Next Article in Journal
Functional Analysis of FgNahG Clarifies the Contribution of Salicylic Acid to Wheat (Triticum aestivum) Resistance against Fusarium Head Blight
Next Article in Special Issue
The Beneficial and Debilitating Effects of Environmental and Microbial Toxins, Drugs, Organic Solvents and Heavy Metals on the Onset and Progression of Multiple Sclerosis
Previous Article in Journal
Prey Lysate Enhances Growth and Toxin Production in an Isolate of Dinophysis acuminata
Previous Article in Special Issue
Why Are Botulinum Neurotoxin-Producing Bacteria So Diverse and Botulinum Neurotoxins So Toxic?
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:

The Incidence of Marine Toxins and the Associated Seafood Poisoning Episodes in the African Countries of the Indian Ocean and the Red Sea

Isidro José Tamele
Marisa Silva
1,4 and
Vitor Vasconcelos
CIIMAR/CIMAR—Interdisciplinary Center of Marine and Environmental Research, University of Porto, Terminal de Cruzeiros do Porto, Avenida General Norton de Matos, 4450-238 Matosinhos, Portugal
Institute of Biomedical Science Abel Salazar, University of Porto, R. Jorge de Viterbo Ferreira 228, 4050-313 Porto, Portugal
Department of Chemistry, Faculty of Sciences, Eduardo Mondlane University, Av. Julius Nyerere, n 3453, Campus Principal, Maputo 257, Mozambique
Department of Biology, Faculty of Sciences, University of Porto, Rua do Campo Alegre, 4619-007 Porto, Portugal
Author to whom correspondence should be addressed.
Toxins 2019, 11(1), 58;
Submission received: 27 November 2018 / Revised: 10 January 2019 / Accepted: 10 January 2019 / Published: 21 January 2019
(This article belongs to the Special Issue Toxins:10th Anniversary)


The occurrence of Harmful Algal Blooms (HABs) and bacteria can be one of the great threats to public health due to their ability to produce marine toxins (MTs). The most reported MTs include paralytic shellfish toxins (PSTs), amnesic shellfish toxins (ASTs), diarrheic shellfish toxins (DSTs), cyclic imines (CIs), ciguatoxins (CTXs), azaspiracids (AZTs), palytoxin (PlTXs), tetrodotoxins (TTXs) and their analogs, some of them leading to fatal outcomes. MTs have been reported in several marine organisms causing human poisoning incidents since these organisms constitute the food basis of coastal human populations. In African countries of the Indian Ocean and the Red Sea, to date, only South Africa has a specific monitoring program for MTs and some other countries count only with respect to centers of seafood poisoning control. Therefore, the aim of this review is to evaluate the occurrence of MTs and associated poisoning episodes as a contribution to public health and monitoring programs as an MT risk assessment tool for this geographic region.
Key Contribution: The scarcity of MT data along African countries of the Indian Ocean and the Red Sea suggests the need for further studies and the creation of specific monitoring programs of MTs, particularly for dinoflagellates and diatoms since these constitute the phytoplankton that produces fatal MTs.

1. Introduction

The occurrence of Harmful Algal Blooms (HABs) in marine ecosystems can be one of the great threats to public health due to their capacity to produce marine toxins (MTs) as secondary metabolites [1,2,3,4,5,6,7,8,9,10,11,12,13,14]. MTs can be accumulated by distinct marine organisms such as fish, mollusks and crustaceans [15,16,17,18,19,20,21,22,23,24] which are the basic diet of coastal human populations. Suspected or confirmed episodes of human poisoning caused by MTs have been reported worldwide in the last century [20,21,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48]. The occurrence of episodes of human poisoning occurs via ingestion of contaminated marine food due to the lack of monitoring programs in some countries or violations of national health authorities’ regulations imposing the closure of harvesting areas and seafoodcommercialization [18,20,26,35,39,45,47,49]. Despite the ideal environmental conditions for theformation of blooms in this geographical area, there are insufficient data related to their occurrence and toxin production [50]. This review analyses the occurrence of MTs and their producers along the African Indian and the Red Sea coasts (from Egypt to South Africa) and associated human poisoning episodes. The existence of monitoring programs of MTs will be also highlighted and finally, some suggestions for the control and prevention of marine toxins in this area will be presented.

2. Marine Toxins and Their Producers

Chemically, toxins can be grouped according to their polarity, lipophilic and hydrophilic. Concerning MT monitoring, analysis and quantification methods in seafood are described in Table 1, including bioassays, immunoassays, and analytical chemistry methods. The bioassay methods (Mouse Bioassay (MBA), Rat Bioassay (RBA)) are no longer in use due to ethical reasons according to Directive 86/609/EEC [51] and procedural variation [52] (e.g., use of different extraction solvents and consequently shortcomings). Chemical methods, mainly liquid chromatography coupled to mass spectrometry, are considered as the most promising since they are fully validated and standardized to replace bioassays in many organizations worldwide. Further information related to each toxin group such as syndromes, producers, common vectors, symptoms, detections methods in seafood, limit of detection (LOD) and quantification (LOQ) and permitted limit used in some parts of the world is also described in Table 1.

2.1. Lipophilic Toxins

Lipophilic toxins are lipid soluble toxins and this group comprises okadaic acid (OA), ciguatoxins (CTX), cyclic imines (CIs) [spirolides (SPXs), gymnodimines (GYMs), pinnatoxins (PnTXs) and pteriatoxins (PtTXs)], brevetoxins (PbTxs), pectenotoxins (PTXs), yessotoxins (YTXs) and azaspiracids [AZAs], Table 1.

2.1.1. Okadaic Acid and Analogs

Okadaic acid (OA)and their analogs, dinophysistoxins-1, -2 and -3 (DTXs) (Figure 1), are polyethers produced by dinoflagellates: Prorocentrum spp. [8], Dinophysis spp. [2,6,9,10,15,53,54] and Phalacroma rotundatum [55] (Table 1).These polyethers are frost-resistant and heat-stable and consequently, their toxicity is not affected by the cooking procedures in water (they are stable at <150 °C) [56]. The OA group is responsible for the diarrheic shellfish poisoning syndrome (DSP), with OA being the main representative of DSP toxins. Okadaic acid (OA)and its analogs act as inhibitors of the serine/threonine phosphoprotein phosphatases 1,22B,4,5 types [57,58].

2.1.2. Ciguatoxins

Ciguatoxins (CTXs) (Figure 2A) are a group of toxins produced by tropical and subtropical dinoflagellates species: Gambierdiscus toxicus and Fukuyoa spp. [59,60] (Table 1) mainly found in the Pacific, Caribbean and the Indian Ocean regions [P-CTX, C-CTX and I-CTX, respectively]. CTXs are lipid-soluble polyethers with 13-14 rings fused by ether linkages into a rigid ladder-like structure [60]. To date, the structures of20 P-CTXs, 10 C-CTXSand 4 I-CTXs analogs have been fully identified and the most reported include P-CTX-1, P-CTX-2, P-CTX-3, P-CTX-3C [61,62,63,64,65,66,67], gambiertoxin [GbTXs, namely, P-CTX-4A and P-CTX-4B] [68], C-CTX-1, C-CTX-2 [67,69], I-CTX-1, I-CTX-2, I-CTX-3 and I-CTX-4 [70,71] mostly in predatory fish and gastropods [20,21,23,66,69,72,73,74]. The major analog of each group of CTXsis P-CTX-1. C-CTX-1, C-CTX-2, I-CTX1, and I-CTX-2. The chemical structure of the last two (I-CTXs) have the same molecular weight and similar structures as C-CTX-1 [62,67,70,71]. CTXs are odorless and tasteless heat-stable molecules and are not affected when subjected to water cooking, freezing and acid or basic conditions, though they suffer structural alterations by oxidation [60]. CTXs and Maitotoxin (MTX) (Figure 2B) (produced by Gambierdiscus spp. [68]) were the first group of toxins reported to be responsible for ciguatera shellfish poisoning (CFP) [23]. The mechanism of action of CTX and analogs is to elevate calcium ion concentration and activate non-selective cation channels in cells causing neurologic effects in humans [75].

2.1.3. Cyclic Imines

Cyclic imines (CI) (Figure 3) are toxins produced by dinoflagellates: SPXs: Alexandrium spp. [1,76], GYMs: Gymnodium spp. [77], PnTXs: Vulcanodinium rugosum [78] and PtTXs: biotransformation from PnTXs via metabolic and hydrolytic transformation in shellfish [1,5,77,78,79] (Table 1). CIs are a heterogenous group composed ofspirolides (SPXs), gymnodimines (GYMs), pinnatoxins (PnTXs) and pteriatoxins (PtTXs) and more than 24 structural analogs have been described to date [80].
Regarding chemical properties, these toxins are a group of macrocyclic compounds that have in common an imine functional group and spiro-linked ether moieties in their structure [80]. They are colorless amorphous solid macrocyclic compounds with imine and spiro-linked ether moieties [80], considerably soluble in organic solvents such as methanol, acetone, chloroform and ethyl acetate [5,80]. CIs are neurotoxins and actby inhibiting the nicotinic and muscarinic acetylcholine receptors (mAChR and nAChR, respectively) in the nervous system and at the neuromuscular junction [81]. CI bioactivity seems to depend on the imine functional group since the hydrolysis of spirolides A–D produce spirolide E and F with a keto-amine structure that is fully inactive [81]. To date, there are no regulations for CIs and no common symptoms can be recognized [82].

2.1.4. Brevetoxins

Brevetoxins (PbTxs) (Figure 4) are cyclic polyethers produced by dinoflagellates: Karenia spp. [4,16,87] (Table 1). There are two known types of BTXs, named type A and type B (also called type 1(PbTx-1) and type 2 (PbTx-2), respectively). The difference between two types of PbTxs consists in a few transfused rings that are ten for PbTx-1 and eleven for PbTx-2. The main analogs include PbTx-3, PbTx-6, PbTx-9, PbTx-B1, PbTx-B2, S-desoxy-PbTx-B2, PbTx-B3, PbTx-B4, and PbTx-B5 [44,88,89,90,91,92,93,94]. PbTxs are lipid-soluble cyclic polyether consisting of 10 to 11 transfused rings [95], stable and resistant to heat and steam autoclaving [96]. PbTxs cause neurotoxic shellfish poisoning (NSP) and actby binding with high affinity to receptor site 5 of the voltage-gated sodium channels (NaV) in cell membranes, and lactone is important for the toxin activity [97]. PbTxs are regulated in USA [98], New Zealand, and Australia [99,100] (Table 1).

