Comparison of microcystin synthetase genes.
Abstract
Although cyanobacteria are essential microorganisms on earth, some cyanobacteria produce toxins known as cyanotoxins, threatening humans and animals’ health. Hence, it is imperative to rapidly and accurately identify those toxic cyanobacteria. Unfortunately, traditional microscopic methods have limitations for accurate identification due to the lack of discernable morphological difference between toxic and non-toxic strains within the same cyanobacterial species or genus. In contrast, their genetic profiles are inherently conserved; therefore, nucleic acid-based assays can be more reliable for precise identification. Furthermore, molecular assays can provide high throughput and significantly reduce the turnaround time of test results. Such advantages make those assays a preferred method for rapid detection and early warning of potential toxicity. Toxigenic cyanobacterial species have synthetase genes (DNAs) for toxin production, which can be excellent marker genes. Numerous molecular assays targeting cyanotoxin synthetase genes have been developed for the identification of toxigenic cyanobacteria at various taxonomic levels. Polymerase chain reaction (PCR)-based assays are the most prevailing. Among different versions of PCR assays, the real-time quantitative PCR can be utilized to quantify the genes of interest in samples, fulfilling the purpose of both taxonomic recognition and biomass estimation. Reverse transcription (RT)-PCR assays can be used to detect transcripts (i.e., mRNAs) from toxin synthetase genes, probably enhancing the predictive value of PCR detection for toxin production from observed cyanobacterial species. Nevertheless, the utility of toxin synthetase gene- or its transcript-based PCR assays for routine cyanotoxin monitoring needs to be further evaluated on a large scale.
Keywords
- cyanobacteria
- cyanotoxins
- toxin synthetase genes
- molecular techniques
- polymerase chain reaction
1. Introduction
Cyanobacteria are essential microorganisms on earth as they produce oxygen and account for a large part of primary aquatic productivity. Simultaneously, some freshwater cyanobacteria can produce various toxins, named cyanotoxins, some of which are potently poisonous to humans and animals. A well-known cyanotoxicosis in humans was reported from Brazil in association with medical malpractice in 1996. In this incident, 126 patients in a hemodialysis unit were affected, and 60 of them died due to using microcystin-contaminated water from a local reservoir. A cyanobacterial bloom was found in that reservoir concurrently [1]. Besides, there have been reports concerning human cyanotoxin poisoning by drinking water or via injury after contacting recreational water [2]. Apart from humans, numerous animal poisoning cases have also taken place because they can reach the unprocessed natural water directly so that the risk of being poisoned becomes higher. These cases involve livestock, pets, and wildlife [3, 4, 5, 6, 7, 8, 9, 10].
Cyanobacterial blooms occurred more frequently in recent years, which may have been attributed to the aggravating eutrophication in freshwater and global warming. As such, cyanotoxin poisoning incidents have also been increasingly reported. Nowadays, freshwater cyanobacterial blooms have broader geographical and temporal impacts on local water bodies that act as vital municipal or agricultural water supplies. With the possibility of cyanotoxin contamination, humans and animals residing in surrounding areas continue to be threatened. Therefore, testing for toxic cyanobacteria or cyanotoxins is imperative for detection and preventive measures.
Although cyanobacteria can be observed under a microscope, their toxigenicity cannot be determined by microscopy because the toxigenic cyanobacteria do not have unique morphological characteristics. Some laboratories have adopted a testing strategy that combines microscopic observation and cyanotoxin detection to indicate the existence of toxigenic cyanobacteria in samples. Although this strategy may seem reasonable and pragmatic, it needs collaboration between chemical analysts and microalgal biologists to reach an agreement on the conclusion. Furthermore, it neglects the complex phenomena of the same toxin production by different species or genera, leading to an incorrect judgment of the truly culpable toxin producers.
Cyanotoxin testing has been in place. Yet, available tests have shortcomings. For example, commercial enzyme-linked immunosorbent assays (ELISAs) have been widely employed in water testing for cyanotoxins. However, it still has issues, such as low sensitivity [11] or inaccuracy. Erroneous detection is due to the cross-reactivity of isomorphic substances with targets. False-positive results can occur in a worst-case scenario [12]. The high performance liquid chromatography (HPLC) and liquid chromatography-mass spectrometry (LC–MS) are the most accurate analytical methods and have been often employed in cyanotoxin testing [11, 13, 14, 15, 16]. But they require exquisite instruments and complicated operations, making them not as affordable as ELISA-based testing. Aside from these limitations, chemical testing can only tell the presence and/or quantity of cyanotoxins without identifying the toxin producer(s). However, it is crucial to recognize the existence of toxigenic cyanobacteria in water bodies for monitoring and early warning of cyanotoxin poisoning incidents.
It is known that cyanotoxin synthesis is catalyzed by a string of relevant enzymes encoded by toxin synthetase genes [17, 18, 19, 20, 21, 22, 23]. Lack of essential genes for forming a toxin backbone or disruption of the enzymatic cascade toward toxin production results in the failure of toxin synthesis. Therefore, the detection of toxin synthetase genes in samples by a molecular test can disclose the presence or absence of toxigenic cyanobacteria. In this chapter, we review the application of molecular techniques, particularly PCR-based assays, for detecting toxigenic cyanobacteria in freshwater.
2. General genomic organization of toxigenic cyanobacteria
Like other bacteria, cyanobacteria often have one circular chromosome and a few plasmids that consist of the whole genome. The cyanobacterial chromosome is a few megabases in size and contains most of the genes, while plasmids play a role in transferring DNA elements. Compared to the eukaryotic microalgae, the cyanobacterial genome is highly compressed but still contains all genes essential for aquatic and photosynthetic life. Some species even have genes that can facilitate competitive superiority in the environment. For example, gas vesicle genes in
Cyanotoxin synthetase genes often cluster together in the genome and constitute one or more operons that are transcribed in identical or opposite directions [19, 21, 22, 23, 26]. The reason for such an arrangement is likely that the transcription can be well regulated so that all pertaining genes are transcribed simultaneously. This process may ensure that all necessary enzymes/proteins are present for subsequent toxin synthesis. The whole-genome sequencing of toxic cyanobacteria to date has demonstrated only a single copy of the toxin gene cluster in the cyanobacterial genome [27, 28, 29]. The toxin synthetase genes have conserved sequences encoding conserved domains/motifs in the corresponding proteins with specific functions during toxin syntheses, such as polyketide synthesis, adenylation, and methylation. The genes are always clustered closely with whose proteins conduct successive functions in a cascade reaction. It should be reiterated that the synthetase genes are indispensable for toxin production, making them the ideal targets for molecular detection.
