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Cylindrospermopsin and Saxitoxin Synthetase Genes in Cylindrospermopsis raciborskii Strains from Brazilian Freshwater

  • Caroline Hoff-Risseti,

    Affiliations Center for Nuclear Energy in Agriculture, University of São Paulo, Piracicaba, São Paulo, Brazil, Faculty of Pharmaceutical Sciences, University of São Paulo, São Paulo, São Paulo, Brazil

  • Felipe Augusto Dörr,

    Affiliation Faculty of Pharmaceutical Sciences, University of São Paulo, São Paulo, São Paulo, Brazil

  • Patricia Dayane Carvalho Schaker,

    Affiliation Center for Nuclear Energy in Agriculture, University of São Paulo, Piracicaba, São Paulo, Brazil

  • Ernani Pinto,

    Affiliation Faculty of Pharmaceutical Sciences, University of São Paulo, São Paulo, São Paulo, Brazil

  • Vera Regina Werner,

    Affiliation Natural Sciences Museum, Zoobotanical Foundation of Rio Grande do Sul, Porto Alegre, Rio Grande do Sul, Brazil

  • Marli Fatima Fiore

    Affiliation Center for Nuclear Energy in Agriculture, University of São Paulo, Piracicaba, São Paulo, Brazil

Cylindrospermopsin and Saxitoxin Synthetase Genes in Cylindrospermopsis raciborskii Strains from Brazilian Freshwater

  • Caroline Hoff-Risseti, 
  • Felipe Augusto Dörr, 
  • Patricia Dayane Carvalho Schaker, 
  • Ernani Pinto, 
  • Vera Regina Werner, 
  • Marli Fatima Fiore


The Cylindrospermopsis raciborskii population from Brazilian freshwater is known to produce saxitoxin derivatives (STX), while cylindrospermopsin (CYN), which is commonly detected in isolates from Australia and Asia continents, has thus far not been detected in South American strains. However, during the investigation for the presence of cyrA, cyrB, cyrC and cyrJ CYN synthetase genes in the genomes of four laboratory-cultured C. raciborskii Brazilian strains, the almost complete cyrA gene sequences were obtained for all strains, while cyrB and cyrC gene fragments were observed in two strains. These nucleotide sequences were translated into amino acids, and the predicted protein functions and domains confirmed their identity as CYN synthetase genes. Attempts to PCR amplify cyrJ gene fragments from the four strains were unsuccessful. Phylogenetic analysis grouped the nucleotide sequences together with their homologues found in known CYN synthetase clusters of C. raciborskii strains with high bootstrap support. In addition, fragments of sxtA, sxtB and sxtI genes involved in STX production were also obtained. Extensive LC-MS analyses were unable to detect CYN in the cultured strains, whereas the production of STX and its analogues was confirmed in CENA302, CENA305 and T3. To our knowledge, this is the first study reporting the presence of cyr genes in South American strains of C. raciborskii and the presence of sxt and cyr genes in a single C. raciborskii strain. This discovery suggests a shift in the type of cyanotoxin production over time of South American strains of C. raciborskii and contributes to the reconstruction of the evolutionary history and diversification of cyanobacterial toxins.


The cyanobacterial genus Cylindrospermopsis (Woloszynska) Seenayya and Subba Raju [1] belongs to the order Nostocales, family Nostocaceae [2], [3]. To date, 10 species have been described, and all of them have been found in the phytoplankton community of freshwater environments [3]. C. raciborskii has been designated as the type species and the C. raciborskii strain AWT205, which was isolated from an ornamental lake (Oatley Pond) in Sydney, Australia [4], [5], as the recognized type strain. C. raciborskii has received attention in the last decade due to its frequent dominance in freshwater blooms and its ability to synthesize cyanotoxins. The production of cylindrospermopsin (CYN) and saxitoxin (STX) by this cyanobacterial species has been reported, but in distinct strains [6], [7]. Both toxins are synthesized on large modular non-ribosomal peptide synthetase (NRPS) and polyketide synthase (PKS) enzyme complexes [8]. CYN is a cyclic sulfated guanidine alkaloid that inhibits glutathione, cytochrome P450 and protein synthesis, causing injury and cell necrosis mainly in liver, kidneys, thymus and heart of vertebrates [9][11]. The gene cluster for the biosynthesis of cylindrospermopsin (cyr) in C. raciborskii strains spans 42-kb encoding 15 open-reading frames (cyrA-O) [12]. STX is also an alkaloid but acts as a neurotoxin that blocks neuronal sodium, potassium and calcium channels, affecting the propagation of nerve impulses resulting in neuromuscular paralysis [13], [14]. Thereby, these two cyanotoxins found in aquatic environments are a potential risk for human health.

Various strains of C. raciborskii are found colonizing eutrophic reservoirs in tropical, subtropical and temperate regions [15][19]. In Brazil, C. raciborskii is the main species found in freshwater bodies [7], [16], [20][31], although the occurrence of C. phillipinensis, C. catemaco and C. acuminato-crispa has also been reported [30], [32], [33]. The most studied Brazilian strain of Cylindrospermopsis is C. raciborskii T3, isolated in 1997 from the Taquacetuba branch of the Billings reservoir in São Paulo municipality [34]. Surprisingly, this strain was found to produce the neurotoxin saxitoxin and some of its derivatives [7]. Unlike the South American isolates, the C. raciborskii strains originated from Australia and Asia were found to synthesize the alkaloid cylindrospermopsin [6], [35][39]. The elucidation of the saxitoxin biosynthesis gene cluster (sxt) was based on the C. raciborskii T3 genomic DNA and comprises approximately 35-kb encoding for 31 open reading frames [40]. Chemical analyses conducted on other C. raciborskii Brazilian isolates also confirmed the production of saxitoxin and its analogues [7], [41], [42]. Although there are reports on the occurrence of CYN in environmental samples [43], [44], the production of this toxin by Brazilian C. raciborskii isolates has not been confirmed. In order to better understand this, four C. raciborskii strains were investigated for the presence of genes associated with the biosynthesis of CYN and SXT analogues. In this way, the cyrA, cyrB, cyrC and cyrJ genes, considered as being exclusives of CYN producers [12], [45], [46], were assessed, as well as sxtA, sxtB and sxtC genes. Both cyanotoxin groups were also evaluated by LC-MS analysis, and all strains were identified by both morphological and molecular analyses.


Cyanobacterial strains and morphological identification

The Brazilian cyanobacterial strains used in this study are shown in Table 1. The strains CENA302, CENA303 and CENA305 were isolated from water samples by transferring single Cylindrospermopsis trichomes using a sterilized Pasteur pipette to sterile test tubes containing 9 mL of ASM-1 [47] liquid medium. The trichomes were repeatedly transferred into new ASM-1 medium until a pure culture was established. Cycloheximide to a final concentration of 75 mg L−1 was added to inhibit eukaryotic cell growth. The cells were grown under a 14∶10 light-dark photoperiod with white fluorescent light (40±5 µmol photons m−2 s−1) at 25±1°C. Moreover, fluorescence microscopy studies (AxioSkop 2, Carl Zeiss, Jena, Germany, equipped with digital camera AxioCamMR3, AxioVision program, Rel. 4.6 software) of cyanobacterial cultures were also performed to confirm the absence of other cyanobacteria. Morphology was evaluated using reference literature [3], [48].

Table 1. Brazilian isolates of Cylindrospermopsis raciborskii used in this study.