2.1.5. Pectenotoxin Group

Pectenotoxins (PTXs) (Figure 5) are lipophilic polyethers produced by several dinoflagellate species [101] (Table 1). They contain spiroketal, bicyclic ketal, cyclic hemiketals, and oxolanes in their structure. To date, more than 15 PTX analogs have been documented and many are derived through biotransformation of PTX2 in marine organism metabolism such as bivalve mollusks [102]. The most reported analogs include PTX1, epi-PTX1, PTX2, PTX2 seco acid (PTX2 SA), 7-epi-PTX2 seco acid (7-epi-PTX2 SA), PTX3, PTX4, PTX6, epi-PTX6, PTX7, PTX11 (34S-hydroxy-PTX2) [6,101,103,104,105]. PTXs are heat-stable and unstable under alkaline conditions [103]. PTX and analogs alter actin-based structures [103,106] causing cell death and apoptosis [107]. PTXs co-occur with the OA—group and contribute to DSP in humans [108].

2.1.6. Yessotoxins

Yessotoxins (YTXs) (Figure 6) are produced by dinoflagellates species: Protoceratium reticulatum [4,109], Lingulodinium polyhedral [4] and Gonyaulax polyhedra [4] (Table 1). They are a heat-stable polyether, with eleven transfused ether rings, an unsaturated side chain, and two sulfate esters [110]. To date, more than 90 YTX analogues have been isolated [102] and only YTX, 45-hydroxyYTX, carboxylic, 1a-homoYTX, 45,46,47-trinorYTX, ketoYTX, 40-epi-ketoYTX, 41a-homoYTX, 9Me-41a-homoYTX, 44,55-dihydroxyYTX, 45-hydroxy-1a-homoYTX, carboxy-1a-homoYTX [111] have been fully identified [111]. The mechanism of action of YTX and their analogs is not fully understood; however, they are involved in phosphodiesterase activation [112] and modulation of calcium migration at several levels [113], alteration of protein disposal [114], cell change shape [115], apoptosis and cell death [116]. To date, there are no reports of human illness associated with YTXs [111].

2.1.7. Azaspiracids

Azaspiracids (AZAs) (Figure 7) are toxins produced by dinoflagellates: Azadinium spinosum [117] and Protoperidinum crassipes [118] (Table 1). They are colorless, odorless and amorphous solids of toxins containing a heterocyclic amine, a unique tri-spiro-assembly and an aliphatic carboxylic acid in their structures [117,119,120,121,122,123,124]. Around 21 compounds of AZAs are well known and documented [117,119,120,121,122,123,124] of which AZA, AZA2, AZA3, AZA4, and AZA5 are the most prevalent ones based on occurrence and toxicity in humans. AZAs are responsible for the AZP syndrome (Table 1) and their mechanism of action is the inhibition of hERG voltage-gated potassium channels [125].

2.2. Hydrophilic Toxins

Hydrophilic Toxins are polar soluble compounds and they include domoic acid (DA) and analogs, Paralytic Shellfish Toxins (PSTs), tetrodotoxins (TTXs) and palytoxins (PlTXs).

2.2.1. Domoic Acid and Analogs

Domoic acid (DA) (Figure 8) and analogs are polar cyclic amino acid toxins of diatom origin Pseudo-nitzschia spp. [126] and red algae: Chondria armata [127] (Table 1). They present three carboxylic acid groups and the most reported DA analogs include epi-domoic acid (epi-DA), domoic acid C5′-diastereomer and isodomoic acids A, B, C, D, E, F, G and H [iso-DA A-H] [128,129]. DA is the representative molecule of the DA-group that is responsible for amnesic shellfish poisoning (ASP) syndrome [130]. The characteristic symptomology of ASP is detailed in Table 1.

2.2.2. Paralytic Shellfish Toxins.

Paralytic shellfish toxins (PSTs) (Figure 9) are water-soluble tetrahydropurine toxins produced mainly by dinoflagellates Alexandrium spp. [2,3,7], Gymnodinium catenatum [3], Pyrodinium bahamense [3] and by cyanobacteria Trichodesmium erythraeum [131] except M (Figure 9) toxins that are Mytilus spp. metabolism products [132]. This group is composed of several analogs and they are prone to various conversions depending on pH (Figure 9), being divided into several groups: carbamoyl (saxitoxin (STX), neosaxitoxin (NeoSTX) and gonyautoxins (GTX1-4)) decarbamoyl [dc-](dcSTX, dcNeoSTX, dcGTX1-4), Nsulfo-carbamoyl [GTX5-6, C1-4], hydroxylated saxitoxins [M1-4] [133,134,135] and benzoyl toxins (GC1-3) [135]. Their heat stability is pH dependent (except for Nsulfo-carbamoyl components) [136]. STX and analogs act by binding to Nav and consequently blocking ion conductance in nerves and muscles fibers leading to paralysis [137]. Symptoms resulting from PSTs poisoning are described in Table 1.

2.2.3. Tetrodotoxins

Tetrodotoxins (TTXs) (Figure 10) are toxins produced by bacteria in marine environments: Serratia marcescens, Vibrio spp. [83], Aeromonas sp. [138], Microbacterium arabinogalactanolyticum [139], Pseudomonas sp. [140], Shewanella putrefaciens [141], Alteromonas sp. [142], Pseudoalteromonas ssp. [143], and Nocardiopsis dassonvillei [144] (Table 1). They are colorless, crystalline-weak basic compounds with one positively charged guanidinium group and a pyrimidine ring [145,146]. TTXpoisoning has been recognized since ancient Egyptian times [42]. To date, TTX is considered an extremely potent emergent toxin in the Atlantic Ocean [83] and acts by binding to Nav on the surface of nerve cell membranes blocking the cellular communication and causing death by cardio-respiratory paralysis [147]. Several poisoning incidents have reported in Asia [Japan is the most affected country] [148], the Mediterranean Sea and the Indian Ocean [35]. TTX is usually concentrated in the ovaries, liver, intestines, and skin ofits principal vector [puffer fish] [42]. To date, the structures of 26 analogs of TTX have been fully elucidated but their relative toxicity and occurrence are not yet fully known [145,146] except for 12compounds, namely, TTX, 11-oxoTTX, 11-deoxyTTX, 11-norTTX-6[R]-ol, 11-norTTX-6[S]-ol, 4-epiTTX, 4,9-anhydroTTX, 5,6,11-trideoxyTTX. [131], 4-CysTTX, 5-deoxyTTX, 5,11-dideoxyTTX, and 6,11-dideoxyTTX [149,150,151,152].

2.2.4. Palytoxin

Palytoxin (PlTX) and its derivatives (Figure 11) are toxins produced by marine zoanthids Palythoa spp., dinoflagellates: Ostreopsis ovata. [153,154,155] and possibly by cyanobacteria: Trichodesmium sp. [156] (Table 1). These polyhydroxylated toxins have both lipophilic and hydrophilic properties [157] with a partial unsaturated aliphatic backbone containing cyclic ethers, 64 chiral centers, 40–42 hydroxyl and 2 amide groups [157]. Among PlTX analogs, known are: isobaric PlTX, ostreocin-D, ovatoxin [a to f], mascarenotoxins, ostreotoxin-1 and 2, homopalytoxin, bishomopalytoxin, neopalytoxin, deopalytoxin and 42-hydroxypalytoxin and their molecular weights range from 2659 to 2680 DA [158,159,160]. PlTX and analogs act on Na+, K+ -ATPase pumps molecules in the cell membrane [161] and the loss of intracellular contents into the blood plasma and consequent injury causing rhabdomyolysis, among other signs, are the most reported as signs of PlTX poisoning [161].

2.3. Marine Cyanotoxins

Most marine toxins reported are produced mainly by microalgae (composed basically by dinoflagellates, diatoms, and marine bacteria), while cyanobacteria are reported as toxin producers in fresh, brackish waters and terrestrial habitats. Recently, cyanotoxins typical from freshwater have been identified in the marine environment [162]. Thus, this section will be focused on the description of the most reported marine cyanotoxins involved in seafood poisoning, their producers and mode of action (Table 1).
One of the most relevant groups of marine cyanotoxins is themicrocystin group (MCs) [163] (Figure 12). MCs are produced by cyanobacteria of genus Pseudoanabaena, Phormidium, Spirilia [164], Leptolyngbya, Oscillatoria, Geitlerinema [165], Trichodesmium [166] and Synechococcus [167] and their occurrence have been reported in many parts of the world, namely: the central Atlantic coast of Portugal [168], Canary Islands Archipelago [166], Brazilian coast [169], Amvrakikos Gulf (Greece) [167] and Indian Ocean [170]. To date, MCs is regulated in freshwater habitats but should be extended to the marine environments since there are reports of these hepatotoxins in marine environments [162].
Other reported marine cyanotoxins [in parenthesis is indicated their producers] (Figure 13) are aplysiatoxin (AT) [171] (Figure 13a), debromoaplysiatoxin (DAT) [171] (Figure 13) (algae Gracilaria coronopifolia [172] and cyanobacteria Lyngbya majuscule [171]), kalkitoxin (KTX) (cyanobacteria Lyngbyamajuscula [173]) (Figure 13b), lyngbyatoxins (LA, LB and LC) (cyanobacteria Lyngbya majuscule [174]) [Figure 13c], cylindrospermopsins (CYNs) (cyanobacteria Cylindrospermopsis raciborskii [175]) (Figure 13d), jamaicamides (JCDs) (Cyanobacteria Lyngbya majuscule [176]) (Figure 13e), anatoxins (ANTX) (cyanobacteria Hydrocoleum lyngbyaceum [177]) [178] (Figure 13f) andantillatoxins (ATX) (cyanobacteria Lyngbya majuscule [179]) (Figure 13g). The mechanism of action anddetection methods are presented in Table 1.
Recent studies indicate Homoanatoxin-a (HANTX, a derivative of anatoxin-a) produced by the cyanobacteria Hydrocoleum sp. and Trichodesmium sp. which co-occur with G. toxicus, may be the causative toxin of CFP [43] (rather than CTXs). This evidence suggests further studies to clarify marine cyanotoxins responsible for CFP and their mechanism of action [178]. The reports of seafood poisoning involving marine cyanotoxins are very scarce and consequently, there is no specific symptomology that can be related to marine cyanotoxin human poisoning.