Cyanotoxins are traditionally named after the first identified toxin-producing genus, as in the case of microcystin (
3. Cyanotoxins and toxin biosynthesis
3.1 Microcystin
Microcystin is the most common cyanotoxin implicated in human and animal poisoning incidents [36, 37, 38]. It is a hepatotoxin and thus can cause severe impairment in the liver when ingested by the casualties. The toxin is known to be produced by several genera of cyanobacteria, such as
Microcystin is a cyclic heptapeptide that inhibits the eukaryotic protein phosphatase type 1 and 2A in humans and animals by forming an irreversible covalent bond to a cysteine in the catalytic domain of these enzymes. It consists of the following amino acids: D-alanine, X, D-MeAsp (D-erythro-ß-methyl-aspartic acid), Z, Adda ((2S,3S,8S,9S)-3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4,6-dienoic acid), D-glutamic acid, and Mdha (N-methyldehydroalanine). X and Z represent variable L amino acids. It has reportedly over 80 variants, mostly differing in amino acids at the positions X and Z [39].
Microcystin is a non-ribosomal oligopeptide, which means unlike most of the peptides and proteins, it is not synthesized by cellular ribosomes. The enzymes responsible for its synthesis contain the non-ribosomal peptide synthetases (NRPS) and polyketide synthases (PKS) modules as well as tailoring functional domains. All the enzymes are the protein products encoded by the microcystin synthetase genes (
Gene | Size (bp)1 | Encoded domain or function2 | Existence in different genera | ||
---|---|---|---|---|---|
8838 | NRPS, C, NMT, E | yes | yes | yes | |
6318 | NRPS, A, T, C | yes | yes | yes | |
3876 | NRPS, C, A, TE | yes | yes | yes | |
11721 | PKS, KS, AT, KR, DH, ACP, CM | yes | yes | yes | |
10464 | PKS, NRPS, KS, AT, ACP, CM, AMT | yes | yes | yes | |
756 | Racemase | yes | yes | no | |
7896 | NRPS, PKS, KS, AT, CM, DH, KR, ACP | yes | yes | yes | |
1617 | Transporter | yes | yes | yes | |
1014 | Dehydrogenase | yes | yes | no | |
837 | OM | yes | yes | yes | |
< 1000 | TE | no | no | yes |
Per annotation of
3.2 Anatoxin-a
The cyanobacterial alkaloid anatoxin-a has been found in different genera, such as
Although anatoxin-a doesn’t look structurally complicated, its synthesis still requires a cascade of many enzymes whose genes known as anatoxin-a synthetase genes (
Gene | Size (bp)1 | Encoded domain or function2 | Existence in different genera | ||
---|---|---|---|---|---|
750 | TE | yes | yes | yes | |
1143 | Proline-ACP oxidase | yes | yes | yes | |
1596 | Proline adenylation | yes | yes | yes | |
273 | Acyl carrier | yes | yes | yes | |
6438 | PKS, KS, AT, DH, ER, KR, ACP | yes | yes | yes | |
5619 | PKS, KS, AT, DH, KR, ACP | yes | yes | yes | |
4896 | PKS, KS, AT, CM, ACP | yes | yes | yes | |
< 1000 | Transposase | no | yes | yes | |
< 2000 | Transporter | yes | yes | yes | |
723 | Cyclase | yes | yes | yes | |
<1000 | Reductase | no | no | yes |
To start the anatoxin-a synthesis, AnaC activates and tethers the precursor proline to AnaD, which covalently combines with the proline. Then AnaB dehydrogenates the heterocyclic ring of proline to form a “C=N” double bond. AnaE introduces a carbonyl group into its connection with the heterocycle passed from AnaD. Then AnaJ catalyzes a cyclization step to form the characteristic bicyclic ring structure of anatoxin-a by connecting the heterocyclic ring with the backbone. At the same time, the growing chain is bound to the acyl carrier protein domain of AnaF. Finally, the bicyclic thioester is transferred to AnaG for chain extension by adding an acyl group, followed by the enzymatic reaction of AnaA to break the single “SCO-C” covalent bond connecting the enzyme (AnaG) and final product for the completion and releasing of anatoxin-a. Similar to its counterpart McyH in microcystin-producing cyanobacteria, AnaI transports the toxin through the cytomembrane. The rest of the Ana proteins are not commonly shared across different genera and have their own functions. AnaH is a transposase only found in
3.3 Cylindrospermopsin
Cylindrospermopsin can be produced by various cyanobacterial genera, such as
Cylindrospermopsin is synthesized via a string of NRPS/PKS reactions conducted by up to over a dozen Cyr proteins (Table 3, Figure 3). The cylindrospermopsin synthetase genes (
Gene | Size (bp)1 | Encoded domain or function2 | Existence in different genera | ||
---|---|---|---|---|---|
1176 | AMT | yes | yes | yes | |
8754 | NRPS, PKS, PCP, KS, AT, DH, MT, KR, ACP | yes | yes | yes | |
5005 | PKS, KS, AT, KR, ACP | yes | yes | yes | |
5631 | PKS, KS, AT, DH, KR, ACP | yes | yes | yes | |
5667 | PKS, KS, AT, DH, KR, ACP | yes | yes | yes | |
4074 | PKS, KS, AT, ACP | yes | yes | yes | |
1437 | Uracil ring formation | yes | yes | yes | |
1431 | Uracil ring formation | yes | yes | yes | |
831 | Hydroxylation | yes | yes | yes | |
780 | Sulfotransferase | yes | yes | yes | |
1398 | Exporter | yes | yes | yes | |
750 | Transposase | yes | no | no | |
318 | Transposase | yes | no | no | |
600 | Adenylylsulfate kinase | yes | no | yes | |
1548 | Regulator | yes | no | yes | |
152 | ATP-grasp protein | no | no | yes | |
404 | Transposase | no | no | yes | |
299 | Transposase | no | no | yes |
As
3.4 Nodularin
Nodularin is a cyclic pentapeptide and has the identical chemical structure as microcystin except the lack of D-alanine and the amino acid at position X. The mechanism of its toxicity is the same as microcystin’s, i.e., inhibiting the eukaryotic protein phosphatase catalytic subunit type 1 and 2A and leading to severe liver damage. Different from the three aforementioned cyanotoxins, nodularin is solely found in
Gene | Size (bp)1 | Encoded domain or function2 |
---|---|---|
2607 | NRPS, A, NM, PCP, C | |
1299 | C, A, PCP, TE | |
2640 | NRPS, PKS, A, PCP, KS, AT, CM, KR, ACP | |
3872 | PKS, KS, AT, CM, DH, KR, ACP | |
927 | OM | |
3475 | PKS, NRPS, KS, AT, CM, ACP, AMT, C, A, PCP | |
235 | Racemase | |
341 | D-3-phosphoglycerate dehydrogenase | |
601 | ABC transporter |
Nodularin synthesis is conducted putatively according to the annotated functions of each Nda protein. NdaC activates the starter unit as phenylalanine or phenylacetate, and then NdaE catalyzes the transfer of a methyl group to the growing chain. NdaD is involved in two further polyketide extension steps, and NdaF facilitates the final round of polyketide extension and the biosynthesis of Adda. Next, epimerization of L-glutamic acid is catalyzed by NdaG, followed by the peptide condensation carried out by NdaA and NdaB. During the condensation, NdaH participates in the conversion of N-methyl-L-threonine (MeThr) to N-methyldehydrobutyrine (MeDhb) with a cofactor nicotinamide adenine dinucleotide (NADH). Finally, the mature peptide chain is cyclized by NdaB and released from the enzyme-substrate complex. As an ABC-transporter, NdaI is responsible for the transmembrane transportation of nodularin for extracellular excretion.
