Download Ecological and molecular investigations of cyanotoxin production

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. A Guide to their Public Health Consequences, Monitoring and Management (Chorus, I. and Bartram, J., Eds.),
pp. 41^111. E and FN Spoon, London.
[3] Tillett, D., Dittmann, E., Erhard, M., von Doehren, H., Boerner, R.
and Neilan, B.A. (2000) Structural organisation of microcystin biosynthesis in M. aeruginosa PCC7806: An integrated peptide-polyketide synthetase system. Chem. Biol., in press.
[4] Kaebernick, M., Neilan, B.A., Boerner, T. and Dittmann, E. (2000)
Light and the transcriptional response of the microcystin biosynthetic
gene cluster. Appl. Environ. Microbiol. 66, 3387^3392.
[5] Marahiel, M.A., Nakano, M.M. and Zuber, P. (1993) Regulation of
peptide antibiotic production in Bacillus. Mol. Microbiol. 7, 631^636.
[6] Carmichael, W.W. (1992) Cyanobacterial secondary metabolites ^ the
cyanotoxins. J. Appl. Bacteriol. 72, 445^459.
[7] Rinehart, K.L., Namikoshi, M. and Choi, B.W. (1994) Structure and
biosynthesis of toxins form blue-green algae (cyanobacteria). J. Appl.
Phycol. 6, 159^176.
[8] deSilva, E.D., Williams, D.E., Anderson, V., Klix, H., Holmes,
C.F.B. and Allen, T.M. (1992) Motuporin, a potent protein phoshatase inhibitor isolated from the Papua New Guinea sponge Theonella
swinhoei Gray. Tetrahedron Lett. 33, 1561^1564.
[9] Ohtani, I., Moore, R.E. and Runnegar, M.T.C. (1992) Cylindrospermopsin, a potent hepatotoxin from the blue-green alga Cylindrospermopsis raciborskii. J. Am. Chem. Soc. 114, 7941^7942.
[10] Runnegar, M., Shou-Ming, K., Ya-Zhen, Z. and Shelly, C. (1995)
Inhibition of reduced glutathione synthesis by cyanobacterial alkaloid
cylindrospermopsin in cultured rat hepatocytes. Biochem. Pharmacol.
49, 219^255.
[11] Terao, K., Ohmori, S., Igarashi, K., Ohtani, I., Watanabe, M., Harada, K., Ito, E. and Watanabe, M. (1994) Electron microscopic studies on experimental poisoning in mice induced by cylindrospermopsin
isolated from blue-green algae Umezakia Natans. Toxicon 32, 833^
843.
[12] Shimizu, Y. (1996) Microalgal metabolites: a new perspective. Annu.
Rev. Microbiol. 50, 431^465.
[13] Humpage, A.R., Rositano, J., Bretag, A.H., Brown, R., Baker, P.D.,
Nicholson, B.C. and Ste¡ensen, D.A. (1994) Paralytic shell¢sh poisons from Australian cyanobacterial blooms. Aust. J. Mar. Freshw.
Res. 45, 461^471.
[14] Pomati, F., Sacchi, S., Rossetti, C., Giovannardi, S., Onondera, H.,
Oshima, Y. and Neilan, B.A. (2000) The freshwater cyanobacterium
Planktothrix sp. FP1: Molecular identi¢cation and detection of paralytic shell¢sh poisoning toxins. J. Phycol. 36, 553^562.
[15] Utkilen, H. and Gjolme, N. (1995) Iron-stimulated toxin production
in Microcystis aeruginosa. Appl. Environ. Microbiol. 61, 797^800.
[16] Lukac, M. and Aegerter, R. (1993) In£uence of trace metals on
growth and toxin production of Microcystis aeruginosa. Toxicon
31, 293^305.
[17] Oh, H.-M., Lee, S.J., Jang, M.-H. and Yoon, B.-D. (2000) Micro-
FEMSEC 1186 27-2-01
[22]
[23]
[24]
[25]
[26]
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
cystin production by Microcystis aeruginosa in a phosphorous-limited
chemostat. Appl. Environ. Microbiol. 66, 176^179.
Rapala, J., Sivonen, K., Luukkainen, R. and Niemelae, S.I. (1993)
Anatoxin-a concentration in Anabaena and Aphanizomenon under
di¡erent environmental conditions and comparison of growth by
toxic and non-toxic Anabaena strains ^ a laboratory study. J. Appl.
Phycol. 5, 581^591.