3. Incidence of Harmful Algal Blooms MarineToxins and Consequent Poisoning Incidents along African Indian and the Red Sea Coasts

The main geographical focus of this review is the African Indian and the Red Sea coasts, including surrounding islands (Figure 14). The marine environment of this area is understudied due to a lack of monitoring infrastructure. There is a high rate of poverty in local communities, and the local population is vulnerable to natural disasters [including HABs, tropical storms]. The exponential increase in population accompanied by industrialization and climate change contributes to eutrophication in coastal areas [295,296]. This study area is characterized as subtropical to tropical climate with a water temperature above 20 °C [297]. Eutrophication and the transportation of cysts [through maritime traffic] are considered the main factors contributing to large phytoplankton blooms, including those comprised of HAB species and/or pathogenic bacteria [295,296]. Countries with monitoring programs of marine environments related to control of seafood poisoning are listed in Table 2. A few of these programs have noted the presence of MTs (Figure 14) and HAB species [dinoflagellates, cyanobacteria, diatoms], some of which [HAB species] were detected/confirmed by microscopic techniques and some confirmed by partial 16 S rRNA genes analysis [12,13,298,299,300,301,302,303,304,305,306,307,308,309,310,311,312,313,314,315,316,317,318,319,320,321,322,323].

3.1. South Africa

The occurrence of species of phytoplankton including MTs-producing HABs has been reported in coastal waters of South Africa through scientific reports and environmental monitoring programmes since 2011 [324]. Reported producer species include cyanobacteria (Microcystisaeruginosa, Oscillatoria sp., Trichodesmium sp.), dinoflagellates (Dinophysisacuminata, D. rotundata, Alexandrium catenella, A. minutum, Gymnodinium sp., Prorocentrum sp., Gambierdiscustoxicus, Ostreopsis siamensis, O. ovata, P. lima, P. concavum), diatoms (Pseudo-nitzschia multiseries) [19,305,309,315,331,332,333] and bacteria (Vibrio parahaemolyticus) [298]. Seafood poisoning cases were also reported in South Africa caused by PSTs, DSPs, PlTXs and GYM [19,216,309,334] (Table 3) after the consumption of mussels (Donax serra, Perna perna and Chloromytilus meridionalis) (Table 4) [37]. To minimize seafood poisoning by MTs, South Africa has implemented, through the Department of Agriculture, a program for MT monitoring in molluscan shellfish on all coasts (South African Molluscan Shellfish Monitoring and Control Programme) [324] (Table 2). This program was created based on the regulations of the European Commission (EC) Regulation, namely: Commission Regulation (EC) No 2074/2005, No 853/2004 and No 15/2011 where limit values are described for MTs and analytical techniques are advised to monitor shellfish [324].
Due to the absence of legislation regarding CTXs, currently, there is an absence of monitoring programs regarding this group in South Africa.Since the Indian Ocean is considered an endemic site of CTXs, this is a matter of major importance.

3.2. Mozambique

Studies related to HAB occurrence in Mozambique are very scarce and the few published works indicate the occurrence of dinoflagellates of the genus Alexandrium [313] and species of cyanobacteria (Phormidium ambiguum, Lyngbya majuscula, and Lyngbya cf. putealis) [307]. To date, due to the absence of a Monitoring Program and trained health staff to recognize specific symptoms of seafood poisoning in humans, there are no records of published data of MT occurrence or reports of seafood poisoning cases in this country.

3.3. Tanzania

Published studies indicate the occurrence of cyanobacteria, namely: Pseudanabaena sp., Spirulina labyrinthiformis, Spirulina sp., Leptolyngbya sp., Phormidium sp., Oscillatoria sp., Lyngbyaaestuarii, Lyngbya sp., Lyngbya majuscula, Nodularia sp., Synechococcus sp., Microcystis sp.; Dinoflagellates: Gambierdiscus toxicus, Procentrum sp. and diatoms: Pseudo-nitzschia sp., Pseudo-nitzschia pungens, P. seriata and P. cuspidate [335,336,337,338,339,340,341]. Data related to MTs and seafood poisoning episodes are very scarce in Tanzania. In 2003, the Tanzanian government created guidelines for investigation and control of foodborne diseases and the regulatory institution is the Tanzania Food and Drugs Authority (TFDA) (Table 2) [325]. The main objective of TFDA is to regulate matters related to food quality and safety for consumers through the dissemination of the information related to causative agents, latency period [duration], principal symptoms, typical vectors, and prevention of poisoning as measures of public health protection [325]. Among several foodborne disease sources, MTs such as CTXs, TTXs, DA, and PSTs are described by TFDA. The creation of alert and monitoring programs is an effective way to prevent poisoning episodes caused by MT-contaminated seafood.

3.4. Kenya

In order to reduce the cases of seafood poisoning caused by MTs, the Kenya Marine and Fisheries Research has carried out projects funded by governmental and non-governmental institutions for monitoring levels of HABs and their toxins (Table 2) in coastal waters and shellfish as well as the possible transfer in the trophic food web [326].Since October 2017, there is an ongoing project called: The occurrence and distribution of HABS in East and South Africa (BIOTOXINS Research Project] funded by National Commission for Science, Technology and Innovation (NACOSTI) at Mombasa Research Center [326]. This project will cover a period of 2 years, which is not enough for long-termmonitoring. In these coastal waters were reported to occur several species of diatoms: Nitzschia sp., N. closterium, N. longisigma, N. sigma, Pseudo-nitzschia sp. Guinardia sp., G. striata, G.delicatula, Skeletonema sp, Leptocylindrus sp., Rhizosolenia sp., Cerataulina sp., Coscinodiscus sp., Thalassiosira sp., Corethron sp., C. criopilum, C. cenofemus and Chaetoceros sp.; dinoflagellates: Alexandrium sp., Dinophysis sp., D. caudata, Gambierdiscus sp., G. toxicus, Gonyaulax sp., Gymnodinium sp., Gyrodinium sp., Ostreopsis sp., Peridinium sp., Prorocentrum sp., Ceratium sp., C. fusus, C. furca, Noctiluca sp., N. scintillans, Protoperidinium sp., Scrippsiella sp. and S. trochoidea [301,310]. Cyanobacteria were also reported: Lyngbya sp., Oscillatoria sp., Fischerella epiphytica, Anabaena sp., Nodularia spumigena, Umezakia natans, Aphanizomenon flos-aquae, Microcystis aeruginosa and Trichodesmium sp. [342].

3.5. Madagascar

Madagascar is the country with more records of published data regarding MT occurrence (Figure 14) and consequently, many reported cases of seafood poisoning [36,47,49,343]. The seafood poisoning cases in Madagascar have been registered since 1930 mainly after the consumption of fish of the family Sphyrnidae, Cacharinidae, Clupeidae (herrings, sardines), and marine turtles species (Eretmochelys imbricata and Chelonia mydas) [36,47,49,343]. The main marine poisoning causative agents reported are CTXs, TTXs, and PlTXs [18,344] (Table 4). To reduce the number of seafood poisoning events, the MadagascarMinistry of Health has created a Seafood Poisoning National Control Program (Table 2) based on the setting of an epidemiological surveillance network, prevention of the communities through educational programs and the development of research on marine eco-environment [327].

3.6. Indian Ocean French Islands

Mayotte, Europa, Banc du Geyser, Bassas da India, Glorioso, Juan de Nova, Reunion and Tromelin islands administratively make part in the French government but since they are in the Indian Ocean, were considered for the present study. In these islands, there are reports of the occurrence of HABs and cases of seafood poisoning linked to MTs. The reported HAB forming species include: dinoflagellates (Prorocentrum lima, P. convacum, Ostreopsis ovata, Gambierdiscus toxicus, Alexandrium spp.), cyanobacteria (Hydrocoleum sp., Lyngbya majuscula, Phormidium sp., Leptolyngbya sp. and Oscillatoria sp.) [70,300,317,319,345]. The recorded human intoxications were due to DSTs and TTXs [35,328] (Table 4). Centers of Disease for control and Preventing is the organization responsible for National Biomonitoring Program of toxins (PSTs) in these islands [35,328] (Table 2).

3.7. Mauritius

In Mauritius there are registered cases of seafood poisoning caused mainly by CTXs [346] after the consumption of reeffish (Lutjanus sebae) [70,71,71] (Table 4). The Ministry of Ocean Economy, Marine Resources, Fisheries and Shipping of Mauritius is the institute responsible for themonitoring of HABs (Table 2) [347,348], developing several activities and reporting the principal vectors species involved in seafood poisoning, namely: fish (Variola louti, Plectroponus maculatus, ceragidae, Vieille loutre, V. plate, V. cuisinier, Lutjanus gibbus, L. sebae, L. monostigmus, L. bohar, Anyperodon leucogramnicus, Harengula ovalis, Sphyraena barracuda, Synancela verrucose, Remora remora, Lactoria carnuta, Diodon hystrix), turtles (Eretmochelys imbricate), crabs (Carpillus maculatus), sea-urchins (Echinothrix sp.) and bivalves (Tridaena sp.) [348].
HAB producers recorded in Mauritius include several dinoflagellates species (Ostreopsis mascarenensis, Gambierdiscus toxicus Adachi & Fukuyo, Ostreopsis ovata Fukuyo, Ostreopsis siamensis, O. mascarenensis, Prorocentrum lima, P. concavum, P. hoffmanianum, Amphidinium sp., A. carterae, Coolia sp., Sinophysis sp., Gymnodinium sp., Gonyaulax sp., and Alexandrium sp.), diatoms (Pseudo-nitzschia sp.) and cyanobacteria (Phormidium sp., Oscillatoria sp. and Lyngbya sp., Phormidium sp., Oscillatoria sp. and Lyngbya sp.) [308].

3.8. The Archipelago of Comoros

Published data of the archipelago of Comoros indicate the occurrence of Gambierdiscus toxicus, G. yasumotoi, G. belizeanus, Prorocentrum arenarium, P. maculosum, P. belizeanum, P. lima, P. mexicanum, P. hoffmanianum, P. concavum, P. emarginatum, P. elegans, P. sp., Ostreopsis caribbeanus, O. mascarenensis, O. ovata O. heptagona, O. labens, O. siamensis, O. lenticularis, O. marinus, Cooliamonotis, C. tropicalis, Sinophysis microcephalus, S. canaliculate and Amphidiniopsis sp. [10,300]. Suspected seafood poisoning episodes linked to MTs were registered in the archipelago of Comoros after the consumption of turtle Eretmochelys imbricate with symptomatology similar to CFP [26], suggesting the presence of CTXs (Table 4).