4. PCR detection of toxic cyanobacteria
PCR-based assays have been most commonly utilized in molecular identification studies because the assays are able to recognize targets accurately. The assays incorporate oligonucleotide primers explicitly designed for complementary sequences of the target gene(s). Two types of PCR methods have been used: conventional gel-based PCR and real-time PCR. In general, the real-time PCR has higher sensitivity (i.e., detect a low amount of the target) than the conventional PCR. The real-time PCR also offers better specificity than the conventional PCR since it uses an additional oligonucleotide known as a probe, which is complementary to sequences between primer-binding sequences.
Furthermore, the real-time PCR allows estimating the number of the intended target in samples when performed with standards with a known copy number of the target sequences. This procedure is referred to as quantitative real-time PCR (qPCR). In addition, reverse transcription (RT)-PCR or RT-qPCR platforms have been utilized for specifically detecting transcripts (i.e., mRNAs) from the target genes of cyanobacteria. Typically, PCR can be completed within one or two hours, much shorter than the traditional analytical methods and microscopy mentioned above.
4.1 Microcystin-producing cyanobacteria
The molecular identification of microcystin-producing cyanobacteria has been conducted using nearly all
Although most publications have been concerned about toxic/toxigenic
The rest of the
With increased bioinformatic data related to
There are a few unidentified open reading frames (ORFs) flanking the
4.2 Anatoxin-a-producing cyanobacteria
The
Like
4.3 Cylindrospermopsin-producing cyanobacteria
Molecular detection of cylindrospermopsin-producing cyanobacteria has been mostly reported for
Multi-generic detection of cylindrospermopsin-producing cyanobacteria was reported as well. Campo et al. found that
There are also a few ORFs flanking the
4.4 Nodularin-producing cyanobacteria
Since
5. Other cyanotoxins and PCR detection of the toxic cyanobacteria
Apart from the four most commonly reported cyanotoxins mentioned above, there are a few other cyanotoxins, such as saxitoxin, lyngbyatoxin, guanitoxin, β-N-methylamino-L-alanin (BMAA), aplysiatoxin, and lipopolysaccharide [18, 77, 78]. Hitherto, only the gene clusters for the biosynthesis of saxitoxin and lyngbyatoxin have been characterized.
Saxitoxin belongs to the group of carbamate alkaloid toxins composed of a tetrahydropurine group and two guanidinium moieties [79] and can also be produced by marine phytoplankton [80]. It can cause paralytic shellfish poisoning syndrome and afflict human health via bioaccumulation. At least 30 clustered saxitoxin synthesis genes (
The
Lyngbyatoxin is characterized as a potent skin irritant produced by
No literature regarding molecular detection of cyanobacteria producing the rest of the toxins mentioned above could be searched. It is most likely because there are few reports as to the molecular mechanisms of their biosynthesis. Nevertheless, it is worthwhile to briefly introduce guanitoxin, previously known as anatoxin-a(S), to emphasize its difference from anatoxin-a. Guanitoxin was recently renamed due to its structural and toxicological disparities from anatoxin-a [77]. It is a guanidino organophosphate neurotoxin that irreversibly inhibits acetylcholinesterase’s active site, leading to excess acetylcholine, which causes severe salvation and chromodacryorrhea, so-called “bloody tears” before respiratory arrest [84]. Up to now, it was only found in planktonic
6. Perspectives
As various cyanobacterial genera can produce the same cyanotoxin, the development of toxigenic cyanobacteria identification needs to be multi-generic detection. Furthermore, as many genes for different toxins have sequences for the same conserved domains, designing PCR methods for all the cyanobacteria producing multiple toxins would be ideal.
Although most publications have focused on the selected cyanotoxins and their producers, more attention should be paid to other cyanotoxins and producers due to their potential of posing a significant threat to animal and human health. However, many cyanotoxin-producing cyanobacteria still lack bioinformation for the synthesis-related genes (e.g., guanitoxin), and it is thereby urgent to make further exploration to enrich the gene pools and their sequences so that a much more comprehensive understanding of the molecular mechanisms and the development of nucleic acid-based identification methods can be facilitated.
With the technical advance in PCR, researchers have been able to develop multiplex PCR methods in which many cyanotoxin biosynthesis genes can be detected simultaneously. For example, Ouahid et al. devised a multiplex PCR assay to detect six
Cyanobacteria with cyanotoxin synthetase genes in their genome are clearly equipped with the ability of toxin production. However, transcription of toxin biosynthesis genes is triggered by various environmental factors [88, 89, 90]; hence, toxin production is not consistently ongoing. It means the presence of genes itself may not always translate into the appearance of toxins unless they are inter- or extra-cellularly accumulated and detectable. Furthermore, the significant positive correlation between gene copies and toxin levels is still controversial, as described in this chapter and another review [91]. Instead, the presence of mRNA transcripts from cyanotoxin synthase genes may be more closely associated with toxin production. Consequently, cDNA detection is justifiable to indicate an ongoing toxin synthesis, which is more critical and useful for monitoring the toxin-producing cyanobacteria. For this purpose, genes located at the end of operons should be good candidates for two reasons. One, primers designed from those genes can be directly used in cDNA testing like other genes because cyanobacteria lack introns. Two, the appearance of those genes in cDNA form signifies the successful cascade transcription of the clustered genes, gearing up all pertinent proteins for toxin synthesis. For example,
Although qPCR is preferred due to its many advantages, conventional PCR should also be considered for assessing the presence or absence of toxigenic cyanobacteria in water samples, as previously reported [49, 56]. In addition, the simplicity and cheaper operation may make conventional PCRs a more cost-effective tool for molecular detection of toxigenic cyanobacteria in comparison to qPCRs.
