entire gene cluster (mcyA-J) has since been identi¢ed and sequenced [3]. A 750-bp promoter region separates the mcyABC operon from mcyD-J, the latter being reversely oriented (Fig. 2). Polyketide synthase domains, identi¢ed as part of mcyDE and G, are in agreement with biochemical data suggesting the requirement of a polyketide synthase for the production of the fatty acid side chain of Adda [3] (Fig. 2). Mutagenesis of polyketide domains provided evidence that the microcystin biosynthetic pathway proceeds via a multienzyme complex consisting of both peptide synthetase and polyketide synthase modules [3]. Gene clusters, encoding the cooperation between peptide synthetases and polyketide synthases have also been identi¢ed in Proteus mirabilis [44] and in the production of coronatine by Pseudomonas syringae [45]. Studies are currently under way to explore the regulation of both the Cyaan Magenta Geel Zwart M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 peptide synthetase and polyketide synthase gene clusters of microcystin, aiming to identify the overall stimuli for toxin production. 7. Regulation and function of other non-ribosomal peptide synthetases (NRPs) Non-ribosomal synthesis of peptides, via a type of multifunctional enzyme is by no means restricted to cyanobacteria. Peptide synthetase genes are highly conserved across a wide array of bacterial genera, fungi and actinomycetes. They are commonly involved in the biosynthesis of peptide antibiotics, exotoxins, siderophores and other bioactive compounds. The genetic regulation and possible function of some of these genes has been studied extensively, and may provide some correlation to the molecular regulation of NRP toxins in cyanobacteria and possibly the function of these compounds. Probably the most investigated peptide synthetase genes, are the gene clusters tycABC, grsTAB and srfABCD, of Bacillus spp., producing the peptide antibiotics tyrocidin and gramicidin S, and the lipopeptide surfactin, respectively. Activation and regulation of these genes is believed to be controlled primarily at the level of transcriptional initiation, possibly via a bacterial two-component system [5]. These systems are involved in the transduction of information regarding the status of the environment from an activator (sensor kinase) to a receiver protein via phosphorylation [46]. Tyrocidin synthesis is under the control of a negative regulator protein (AbrB) which interacts directly with sequences near the tycABC operon [5]. During the onset of stationary growth, the transcriptional activator protein (SpoA) represses abrB transcription, reducing AbrB expression and subsequently its inhibition of tycABC transcription [5]. Modi¢ed two-component regulatory systems also appear to be responsible for the regulated production of syringomycin and corantine (COR) by P. syringae [45,47]. COR consists of two distinct moieties, coronamic acid (CMA) and coronafacic acid (CFA), which are produced via a peptide synthetase and polyketide synthase, respectively [45,48]. As in the microcystin gene clusters (mcyABC and mcyD-H), the structural genes for the peptide synthetase, CMA (cmaABTU) and polyketide synthase, CFA (cfa) biosynthesis, are separated by a regulatory region [47]. This region contains three genes corP, corS and corR, whose products encode two response regulators (CorP and CorR) and one sensor histidine kinase (CorS), as part of a putative two-component regulatory system [47]. Both, two-component systems, and protein phosphorylation and dephosphorylation in response to environmental stimuli, have been found in cyanobacteria. Putative response regulators and sensory kinases have been identi¢ed from Anabaena sp. [49], Nostoc sp. [50], Fremyella diplosiphon [51], and Synechococcus [52]. To FEMSEC 1186 27-2-01 7 date, no such regulatory proteins have been identi¢ed as part of the microcystin biosynthetic gene cluster. An interesting comparison may be drawn between the production of the siderophore, exochelin (Mycobacterium sp.) and microcystin. The exochelin gene cluster is made up of distinct operons, encoding peptide synthetase genes (fxbB, fxbC), a putative formyltransferase (fxbA transcribed in opposite orientation) and an ABC transporter (exiT) [53]. The expression of fxbA is regulated by a repressor protein, IdeR (homologous to the DtxR iron regulator of Corynebacterium diphtheriae), whose binding ability is dependent on the presence of divalent metals [54]. This system thus leads to the production of exochelin under iron-de¢cient conditions. Similarly, ecological studies have indicated increased microcystin production under low iron concentrations [16]. However, unlike microcystin which is believed to remain intracellular [36], exochelin is secreted. The putative ABC transporter polypeptide encoded by exiT, is hypothesised to be responsible for exochelin export and provides the ¢rst example of a potential siderophore exporter protein [53]. A putative ABC transporter gene has also recently been identi¢ed downstream of mcyG [3] indicating possible intracellular or extracellular microcystin transport. The association of ABC transporters with peptide synthetase genes, is not uncommon. ABC-type transport genes have also been identi¢ed associated with the surfactin synthetase operon in Bacillus subtilis, the syringomycin synthetase operon of P. syringae, a putative non-ribosomal peptide/polyketide operon involved in swarming of P. mirabilis, and in many antibiotic-producing actinomycetes. 8. Conclusions and future directions Over the past 20 years, cyanobacterial research has advanced from ecological studies on bloom formation, the development of toxin analysis techniques and toxin pharmacology in higher animals, to the realisation of the importance of this knowledge in microbial ecology and public health. Since then, chemical structure determinations and ecological studies of environmental parameters a¡ecting toxin production, have provided some clues to the regulation and function of some of these toxins. To date, laboratory-based ecological investigations have focused on the isolation of individual environmental e¡ectors, which is a crucial step in the identi¢cation of toxin regulation mechanisms. However, such results can not be extrapolated to the real environment, where a greater array of factors exist and possibly control toxin production. Advances in molecular biology have led to the identi¢cation of some of the genes responsible for cyanotoxin production. Regulatory studies of these genes are still in their early stages, guided by both the results from previous ecological investigations, and the comparison to similar biosynthetic systems in other bacteria. Molecular studies Cyaan Magenta Geel Zwart 8 M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9 have the potential to identify the individual environmental factors which a¡ect gene transcription and expression directly, opposed to those factors that may a¡ect other cellular processes which lead to changes in toxin production. Thus, through a combination of ecological and molecular research, it is hoped that experimental data may ultimately be extrapolated to the environment not only to understand the timing of toxin production but also the purpose for it. [18] [19] [20] References [21] [1] Carmichael, W.W. (1994) The toxins of cyanobacteria. Sci. Am. 270, 78^86. [2] Sivonen, K. and Jones, G. (1999) Cyanobacterial toxins. In: Toxic Cyanobacteria in Water. 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