3.9. Somalia and Seychelles

There are no published studies related to the occurrence of HABs and MTs in Somalia and Seychelles. While there are no published reports of HABs or MTs in Somalia and Seychelles waters, the proximity to other countries with such reports and currents in the area suggest that investigations are necessary to avoid potential seafood poisoning events [62].

3.10. Mediterranean and Red Sea (Djibouti, Eritrea, Sudan, Egypt)

Several research works related to MTs are carried out in the Red Sea but are very limited on the African coast. Saudi Arabia is the country with the most published studies related to the occurrence of HABs along the Red Sea [13,308,311,316,321,322,352,353]. The Dinoflagellates (Alexandrium sp., Dinophysis sp., Prorocentrum sp., Pyrodinium sp., Gymnodinium sp.), cyanobacteria (Lyngbya sp., Oscillatoria sp., Trichodesmium sp.) and diatoms (Pseudonitzschia spp.) are the most reported marine producer species [13,308,311,316,321,322,352,353]. The bacteria Vibrio paraehemolyticus, producer of TTX, was detected in shrimp (Penaeus latisulcatus) in the Suez Gulf [299]. MTs reported in the Red Sea, mainly the Egyptian coast, described in Table 3 and Table 4, include CTXs, TTXs, PSTs detected in puffer fish such as Pleuranacanthus sceleratus and Lagocephalus sceleratus [13,316,349,350,351,352,353]. Cases of seafood poisoning caused by CTXs and TTXs were reported, and according to the Poison Control Center, affiliated with Ain Shams University (Cairo, Egypt), CTXs are the third most responsible agents that induce food poisoning in Egypt [354]. Puffer fish poisoning has been recorded since ancient Egyptian times [42]. In Egypt, there is monitoring ofHABs in aquatic ecosystems since 1994 when Egypt became a member of the Convention on Biological Diversity although the Nature Conservation Sector, Egyptian Environment Affairs Agency and the Ministry of State for Environmental Affairs (Table 2) are focal points [330]. There are no reports of HABs and MT occurrence in coastal areas of Djibouti, Eritrea, and Sudan.

4. Final Considerations and Recomendations

African Indian Ocean and the Red Sea coasts have a subtropical and tropical climate, considered optimal for the development and transportation of several HAB-forming species, and consequently, the production of MTs. Paradoxically, studiesrelated to the occurrence and incidence of HABs and MTs are very limited, from South Africa to Egypt. From a few data available in this zone, most describe only the genus and not the full species, making it very difficult to evaluate the occurrence of the toxic species. The most reported HAB phytoplanktons in this region are cyanobacteria, followed by dinoflagellates, and diatoms as potential MT producers. Relative to MTs, the most reported and involved in seafood poisoning episodes include CTXs, PSTs, and TTXs. The scarcity of the data related to MTs suggests the need for further studies and the creation of specific monitoring programs of HABs, particularly for dinoflagellates and diatoms since these constitute the phytoplankton that produces more fatal MTs, though in recent years several genera of bacteria have been described as producers of a potent group of marine toxins, TTXs, which have already been detected on the African coasts of the Indian Ocean and Red Sea. The main MTs that must be monitored in shellfish are presented in Table 5. Analytical techniques such as LC-MS/MS are advised and recommended as determination and quantification methods due to their higher reproducibility, specificity, sensitivity and capacity to discriminate analogs of given toxins in the sample. The permitted limit of a toxin in shellfish can be adopted from other countries as an example to follow such as the EU region, USA, Japan, Australia, and New Zealand.
For the success of the MT monitoring programs, the integration and intercollaboration of environmental, public health and researches institutions and universities of the all African Countries of the Indian Ocean and the Red Sea is crucial.

Author Contributions

Conceptualization, V.V. and M.S.; Writing-Original Draft Preparation, I.J.T.; Writing-Review & Editing, I.J.T, M.S. and V.V.; Supervision, V.V. and M.S.


This research was supported by the project Alertox-Net [EAPA-317-2016] of the Interreg Atlantic Area Program funded by the European Regional Development Fund and by the Portuguese Foundation of Science and Technology [FCT] project UID/Multi/04423/2013.