The C. raciborskii strain T3 was isolated in 1997 by Dr. Pedro A. Zagatto (CETESB, São Paulo, Brazil) from Billings reservoir but in the Taquacetuba branch, São Paulo, São Paulo State, and was deposited in the culture collection of the Botanic Institute of São Paulo (CCIBt), Brazil. This strain was originally obtained from the CCIBt on June 4, 2003 and has since been maintained in culture in our lab. The Australian C. raciborskii strain CYP011K, originally isolated by Dr. Peter Baker from the Julius Lake, Mount Isa, Queensland, was obtained from the laboratory of Prof. Sandra M.F.O. Azevedo (Federal University of Rio de Janeiro, RJ, Brazil). This strain is known as a CYN producer and therefore was considered as a reference sample.

DNA extraction, PCR amplification, sequencing and phylogeny

A total of 4.5 mL of cyanobacterial liquid culture was collected at the final exponential growth phase and concentrated by centrifugation (5 min at 13,000 × g). Total genomic DNA was extracted from the pellet using a modified CTAB (cetyl-trimethyl-ammonium bromide)-based extraction method adapted for cyanobacteria [49].

The presence of genes involved in the biosynthesis of cylindrospermopsin and saxitoxin were investigated using the specific primer sets (Table S1). All of the PCR reactions were performed in a 25 µL reaction volume containing 1X PCR buffer, 1.5 U Platinum® Taq DNA polymerase (Life Technologies, Carlsbad, CA, USA), 3.0 mM MgCl2, 200 µM dNTP, 0.2 µM of each primer and 10 ng of genomic DNA. Thermal cycling was performed in a Techne TC-412 Thermal Cycler (Bibby Scientific Limited, Stone, Staffordshire, England) using the designed primers (for sxt genes) under the following conditions: sxtA4 (C-terminal domain), 94 °C for 5 min, followed by 35 cycles of 94 °C for 30 s, 61 °C for 30 s, 72 °C for 30 s, and final extension of 72 °C for 7 min; sxtI, 94°C for 5 min, followed by 35 cycles of 94°C for 30 s, 61 °C for 30 s, 72 °C for 90 s, and final extension of 72°C for 7 min; sxtB, 94 °C for 3 min, followed by 35 cycles of 94 °C for 30 s, 53 °C for 1 min, 72 °C for 1 min, and a final extension of 72 °C for 7 min. Thermal cycling conditions for cyrJ gene amplification were 94 °C for 3 min, followed by 30 cycles of 94 °C for 20 s, 53 °C for 1 min, 72 °C for 1 min, and a final extension of 72 °C for 7 min, adapted from Mazmouz et al., [50]. The cyrA, cyrB and cyrC fragments were amplified using the conditions previously described by the authors [51], with the exception of annealing temperature that was 57 °C.

The amplified gene fragments were cloned using pGEM®-T Easy Vector Systems (Promega, Madison, WI, USA). Competent Escherichia coli DH5α cells were transformed, and recombinant plasmids were purified from white colonies by the alkaline lysis method [52]. The cloned PCR products were sequenced using the DYEnamic ET Terminator Cycle Sequencing (GE Healthcare, Little Chalfont, Buckinghamshire, England) using the T7 and SP6 primer sites of the vector. The cycle sequencing reaction was performed with a Techne TC-412 (Bibby Scientific Limited) for 25 cycles of 95°C for 20 s, 52°C for 15 s, and 60°C for 1 min. After the completion of the reaction, a 75% isopropanol wash followed by a 70% ethanol wash was performed to remove residual dye terminators. The purified reaction mixtures were reconstituted in HiDi formamide (Applied Biosystems/Life Technologies, Foster City, CA, USA), and the samples were analyzed in an ABI PRISM 3100 Genetic Analyzer (Applied Biosystems/Life Technologies).

The 16S rRNA genes were PCR amplified using the specific primers listed in Table S1 and PCR amplification and sequencing were performed as described previously [53].

The partial nucleotide sequences of the cyr and sxt genes obtained in this study and reference sequences retrieved from GenBank were aligned, refined, and used to generate phylogenetic trees. Trees were reconstructed with maximum-likelihood (ML) and neighbor-joining (NJ) algorithms implemented by the MEGA version 5.0 program package [54] using the Tamura-Nei and ρ-distance models, respectively. The stability of the phylogenetic relationships was assessed by bootstrapping (1,000 replicates). Moreover, the gene sequences were analyzed by BLASTX (NCBI GenBank database) to compare each of the nucleotide sequence with its translated proteins in all possible reading frames. After identifying the correct reading frame, the nucleotide sequence was translated into protein using the translate tool from ExPASy Proteomic Server (Swiss Institute of Bioinformatics).

Nucleotide sequences generated in this study have been deposited in the NCBI GenBank database under the following accession numbers: JX175238 to JX175243 and KC894581 to KC894586 for the cyr genes; JX175232 to JX175237 and KC894587 to KC894589 for sxt genes; JQ707291 to JQ707296 for 16S rRNA genes.

Chromatographic analyses

Culture samples of the C. raciborskii strains CENA302, CENA303, CENA305 T3 and CYP011K (20 days old) were filtered through Millipore glass fiber filters (Millipore, Milford, MA, USA) to harvest cells. Filters were frozen and later extracted with 0.05 M acetic acid by vortexing and cell disruption by sonication in a water bath for 15 min (2 cycles). After centrifugation (10,000 × g for 10 min), supernatants were recovered and filtered (0.45 µm, PVDF, Millipore) into appropriate vials. To assess the intra- and extracellular content of CYN and STX, 30 mL of homogeneous whole cultures were frozen and freeze-dried. After reconstitution and extraction in 1 mL 0.05 M acetic acid, samples were centrifuged (10,000 × g for 10 min) and filtered into appropriate vials.

Chromatographic analyses were performed with a Shimadzu Prominence (Kyoto, Japan) liquid chromatography system equipped with a diode array detector (SPD-M20A) and coupled to an ion trap mass spectrometer (Esquire HCT, Bruker Daltonics, Billerica, MA, USA). Separation was achieved at 0.2 mL min−1 in a Synergy Hydro column (150×2.0 mm, 4 µm; Phenomenex, CA, USA) under gradient elution of (A) water and (B) acetonitrile-water 90/10, both containing 5 mM ammonium formate and 0.01% formic acid. A linear gradient with a total run time of 35 min was used as follows: 1 to 30% solution B in 15 min, 30 to 90% B in 5 min, 90% B for 2 min, 90 to 1% B in 1 min and 1% B for 12 min. Absorption spectra were acquired in the 200-600 nm wavelength range. Detector effluent was split 1∶4 before entering the mass spectrometer through an electrospray source operated in the positive mode at 4 KV. Analyses were performed using nitrogen as nebulizing (35 psi) and drying gas (5 mL min−1, 300°C) and helium as buffer gas (4×10−6 mbar). Initial survey runs were taken by scanning the m/z range 50–800. In addition, multiple reaction monitoring (MRM) experiments using precursor ions at m/z 416 (CYN) and 400 (7-deoxy-CYN) were performed. For compound identification, product ion spectra were acquired and the fragmentation behavior analyzed according to Dörr et al., [55]. Because a standard solution of 7-epi-CYN was not available, efforts were not undertaken to separate this isomer, and therefore the single chromatographic band at 6.7 min for m/z 416 was assumed as CYN. A commercially available standard of CYN (Abraxis, Warminster, PA, USA) was also used for identity confirmation.

The saxitoxin analogues were investigated by three complementary analytical methods: two post-column oxidation methods with fluorescence detection (HPLC-FD), according to Diener et al., [56], [57], and a method based on hydrophilic interaction liquid chromatography coupled to mass spectrometry (HILIC-MS) following the recommendations of Soto-Liebe et al., [58]. Briefly, the compounds were separated on a Shimadzu Prominence (Kyoto, Japan) liquid chromatography system equipped with a post-column reaction oven and a fluorescence detector (RF-10AX). For HILIC-MS analyses, the chromatography system was coupled to an ion trap mass spectrometer (Esquire HCT, Bruker Daltonics, Billerica, MA, USA) through an electrospray ionization source. The identity of STX derivatives was confirmed by the fragmentation behavior of the [M+H]+ as well as the [M-H] ions [59]. Commercially available standards of STX derivatives (National Research Council/Institute for Marine Biosciences, Halifax, NS, Canada) were employed in HPLC-FD experiments for compound identification.