Besides PCR-based assays, there are other molecular technologies applicable to the identification and/or characterization of toxigenic cyanobacteria. A noteworthy method is the next-generation sequencing (NGS) technology. The technology has been widely used to identify previously unrecognized agents, non-culturable microorganisms, and/or variants because of its advanced and hypothesis-free sequencing ability [92] and has been applied to cyanobacteria research. Although most NGS studies have been investigations of taxonomic diversities using representative cyanobacterial genetic markers such as 16S rDNA [93, 94], the potential toxigenicity of cyanobacteria can be disclosed by sequencing the pooled libraries of toxin biosynthesis associated genes. Casero et al. revealed the existence of multiple toxigenic taxa in a summer bloom in a Spanish reservoir using
7. Conclusions
Nowadays, freshwater cyanobacterial blooms are seen more frequently than ever before because of increased eutrophication of their habitats and climate changes (e.g., global warming), which are utterly favorable to the overgrowth of cyanobacteria. Even though toxic cyanobacterial species are not always the mere culprit for these ecological disasters, they are often the dominant organisms and cause more destructive consequences because they can produce potent cyanotoxins into the water. There is no doubt that the toxic freshwater cyanobacteria pose a grave threat to human and animal health, agricultural production, tourism, to name a few. Hence, advancing techniques and technologies for rapid and reliable identification and monitoring of toxic cyanobacteria is an inevitable mission for healthcare, economy, and environmental conservation. To date, molecular assays, especially PCR-based tests, have been employed in toxic cyanobacterial identification, but their utilization should be further expanded into large-scale and long-term detection tasks and routine monitoring programs for not only the acute poisoning incidents but also the chronic impacts and preventative measures.
Acknowledgments
The authors disclosed receipt of the following financial support for the publication of this chapter: the manuscript compilation was supported in part by funding from the Innovative Swine Industry Enhancement Grant Program by the Iowa Attorney General’s Office, Iowa State University (ISU) Health Research Initiative, and ISU Veterinary Diagnostic Laboratory Research Support Fund.
Thanks
The authors are sincerely grateful to the people who have made their contributions to the pertaining studies that are helpful for writing the chapter, including, but not limited to, Dr. Steve Ensley, Dr. Hyun-Joong Kim, Dr. Christopher Filstrup, Dr. Baoqing Guo, Dr. Paula Imerman, Dr. Grace Wilkinson, Dwayne Schrunk, and Amy Curtis.
References
- 1.
Domingos P, Rubim TK, Molica RJR, Azevedo SM, Carmichael WW. First report of microcystin production by picoplanktonic cyanobacteria isolated from a northeast Brazilian drinking water supply. Environ Toxicol. 1999;14:31-35. DOI: 10.1002/(SICI)1522-7278(199902)14:1<31::AID-TOX6>3.0.CO;2-B - 2.
Falconer IR. An overview of problems caused by toxic blue–green algae (cyanobacteria) in drinking and recreational water. Environ Toxicol. 1999;14:5-12. DOI: 10.1002/(SICI)1522-7278(199902)14:1<5::AID-TOX3>3.0.CO;2-0 - 3.
Falconer IR, Burch MD, Steffensen DA, Choice M, Coverdale OR. Toxicity of the blue-green alga (cyanobacterium) Microcystis aeruginosa in drinking water to growing pigs, as an animal model for human injury and risk assessment. Environ Toxic Water. 1994;9:131-139. DOI: 10.1002/tox.2530090209 - 4.
Zaccaroni A, Scaravelli D. Toxicity of fresh water algal toxins to humans and animals. In: Evangelista V, Barsanti L, Frassanito AM, Passarelli V, Gualtieri P, editors. Algal Toxins: Nature, Occurrence, Effect and Detection. Dordrecht: Springer; 2008. p. 45-89. DOI: 10.1007/978-1-4020-8480-5_3 - 5.
Carmichael WW, Boyer GL. Health impacts from cyanobacteria harmful algae blooms: Implications for the North American Great Lakes. Harmful Algae. 2016;54:194-212. DOI: 10.1016/j.hal.2016.02.002 - 6.
Paerl HW, Otten TG. Harmful cyanobacterial blooms: causes, consequences, and controls. Microb Ecol. 2013;65(4):995-1010. doi: 10.1007/s00248-012-0159-y - 7.
Miller MA, Kudela RM, Mekebri A, Crane D, Oates SC, Tinker MT, et al. Evidence for a novel marine harmful algal bloom: cyanotoxin (microcystin) transfer from land to sea otters. PLoS One. 2010;5:e12576. DOI: 10.1371/journal.pone.0012576. - 8.
Backer LC, Landsberg JH, Miller M, Keel K, Taylor TK. Canine cyanotoxin poisonings in the United States (1920s-2012): review of suspected and confirmed cases from three data sources. Toxins. 2013;5:1597-1628. DOI: 10.3390/toxins5091597 - 9.
Dittmann E, Wiegand C. Cyanobacterial toxins--occurrence, biosynthesis and impact on human affairs. Mol Nutr Food Res. 2006;50:7-17. DOI: 10.1002/mnfr.200500162. PubMed PMID: 16304634 - 10.
Stewart I, Seawright AA, Shaw GR. Cyanobacterial poisoning in livestock, wild mammals and birds – an overview. In: Hudnell HK, editor. Cyanobacterial Harmful Algal Blooms: State of the Science and Research Needs. New York: Springer; 2008. p. 613-637. DOI: 10.1007/978-0-387-75865-7_28 - 11.
Mathys W, Surholt B. Analysis of microcystins in freshwater samples using high performance liquid chromatography and an enzyme-linked immunosorbent assay. Int J Hyg Envir Heal. 2004;207:601-605. DOI: 10.1078/1438-4639-00334 - 12.