We acknowledge the project EMERTOX [grant 734748], funded by H2020-MSCA-RISE 2016.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Cembella, A.D.; Lewis, N.I.; Quilliam, M.A. The marine dinoflagellate Alexandrium ostenfeldii [Dinophyceae] as the causative organism of spirolide shellfish toxins. Phycologia 2000, 39, 67–74. [Google Scholar] [CrossRef]
  2. MacKenzie, L.; de Salas, M.; Adamson, J.; Beuzenberg, V. The dinoflagellate genus Alexandrium [Halim] in New Zealand coastal waters: Comparative morphology, toxicity and molecular genetics. Harmful Algae 2004, 3, 71–92. [Google Scholar] [CrossRef]
  3. Beppu, R.; Nojima, K.; Tsuruda, S.; Gomez-Delan, G.; Barte-Quilantang, M.; Taniyama, S.; Sagara, T.; Nishio, S.; Takayama, H.; Miyazawa, K.; et al. Occurrence of PSP-producing dinoflagellate Alexandrium tamiyavanichii in Bingo-Nada, the central coastal water of the Seto Inland Sea, Hiroshima Prefecture, Japan. Mar. Pollut. Bull. 2008, 56, 758–763. [Google Scholar] [CrossRef] [PubMed]
  4. Wang, D.-Z. Neurotoxins from marine dinoflagellates: A brief review. Mar. Drugs 2008, 6, 349–371. [Google Scholar] [CrossRef] [PubMed]
  5. Seki, T.; Satake, M.; Mackenzie, L.; Kaspar, H.F.; Yasumoto, T. Gymnodimine, a new marine toxin of unprecedented structure isolated from New Zealand oysters and the dinoflagellate, Gymnodinium sp. Tetrahedron Lett. 1995, 36, 7093–7096. [Google Scholar] [CrossRef]
  6. Draisci, R.; Lucentini, L.; Giannetti, L.; Boria, P.; Poletti, R. First report of pectenotoxin-2 [PTX-2] in algae [Dinophysis fortii] related to seafood poisoning in Europe. Toxicon 1996, 34, 923–935. [Google Scholar] [CrossRef]
  7. Martin, J.L.; Hanke, A.R.; LeGresley, M.M. Long term phytoplankton monitoring, including harmful algal blooms, in the Bay of Fundy, eastern Canada. J. Sea Res. 2009, 61, 76–83. [Google Scholar] [CrossRef]
  8. Jeffrey, L.C. Identification of DTX-4, a new water-soluble phosphatase inhibitor from the toxic dinoflagellate Prorocentrum lima. J. Chem. Soc. Chem. Commun. 1995, 597–599. [Google Scholar] [CrossRef]
  9. MacKenzie, L.; Beuzenberg, V.; Holland, P.; McNabb, P.; Suzuki, T.; Selwood, A. Pectenotoxin and okadaic acid-based toxin profiles in Dinophysis acuta and Dinophysis acuminata from New Zealand. Harmful Algae 2005, 4, 75–85. [Google Scholar] [CrossRef]
  10. Ten-Hage, L.; Turquet, J.; Quod, J.P.; Couté, A. Coolia areolata sp. nov. [Dinophyceae], a new sand-dwelling dinoflagellate from the southwestern Indian Ocean. Phycologia 2000, 39, 377–383. [Google Scholar] [CrossRef]
  11. Ten-Hage, L.; Delaunay, N.; Pichon, V.; Couté, A.; Puiseux-Dao, S.; Turquet, J. Okadaic acid production from the marine benthic dinoflagellate Prorocentrum arenarium Faust [Dinophyceae] isolated from Europa Island coral reef ecosystem [SW Indian Ocean]. Toxicon 2000, 38, 1043–1054. [Google Scholar] [CrossRef]
  12. Pitcher, G.C.; Cembella, A.D.; Krock, B.; Macey, B.M.; Mansfield, L.; Probyn, T.A. Identification of the marine diatom Pseudo-nitzschia multiseries [Bacillariophyceae] as a source of the toxin domoic acid in Algoa Bay, South Africa. Afr. J. Mar. Sci. 2014, 36, 523–528. [Google Scholar] [CrossRef]
  13. Mohamed, Z.A.; Al-Shehri, A.M. Biodiversity and toxin production of cyanobacteria in mangrove swamps in the Red Sea off the southern coast of Saudi Arabia. Bot. Mar. 2015, 58, 23–34. [Google Scholar] [CrossRef]
  14. Lenoir, S.; Ten-Hage, L.; Turquet, J.; Quod, J.; Bernard, C.; Hennion, M. First evidence of palytoxin analogues from an Ostreopsis mascarenensis (Dinophyceae) benthic bloom in Southwestern Indian Ocean. J. Phycol. 2004, 40, 1042–1051. [Google Scholar] [CrossRef]
  15. Jørgensen, K.; Andersen, P. Relation between the concentration of Dinophysis acuminata and diarrheic shellfish poisoning toxins in blue mussels [Mytilus edulis] during a toxic episode in the Limfjord [Denmark], 2006. J. Shellfish Res. 2007, 26, 1081–1087. [Google Scholar] [CrossRef]
  16. Landsberg, J.H.; Flewelling, L.J.; Naar, J. Karenia brevis red tides, brevetoxins in the food web, and impacts on natural resources: Decadal advancements. Harmful Algae 2009, 8, 598–607. [Google Scholar] [CrossRef]
  17. El-Sayed, M.; Yacout, G.A.; El-Samra, M.; Ali, A.; Kotb, S.M. Toxicity of the Red Sea pufferfish Pleuranacanthus sceleratus “El-Karad. ” Ecotoxicol. Environ. Saf. 2003, 56, 367–372. [Google Scholar] [CrossRef]
  18. Onuma, Y.; Satake, M.; Ukena, T.; Roux, J.; Chanteau, S.; Rasolofonirina, N.; Ratsimaloto, M.; Naoki, H.; Yasumoto, T. Identification of putative palytoxin as the cause of clupeotoxism. Toxicon 1999, 37, 55–65. [Google Scholar] [CrossRef]
  19. Pitcher, G.C.; Krock, B.; Cembella, A.D. Accumulation of diarrhetic shellfish poisoning toxins in the oyster Crassostrea gigas and the mussel Choromytilus meridionalis in the southern Benguela ecosystem. Afr. J. Mar. Sci. 2011, 33, 273–281. [Google Scholar] [CrossRef]
  20. Diogène, J.; Reverté, L.; Rambla-Alegre, M.; Río, V.; Iglesia, P.; Campàs, M.; Palacios, O.; Flores, C.; Caixach, J.; Ralijaona, C.; et al. Identification of ciguatoxins in a shark involved in a fatal food poisoning in the Indian Ocean. Sci. Rep. 2017, 7, 8240. [Google Scholar] [CrossRef]
  21. Habermehl, G.G.; Krebs, H.C.; Rasoanaivo, P.; Ramialiharisoa, A. Severe ciguatera poisoning in Madagascar: A case report. Toxicon 1994, 32, 1539–1542. [Google Scholar] [CrossRef]
  22. Pitcher, G.C.; Franco, J.M.; Doucette, G.J.; Powell, C.L.; Mouton, A. Paralytic Shellfish Poisoning in the abalone Haliotis midae on the West Coast of South Africa. J. Shellfish Res. 2001, 20, 895–904. [Google Scholar]
  23. Silva, M.; Rodriguez, I.; Barreiro, A.; Kaufmann, M.; Neto, A.I.; Hassouani, M.; Sabour, B.; Alfonso, A.; Botana, L.M.; Vasconcelos, V. First report of ciguatoxins in two starfish species: Ophidiaster ophidianus and Marthasterias glacialis. Toxins 2015, 7, 3740–3757. [Google Scholar] [CrossRef] [PubMed]
  24. Vale, P.; de M Sampayo, M.A. First confirmation of human diarrhoeic poisonings by okadaic acid esters after ingestion of razor clams [Solen marginatus] and green crabs [Carcinus maenas] in Aveiro lagoon, Portugal and detection of okadaic acid esters in phytoplankton. Toxicon 2002, 40, 989–996. [Google Scholar] [CrossRef]
  25. Ahmed, S. Puffer fish tragedy in Bangladesh: An incident of Takifugu oblongus poisoning in Degholia, Khulna. Afr. J. Mar. Sci. 2006, 28, 457–458. [Google Scholar] [CrossRef]
  26. Mbaé, S.B.A.; Mlindassé, M.; Mihidjaé, S.; Seyler, T. Food-poisoning outbreak and fatality following ingestion of sea turtle meat in the rural community of Ndrondroni, Mohéli Island, Comoros, December 2012. Toxicon 2016, 120, 38–41. [Google Scholar] [CrossRef] [PubMed]
  27. Yong, Y.S.; Quek, L.S.; Lim, E.K.; Ngo, A. A case report of puffer fish poisoning in Singapore. Case Rep. Med. 2013. [Google Scholar] [CrossRef]
  28. Hwang, P.-A.; Tsai, Y.-H.; Lu, Y.-H.; Hwang, D.-F. Paralytic toxins in three new gastropod [Olividae] species implicated in food poisoning in southern Taiwan. Toxicon 2003, 41, 529–533. [Google Scholar] [CrossRef]
  29. Rafiqui Islam, M.; Chowdhury, F.R.; Das, S.K.; Rahman, S.; Mahmudur, M.D.; Amin, M.D.R. Outbreak of Puffer Fish Poisoning in Dhaka City. J. Med. 2018, 19, 30–34. [Google Scholar] [CrossRef]
  30. Field, J. Puffer fish poisoning. Emerg. Med. J. 1998, 15, 334–336. [Google Scholar] [CrossRef]
  31. Chopra, S.A. A case of fatal puffer-fish poisoning in a Zanzibari fisherman. East Afr. Med. J. 1967, 44, 493–496. [Google Scholar]
  32. Ellis, R.; Jelinek, G.A. Never eat an ugly fish: Three cases of tetrodotoxin poisoning from Western Australia. Emerg. Med. 1997, 9, 136–142. [Google Scholar] [CrossRef]
  33. Ghose, A.; Ahmed, H.; Basher, A.; Amin, M.R.; Sayeed, A.A.; Faiz, M.A. Tetrodotoxin poisoning in Blangadesh: A case study. J. Med. Toxicol. 2008, 4, 216. [Google Scholar]
  34. Halstead, B.W.; Cox, K.W. An investigation on fish poisoning in Mauritius. Proc. R. Soc. Arts Sci. Maruritius 1973, 4, 1–26. [Google Scholar]
  35. Puech, B.; Batsalle, B.; Roget, P.; Turquet, J.; Quod, J.-P.; Allyn, J.; Idoumbin, J.P.; Chane-Ming, J.; Villefranque, J.; Mougin-Damour, K.; et al. Family tetrodotoxin poisoning in Reunion Island [Southwest Indian Ocean] following the consumption of Lagocephalus sceleratus [Pufferfish]. Bull. Soc. Pathol. Exot. 2014, 107, 79–84. [Google Scholar] [CrossRef] [PubMed]
  36. Ribes, G.C.; Ramarokoto, S.; Rabearintsoa, S.; Robinson, R.; Ranaivoson, G.; Rakotonjanabelo, L.A.; Rabeson, D. Seafood poisoning in Madagascar: Current state of knowledge and results of a retrospective study of the inhabitants of coastal villages [Internet]. Sante 1999, 9, 235–241. Available online: (accessed on 20 June 2018). [PubMed]
  37. Grindley, J.R.; Sapeika, N. The cause of mussel poisoning in South Africa. S. Afr. Med. J. 1969, 43, 275–279. [Google Scholar]
  38. Linlawan, S.; Suteparuk, S. Puffer fish poisoning from illicit fish trading in Bangkok, Thailand. J. Med. Toxicol. 2008, 4, 215. [Google Scholar]
  39. Popkiss, M.E.; Horstman, D.A.; Harpur, D. Paralytic shellfish poisoning. A report of 17 cases in Cape Town. S. Afr. Med. J. Suid-Afrikaanse Tydskr vir Geneeskd 1979, 55, 1017–1023. [Google Scholar]