Enzyme-linked immunosorbent assay (ELISA)

The presence of STX and CYN were tested with the saxitoxin and cylindrospermopsin ELISA kit (Abraxis LLC, PA, USA). All analyses were performed in accordance with the manufacturer’s instructions.


Cylindrospermopsin (cyr) and saxitoxin (sxt) genes

The specific primer set (CYLAT-R/CYLAT-F), targeting the gene encoding the amidinotransferase enzyme involved in the biosynthesis of CYN, successfully amplified cyrA genes from the genomes of four Brazilian C. raciborskii strains (CENA302, CENA303, CENA305 and T3) and also from the Australian strain C. raciborskii CYP011K. These PCR products were sequenced and the almost entire cyrA gene sequences were obtained from the four Brazilian C. raciborskii strains (Table 2). However, the cyrA sequence of the T3 strain showed two nucleotide deletions in the position 525 and 1054 (Figure S1), causing a frameshift mutation. As a result, stop codons are replacing amino acid codons and hence the deduced protein sequence was truncated. The identities of the four cyrA sequences with homologous gene sequences from other CYN-producing closely related strains (C. raciborskii AWT205, C. raciborskii CS505, C. raciborskii CYP011K, R. curvata CHAB1150) were high and were larger than 98.8% (Table S2). However, lower identities were observed with Aphanizomenon ovalisporum (varying 95.5 – 95.8%) and Oscillatoria sp. strain PCC 6506 (varying 86.8 – 87.1%). In addition, the predicted protein functions and domains agreed with proteins from C. raciborskii AWT205 and CS505 (Figure S2), further supporting the identity of the sequences obtained as a CYN synthetase gene. The phylogenetic analysis grouped the cyrA sequences of C. raciborskii Brazilian strains together with other homologous nucleotide sequences from CYN-producing cyanobacteria with highly supported bootstrap value (100% ML and NJ) (Figure 1A). The amidinotransferase enzymes from other organisms grouped separately according to their biosynthetic pathway. Thus, sequences of the sxtG gene, that also encodes an amidinotransferase but is involved in the STX biosynthesis pathway, formed a fully supported clade distantly related to cyrA clade.

Figure 1. Maximum likelihood phylogenetic trees of cyrA (A), cyrB (B) and cyrC (C).

The C. raciborskii Brazilian strains used in this study are shown in bold. Bootstrap test (1,000 resamplings) was performed and values >50% for ML and NJ analyses are shown over the nodes. Cyanobacterial taxa are shown in blue, other organisms in red. (+) CYN-producing and (–) CYN-non-producing strains; (++) STX-producing and (––) STX-non-producing strains.

Table 2. Size of cylindrospermopsin and saxitoxin gene sequences obtained from the Cylindrospermopsis raciborskii strains.

Three Brazilian strains (CENA302, CENA303 and T3) and also the Australian CYP011K strain showed positive results for the presence of a region of cyrB gene that encodes an adenylation domain of a NRPS. However, after sequencing the PCR products, the cyrB gene fragment obtained from the C. raciborskii T3 showed no identity with sequences deposited in the GenBank, an indicative of non-specific PCR amplification. The percentage of identity between partial cyrB adenylation domain of CENA302 and CENA303 was 99.4%, and ranged from 84.6 to 99.6% with other cyrB sequences retrieved from GenBank (Table S2). The phylogenetic analysis grouped the cyrB sequences of the two C. raciborskii Brazilian strains together with other homologous nucleotide sequences from CYN-producing cyanobacteria with highly supported bootstrap value (99% ML and NJ) (Figure 1B).

PCR products of a region of the cyrC gene that encodes a ketosynthase domain of a PKS were obtained from the genomes of the four C. raciborskii Brazilian strains as well as from the Australian CYP011K strain. However, after sequencing the amplicons, the sequences of the strains CENA305 and T3, with 99.0% identity between them, showed the highest identity (76.5% and 76.8%, respectively) with the phosphopantothenoylcysteine decarboxylase/phosphopantothenate-cysteine ligase of the Anabaena sp. 90. In the phylogenetic reconstruction of the partial cyrC sequences of the C. raciborskii Brazilian strains CENA302 and CENA303 a highly supported clade (bootstrap value of 99% ML and NJ) was formed with other homologous nucleotide sequences from CYN-producing cyanobacteria (Figure 1C).

Several attempts to amplify fragments of the cyrJ gene from all four Brazilian C. raciborskii strains were unsuccessful. Only in the Australian strain C. raciborskii CYP011K (CYN producer used as reference) partial cyrJ gene was successfully amplified.

The three sxt gene fragments (sxtA4, sxtB and sxtI) investigated were PCR amplified and sequenced from the genomes of almost all studied cyanobacteria, with the exception of the Australian CYN producer strain CYP011K, in which none of STX genes were detected (Table 2). BLAST analysis showed that the gene sequences of the sxtA4 (8-amino-7-oxanonanoate synthase - AONS), sxtB (cytidine deaminase) and sxtI (O-carbamoyltransferase) from C. raciborskii CENA302, CENA303 and 305 strains had high similarity with the corresponding sequences from the STX-producing Brazilian strains C. raciborskii T3 and Raphidiopsis brookii D9 (Table S3). In the phylogenetic analysis, the sxt sequences clustered with other sequences of STX-producing cyanobacterial strains with supported bootstrap (100% ML and NJ) for sxtA4, (99% ML and NJ) for sxtB and sxtI, (Figure 2). The sxt gene sequences from T3 were identified previously [40]. The sxt genes were also compared with sequences that encode for homologous enzymes from other organisms and demonstrate phylogenetic differences according to its biosynthetic pathway. The predicted SxtA4, SxtB and SxtI protein functions and domains agreed with those homologous proteins from T3 and D9 (Table S3).

Figure 2. Maximum likelihood phylogenetic trees of sxtA (A), sxtB (B) and sxtI (C).

The C. raciborskii Brazilian strains used in this study are shown in bold. Bootstrap test (1,000 resamplings) was performed and values >50% for ML and NJ analyses are shown over the nodes. Cyanobacterial taxa are shown in blue, other organisms in red. (+) STX-producing; (–) STX-non-producing strains.

Chemical and Enzyme Immunoassay analyses of CYN and STX

Despite several attempts to detect CYN and 7-deoxy-CYN derivatives in Brazilian strains, neither compound could be identified. As expected, both compounds were detected in the Australian CYP011K strain in the intra- and extracellular fractions (Figure 3). ELISA immunoassays were unable to detect CYN in the Brazilian strains of C. raciborskii, but showed positive results for Australian strain CYP011K. The lack of cyrJ amplification prompted the search for possible CYN derivatives missing the sulfate group at C-12, but all efforts failed to detect such a compound in the Brazilian isolates.

Figure 3. LC-UV-MS analysis of a freeze-dried culture sample of C. raciborskii CYP011K.

(A) UV trace at 262 nm and extracted ion chromatograms for CYN (m/z 418) and 7-deoxy-CYN (m/z 400). Product ion spectra and UV absorption spectra are depicted for CYN (B and C) and 7-deoxy-CYN (D and E).