Rapala, J., Sivonen, K., Lyra, C. and Niemelae, S.I. (1997) Variation
of microcystins, cyanobacterial hepatotoxins, in Anabaena spp. as a
function of growth stimuli. Appl. Environ. Microbiol. 63, 2206^2212.
Lehtimaeki, J., Moisander, P., Sivonen, K. and Kononen, K. (1997)
Growth, nitrogen ¢xation and nodularin production by two Baltic
Sea cyanobacteria. Appl. Environ. Microbiol. 63, 1647^1656.
Saker, M.L., Neilan, B.A. and Gri¤ths, D.J. (1999) Two morpholgical forms of Cylindrospermopsis raciborskii (cyanobacteria) isolated
from Solomon dam, Palm Island, Queensland. J. Phycol. 35, 599^
606.
Rapala, J. and Sivonen, K. (1998) Assessment of environmental conditions that favour hepatotoxic and neurotoxic Anabaena spp. strains
in culture under light-limitation and di¡erent temperatures. Microb.
Ecol. 36, 181^192.
Anderson, D.M., Kulis, D.M., Sullivan, J.J., Hall, S. and Lee, C.
(1990) Dynamics and physiology of saxitoxin production by the dino£agellates Alexandrium spp. Mar. Biol. 104, 511^524.
Orr, P.T. and Jones, G.J. (1998) Relationship between microcystin
production and cell division rates in nitrogen limited Microcystis
aeruginosa cultures. Limnol. Oceanogr. 43, 1604^1614.
Walsh, K., Jones, G.J. and Dunstan, R.H. (1997) E¡ect of irradiance
on fatty acid, carotenoid, total protein composition and growth of
Microcystis aeruginosa. Phytochemistry 44, 817^824.
Tencalla, F. and Dietrich, D. (1997) Biochemical characterization of
microcystin toxicity in rainbow trout (Oncorhynchus mykiss). Toxicon
35, 583^595.
DeMott, W.R. (1991) E¡ects of toxic cyanobacteria and puri¢ed
toxins on the survival and feeding of a copepod and three species
of Daphnia. Limnol. Oceanogr. 36, 1346^1357.
Burns, C.W. (1987) Insights into zooplankton^cyanobacteria interactions derived from enclosure studies. N.Z.L. J. Mar. Freshw. Res. 21,
477^482.
Rohrlack, T., Dittmann, E., Henning, M., Boerner, T. and Kohl,
J.-G. (1999) Role of microcystins in poisoning and food ingestion
inhibition of Daphnia galeata caused by the cyanobacterium Microcystis aeruginosa. Appl. Environ. Microbiol. 65, 737^739.
Paerl, H.W. and Millie, D.F. (1996) Physiological ecology of toxic
aquatic cyanobacteria. Phycologia 35, 160^167.
Keating, K.I. (1978) Blue-green algal inhibition of diatom growth:
Transition from mesotrophic to eutrophic community structure. Science 199, 971^973.
Zhang, C.C. (1993) A gene encoding a protein related to eukaryotic
protein kinases from the ¢lamentous heterocystous cyanobacterium
Anabaena PCC 7120. Proc. Natl. Acad. Sci. USA 90, 11840^11844.
Zhang, C.C., Friry, A. and Peng, L. (1998) Molecular and genetic
analysis of two closely linked genes that encode, respectively, a protein phosphatase 1/2A/2B homolog and a protein kinase homolog in
the cyanobacterium Anabaena sp. Strain PCC7120. J. Bacteriol. 180,
2616^2622.
Potts, M., Sun, H., Mockaitis, K., Kennelly, P.J., Reed, D. and
Tonks, N.K. (1993) A protein-tyrosine/serine phosphatase encoded
by the genome of the cyanobacterium Nostoc commune UTEX 584.
J. Biol. Chem. 268, 7632^7635.
Shi, L., Carmichael, W.W. and Kennelly, P.J. (1999) Cyanobacterial
PPP family protein phosphatases possess multifunctional capabilities
and are resistant to microcystin-LR. J. Biol. Chem. 274, 10039^
10046.
Shi, L., Carmichael, W.W. and I, M. (1995) Immuno-gold localization of hepatotoxins in cyanobacterial cells. Arch. Microbiol. 163, 7^
15.
Cyaan Magenta Geel Zwart
M. Kaebernick, B.A. Neilan / FEMS Microbiology Ecology 35 (2001) 1^9
[37] Humble, A., Gadd, G.M. and Codd, G.A. (1994) Polarographic analysis of the interactions between cyanobacterial microcystin (hepatotoxin) variants and metal cations. In: VIII International Symposium
on Phototrophic Prokaryotes, 82 pp., Florence.