  40. Mann, N.M.; Winship, W.S. Paralytic mussel poisoning in Natal. S. Afr. Med. J. 1958, 32, 548–549. [Google Scholar]
  41. Ravaonindrina, N.; Andriamaso, T.H.; Rasolofonirina, N. Puffer fish poisoning in Madagascar: Four case reports. Arch. Inst. Pasteur Madag. 2001, 67, 61–64. [Google Scholar]
  42. Jong, E.C. Fish and shellfish poisoning: Toxic syndromes. In The Travel and Tropical Medicine Manual; Jong, E.C., Sanford, C., Eds.; W.B. Saunders: Edinburgh, 2008; pp. 474–480. [Google Scholar]
  43. Laurent, D.; Kerbrat, A.-S.; Darius, H.T.; Girard, E.; Golubic, S.; Benoit, E.; Sauviat, M.-P.; Chinain, M.; Molgo, J.; Pauillac, S.; et al. Are cyanobacteria involved in Ciguatera Fish Poisoning-like outbreaks in New Caledonia? Harmful Algae 2008, 7, 827–838. [Google Scholar] [CrossRef]
  44. Ishida, H.; Muramatsu, N.; Nukaya, H.; Kosuge, T.; Tsuji, K. Study on neurotoxic shellfish poisoning involving the oyster, Crassostrea gigas, in New Zealand. Toxicon 1996, 34, 1050–1053. [Google Scholar] [CrossRef]
  45. Boisier, P.; Ranaivoson, G.; Rasolofonirina, N.; Roux, J.; Chanteau, S.; Takeshi, Y. Fatal mass poisoning in Madagascar following ingestion of a shark [Carcharhinus leucas]: Clinical and epidemiological aspects and isolation of toxins. Toxicon 1995, 33, 1359–1364. [Google Scholar] [CrossRef]
  46. F.E.R. Paralytic shellfish poisoning in eastern canada: Prackash, A., Medcof, J. C. And Tennant, A.D. Fisheries research board of canada, bull. 71, ottawa, 1971, 88 p. Toxicon 1973, 11, 209–210. [Google Scholar] [CrossRef]
  47. Ranaivoson, G.; de Ribes Champetier, G.; Mamy, E.R.; Jeannerod, G.; Razafinjato, P.; Chanteau, S. Mass food poisoning after eating sea turtle in the Antalaha district. Arch. Inst. Pasteur Madag. 1994, 61, 84–86. [Google Scholar]
  48. Islam, Q.T.; Razzak, M.A.; Islam, M.A.; Bari, M.I.; Basher, A.; Chowdhury, F.R.; Sayeduzzaman, A.B.; Ahasan, H.A.; Faiz, M.A.; Arakawa, O.; et al. Puffer fish poisoning in Bangladesh: Clinical and toxicological results from large outbreaks in 2008. Trans. R. Soc. Trop. Med. Hyg. 2011, 105, 74–80. [Google Scholar] [CrossRef]
  49. Champetier, D.R.G.; Rasolofonirina, R.N.; Ranaivoson, G.; Razafimahefa, N.; Rakotoson, J.D.; Rabeson, D. Intoxication by marine animal venoms in Madagascar [ichthyosarcotoxism and chelonitoxism]: Recent epidemiological data. Bull. Soc. Pathol. Exot. 1997, 90, 286–290. [Google Scholar]
  50. Hallegraeff, G.M. A review of harmful algal blooms and their apparent global increase. Phycologia 1993, 32, 79–99. [Google Scholar] [CrossRef]
  51. Council of the European Union; Council Directive 86/609/EEC of 24 November 1986 on the approximation of laws, regulations and administrative provisions of the Member States regarding the protection of animals used for experimental and other scientific purposes. Off. J. Eur. Commun. 1986, 29, L358.
  52. Regulation, C. COMMISSION REGULATION [EU] No 15/2011 of 10 January 2011 amending Regulation [EC] No 2074/2005 as regards recognised testing methods for detecting marine biotoxins in live bivalve molluscs. Off. J. Eur. Commun. 2011, 50, 3–4. [Google Scholar]
  53. Spatharis, S.; Dolapsakis, N.P.; Economou-Amilli, A.; Tsirtsis, G.; Danielidis, D.B. Dynamics of potentially harmful microalgae in a confined Mediterranean Gulf—Assessing the risk of bloom formation. Harmful Algae 2009, 8, 736–743. [Google Scholar] [CrossRef]
  54. Raho, N.; Pizarro, G.; Escalera, L.; Reguera, B.; Marín, I. Morphology, toxin composition and molecular analysis of Dinophysis ovum Schütt, a dinoflagellate of the “Dinophysis acuminata complex”. Harmful Algae 2008, 7, 839–848. [Google Scholar] [CrossRef]
  55. Caroppo, C.; Congestri, R.; Bruno, M. On the presence of Phalacroma rotundatum in the southern Adriatic Sea [Italy]. Aquat. Microb. Ecol. 1999, 17, 301–310. [Google Scholar] [CrossRef]
  56. McCarron, P.; Kilcoyne, J.; Hess, P. Effects of cooking and heat treatment on concentration and tissue distribution of okadaic acid and dinophysistoxin-2 in mussels [Mytilus edulis]. Toxicon 2008, 51, 1081–1089. [Google Scholar] [CrossRef] [PubMed]
  57. Tanti, J.-F.; Gremeaux, T.; Van Obberghen, E.; Le Marchand-Brustel, Y. Effects of okadaic acid, an inhibitor of protein phosphatases-1 and-2A, on glucose transport and metabolism in skeletal muscle. J. Biol. Chem. 1991, 266, 2099–2103. [Google Scholar] [PubMed]
  58. Louzao, M.C.; Vieytes, M.R.; Botana, L.M. Effect of okadaic acid on glucose regulation. Mini Rev. Med. Chem. 2005, 5, 207–215. [Google Scholar] [CrossRef]
  59. Yasumoto, T.; Seino, N.; Murakami, Y.; Murata, M. Toxins produced by benthic dinoflagellates. Biol. Bull. 1987, 172, 128–131. [Google Scholar] [CrossRef]
  60. Naoki, H.; Fujita, T.; Cruchet, P.; Legrand, A.M.; Igarashi, T.; Yasumoto, T. Structural determination of new ciguatoxin congeners by tandem mass spectrometry. In International IUPAC Symposium on Mycotoxins and Phycotoxins Ponsen & Looyen; Ponsen and Looijen: Wageningen, The Netherlands, 2001; pp. 475–482. [Google Scholar]
  61. Lewis, R.J.; Sellin, M.; Poli, M.A.; Norton, R.S.; MacLeod, J.K.; Sheil, M.M. Purification and characterization of ciguatoxins from moray eel [Lycodontis javanicus, Muraenidae]. Toxicon 1991, 29, 1115–1127. [Google Scholar] [CrossRef]
  62. Lewis, R.J. The changing face of ciguatera. Toxicon 2001, 39, 97–106. [Google Scholar] [CrossRef]
  63. Lehane, L.; Lewis, R.J. Ciguatera: Recent advances but the risk remains. Int. J. Food Microbiol. 2000, 61, 91–125. [Google Scholar] [CrossRef]
  64. Satake, M.; Murata, M.; Yasumoto, T. The structure of CTX3C, a ciguatoxin congener isolated from cultured Gambierdiscus toxicus. Tetrahedron Lett. 1993, 34, 1975–1978. [Google Scholar] [CrossRef]
  65. Satake, M.; Murata, M.; Yasumoto, T. Gambierol: A new toxic polyether compound isolated from the marine dinoflagellate Gambierdiscus toxicus. J. Am. Chem. Soc. 1993, 115, 361–362. [Google Scholar] [CrossRef]
  66. Satake, M.; Fukui, M.; Legrand, A.-M.; Cruchet, P.; Yasumoto, T. Isolation and structures of new ciguatoxin analogs, 2, 3-dihydroxyCTX3C and 51-hydroxyCTX3C, accumulated in tropical reef fish. Tetrahedron Lett. 1998, 39, 1197–1198. [Google Scholar] [CrossRef]
  67. Pottier, I.; Vernoux, J.-P.; Jones, A.; Lewis, R.J. Characterisation of multiple Caribbean ciguatoxins and congeners in individual specimens of horse-eye jack [Caranx latus] by high-performance liquid chromatography/mass spectrometry. Toxicon 2002, 40, 929–939. [Google Scholar] [CrossRef]
  68. Bagnis, R.; Kuberski, T.; Laugier, S. Clinical observations on 3,009 cases of ciguatera [fish poisoning] in the South Pacific. Am. J. Trop. Med. Hyg. 1979, 28, 1067–1073. [Google Scholar] [CrossRef] [PubMed]
  69. Lewis, R.J.; Vernoux, J.-P.; Brereton, I.M. Structure of Caribbean ciguatoxin isolated from Caranx latus. J. Am. Chem. Soc. 1998, 120, 5914–5920. [Google Scholar] [CrossRef]
  70. Hamilton, B.; Hurbungs, M.; Jones, A.; Lewis, R.J. Multiple ciguatoxins present in Indian Ocean reef fish. Toxicon 2002, 40, 1347–1353. [Google Scholar] [CrossRef]
  71. Hamilton, B.; Hurbungs, M.; Vernoux, J.-P.; Jones, A.; Lewis, R.J. Isolation and characterisation of Indian Ocean ciguatoxin. Toxicon 2002, 40, 685–693. [Google Scholar] [CrossRef]
  72. Hokama, Y.; Abad, M.A.; Kimura, L.H. A rapid enzyme-immunoassay for the detection of ciguatoxin in contaminated fish tissues. Toxicon 1983, 21, 817–824. [Google Scholar] [CrossRef]
  73. Panel, E.C. Scienti fi c opinion on marine biotoxins in shell fi sh-emerging toxins: Ciguatoxin-group toxins. EFSA Panel Contam. Food Chain EFSA J. 2010, 8, 1627–1638. [Google Scholar]
  74. Pottier, I.; Vernoux, J.P.; Jones, A.; Lewis, R.J. Analysis of toxin profiles in three different fish species causing ciguatera fish poisoning in Guadeloupe, French West Indies. Food Addit. Contam. 2002, 19, 1034–1042. [Google Scholar] [CrossRef]
  75. Mello, F.D.; Braidy, N.; Marcal, H.; Guillemin, G.; Nabavi, S.M.; Neilan, B.A. Mechanisms and effects posed by neurotoxic products of cyanobacteria/microbial eukaryotes/dinoflagellates in algae blooms: A review. Neurotox. Res. 2018, 33, 153–167. [Google Scholar] [CrossRef] [PubMed]
  76. Touzet, N.; Franco, J.M.; Raine, R. Morphogenetic diversity and biotoxin composition of Alexandrium [Dinophyceae] in Irish coastal waters. Harmful Algae 2008, 7, 782–797. [Google Scholar] [CrossRef]
  77. Miles, C.O.; Wilkins, A.L.; Stirling, D.J.; MacKenzie, A.L. Gymnodimine C, an isomer of gymnodimine B, from Karenia selliformis. J. Agric. Food Chem. 2003, 51, 4838–4840. [Google Scholar] [CrossRef] [PubMed]
  78. Nézan, E.; Chomérat, N. Vulcanodinium rugosum gen. et sp. nov. [Dinophyceae], un nouveau dinoflagellé marin de la côte méditerranéenne française. Cryptogam. Algol. 2011, 32, 3–18. [Google Scholar] [CrossRef]
  79. Selwood, A.I.; Miles, C.O.; Wilkins, A.L.; van Ginkel, R.; Munday, R.; Rise, F.; McNabb, P. Isolation, structural determination and acute toxicity of pinnatoxins E, F and G. J. Agric. Food Chem. 2010, 58, 6532–6542. [Google Scholar] [CrossRef] [PubMed]
  80. Krock, B.; Tillmann, U.; John, U.; Cembella, A. LC-MS-MS aboard ship: Tandem mass spectrometry in the search for phycotoxins and novel toxigenic plankton from the North Sea. Anal. Bioanal. Chem. 2008, 392, 797–803. [Google Scholar] [CrossRef]