Different chromatographic methods were employed to evaluate the production of saxitoxin analogues in the isolated strains. The post-column oxidation methods with fluorescence detection allowed the identification of the epimers GTX2/GTX3, STX and dcSTX in CENA302 (Figure 4A), STX, dcSTX, NEO and dcNEO in CENA305 (Figure 4B), as well as NEO, STX and dcSTX in strain T3 (Figure 4D). These results were further confirmed by HILIC-MS analyses. ELISA immunoassay provided positive results for STX in the strains CENA302, CENA305 and T3. None of the employed methods was able to detected STX analogues in the strains CENA303 and CYP011K.

Figure 4. HILIC-FD chromatograms of STX variants in Brazilian C. raciborskii strains.

(A) commercial standards; (B) C. raciborskii CENA302; (C) C. raciborskii CENA305; (D) C. raciborskii T3. Chromatograms acquired according to Diener et al. (2007). *denotes unspecific peaks.

Morphological and 16S rRNA gene phylogenetic analyses

Morphological analysis of the isolated strains obtained in this study showed that they belong to the order Nostocales, genus Cylindrospermopsis. The filamentous strains presented typical characteristics of the Cylindrospermopsis raciborskii species (Figure 5) such as: subsymmetrical trichomes with cylindrical cells but attenuated and pointed at the ends of trichomes; pale blue-green or yellowish color; presence of aerotopes; conical terminal heterocytes developed after asymmetrical division of the end cells; and elongated oval akinetes situated adjacent to the heterocyte or the terminal vegetative cell [48], [3].

Figure 5. Brigthfield micrograph of C. raciborskii isolates showing details of trichome morphology, vegetative cells, heterocytous (H) and akinete (A).

A-B) C. raciborskii CENA302; C-D) C. raciborskii CENA303; E-F) C. raciborskii CYP011K; G-H) C. raciborskii T3.

The nearly complete 16S rRNA gene sequences of C. raciborskii strains showed high similarity with sequences of C. raciborskii from GenBank (Table S4). The identities between the sequences of the four Brazilian C. raciborskii strains obtained in this study ranged from 99.1 to 100%. Moreover, these four 16S rRNA sequences displayed similarities varying between 99.2 to 99.6% with the sequence of the type strain C. raciborskii AWT205. The 16S rRNA sequences of the CENA302, CENA305 and T3 strains isolated from the Billings reservoir also showed high similarity (99.3%, 99.7% and 99.8%, respectively) with that of the R. brookii D9 found in the same reservoir. The 16 rRNA gene sequence of the C. raciborskii CYP011K showed 99.6 to 99.8% similarities with other sequences of C. raciborskii Australian strains (QHSS/NR/CYL/03, 05E and 23B).

In the phylogenetic tree, the 16S rRNA gene sequences of the C. raciborskii strains fall within a highly supported (bootstrap values of 99 and 100% for ML and NJ algorithms, respectively) major clade containing sequences of planktonic members of C. raciborskii isolated from several countries and an internal separated clade with members of Raphidiopsis genus (Figure 6). Within this major clade, the Brazilian and Australian strains formed distinct clades according to their origin but with low supported bootstrap. Strains of North American, African, European and Asian origin were mixed in other internal clades. The evolutionary relatedness of the four cyanobacterial strains (C. raciborskii AWT205, C. raciborskii CS-505, Aphanizomenon sp. 10E6 and Oscillatoria sp. PCC 6506) with the cyr clusters already described can be visualized in the phylogenetic tree. The phylogeny of the polyphyletic genus Aphanizomenon that possess several CYN and STX producer strains was also given in the phylogenetic tree. After revision of several genera of cyanobacteria, new designation for some of them was adopted and it is also shown in the phylogenetic tree.

Figure 6. Maximum likelihood phylogenetic tree based on the 16S rRNA gene sequences showing the relationships of the studied cyanobacteria (in bold).

Bootstrap test (1,000 resamplings) was performed and values >50% for ML and NJ analyses are shown over the nodes. Branch lengths are proportional to the number of substitutions per site (see scale bar). Taxon name in red or blue denotes STX or CYN producer strains, respectively.


In this study, cylindrospermopsin synthetase (cyr) genes were PCR amplified and sequenced for the first time from the genomes of four non-CYN-producing C. raciborskii strains isolated from Brazilian freshwater environments. Of the four cyr genes investigated and considered essential for the synthesis of CYN [12], [45], [46], three were amplified and sequenced from the genomes of two C. raciborskii Brazilian strains (CENA302 and CENA303). In other two Brazilian strains (CENA305 and T3) only the almost complete cyrA gene sequence was found. In addition, the four cyr genes were sequenced from the CYN-producing Australian strain C. raciborskii CYP011K. This Australian strain produces two CYN analogues, therefore must encode at least 11 genes (cyrA, B, C, D, E, F, G, H, I, J and K), that appear to be directly implicated in the synthesis of CYN according to the four cyr clusters already found in different cyanobacterial CYN-producing genera [60].

The almost complete cyrA sequences obtained from the four non-CYN-producing Brazilian C. raciborskii strains (CENA302, CENA303, CENA305 and T3) resemble those identified in the C. raciborskii strains AWT205 and CS-505 [12], [61]. The cyrA amplification product of T3 strain was achieved using more stringent annealing temperature than that recommended [51]. However, the truncated cyrA sequence of the T3 with the absence of two nucleotides indicated that this gene underwent reduction and was inactivated. The cyrA gene identified in the type strain C. raciborskii AWT205 is 1,176 bp long and encodes an L–arginine:glycine amidinotransferase enzyme responsible for the formation of guanidinoacetate and ornithine from L–arginine and glycine, the first step in the CYN biosynthesis [12], [46]. Amidinotransferases are a monophyletic group of enzymes distributed among vertebrates, plants and prokaryotes [46]. Furthermore, these enzymes are involved in biosynthesis of the neurotoxin saxitoxin in cyanobacteria (SxtG, L–arginine/L–lysine amidinotransferase) and dinoflagellates [40], [62]. Despite a high level of conservation with regard to residues involved in catalysis and substrate binding, these enzymes have different substrate specificities and kinetic mechanisms [63]. The CyrA amidinotransferase was shown to be unique to the metabolic pathway for biosynthesis of CYN [12], [46]. In this manner, the cyrA gene sequences obtained in this study formed a fully supported clade in phylogenetic analysis, together with other cyrA sequences of CYN-producing cyanobacterial strains retrieved from GenBank, but distantly related to the also fully supported clade containing sxtG sequences. This topology is in agreement with cyrA and sxtG phylogenetic reconstruction performed by Orr et al., [62], who defined the cluster containing the cyrA sequences as amidinotransferase 2 and that with sxtG sequences as amidinotransferase 1.

The CyrB enzyme (mixed NRPS-PKS) catalyzes the second reaction in the proposed biosynthetic pathway, incorporating an acetate unit into guanidinoacetate [12]. In the present study, the primer set used for detection of cyrB gene targeted the NRPS adenylation domain, which is responsible for amino acid recognition and activation. Therefore, this adenylation domain of CyrB uses the guanidinoacetate as a substrate for subsequent polyketide extensions. The two partial sequences of cyrB adenylation domain found in the genomes of two non-CYN-producing C. raciborskii Brazilian strains (CENA302 and CENA303) had high identities to other C. raciborskii cyrB adenylation domains and formed a fully supported clade in phylogenetic analysis. The partial cyrB amplification product of T3 strains was also achieved using more stringent annealing temperature than that recommended [51]. Nevertheless, the partial cyrB sequence obtained for T3 showed to be nonspecific amplification that led to a false-positive result. It is worth noting that a study applying neutrality test to the adenylation domain sequences of uncultured A. ovalisporum-like cyrB obtained from environmental samples indicated that it is under purifying selection. Purifying selection makes sure that deleterious mutations cannot take over a population and that any improved structures, once fixed in a population, are maintained as long as they are needed [64]. Furthermore, the cyrB gene of the CYN-producing Oscillatoria sp. PCC 6506 showed moderate identity (79%) to the one characterized in the toxic A. ovalisporum [50]. This Oscillatoria sp. PCC 6506 gene possesses a 150-bp-long GC rich fragment containing repeated sequences which encodes for a proline rich motif (PPLP) repetition that might function as a linker between the ketoreductase and methyltransferase domains of this NRPS-PKS hybrid enzyme. It is thus very likely that the cyrB from distinct cyanobacterial genera is evolutionarily related but that it substantially diverged from a common ancestor. The presence of cyrB sequence homologous to cyrB of the CYN producers was reported before for the non-toxic strain C. raciborskii Hung1 (Hungarian), however, cyrA and cyrC genes were not detected [51].