Brown A, Foss A, Miller MA, Gibson Q. Detection of cyanotoxins (microcystins/nodularins) in livers from estuarine and coastal bottlenose dolphins ( Tursiops truncatus ) from Northeast Florida. Harmful Algae. 2018;76:22-34. DOI: 10.1016/j.hal.2018.04.011 - 13.
Melvin A. Determination of microcystin-LR in municipal water using HPLC-UV/Vis. In: Proceedings of the National Conference on Undergraduate Research (2017 NCUR); 6-8 April 2017; Memphis - 14.
Shamsollahi HR, Alimohammadi M, Nabizadeh R, Nazmara S, Mahvi AH. Measurement of microcystin -LR in water samples using improved HPLC method. Global Journal of Health Science. 2014;7:66-70. DOI: 10.5539/gjhs.v7n2p66 - 15.
Robillot C, Vinh J, Puiseux-Dao S, Hennion M-C. Hepatotoxin production kinetics of the cyanobacterium Microcystis aeruginosa PCC 7820, as determined by HPLC−mass spectrometry and protein phosphatase bioassay. Environ Sci Technol. 2000;34:3372-3378. DOI: 10.1021/es991294v - 16.
Lu N, Sun J, Kong F, Zhang D. Statistical analysis of a HPLC method for microcystins determination in drinking water. In: Proceedings of the International Conference on Electric Technology and Civil Engineering (ICETCE); 22-24 April 2011; Lushan. New York: IEEE; 2011. p. 4469-4470 - 17.
Christiansen G, Fastner J, Erhard M, Borner T, Dittmann E. Microcystin biosynthesis in Planktothrix : genes, evolution, and manipulation. J Bacteriol. 2003;185:564-572. DOI: 10.1128/jb.185.2.564-572.2003 - 18.
Dittmann E, Fewer DP, Neilan BA. Cyanobacterial toxins: biosynthetic routes and evolutionary roots. FEMS Microbiol Rev. 2013;37:23-43. DOI: 10.1111/j.1574-6976.2012.12000.x - 19.
Rantala-Ylinen A, Kana S, Wang H, Rouhiainen L, Wahlsten M, Rizzi E, et al. Anatoxin-a synthetase gene cluster of the cyanobacterium Anabaena sp. strain 37 and molecular methods to detect potential producers. Appl Enviro Microb. 2011;77:7271-7278. DOI: 10.1128/AEM.06022-11 - 20.
Mazmouz R, Chapuis-Hugon F, Mann S, Pichon V, Mejean A, Ploux O. Biosynthesis of cylindrospermopsin and 7-epicylindrospermopsin in Oscillatoria sp. strain PCC 6506: identification of thecyr gene cluster and toxin analysis. Appl Enviro Microb. 2010;76:4943-4949. DOI: 10.1128/AEM.00717-10 - 21.
Mihali TK, Kellmann R, Neilan BA. Characterisation of the paralytic shellfish toxin biosynthesis gene clusters in Anabaena circinalis AWQC131C andAphanizomenon sp. NH-5. BMC Biochem. 2009;10:8. DOI: 10.1186/1471-2091-10-8 - 22.
Mihali TK, Kellmann R, Muenchhoff J, Barrow KD, Neilan BA. Characterization of the gene cluster responsible for cylindrospermopsin biosynthesis. Appl Enviro Microb. 2008;74:716-722. DOI: 10.1128/AEM.01988-07 - 23.
Moffitt MC, Neilan BA. Characterization of the nodularin synthetase gene cluster and proposed theory of the evolution of cyanobacterial hepatotoxins. Appl Enviro Microb. 2004;70:6353-6362. doi: 10.1128/AEM.70.11.6353-6362.2004 - 24.
Beard S, Davis P, Iglesias-Rodrıguez D, Skulberg O, Walsby A. Gas vesicle genes in Planktothrix spp. from Nordic lakes: strains with weak gas vesicles possess a longer variant ofgvpC . Microbiology. 2000;146:2009-2018. DOI: 10.1099/00221287-146-8-2009 - 25.
Yang J, Deng X, Xian Q, Qian X, Li A. Allelopathic effect of Microcystis aeruginosa onMicrocystis wesenbergii : microcystin-LR as a potential allelochemical. Hydrobiologia. 2014;727:65-73. DOI: 10.1007/s10750-013-1787-z - 26.
Tillett D, Dittmann E, Erhard M, Dohren HV, Borner T, Neilan BA. Structural organization of microcystin biosynthesis in Microcystis aeruginosa PCC7806: an integrated peptide–polyketide synthetase system. Chem Biol. 2000;7:753-764. DOI: 10.1016/S1074-5521(00)00021-1 - 27.
Brown NM, Mueller RS, Shepardson JW, Landry ZC, Morre JT, Maier CS, et al. Structural and functional analysis of the finished genome of the recently isolated toxic Anabaena sp. WA102. BMC Genomics. 2016;17:457. DOI: 10.1186/s12864-016-2738-7 - 28.
Wang H, Sivonen K, Rouhiainen L, Fewer DP, Lyra C, Rantala-Ylinen A, et al. Genome-derived insights into the biology of the hepatotoxic bloom-forming cyanobacterium Anabaena sp. strain 90. BMC Genomics. 2012;13:613. DOI: 10.1186/1471-2164-13-613 - 29.
Rounge TB, Rohrlack T, Nederbragt AJ, Kristensen T, Jakobsen KS. A genome-wide analysis of nonribosomal peptide synthetase gene clusters and their peptides in a Planktothrix rubescens strain. BMC Genomics. 2009;10:396. DOI: 10.1186/1471-2164-10-396 - 30.
Buratti FM, Manganelli M, Vichi S, Stefanelli M, Scardala S, Testai E, et al. Cyanotoxins: producing organisms, occurrence, toxicity, mechanism of action and human health toxicological risk evaluation. Arch Toxicol. 2017;91:1049-1130. DOI: 10.1007/s00204-016-1913-6 - 31.
Rouhiainen L, Vakkilainen T, Siemer BL, Buikema W, Haselkorn R, Sivonen K. Genes coding for hepatotoxic heptapeptides (microcystins) in the cyanobacterium Anabaena strain 90. Appl Enviro Microb. 2004;70:686-692. DOI: 10.1128/aem.70.2.686-692.2004 - 32.
Mejean A, Paci G, Gautier V, Ploux O. Biosynthesis of anatoxin-a and analogues (anatoxins) in cyanobacteria. Toxicon. 2014;91:15-22. DOI: 10.1016/j.toxicon.2014.07.016 - 33.