  81. Gill, S.; Murphy, M.; Clausen, J.; Richard, D.; Quilliam, M.; MacKinnon, S.; LaBlanc, P.; Mueller, R.; Pulido, O. Neural injury biomarkers of novel shellfish toxins, spirolides: A pilot study using immunochemical and transcriptional analysis. Neurotoxicology 2003, 24, 593–604. [Google Scholar] [CrossRef]
  82. Lawrence, J.; Loreal, H.; Toyofuku, H.; Hess, P.; Iddya, K. Assessment and Management of Biotoxin Risks in Bivalve Molluscs. 2011. Available online: (accessed on 10 November 2018).
  83. Silva, M.; Pratheepa, V.K.; Botana, L.M.; Vasconcelos, V. Emergent toxins in North Atlantic temperate waters: A challenge for monitoring programs and legislation. Toxins 2015, 7, 859–885. [Google Scholar] [CrossRef]
  84. Cembella, A.; Krock, B. Cyclic Imine Toxins: Chemistry, Biogeography, Biosynthesis and Pharmacology. In Seaf Freshw toxins Pharmacol Physiol Detect; Botana, L.M., Ed.; CRC Press: Boca Raton, FL, USA, 2007; pp. 561–580. [Google Scholar]
  85. Rundberget, T.; Aasen, J.A.B.; Selwood, A.I.; Miles, C.O. Pinnatoxins and spirolides in Norwegian blue mussels and seawater. Toxicon 2011, 58, 700–711. [Google Scholar] [CrossRef] [PubMed]
  86. Otero, P.; Alfonso, A.; Alfonso, C.; Vieytes, M.R.; Louzao, M.C.; Botana, A.M.; Botana, L.M. New protocol to obtain spirolides from Alexandrium ostenfeldii cultures with high recovery and purity. Biomed. Chromatogr. 2010, 24, 878–886. [Google Scholar] [PubMed]
  87. Watkins, S.M.; Reich, A.; Fleming, L.E.; Hammond, R. Neurotoxic shellfish poisoning. Mar. Drugs 2008, 6, 431–455. [Google Scholar] [CrossRef] [PubMed]
  88. Abraham, A.; Plakas, S.M.; Wang, Z.; Jester, E.L.E.; El Said, K.R.; Granade, H.R.; Henry, M.S.; Blum, P.C.; Pierce, R.H.; Dickey, R.W. Characterization of polar brevetoxin derivatives isolated from Karenia brevis cultures and natural blooms. Toxicon 2006, 48, 104–115. [Google Scholar] [CrossRef] [PubMed]
  89. Dickey, R.; Jester, E.; Granade, R.; Mowdy, D.; Moncreiff, C.; Rebarchik, D.; Robl, M.; Musser, S.; Poli, M. Monitoring brevetoxins during a Gymnodinium breve red tide: Comparison of sodium channel specific cytotoxicity assay and mouse bioassay for determination of neurotoxic shellfish toxins in shellfish extracts. Nat. Toxins 1999, 7, 157–165. [Google Scholar] [CrossRef]
  90. Ishida, H.; Nozawa, A.; Hamano, H.; Naoki, H.; Fujita, T.; Kaspar, H.F.; Tsuji, K. Brevetoxin B5, a new brevetoxin analog isolated from cockle Austrovenus stutchburyi in New Zealand, the marker for monitoring shellfish neurotoxicity. Tetrahedron Lett. 2004, 45, 29–33. [Google Scholar] [CrossRef]
  91. Murata, M.; Legrand, A.M.; Ishibashi, Y.; Fukui, M.; Yasumoto, T. Structures and configurations of ciguatoxin from the moray eel Gymnothorax javanicus and its likely precursor from the dinoflagellate Gambierdiscus toxicus. J. Am. Chem. Soc. 1990, 112, 4380–4386. [Google Scholar] [CrossRef]
  92. Morohashi, A.; Satake, M.; Murata, K.; Naoki, H.; Kaspar, H.F.; Yasumoto, T. Brevetoxin B3, a new brevetoxin analog isolated from the greenshell mussel Perna canaliculus involved in neurotoxic shellfish poisoning in New Zealand. Tetrahedron Lett. 1995, 36, 8995–8998. [Google Scholar] [CrossRef]
  93. Plakas, S.M.; El Said, K.R.; Jester, E.L.E.; Granade, H.R.; Musser, S.M.; Dickey, R.W. Confirmation of brevetoxin metabolism in the Eastern oyster [Crassostrea virginica] by controlled exposures to pure toxins and to Karenia brevis cultures. Toxicon 2002, 40, 721–729. [Google Scholar] [CrossRef]
  94. Wang, Z.; Plakas, S.M.; El Said, K.R.; Jester, E.L.E.; Granade, H.R.; Dickey, R.W. LC/MS analysis of brevetoxin metabolites in the Eastern oyster [Crassostrea virginica]. Toxicon 2004, 43, 455–465. [Google Scholar] [CrossRef]
  95. Baden, D.G. Brevetoxins: Unique polyether dinoflagellate toxins. FASEB J. 1989, 3, 1807–1817. [Google Scholar] [CrossRef] [PubMed]
  96. Poli, M.A. Laboratory procedures for detoxification of equipment and waste contaminated with brevetoxins PbTx-2 and PbTx-3. J. Assoc. Off. Anal. Chem. 1988, 71, 1000–1002. [Google Scholar] [PubMed]
  97. Baden, D.G.; Bourdelais, A.J.; Jacocks, H.; Michelliza, S.; Naar, J. Natural and derivative brevetoxins: Historical background, multiplicity, and effects. Environ. Health Perspect. 2005, 113, 621–625. [Google Scholar] [CrossRef] [PubMed]
  98. U.S. FDA [United States Food and Drug Administration]. Fish and Fisheries Products Hazards and Controls Guidance, 3rd ed.; Appendix 5—FDA & EPA Safety Levels in Regulations and Guidance; June 2001. Available online: 091782.htm (accessed on 24 July 2018 ).
  99. FSANZ [Food Standards Australia New Zealand]. Food Standard Code, Incorporating Amendments up to and Including Amendment 116, Standard 4.1.1, Primary Production and Processing Standards, Preliminary provisisons, Standard 1.4.1, Contaminants and Natural toxicants, [Internet]. 2010. Available online: (accessed on 24 July 2018).
  100. NZFSA (New Zealand Food). Animal products [specification for Bivalve Molluscan Shellfish]. 2006. Available online: (accessed on 24 July 2018).
  101. Miles, C.O.; Wilkins, A.L.; Munday, R.; Dines, M.H.; Hawkes, A.D.; Briggs, L.R.; Sandvik, M.; Jensen, D.J.; Cooney, J.M.; Holland, P.T.; et al. Isolation of pectenotoxin-2 from Dinophysis acuta and its conversion to pectenotoxin-2 seco acid, and preliminary assessment of their acute toxicities. Toxicon 2004, 43, 1–9. [Google Scholar] [CrossRef] [PubMed]
  102. Miles, C.O. Pectenotoxins. In Phycotoxins Chemistry Biochemistry; Botana, L.B., Alfonso, A., Eds.; Wiley-Blackwell: Hoboken, NJ, USA, 2007; pp. 159–186. [Google Scholar]
  103. Allingham, J.S.; Miles, C.O.; Rayment, I. A structural basis for regulation of actin polymerization by pectenotoxins. J. Mol. Biol. 2007, 371, 959–970. [Google Scholar] [CrossRef] [PubMed]
  104. Sasaki, K.; Wright, J.L.C.; Yasumoto, T. Identification and characterization of pectenotoxin [PTX] 4 and PTX7 as spiroketal stereoisomers of two previously reported pectenotoxins. J. Org. Chem. 1998, 63, 2475–2480. [Google Scholar] [CrossRef]
  105. Suzuki, T.; Walter, J.A.; LeBlanc, P.; MacKinnon, S.; Miles, C.O.; Wilkins, A.L.; Munday, R.; Beuzenberg, V.; MacKenzie, A.L.; Jensen, D.J.; et al. Identification of pectenotoxin-11 as 34 S-hydroxypectenotoxin-2, a new pectenotoxin analogue in the toxic dinoflagellate Dinophysis acuta from New Zealand. Chem. Res. Toxicol. 2006, 19, 310–318. [Google Scholar] [CrossRef]
  106. Zhou, Z.; Komiyama, M.; Terao, K.; Shimada, Y. Effects of pectenotoxin-1 on liver cells in vitro. Nat. Toxins 1994, 2, 132–135. [Google Scholar] [CrossRef]
  107. Cañete, E.; Diogène, J. Comparative study of the use of neuroblastoma cells [Neuro-2a] and neuroblastoma× glioma hybrid cells [NG108-15] for the toxic effect quantification of marine toxins. Toxicon 2008, 52, 541–550. [Google Scholar] [CrossRef]
  108. Toyofuku, H. Joint FAO/WHO/IOC activities to provide scientific advice on marine biotoxins. Mar. Pollut. Bull. 2006, 52, 1735–1745. [Google Scholar] [CrossRef]
  109. Loader, J.I.; Hawkes, A.D.; Beuzenberg, V.; Jensen, D.J.; Cooney, J.M.; Wilkins, A.L.; Fitzgerald, J.M.; Briggs, L.R.; Miles, C.O. Convenient large-scale purification of yessotoxin from Protoceratium reticulatum culture and isolation of a novel furanoyessotoxin. J. Agric. Food Chem. 2007, 55, 11093–11100. [Google Scholar] [CrossRef] [PubMed]
  110. Samdal, I.A. Yessotoxins in algae and mussels: Studies on its sources, disposition, and levels. uitgever niet vastgesteld. 2005. Available online: (accessed on 10 June 2018).
  111. EFSA. Opinion of the Scientific Panel on Contaminants in the Food chain on a request from the European Commission on marine biotoxins in shellfish—Yessotoxin group. EFSA J. 2008, 907, 1–62. [Google Scholar]
  112. Alfonso, A.; de la Rosa, L.; Vieytes, M.R.; Yasumoto, T.; Botana, L.M. Yessotoxin, a novel phycotoxin, activates phosphodiesterase activity: Effect of yessotoxin on cAMP levels in human lymphocytes. Biochem. Pharmacol. 2003, 65, 193–208. [Google Scholar] [CrossRef]
  113. Malagoli, D.; Casarini, L.; Ottaviani, E. Algal toxin yessotoxin signalling pathways involve immunocyte mussel calcium channels. Cell Biol. Int. 2006, 30, 721–726. [Google Scholar] [CrossRef] [PubMed]
  114. Pierotti, S.; Malaguti, C.; Milandri, A.; Poletti, R.; Rossini, G.P. Functional assay to measure yessotoxins in contaminated mussel samples. Anal. Biochem. 2003, 312, 208–216. [Google Scholar] [CrossRef]
  115. Malagoli, D.; Ottaviani, E. Yessotoxin affects fMLP-induced cell shape changes in Mytilus galloprovincialis immunocytes. Cell Biol. Int. 2004, 28, 57–61. [Google Scholar] [CrossRef] [PubMed]
  116. Dell’Ovo, V.; Bandi, E.; Coslovich, T.; Florio, C.; Sciancalepore, M.; Decorti, G.; Sosa, S.; Lorenzon, P.; Yasumoto, T.; Tubaro, A. In vitro effects of yessotoxin on a primary culture of rat cardiomyocytes. Toxicol. Sci. 2008, 106, 392–399. [Google Scholar] [CrossRef]
  117. Tillmann, U.; Elbrächter, M.; Krock, B.; John, U.; Cembella, A. Azadinium spinosum gen. et sp. nov. [Dinophyceae] identified as a primary producer of azaspiracid toxins. Eur.J. Phycol. 2009, 44, 63–79. [Google Scholar] [CrossRef]