The CyrC, a polyketide synthase, is the subsequent enzyme proposed in the biosynthesis of CYN. It is responsible for the incorporation of a further acetate unit into guanidinoacetate, while a subsequent keto reduction provides the next intermediate [12]. The primer set used in this study targeted a region of the ketosynthase domain of the CyrC, which was highly similar to other C. raciborskii cyrC ketosynthase domain and formed a fully supported clade in phylogenetic analysis.

The cyrJ gene (780 bp long in the C. raciborskii AWT205) encodes a sulfotransferase, a tailoring enzyme responsible for the sulfation of the CYN [12]. The three structural variants of CYN described so far (CYN, 7-epi-CYN and 7-deoxy-CYN) are sulfated [65]-[67], therefore, the CYN-producing cyanobacteria must have the cyrJ gene. The lack of cyrJ gene amplification in all four Brazilian C. raciborskii strains indicates absence of this gene in the genomes of these cyanobacteria and supports the negative results of chemical analyses. These data also corroborated other studies indicating that cyrJ gene is only present in the genome of CYN-producing cyanobacteria [12], [50], [68]. It remains to be shown if the other nine genes (cyrD, E, F, G, H, I, J and K), that appear to be directly implicated in the synthesis of CYN, are present in these Brazilian C. raciborskii strains.

The importance of this finding is that, to date, only the saxitoxin genes have been reported in C. raciborskii strains isolated from Brazilian freshwater [40], [51] and this study demonstrates for the first time that Brazilian STX-producing C. raciborskii strains also carry fragments of cyr genes. These cyr genes could represent either a remnant or an otherwise ancestral intermediate of a functional CYN gene cluster. The acquisition/loss of cyanotoxin genes among cyanobacteria is not yet understood and vertical and horizontal gene transfer have been suggested for cyr and sxt gene clusters [50], [51], [60], [69][72]. These presumptions may be further elucidated with the increasing discovery of new cyanotoxin producers and their gene clusters.

The selective pressures in Brazilian environments that favored STX-producing C. raciborskii over CYN-producing strains are unknown. The physiological role of these secondary metabolites is still unclear, despite their effects on animal cells being relatively well understood. Recently, Soto-Liebe et al., [73] suggested STX analogues as protective compounds against elevated salt concentration in the environment. It remains to be demonstrated if STX variants produced by Brazilian C. raciborskii strains are replacing a possible cellular function of CYN.

Although the existence of CYN in Brazilian freshwater blooms has been known for several years, C. raciborskii strains that synthesize CYN have not been isolated so far. Thus, it is likely that other cyanobacterial genera are responsible for CYN production in Brazilian environments. In a similar way, no CYN-producing strains of C. raciborskii have been found in North America and Europe [74][76]. So far, in North America only Aphanizomenon strains were found to produce CYN [76], [77], while in Europe strains of Aphanizomenon [78], [79] and Anabaena [80] were identified as CYN-producers.

The presence of partial sxt gene sequences (sxtA4, sxtB and sxtI) in the CENA302, CENA303 and CENA305 strains with high similarities to those of other STX-producing strains already described, as well as the detection of some congeners by chromatographic analyses in the strains CENA302 and CENA305, corroborate previous findings that the Brazilian C. raciborskii strains produce these neurotoxins [7], [41], [42]. Moustafa and collaborators [70] showed that SxtA is comprised of two distinct regions and resulted from the fusion of two proteins acquired from different bacterial sources. The C-terminal region encodes an enzyme that presents significant identity to class I and II aminotransferases from actinobacteria. This region includes the SxtA4 catalytic domain and presents high similarity to AONS [40], [70]. Our phylogenetic analyses demonstrated that sxtA4 gene sequences from cyanobacteria strains are highly conserved and different from the actinobacteria Frankia sp. suggesting that AONS from cyanobacteria is involved only in the STX biosynthesis.

The strain CENA302 was found to produce the epimers GTX2/GTX3, STX and dcSTX while CENA305 produces NEO, STX and dcSTX. Low levels of dcNEO (decarbamoyl neosaxitoxin) congener were detected only by HILIC-MS because co-elution hampered its identification in the HPLC-FD methods. A similar toxin profile was reported for C. raciborskii strains T2 and T3 [7]. Coincidentally, these four strains were isolated from the Billings reservoir in southeastern Brazil (subtropical climate) but at different locations. Our results of the STX toxin profile of C. raciborskii T3 are comparable to those obtained by Soto-Liebe et al., [58]. As noted by these authors, available literature on the toxin profile of T3 has been confusing given that several groups have identified different saxitoxin derivatives. In this study, the production of NEO, STX and dcSTX was confirmed by three complementary analytical techniques while dcNEO was detected only by HILIC-MS.

Previous phylogenetic analysis studies using nucleotide sequences of the fast-evolving 16S-23S internal transcribed spacer, nifH and cpcBA-IGS have shown a separation of C. raciborskii strains according to their geographic origin [81][83]. In our study, a phylogenetic tree based on the low-evolving sequences of 16S rRNA gene also showed a geographic separation of Brazilian and Australian strains according to their origin, but with low supported bootstrap. However, strains of North American, African, European and Asian origin were not geographically separated. If geographic isolation, rather than environmental selection, drives diversity, location-specific lineages would arise in different provinces regardless of microhabitat [84]. Nevertheless, studies on global biogeography of the cyanobacterial genera Chroococcidiopsis [84] and Microcystis [85] concluded that environmental selection drives diversification. Although some studies have shown geographic distribution of C. raciborskii strains, a larger number of nucleotide sequences from a variety of regions of different continents are needed to better understand the distribution and evolution of this cyanobacterial species.


In this study, we identified and sequenced partial cyr and sxt synthetase genes of four Brazilian planktonic C. raciborskii strains, CENA302, CENA303, CENA305 and T3. Although the occurrence of both neurotoxins (anatoxin-a and homoanatoxin-a) and cyr genes has already been documented in a single Oscillatoria sp. strain [50], this is the first report to our knowledge of the presence of sxt and cyr genes in C. raciborskii strains. The results obtained here provide the first insight of the presence of CYN genes in C. raciborskii strains from an American country and contribute to the reconstruction of the evolutionary history and diversification of cyanobacterial toxins.

Supporting Information

Figure S1.

Alignment of partial cyrA nucleotide sequences showing the two nucleotide deletions in the position 525 and 1054 (red marks) of the Cylindrospermopsis raciborskii T3.


Figure S2.

Maximum likelihood phylogenetic trees of Cyr (A) and Sxt (B) amino acids sequences. The C. raciborskii strains used in this study are shown in bold. Bootstrap test (1,000 resamplings) was performed and values >50% for ML and NJ analyses are shown over the nodes.


Table S1.

PCR primer sequences used in this study.


Table S2.

Percentage of identities of cyr nucleotide sequences of CYN-non-producing Brazilian strains of C. raciborskii with other sequences from CYN-producing cyanobacterial strains.