[38] Boland, M.P., Taylor, M.F.J.R. and Holmes, C.F.B. (1993) Identi¢cation and characterisation of a type-1 protein phosphatase from
the okadaic acid-producing marine dino£agellate Prorocentrum lima.
FEBS Lett. 334, 13^17.
[39] Arment, A.R. and Carmichael, W.W. (1996) Evidence that microcystin is a thiotemplate product. J. Phycol. 32, 591^597.
[40] Kleinkauf, H. and von Doehren, H. (1990) Nonribosomal biosynthesis of peptide antibiotics. Eur. J. Biochem. 192, 1^15.
[41] Marahiel, M.A. (1992) Multidomain enzymes involved in peptide
synthesis. FEBS Lett. 307, 40^43.
[42] Dittmann, E., Neilan, B.A., Erhard, M., von Doehren, H. and Boerner, T. (1997) Insertional mutagenesis of a peptide synthetase gene
that is responsible for hepatotoxin production in the cyanobacterium
Microcystis aeruginosa PCC 7806. Mol. Microbiol. 26, 779^787.
[43] Nishizawa, T., Asayama, M., Fujii, K., Harada, K. and Shirai, M.
(1999) Genetic analysis of the peptide synthetase genes for a cyclic
heptapeptide microcystin in Microcystis spp. J. Biochem. 126, 520^
526.
[44] Gaisser, S. and Hughes, C. (1997) A locus coding for putative nonribosomal peptide/polyketide synthase functions is mutated in a
swarming-defective Proteus mirabilis strain. Mol. Gen. Genet. 253,
415^427.
[45] Ullrich, M. and Bender, C.L. (1994) The biosynthetic gene cluster for
coronamic acid, an ethylcyclopropyl amino acid, contains genes homologous to amino acid-activating enzymes and thioesterases. J. Bacteriol. 176, 7574^7586.
[46] Stock, J.B., Stock, A.M. and Mottonen, J.M. (1990) Signal transduction in bacteria. Nature 344, 395^400.
[47] Ullrich, M., Penaloza-Vazquez, A., Bailey, A. and Bender, C. (1995)
FEMSEC 1186 27-2-01
[48]
[49]
[50]
[51]
[52]
[53]
[54]
9
A modi¢ed two-component regulatory system is involved in temperature-dependent biosynthesis of the Pseudomonas syringae phytotoxin
coronatine. J. Bacteriol. 177, 6160^6169.
Rangaswamy, V., Ullrich, M., Jones, W., Mitchell, R., Parry, R.,
Reynolds, P. and Bender, C.L. (1997) Expression and analysis of
coronafacate ligase, a thermoregulated gene required for production
of the phytotoxin coronatine in Pseudomonas syringae. FEMS Microbiol. Lett. 154, 65^72.
Schwarz, R. and Grossman, A.R. (1998) A response regulator of
cyanobacteria integrates diverse environmental signals and is critical
for survival under extreme conditions. Proc. Natl. Acad. Sci. USA 95,
11008^11013.
Campbell, E.L., Hagen, K.D., Cohen, M.F., Summers, M.L. and
Meeks, J.C. (1996) The devR gene product is characteristic of receivers of two-component regulatory systems and is essential for heterocyst development in the ¢lamentous cyanobacterium Nostoc sp.
strain ATCC29133. J. Bacteriol. 178, 2037^2043.
Chiang, G.G., Schaefer, M.R. and Grossman, A.R. (1992) Complementation of a red-light-indi¡erent cyanobacterial mutant. Proc.
Natl. Acad. Sci. USA 89, 9415^9419.
Aiba, H., Nagaya, M. and Mizuno, T. (1993) Sensor and regulator
proteins from the cyanobacterium Synechococcus species PCC7942
that belong to the bacterial signal-transduction protein families: implication in the adaptive response to phosphate limitation. Mol. Microbiol. 8, 81^91.
Zhu, W., Arceneaux, J.E., Beggs, M.L., Byers, B.R., Eisenach, K.D.
and Lundrigan, M.D. (1998) Exochelin genes in Mycobacterium
smegmatis: identi¢cation of an ABC transporter and two non-ribosomal peptide synthetase genes. Mol. Microbiol. 29, 629^639.
Dussurget, O. et al. (1999) Transcriptional control of the iron-responsive fxbAgene by the mycobacterial regulator IdeR. J. Bacteriol. 181,
3402^3408.
Cyaan Magenta Geel Zwart