Rantala A, Fewer DP, Hisbergues M, Rouhiainen L, Vaitomaa J, Borner T, et al. Phylogenetic evidence for the early evolution of microcystin synthesis. Proc Natl Acad Sci. 2004;101:568-573. DOI: 10.1073/pnas.0304489101 - 34.
Kellmann R, Michali TK, Neilan BA. Identification of a saxitoxin biosynthesis gene with a history of frequent horizontal gene transfers. J Mol Evol. 2008;6:526-538. DOI: 10.1007/s00239-009-9210-0 - 35.
Beversdorf LJ, Chaston SD, Miller TR, McMahon KD. Microcystin mcyA andmcyE gene abundances are not appropriate indicators of microcystin concentrations in lakes. PLoS One. 2015;10:e0125353. DOI: 10.1371/journal.pone.0125353 - 36.
Pouria S, de Andrade A, Barbosa J, Cavalcanti RL, Barreto VTS, Ward CJ, et al. Fatal microcystin intoxication in haemodialysis unit in Caruaru, Brazil. Lancet. 1998;352:21-26. DOI: 10.1016/s0140-6736(97)12285-1 - 37.
Fitzgerald SD, Poppenga RH. Toxicosis due to microcystin hepatotoxins in three Holstein heifers. J Vet Diagn Invest. 1993;5:651-653. DOI: 10.1177/104063879300500433 - 38.
Classen DM, Schwartz KJ, Madson D, Ensley SM. Microcystin toxicosis in nursery pigs. J Swine Health Prod. 2017;25:198-205. - 39.
Sivonen K, Jones G. Cyanobacterial toxins. In: Chorus I, Bartram J, editors. Toxic Cyanobacteria in Water: A Guide to Their Public Health Consequences, Monitoring and Management. Boca Raton; CRC Press; 1999. p. 43-112. DOI: 10.1201/9781482295061 - 40.
Kaneko T, Nakajima N, Okamoto S, Suzuki I, Tanabe Y, Tamaoki M, et al. Complete genomic structure of the bloom-forming toxic cyanobacterium Microcystis aeruginosa NIES-843. DNA Res. 2007;14:247-256. DOI: 10.1093/dnares/dsm026 - 41.
Méjean A, Dalle K, Paci G, Bouchonnet S, Mann S, Pichon V, et al. Dihydroanatoxin-a is biosynthesized from proline in Cylindrospermum stagnale PCC 7417: isotopic incorporation experiments and mass spectrometry analysis. J Nat Prod. 2016;79:1775-1782. DOI: 10.1021/acs.jnatprod.6b00189 - 42.
Devlin J, Edwards O, Gorham P, Hunter N, Pike R, Stavric B. Anatoxin-a, a toxic alkaloid from Anabaena flos-aquae NRC-44h. Can J Chem. 1977;55:1367-1371. DOI: 10.1139/v77-189 - 43.
Méjean A, Mann S, Maldiney T, Vassiliadis G, Lequin O, Ploux O. Evidence that biosynthesis of the neurotoxic alkaloids anatoxin-a and homoanatoxin-a in the cyanobacterium Oscillatoria PCC 6506 occurs on a modular polyketide synthase initiated by L-proline. J Am Chem Soc. 2009;131:7512-7513. DOI: 10.1021/ja9024353 - 44.
Runnegar MT, Kong S-M, Zhong Y-Z, Lu SC. Inhibition of reduced glutathione synthesis by cyanobacterial alkaloid cylindrospermopsin in cultured rat hepatocytes. Biochem Pharmacol. 1995;49:219-225. DOI: 10.1016/S0006-2952(94)00466-8 - 45.
Runnegar MT, Xie C, Snider BB, Wallace GA, Weinreb SM, Kuhlenkamp J. In vitro hepatotoxicity of the cyanobacterial alkaloid cylindrospermopsin and related synthetic analogues. Toxicol Sci. 2002;67:81-87. DOI: 10.1093/toxsci/67.1.81 - 46.
Humpage AR, Fenech M, Thomas P, Falconer IR. Micronucleus induction and chromosome loss in transformed human white cells indicate clastogenic and aneugenic action of the cyanobacterial toxin, cylindrospermopsin. Mutat Res-Gen Tox En 2000;472:155-161. DOI: 10.1016/S1383-5718(00)00144-3 - 47.
Kiss T, Vehovszky A, Hiripi L, Kovacs A, Vörös L. Membrane effects of toxins isolated from a cyanobacterium, Cylindrospermopsis raciborskii , on identified molluscan neurones. Comp Biochem Phys C. 2002;131:167-176. DOI: 10.1016/S1532-0456(01)00290-3 - 48.
Stuken A, Jakobsen KS. The cylindrospermopsin gene cluster of Aphanizomenon sp. strain 10E6: organization and recombination. Microbiology. 2010;156:2438-2351. DOI: 10.1099/mic.0.036988-0 - 49.
Tillett D, Parker DL, Neilan BA. Detection of toxigenicity by a probe for the microcystin synthetase A gene ( mcyA ) of the cyanobacterial genusMicrocystis : comparison of toxicities with 16S rRNA and phycocyanin operon (Phycocyanin Intergenic Spacer) phylogenies. Appl Environ Microb. 2001;67:2810-2818. DOI: 10.1128/AEM.67.6.2810-2818.2001 - 50.
Kurmayer R, Christiansen G, Chorus I. The abundance of microcystin-producing genotypes correlates positively with colony size in Microcystis sp. and determines its microcystin net production in Lake Wannsee. Appl Environ Microb. 2003;69:787-95. DOI: 10.1128/aem.69.2.787-795.2003 - 51.
Kaebernick M, Neilan BA, Börner T, Dittmann E. Light and the transcriptional response of the microcystin biosynthesis gene cluster. Appl Environ Microb. 2000;66:3387-3392. DOI: 10.1128/AEM.66.8.3387-3392.2000 - 52.
Briand E, Gugger M, Francois JC, Bernard C, Humbert JF, Quiblier C. Temporal variations in the dynamics of potentially microcystin-producing strains in a bloom-forming Planktothrix agardhii (Cyanobacterium) population. Appl Environ Microb. 2008;74:3839-3848. DOI: 10.1128/AEM.02343-07 - 53.