Table S3.

Percentage of identities of sxt nucleotide sequences of C. raciborskii Brazilian strains with other sequences from STX-producing cyanobacterial strains.


Table S4.

The 16S rRNA gene sequences identities among the Brazilian C. raciborskii strains and other sequences of related cyanobacterial strains.



We thank the National Council for Scientific and Technological Development (CNPq) for provided the graduate scholarships to C. Hoff-Risseti and P.D.C. Schaker (142749/2009-5, 132410/2012-5, respectively). We would like to thank E. Crespim for help with primer design. M.F. Fiore would also like to thank CNPq for a research fellowship (306607/2012-3).

Author Contributions

Conceived and designed the experiments: MFF CHR. Performed the experiments: CHR FAD PDCS. Analyzed the data: CHR FAD PDCS EP VRW MFF. Contributed reagents/materials/analysis tools: MFF EP VRW. Wrote the paper: CHR MFF FAD PDCS EP VRW.


  1. 1. Seenayya G, Subba Raju N (1972) On the ecology and systematic position of the algae known as Anabaenopsis raciborskii (Wolosz) Elenkin and a critical evaluation of the forms described under the genus Anabaenopsis. In: Desikachary, TV (1972) Taxonomy and Biology of Blue-green Algae. India: University of Madras. pp 52–57.
  2. 2. Hoffmann L, Komárek J, Kaštovský J (2005) System of cyanoprokaryotes (Cyanobacteria): state in 2004. Algol Stud 117: 95–115.
  3. 3. Komárek J, Hauer T (2013) CyanoDBcz: On-line database of cyanobacterial genera University of South Bohemia and Institute of Botany AS CR. Available: Accessed 25 March 2013.
  4. 4. Hawkins PR, Chandrasena NR, Jones GJ, Humpage AR, Falconer IR (1997) Isolation and toxicity of Cylindrospermopsis raciborskii from an ornamental lake. Toxicon 35: 341–346.
  5. 5. Castenholz RW (2001) Phylum BX Cyanobacteria oxygenic photosynthetic bacteria. In: Boone DR, Castenholz RW, Garrity GM, eds. Bergey’s manual of systematic bacteriology. New York: Springer-Verlag. pp 474–599.
  6. 6. Hawkins PR, Runnegar MTC, Jackson ARB, Falconer IR (1985) Severe hepatotoxicity caused by the tropical cyanobacterium (blue-green-alga) Cylindrospermopsis raciborskii (Woloszynska) Seenaya and Subba Raju isolated from a domestic water-supply reservoir. Appl Environ Microbiol 50: 1292–1295.
  7. 7. Lagos N, Onodera H, Zagatto PA, Andrinolo D, Azevedo SMFQ, et al. (1999) The first evidence of paralytic shellfish toxins in the fresh water cyanobacterium Cylindrospermopsis raciborskii, isolated from Brazil. Toxicon 37: 1359–1373.
  8. 8. Dittmann E, Fewer DP, Neilan BA (2012) Cyanobacterial toxins: biosynthetic routes and evolutionary roots. FEMS Microbiol Rev 37: 23–43.
  9. 9. Terao K, Ohmori S, Igarashi K, Ohtani I, Watanabe MF, et al. (1994) Electron microscopic studies on experimental poisoning in mice induced by cylindrospermopsin isolated from blue-green alga Umezakia natans. Toxicon 32: 833–843.
  10. 10. Humpage AR, Fenech M, Thomas P, Falconer IR (2000) Micronucleus induction and chromosome loss in transformed human white cells indicate clastogenic and aneugenic action of the cyanobacterial toxin, cylindrospermopsin. Mutat Res 472: 155–161.
  11. 11. Froscio SM, Humpage AR, Burcham PC, Falconer IR (2003) Cylindrospermopsin induced protein synthesis inhibition and its dissociation from acute toxicity in mouse hepatocytes. Environ Toxicol 18: 243–251.
  12. 12. Mihali TK, Kellmann R, Muenchhoff J, Barrow KD, Neilan BA (2008) Characterization of the gene cluster responsible for cylindrospermopsin biosynthesis. Appl Environ Microbiol 74: 716–722.
  13. 13. Wang J, Salata JJ, Bennett PB (2003) Saxitoxin is a gating modifier of HERG K+ channels. J Gen Physiol 121: 583–598.
  14. 14. Su Z, Sheets M, Ishida H, Li F, Barry WH (2004) Saxitoxin blocks L-type Ica. J Pharmacol Exp Ther 308: 324–329.
  15. 15. Padisák J (1997) Cylindrospermopsis raciborskii (Woloszynska) Seenayya et Subba Raju, an expanding, highly adaptative cyanobacterium: worldwide distribution and review of its ecology. Arch Hydrobiol 107: 563–593.
  16. 16. Sant’Anna CL, Azevedo MTP (2000) Contribution to the knowledge of potentially toxic Cyanobacteria from Brazil. Nova Hedwigia 71: 359–385.
  17. 17. Briand JF, Robillot C, Quiblier-Lloberas C, Humbert JF, Couté A, et al. (2002) Environmental context of Cylindrospermopsis raciborskii (Cyanobacteria) blooms in a shallow pond in France. Water Res 36: 3183–3192.
  18. 18. Wiedner C, Rücker J, Brüggemann R, Nixdorf B (2007) Climate change affects timing and size of populations of an invasive cyanobacterium in temperate regions. Oecologia 152: 473–484.
  19. 19. Sinha R, Pearson LA, Davis TW, Burford MA, Orr PT, et al. (2012) Increased incidence of Cylindrospermopsis raciborskii in temperate zones - Is climate change responsible? Water Res 46: 1408–1419.
  20. 20. Branco CWC, Senna PAC (1991) The taxonomic elucidation of the Paranoá Lake (Brasilia, Brazil) problem: Cylindrospermopsis raciborskii.. Bull Jard Bot Nat Belg 61: 85–91.
  21. 21. de Souza RCR, Carvalho MC, Truzzi AC (1998) Cylindrospermopsis raciborskii (Wolosz) Seenaya and Subba Raju (Cyanophyceae) dominance and a contribution to the knowledge of Rio Pequeno arm, Billings Reservoir, Brazil. Environ Toxicol Water Qual 13: 73–81.
  22. 22. Komárková J, Laudares-Silva R, Senna PAC (1999) Extreme morphology of Cylindrospermopsis raciborskii (Nostocales, Cyanobacteria) in the Lagoa do Peri, a freshwater coastal lagoon, Santa Catarina, Brazil. Algol Stud 94: 207–222.
  23. 23. Bouvy M, Falcão D, Marinho M, Pagano M, Moura A (2000) Occurrence of Cylindrospermopsis (Cyanobacteria) in 39 Brazilian tropical reservoirs during the 1998 drought. Aquat Microb Ecol 23: 13–27.
  24. 24. Tucci A, Sant’Anna CL (2003) Cylindrospermopsis raciborskii (Woloszynska) Seenayya and Suba Raju (Cyanobacteria): weekly variation and relation with environmental factors in an eutrophic lake, São Paulo, SP, Brazil. Rev Bras Bot 26: 97–112.
  25. 25. Yunes JS, Cunha NT, Proença LA, Monserrat JM (2003) Cyanobacterial neurotoxins from Southern Brazilian Freshwaters. Comm Toxicol 9: 103–115.
  26. 26. Costa IAS, Azevedo SMFO, Senna PAC, Bernardo RR, Costa SM, et al. (2006) Occurrence of toxin-producing cyanobacteria blooms in a Brazilian semiarid reservoir. Braz J Biol 66: 211–219.
  27. 27. Carvalho LR, Pipole F, Werner VR, Laughinghouse HD, de Camargo ACM, et al. (2008) A toxic cyanobacterial bloom in an urban coastal lake, Rio Grande do Sul State, Southern Brazil. Braz J Microbiol 39: 761–769.
  28. 28. Gemelgo MCP, Sant’Anna CL, Tucci A, Barbosa HR (2008) Population dynamics of Cylindrospermopsis raciborskii (Woloszynska) Seenayya & Subba Raju, a Cyanobacteria toxic species, in water supply reservoirs in São Paulo, Brazil. Hoehnea 35: 297–307.
  29. 29. Figueredo CC, Giani A (2009) Phytoplankton community in the tropical lake of Lagoa Santa (Brazil): conditions favoring a persistent bloom of Cylindrospermopsis raciborskii.. Limnol 39: 264–272.
  30. 30. Moschini-Carlos V, Bortoli S, Pinto E, Nishimura PY, de Freitas LG, et al. (2009) Cyanobacteria and cyanotoxin in the Billings reservoir (São Paulo, SP, Brazil). Limnetica 28: 273–282.
  31. 31. Soares MCS, Rocha MIA, Marinho MM, Azevedo SMFO, Branco CWC, et al. (2009) Changes in species composition during annual cyanobacterial dominance in a tropical reservoir: physical factors, nutrients and grazing effects. Aquat Microb Ecol 57: 137–149.
  32. 32. Huszar VLM, Silva LHS, Marinho M, Domingos P, Sant’Anna CL (2000) Cyanoprokariote assemblage in eight productive tropical Brazilian waters. Hydrobiology 424: 67–77.
  33. 33. Couté A, Bouvy M (2004) A new species of the genus Cylindrospermopsis, C acuminato-crispa spec. nova (Cyanophyceae, Nostocales) from Ingazeira reservoir, Northeast Brazil. Algol Stud 113: 57–72.
  34. 34. Zagatto PA (1998) Toxicity of the algae Cylindrospermopsis raciborskii isolated from Billings reservoir, Taquacetuba branch Technical Report, September 1998 Environmental Agency of São Paulo State (CETESB), São Paulo, SP, pp 23.
  35. 35. Saker ML, Thomas AD, Norton JH (1999) Cattle mortality attributed to the toxic cyanobacterium Cylindrospermopsis raciborskii in an outback region of north Queensland. Environ Toxicol 14: 179–182.
  36. 36. Li R, Carmichael WW, Brittain S, Eaglesham GK, Shaw GR, et al. (2001b) First report of the cyanotoxins cylindrospermopsin and deoxycylindrospermopsin from Raphidiopsis curvata (Cyanobacteria). J Phycol 37: 1121–1126.
  37. 37. Wood SA, Stirling DJ (2003) First identification of the cylindrospermopsin-producing cyanobacterium Cylindrospermopsis raciborskii in New Zealand. N Z J Mar Freshw Res 37: 821–828.
  38. 38. Chonudomkul D, Yongmanitchai W, Theeragool G, Kawachi M, Kasai F, et al. (2004) Morphology, genetic diversity, temperature tolerant and toxicity of Cylindrospermopsis raciborskii (Nostocales, Cyanobacteria) strains from Thailand and Japan. FEMS Microbiol Ecol 48: 345–355.
  39. 39. Everson S, Fabbro L, Susan Kinnear S, Wright P (2011) Extreme differences in akinete, heterocyte and cylindrospermopsin concentrations with depth in a successive bloom involving Aphanizomenon ovalisporum (Forti) and Cylindrospermopsis raciborskii (Woloszynska) Seenaya and Subba Raju. Harmful Algae 10: 265–276.
  40. 40. Kellmann R, Mihali TK, Jeon YJ, Pickford R, Pomati F, et al. (2008) Biosynthetic intermediate analysis and functional homology reveal a saxitoxin gene cluster in cyanobacteria. Appl Environ Microbiol 74: 4044–4053.
  41. 41. Molica R, Onodera H, García C, Rivas M, Andrinolo D, et al. (2002) Toxins in the freshwater cyanobacterium Cylindrospermopsis raciborskii (Cyanophyceae) isolated from Tabocas reservoir in Caruaru, Brazil, including demonstration of a new saxitoxin analogue. Phycologia 41: 606–611.
  42. 42. Ferrão-Filho AS, Cunha R, Magalhães VF, Soares MCS, Baptista DF (2007) Evaluation of sub-lethal toxicity of cyanobacteria on the swimming activity of aquatic organisms by image analysis. J Braz Soc Ecotoxicol 2: 93–100.
  43. 43. Carmichael WW, Azevedo SMFO, An JS, Molica RJR, Jochimsen EM, et al. (2001) Human Fatalities from Cyanobacteria: Chemical and Biological Evidence for Cyanotoxins. Environ Health Persp 109: 663–668.
  44. 44. Bittencourt-Oliveira MC, Piccin-Santos V, Kujbida P, Moura AN (2011) Cylindrospermopsin in water supply reservoirs in Brazil determined by immunochemical and molecular methods. J Water Res Protec 3: 349–355.
  45. 45. Schembri MA, Neilan BA, Saint CP (2001) Identification of genes implicated in toxin production in the cyanobacterium Cylindrospermopsis raciborskii. Environ Toxicol 16: 413–421.
  46. 46. Muenchhoff J, Siddiqui KS, Poljak A, Raftery MJ, Barrow KD (2010) A novel prokaryotic L-arginine:glycine amidinotransferase is involved in cylindrospermopsin biosynthesis. FEBS Journal 277: 3844–3860.
  47. 47. Gorham PR, McLahlan JR, Hammer VT, Kim WK (1964) Isolation and culture of toxic strains of Anabaena flos-aquae (Lyngb) de Bréb Verh int Verein. Theor Angew Limnol 15: 796–804.
  48. 48. Cronberg G, Annadotter H (2006) Manual on aquatic cyanobacteria: a photo guide and a synopsis of their toxicology. International Society for the Study of Harmful AlgaeUnited Nations Educational, Scientific and Cultural Organization, Copenhagen. 106 p.
  49. 49. Fiore MF, Moon DH, Tsai SM, Lee H, Trevors JT (2000) Miniprep DNA isolation from unicellular and filamentous cyanobacteria. J Microbiol Meth 39: 159–169.
  50. 50. Mazmouz R, Chapuis-Hugon F, Mann S, Pichon V, Méjean A, et al. (2010) Biosynthesis of cylindrospermopsin and 7-epicylindrospermopsin in Oscillatoria sp strain PCC 6506: identification of the cyr gene cluster and toxin analysis. Appl Environ Microbiol 76: 4943–4949.
  51. 51. Kellmann R, Mills T, Neilan BA (2006) Functional modeling and phylogenetic distribution of putative cylindrospermopsin biosynthesis Enzymes. J Mol Evol 62: 267–280.
  52. 52. Birnboim HC, Doly J (1979) A rapid alkaline extraction procedure for screening recombinant plasmid DNA. Nucleic Acids Res 7: 1513–1518.
  53. 53. Fiore MF, Sant’Anna CL, Azevedo MTP, Komárek J, Kaštovský J (2007) The cyanobacterial genus Brasilonema, gen. nov., a molecular and phenotypic evaluation. J Phycol 43: 789–798.
  54. 54. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, et al. (2011) MEGA5: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol Biol Evol 28: 2731–2739.
  55. 55. Dörr FA, Tomaz JC, Lopes NP, Pinto E (2008) Comparative analysis of the gas-phase reactions of cylindrospermopsin and the difference in the alkali metal cation mobility. Rapid Commun Mass Sp 22: 2015–2020.