Mbedi S, Welker M, Fastner J, Wiedner C. Variability of the microcystin synthetase gene cluster in the genus Planktothrix (Oscillatoriales, Cyanobacteria). FEMS Microbiol Lett. 2005;245:299-306. DOI: 10.1016/j.femsle.2005.03.020 - 54.
Vaitomaa J, Rantala A, Halinen K, Rouhiainen L, Tallberg P, Mokelke L, et al. Quantitative real-time PCR for determination of microcystin synthetase E copy numbers for Microcystis andAnabaena in lakes. Appl Environ Microb. 2003;69:7289-7297. DOI: 10.1128/aem.69.12.7289-7297.2003 - 55.
Ngwa FF, Madramootoo CA, Jabaji S. Comparison of cyanobacterial microcystin synthetase ( mcy ) E gene transcript levels,mcyE gene copies, and biomass as indicators of microcystin risk under laboratory and field conditions. MicrobiologyOpen. 2014;3(4):411-25. doi: 10.1002/mbo3.173 - 56.
Yuan J, Kim H-J, Filstrup CT, Guo B, Imerman P, Ensley S, et al. Utility of a PCR-based method for rapid and specific detection of toxigenic Microcystis spp. in farm ponds. J Vet Diagn Invest. 2020;32:369-381. DOI: 10.1177/1040638720916156 - 57.
Ouahid Y, Perez-Silva G, del Campo FF. Identification of potentially toxic environmental Microcystis by individual and multiple PCR amplification of specific microcystin synthetase gene regions. Environ Toxicol. 2005;20:235-242. DOI: 10.1002/tox.20103 - 58.
Zhang W, Lou I, Ung WK, Kong Y, Mok KM. Analysis of cylindrospermopsin- and microcystin-producing genotypes and cyanotoxin concentrations in the Macau storage reservoir. Hydrobiologia. 2014;741:51-68. DOI: 10.1007/s10750-013-1776-2 - 59.
Kim S-G, Joung S-H, Ahn C-Y, Ko S-R, Boo SM, Oh H-M. Annual variation of Microcystis genotypes and their potential toxicity in water and sediment from a eutrophic reservoir. FEMS Microbiol Ecol. 2010;74:93-102. DOI: 10.1111/j.1574-6941.2010.00947.x - 60.
Joung S-H, Oh H-M, Ko S-R, Ahn C-Y. Correlations between environmental factors and toxic and non-toxic Microcystis dynamics during bloom in Daechung Reservoir, Korea. Harmful Algae. 2011;1:188-193. DOI: 10.1016/j.hal.2010.09.005 - 61.
Hisbergues M, Christiansen G, Rouhiainen L, Sivonen K, Börner T. PCR-based identification of microcystin-producing genotypes of different cyanobacterial genera. Arch Microbiol. 2003;180:402-410. DOI: 10.1007/s00203-003-0605-9 - 62.
Hautala H, Lamminmaki U, Spoof L, Nybom S, Meriluoto J, Vehniainen M. Quantitative PCR detection and improved sample preparation of microcystin-producing Anabaena ,Microcystis andPlanktothrix . Ecotox Environ Safe. 2012;87:49-56. DOI: 10.1016/j.ecoenv.2012.10.008 - 63.
Ye W, Liu X, Tan J, Li D, Yang H. Diversity and dynamics of microcystin—producing cyanobacteria in China's third largest lake, Lake Taihu. Harmful Algae. 2009;8:637-644. DOI: 10.1016/j.hal.2008.10.010 - 64.
Legrand B, Lesobre J, Colombet J, Latour D, Sabart M. Molecular tools to detect anatoxin-a genes in aquatic ecosystems: Toward a new nested PCR-based method. Harmful Algae. 2016;58:16-22. - 65.
Ballot A, Fastner J, Lentz M, Wiedner C. First report of anatoxin-a-producing cyanobacterium Aphanizomenon issatschenkoi in northeastern Germany. Toxicon. 2010;56:964-971. DOI: 10.1016/j.toxicon.2010.06.021 - 66.
Mann S, Cohen M, Chapuis-Hugon F, Pichon V, Mazmouz R, Mejean A, et al. Synthesis, configuration assignment, and simultaneous quantification by liquid chromatography coupled to tandem mass spectrometry, of dihydroanatoxin-a and dihydrohomoanatoxin-a together with the parent toxins, in axenic cyanobacterial strains and in environmental samples. Toxicon. 2012;60:1404-1414. DOI: 10.1016/j.toxicon.2012.10.006 - 67.
Wood SA, Smith FM, Heath MW, Palfroy T, Gaw S, Young RG, et al. Within-mat variability in anatoxin-a and homoanatoxin-a production among benthic Phormidium (cyanobacteria) strains. Toxins. 2012;4:900-912. DOI: 10.3390/toxins4100900 - 68.
Burford MA, Davis TW, Orr PT, Sinha R, Willis A, Neilan BA. Nutrient-related changes in the toxicity of field blooms of the cyanobacterium, Cylindrospermopsis raciborskii . FEMS Microbiol Ecol. 2014;89:135-148. DOI: 10.1111/1574-6941.12341 - 69.
Fergusson KM, Saint CP. Multiplex PCR assay for Cylindrospermopsis raciborskii and cylindrospermopsin-producing cyanobacteria. Environ Toxicol. 2003;18:120-125. DOI: 10.1002/tox.10108 - 70.
Moreira C, Martins A, Azevedo J, Freitas M, Regueiras A, Vale M, et al. Application of real-time PCR in the assessment of the toxic cyanobacterium Cylindrospermopsis raciborskii abundance and toxicological potential. Appl Microbiol Biot. 2011;92:189-197. DOI: 10.1007/s00253-011-3360-x - 71.
Wilson KM, Schembri MA, Baker PD, Saint CP. Molecular characterization of the toxic cyanobacterium Cylindrospermopsis raciborskii and design of a species-specific PCR. Appl Environ Microb. 2000;66:332-338. DOI: 10.1128/AEM.66.1.332-338.2000 - 72.
Marbun YR, Yen HK, Lin T-F, Lin HL, Michinaka A. Rapid on-site monitoring of cylindrospermopsin-producers in reservoirs using quantitative PCR. Sustain Environ Res. 2012;22:143-151. - 73.
Campo E, Agha R, Cirés S, Quesada A, El-Shehawy R. First TaqMan assay to identify and quantify the cylindrospermopsin-producing cyanobacterium Aphanizomenon ovalisporum in water. Adv Microbiol. 2013;3:430-437. DOI: 10.4236/aim.2013.35058 - 74.