  56. 56. Diener M, Erler K, Hiller S, Christian B, Luckas B (2006) Determination of Paralytic Shellfish Poisoning (PSP) toxins in dietary supplements by application of a new HPLC/FD method. Eur Food Res Technol 224: 147–151.
  57. 57. Diener M, Erler K, Christian B, Luckas B (2007) Application of a new zwitterionic hydrophillic interaction chromatography column for determination of paralytic shellfish poisoning toxins. J Sep Sci 30: 1821–1826.
  58. 58. Soto-Liebe K, Murillo AA, Krock B, Stucken K, Fuentes-Valdés JJ, et al. (2010) Reassessment of the toxin profile of Cylindrospermopsis raciborskii T3 and function of putative sulfotransferases in synthesis of sulfated and sulfonated PSP toxins. Toxicon 56: 1350–1361.
  59. 59. Dörr FA, Kovačević B, Maksić ZB, Pinto E, Volmer DA (2011) Intriguing differences in the gas-phase dissociation behavior of protonated and deprotonated gonyautoxin epimers. J Am Soc Mass Spectro 22: 2011–2020.
  60. 60. Jiang Y, Xiao P, Yu G, Sano T, Pan Q, et al. (2012) Molecular basis and phylogenetic implications of deoxycylindrospermopsin biosynthesis in the cyanobacterium Raphidiopsis curvata. Appl Environ Microbiol 78: 2256–2263.
  61. 61. Stucken K, John U, Cembella A, Murillo AA, Soto-Liebe K, et al. (2010) The smallest known genomes of multicellular and toxic cyanobacteria: comparison, minimal gene sets for linked traits and the evolutionary implications. PLoS One 5: e9235.
  62. 62. Orr RJS, Stüken A, Murray SA, Jakobsen KS (2013) Evolutionary acquisition and loss of saxitoxin biosynthesis in dinoflagellates: the second “core” gene, sxtG. Appl Environ Microbiol 79: 2128–2136.
  63. 63. Muenchhoff J, Siddiqui KS, Neilan BA (2012) Identification of two residues essential for the stringent substrate specificity and active site stability of the prokaryotic L-arginine:glycine amidinotransferase CyrA. FEBS Journal 279: 805–815.
  64. 64. Loewe L, Cutter AD (2008) On the potential for extinction by Muller's Ratchet in Caenorhabditis elegans. BMC Evol Biol 8: 125.
  65. 65. Ohtani I, Moore RE, Runnegar MTC (1992) Cylindrospermopsin: A potent hepatotoxin from the blue-green alga Cylindrospermopsis raciborskii. J Am Chem Soc 114: 7942–7944.
  66. 66. Banker R, Teltsch B, Sukenik A, Carmeli S (2000) 7-epicylindrospermopsin, a toxic minor metabolite of the cyanobacterium Aphanizomenon ovalisporum from Lake Kinneret, Israel. J Nat Prod 63: 387–389.
  67. 67. Li R, Carmichael WW, Brittain S, Eaglesham GK, Shaw GR, et al. (2001a) Isolation and identification of the cyanotoxin cylindrospermopsin and deoxy-cylindrospermopsin from a Thailand strain of Cylindrospermopsis raciborskii (Cyanobacteria). Toxicon 39: 973–980.
  68. 68. Ballot A, Ramm J, Rundberget T, Kaplan-Levy RN, Hadas O, et al. (2011) Occurrence of non-cylindrospermopsin producing Aphanizomenon ovalisporum and Anabaena bergii in Lake Kinneret (Israel). J Plankton Res 33: 1736–1746.
  69. 69. Stucken K, Murillo AA, Soto-Liebe K, Fuentes-Valdés JJ, Méndez MA (2009) Toxicity phenotype does not correlate with phylogeny of Cylindrospermopsis raciborskii strains. Syst Appl Microbiol 32: 37–48.
  70. 70. Moustafa A, Loram JE, Hackett JD, Anderson DM, Plumley FG, et al. (2009) Origin of saxitoxin biosynthetic genes in cyanobacteria. PLoS ONE 4: e5758.
  71. 71. Stüken A, Jakobsen KS (2010) The cylindrospermopsin gene cluster of Aphanizomenon sp. strain 10E6: organization and recombination. Microbiol 156: 2438–2451.
  72. 72. Hackett JD, Wisecaver JH, Brosnahan ML, Kulis DM, Anderson DM, et al. (2012) Evolution of Saxitoxin Synthesis in Cyanobacteria and dinoflagellates. Mol BioL Evol 30: 70–78.
  73. 73. Soto-Liebe K, Méndez MA, Fuenzalida L, Krock B, Cembella A, et al. (2012) PSP toxin release from the cyanobacterium R. brookii D9 (Nostocales) can be induced by sodium and potassium ions. Toxicon 60: 1324–1334.
  74. 74. Fastner J, Heinze R, Humpage AR, Mischke U, Eaglesham GK, et al. (2003) Cylindrospermopsin occurrence in two German lakes and preliminary assessment of toxicity and toxin production of Cylindrospermopsis raciborskii (Cyanobacteria) isolates. Toxicon 42: 313–321.
  75. 75. Neilan BA, Saker ML, Fastner J, Törökné A, Burns BP (2003) Phylogeography of the invasive cyanobacterium Cylindrospermopsis raciborskii. Mol Ecol 12: 133–140.
  76. 76. Yilmaz M, Phlips EJ, Szabo NJ, Badylak S (2008) A comparative study of Florida strains of Cylindrospermopsis and Aphanizomenon for cylindrospermopsin production. Toxicon 51: 130–139.
  77. 77. Yilmaz M, Phlips EJ (2011) Diversity of and selection acting on cylindrospermopsin cyrB gene adenylation domain Sequences in Florida. Appl Environ Microbiol 77: 2502–2507.
  78. 78. Preuβel K, Stüken A, Wiedner C, Chorus I, Fastner J (2006) First report on cylindrospermopsin producing Aphanizomenon flos-aquae (Cyanobacteria) isolated from two German lakes. Toxicon 47: 156–162.
  79. 79. Wormer L, Cires S, Carrasco D, Quesada A (2008) Cylindrospermopsin is not degraded by co-occurring natural bacterial communities during a 40-day study. Harmful Algae 7: 206–213.
  80. 80. Spoof L, Berg KA, Rapala J, Lahti K, Lepistö L, et al. (2006) First observation of cylindrospermopsin Anabaena lapponica isolated from the boreal environment (Finland). Environ Toxicol 21: 552–560.
  81. 81. Dyble J, Paerl HW, Neilan BA (2002) Genetic characterization of Cylindrospermopsis raciborskii (Cyanobacteria) isolates from diverse geographic origins based on nifH and cpcBA-IGS nucleotide sequence analysis. Appl Environ Microbiol 68: 2567–2571.
  82. 82. Gugger M, Molica R, Le Berre B, Dufour P, Bernard C (2005) Genetic diversity of Cylindrospermopsis strains (Cyanobacteria) isolated from four continents. Appl Environ Microbiol 71: 1097–1100.
  83. 83. Piccini C, Aubriot L, Fabre A, Amaral V, González-Piana M, et al. (2011) Genetic and eco-physiologycal differences of South American Cylindrospermopsis raciborskii isolates support the hypothesis of multiple ecotypes. Harmful Algae 10: 644–653.
  84. 84. Bahl J, Lau MCY, Smith GJD, Vijaykrishna D, Cary SC, et al. (2011) Ancient origins determine global biogeography of hot and cold desert cyanobacteria. Nat Commun 2: 163.
  85. 85. van Gremberghe I, Leliaert F, Mergeay J, Vanormelingen P, Van der Gucht K, et al. (2011) Lack of phylogeographic structure in the freshwater cyanobacterium Microcystis aeruginosa suggests global dispersal. PLoS One 6: e19561.