Krüger T, Oelmüller R, Luckas B. Comparative PCR analysis of toxic Nodularia spumigena and non-toxicNodularia harveyana (Nostocales, Cyanobacteria) with respect to the nodularin synthetase gene cluster. Eur J Phycol. 2009;44:291-295. DOI: 10.1080/09670260802588434 - 75.
Koskenniemi K, Lyra C, Rajaniemi-Wacklin P, Jokela J, Sivonen K. Quantitative real-time PCR detection of toxic Nodularia cyanobacteria in the Baltic Sea. Appl Environ Microb. 2007;73:2173-2179. DOI: 10.1128/AEM.02746-06 - 76.
Jonasson S, Vintila S, Sivonen K, El-Shehawy R. Expression of the nodularin synthetase genes in the Baltic Sea bloom-former cyanobacterium Nodularia spumigena strain AV1. FEMS Microbiol Ecol. 2008;65:31-39. DOI: 10.1111/j.1574-6941.2008.00499.x - 77.
Fiore MF, de Lima ST, Carmichael WW, McKinnie SM, Chekan JR, Moore BS. Guanitoxin, re-naming a cyanobacterial organophosphate toxin. Harmful Algae. 2020;92:101737. DOI: 10.1016/j.hal.2019.101737 - 78.
Stewart I, Schluter PJ, Shaw GR. Cyanobacterial lipopolysaccharides and human health - a review. Environ Health. 2006;5:1-23. DOI: 10.1186/1476-069X-5-7 - 79.
Schantz EJ, Ghazarossian V, Schnoes HK, Strong F, Springer J, Pezzanite JO, et al. Structure of saxitoxin. J Am Chem Soc. 1975;97:1238-1239. DOI: 10.1021/ja00838a045 - 80.
Gu H. Morphology, phylogenetic position, and ecophysiology of Alexandrium ostenfeldii (Dinophyceae) from the Bohai Sea, China. J Syst Evol. 2011;49:606-616. DOI: 10.1111/j.1759-6831.2011.00160.x - 81.
Kellmann R, Mihali TK, Jeon YJ, Pickford R, Pomati F, Neilan BA. Biosynthetic intermediate analysis and functional homology reveal a saxitoxin gene cluster in cyanobacteria. Appl Environ Microb. 2008;74:4044-4053. DOI: 10.1128/AEM.00353-08 - 82.
Al-Tebrineh J, Mihali TK, Pomati F, Neilan BA. Detection of saxitoxin-producing cyanobacteria and Anabaena circinalis in environmental water blooms by quantitative PCR. Appl Environ Microb. 2010;76:7836-7842. DOI: 10.1128/AEM.00174-10 - 83.
Al-Tebrineh J, Merrick C, Ryan D, Humpage A, Bowling L, Neilan BA. Community composition, toxigenicity, and environmental conditions during a cyanobacterial bloom occurring along 1,100 kilometers of the Murray River. Appl Environ Microb. 2012;78:263-272. DOI: 10.1128/AEM.05587-11 - 84.
Carmichael WW, Gorham PR. Anatoxins from clones of Anabaena flos-aquae isolated from lakes of western Canada. Internationale Vereinigung für Theoretische und Angewandte Limnologie: Mitteilungen. 1978;21:285-295. DOI: 10.1080/05384680.1978.11903972 - 85.
Ouahid Y, Del Campo FF. Typing of toxinogenic Microcystis from environmental samples by multiplex PCR. Appl Microbiol Biot. 2009;85:405. DOI: 10.1007/s00253-009-2249-4 - 86.
Rasmussen JP, Giglio S, Monis P, Campbell R, Saint C. Development and field testing of a real-time PCR assay for cylindrospermopsin-producing cyanobacteria. J Appl Microbiol. 2008;104:1503-1515. DOI: 10.1111/j.1365-2672.2007.03676.x - 87.
Al-Tebrineh J, Pearson LA, Yasar SA, Neilan BA. A multiplex qPCR targeting hepato-and neurotoxigenic cyanobacteria of global significance. Harmful Algae. 2012;15:19-25. DOI: 10.1016/j.hal.2011.11.001 - 88.
Wood SA, Rueckert A, Hamilton DP, Cary SC, Dietrich DR. Switching toxin production on and off: intermittent microcystin synthesis in a Microcystis bloom. Env Microbiology Rep. 2011;3:118-124. DOI: 10.1111/j.1758-2229.2010.00196.x. - 89.
Pimentel JSM, Giani A. Microcystin production and regulation under nutrient stress conditions in toxic Microcystis strains. Appl Environ Microb. 2014;80:5836-5843. DOI: 10.1128/aem.01009-14. - 90.
Gobler CJ, Davis TW, Coyne KJ, Boyer GL. Interactive influences of nutrient loading, zooplankton grazing, and microcystin synthetase gene expression on cyanobacterial bloom dynamics in a eutrophic New York lake. Harmful Algae. 2007;6:119-133. DOI: 10.1016/j.hal.2006.08.003. - 91.
Pacheco AB, Guedes IA, Azevedo SM. Is qPCR a reliable indicator of cyanotoxin risk in freshwater? Toxins. 2016;8:172. DOI: 10.3390/toxins8060172. - 92.
Cao Y, Fanning S, Proos S, Jordan K, Srikumar S. A review on the application of next generation sequencing technologies as applied to food-related microbiome studies. Frontiers in Microbiology. 2017;8:1829. DOI: 10.3389/fmicb.2017.01829. - 93.
Celikkol-Aydin S, Gaylarde CC, Lee T, Melchers RE, Witt DL, Beech IB. 16S Rrna gene profiling of planktonic and biofilm microbial populations in the Gulf of Guinea using Illumina NGS. Marine Environmental Research. 2016;122:105-112. DOI: 10.1016/j.marenvres.2016.10.001. - 94.
Casero MC, Velázquez D, Medino-Cobo M, Quesada A, Cirés S. Unmasking the identity of toxigenic cyanobacteria driving a multi-toxin bloom by high-throughput sequencing of cyanotoxins genes and 16S rRNA metabarcoding. Science of the Total Environment. 2019;665:367-378. DOI: 10.1016/j.scitotenv.2019.02.083. - 95.
Rantala A, Rizzi E, Castiglioni B, de Bellis G, Sivonen K. Identification of hepatotoxin-producing cyanobacteria by DNA-chip. Environmental Microbiology. 2008;10:653-664. DOI: 10.1111/j.1462-2920.2007.01488.x.