  118. James, K.J.; Moroney, C.; Roden, C.; Satake, M.; Yasumoto, T.; Lehane, M.; Furey, A. Ubiquitous ‘benign’alga emerges as the cause of shellfish contamination responsible for the human toxic syndrome, azaspiracid poisoning. Toxicon 2003, 41, 145–151. [Google Scholar] [CrossRef]
  119. Satake, M.; Ofuji, K.; Naoki, H.; James, K.J.; Furey, A.; McMahon, T.; Silke, J.; Yasumoto, T. Azaspiracid, a new marine toxin having unique spiro ring assemblies, isolated from Irish mussels, Mytilus edulis. J. Am. Chem. Soc. 1998, 120, 9967–9968. [Google Scholar] [CrossRef]
  120. Ofuji, K.; Satake, M.; McMahon, T.; James, K.J.; Naoki, H.; Oshima, Y.; Yasumoto, T. Structures of azaspiracid analogs, azaspiracid-4 and azaspiracid-5, causative toxins of azaspiracid poisoning in Europe. Biosci. Biotechnol. Biochem. 2001, 65, 740–742. [Google Scholar] [CrossRef] [PubMed]
  121. Ofuji, K.; Satake, M.; McMahon, T.; Silke, J.; James, K.J.; Naoki, H.; Oshima, Y.; Yasumoto, T. Two analogs of azaspiracid isolated from mussels, Mytilus edulis, involved in human intoxication in Ireland. Nat. Toxins 1999, 7, 99–102. [Google Scholar] [CrossRef]
  122. Rehmann, N.; Hess, P.; Quilliam, M.A. Discovery of new analogs of the marine biotoxin azaspiracid in blue mussels [Mytilus edulis] by ultra-performance liquid chromatography/tandem mass spectrometry. Rapid Commun. Mass Spectrom. 2008, 22, 549–558. [Google Scholar] [CrossRef] [PubMed]
  123. Brombacher, S.; Edmonds, S.; Volmer, D.A. Studies on azaspiracid biotoxins. II. Mass spectral behavior and structural elucidation of azaspiracid analogs. Rapid Commun mass Spectrom. 2002, 16, 2306–2316. [Google Scholar] [CrossRef]
  124. James, K.J.; Sierra, M.D.; Lehane, M.; Magdalena, A.B.; Furey, A. Detection of five new hydroxyl analogues of azaspiracids in shellfish using multiple tandem mass spectrometry. Toxicon 2003, 41, 277–283. [Google Scholar] [CrossRef]
  125. Twiner, M.J.; Doucette, G.J.; Rasky, A.; Huang, X.-P.; Roth, B.L.; Sanguinetti, M.C. The marine algal toxin azaspiracid is an open state blocker of hERG potassium channels. Chem. Res. Toxicol. 2012, 25, 1975–1984. [Google Scholar] [CrossRef] [PubMed]
  126. Bates, S.S.; Trainer, V.L. The ecology of harmful diatoms. In Ecology of Harmful Algae; Springer: New York, NY, USA, 2006; pp. 81–93. [Google Scholar]
  127. Zaman, L.; Arakawa, O.; Shimosu, A.; Onoue, Y.; Nishio, S.; Shida, Y.; Noguchi, T. Two new isomers of domoic acid from a red alga, Chondria armata. Toxicon 1997, 35, 205–212. [Google Scholar] [CrossRef]
  128. Walter, J.A.; Falk, M.; Wright, J.L.C. Chemistry of the shellfish toxin domoic acid: Characterization of related compounds. Can. J. Chem. 1994, 72, 430–436. [Google Scholar] [CrossRef]
  129. Meda, M.; Kodama, T.; Tanaka, T.; Yoshizumi, H.; Takemoto, T.; Nomoto, K.; Fujita, T. Structures of isodomoic acids A, B and C, novel insecticidal amino acids from the red alga Chondria armata. Chem. Pharm. Bull. 1986, 34, 4892–4895. [Google Scholar] [CrossRef]
  130. EFSA CONTAM Panel [EFSA Panel on Contaminants in the Food Chain]; Alexander, J.; Benford, D.; Cockburn, A.; Cravedi, J.P.; Dogliotti, E.; Di Domenico, A.; Fernández-Cruz, M.L.; Fink-Gremmels, J.; Galli, P.F.C.; et al. Scientific opinion of the panel on contaminants in the food chain on a request from the European commission on marine biotoxins in shellfish—Saxitoxin Group. EFSA J. 2009, 1019, 1–76. [Google Scholar]
  131. Alexander, J.; Barregård, L.; Bignami, M.; Brüschweiler, B.; Ceccatelli, S.; Cottrill, B. Scientific opinion on the risks for public health related to the presence of tetrodotoxin [TTX] and TTX analogues in marine bivalves and gastropods. EFSA J. 2017, 15, 4752. [Google Scholar]
  132. Vale, P. Metabolites of saxitoxin analogues in bivalves contaminated by Gymnodinium catenatum. Toxicon 2010, 55, 162–165. [Google Scholar] [CrossRef] [PubMed]
  133. Oshima, Y. Postcolumn derivatization liquid chromatographic method for paralytic shellfish toxins. J. AOAC Int. 1995, 78, 528–532. [Google Scholar]
  134. Vale, P. Complex profiles of hydrophobic paralytic shellfish poisoning compounds in Gymnodinium catenatum identified by liquid chromatography with fluorescence detection and mass spectrometry. J. Chromatogr. A 2008, 1195, 85–93. [Google Scholar] [CrossRef] [PubMed]
  135. Negri, A.; Stirling, D.; Quilliam, M.; Blackburn, S.; Bolch, C.; Burton, I.; Eaglesham, G.; Thomas, K.; Walter, J.; Willis, R. Three novel hydroxybenzoate saxitoxin analogues isolated from the dinoflagellate Gymnodinium catenatum. Chem. Res. Toxicol. 2003, 16, 1029–1033. [Google Scholar] [CrossRef] [PubMed]
  136. Mons, M.P.; Van Egmond, H.P.; Speijers, G.J.A. Paralytic shellfish poisoning: A review. J. Am. Vet. Med. Assoc. 1978, 171, 1178–1180. [Google Scholar]
  137. Boczar, B.A.; Beitler, M.K.; Liston, J.; Sullivan, J.J.; Cattolico, R.A. Paralytic shellfish toxins in Protogonyaulax tamarensis and Protogonyaulax catenella in axenic culture. Plant Physiol. 1988, 88, 1285–1290. [Google Scholar] [CrossRef]
  138. Cheng, C.A.; Hwang, D.F.; Tsai, Y.H.; Chen, H.C.; Jeng, S.S.; Noguchi, T.; Ohwada, K.; Hasimoto, K. Microflora and tetrodotoxin-producing bacteria in a gastropod, Niotha clathrata. Food Chem. Toxicol. 1995, 33, 929–934. [Google Scholar] [CrossRef]
  139. Yu, C.-F.; Yu, P.H.-F.; Chan, P.-L.; Yan, Q.; Wong, P.-K. Two novel species of tetrodotoxin-producing bacteria isolated from toxic marine puffer fishes. Toxicon 2004, 44, 641–647. [Google Scholar] [CrossRef]
  140. Yotsu, M.; Yamazaki, T.; Meguro, Y.; Endo, A.; Murata, M.; Naoki, H.; Yasumoto, T. Production of tetrodotoxin and its derivatives by Pseudomonas sp. isolated from the skin of a pufferfish. Toxicon 1987, 25, 225–228. [Google Scholar] [CrossRef]
  141. Auawithoothij, W.; Noomhorm, A. Shewanella putrefaciens, a major microbial species related to tetrodotoxin [TTX]-accumulation of puffer fish Lagocephalus lunaris. J. Appl. Microbiol. 2012, 113, 459–465. [Google Scholar] [CrossRef] [PubMed]
  142. Hwang, D.F.; Arakawa, O.; Saito, T.; Noguchi, T.; Simidu, U.; Tsukamoto, K.; Shida, Y.; Hashimoto, K. Tetrodotoxin-producing bacteria from the blue-ringed octopus Octopus maculosus. Mar. Biol. 1989, 100, 327–332. [Google Scholar] [CrossRef]
  143. Ritchie, K.B.; Nagelkerken, I.; James, S.; Smith, G.W. Environmental microbiology: A tetrodotoxin-producing marine pathogen. Nature 2000, 404, 354. [Google Scholar] [CrossRef] [PubMed]
  144. Wu, Z.; Xie, L.; Xia, G.; Zhang, J.; Nie, Y.; Hu, J.; Wang, S.; Zhang, R. A new tetrodotoxin-producing actinomycete, Nocardiopsis dassonvillei, isolated from the ovaries of puffer fish Fugu rubripes. Toxicon 2005, 45, 851–859. [Google Scholar] [CrossRef] [PubMed]
  145. Bane, V.; Lehane, M.; Dikshit, M.; O’Riordan, A.; Furey, A. Tetrodotoxin: Chemistry, toxicity, source, distribution and detection. Toxins 2014, 6, 693–755. [Google Scholar] [CrossRef] [PubMed]
  146. Noguch, T.; Arakawa, O. Tetrodotoxin–distribution and accumulation in aquatic organisms, and cases of human intoxication. Mar. Drugs 2008, 6, 220–242. [Google Scholar] [CrossRef]
  147. Vasconcelos, V.; Azevedo, J.; Silva, M.; Ramos, V. Effects of marine toxins on the reproduction and early stages development of aquatic organisms. Mar. Drugs 2010, 8, 59–79. [Google Scholar] [CrossRef]
  148. White, J.; Meier, J. Handbook of Clinical Toxicology of Animal Venoms And Poisons; CRC Press: Boca Raton, FL, USA, 2017. [Google Scholar]
  149. Jang, J.-H.; Lee, J.-S.; Yotsu-Yamashita, M. LC/MS analysis of tetrodotoxin and its deoxy analogs in the marine puffer fish Fugu niphobles from the southern coast of Korea, and in the brackishwater puffer fishes Tetraodon nigroviridis and Tetraodon biocellatus from Southeast Asia. Mar. Drugs 2010, 8, 1049–1058. [Google Scholar] [CrossRef]
  150. Jang, J.; Yotsu-Yamashita, M. Distribution of tetrodotoxin, saxitoxin, and their analogs among tissues of the puffer fish Fugu pardalis. Toxicon 2006, 48, 980–987. [Google Scholar] [CrossRef]
  151. Kudo, Y.; Finn, J.; Fukushima, K.; Sakugawa, S.; Cho, Y.; Konoki, K.; Yotsu-Yamashita, M. Isolation of 6-deoxytetrodotoxin from the pufferfish, Takifugu pardalis, and a comparison of the effects of the C-6 and C-11 hydroxy groups of tetrodotoxin on its activity. J. Nat. Prod. 2014, 77, 1000–1004. [Google Scholar] [CrossRef]
  152. Yotsu-Yamashita, M.; Abe, Y.; Kudo, Y.; Ritson-Williams, R.; Paul, V.J.; Konoki, K.; Cho, Y.; Adachi, M.; Imazu, T.; Nishikawa, T.; et al. First identification of 5, 11-dideoxytetrodotoxin in marine animals, and characterization of major fragment ions of tetrodotoxin and its analogs by high resolution ESI-MS/MS. Mar. Drugs 2013, 11, 2799–2813. [Google Scholar] [CrossRef] [PubMed]
  153. Moore, R.E.; Bartolini, G. Structure of palytoxin. J. Am. Chem. Soc. 1981, 103, 2491–2494. [Google Scholar] [CrossRef]
  154. Ramos, V.; Vasconcelos, V. Palytoxin and analogs: Biological and ecological effects. Mar. Drugs 2010, 8, 2021–2037. [Google Scholar] [CrossRef] [PubMed]
  155. Ukena, T.; Satake, M.; Usami, M.; Oshimay, Y.; Naoki, H.; Fujita, T.; Kan, Y.; Yasumoto, T. Structure elucidation of ostreocin D, a palytoxin analog isolated from the dinoflagellate Ostreopsis siamensis. Biosci. Biotechnol. Biochem. 2001, 65, 2585–2588. [Google Scholar] [CrossRef] [PubMed]
  156. Kerbrat, A.S.; Amzil, Z.; Pawlowiez, R.; Golubic, S.; Sibat, M.; Darius, H.T.; Chinain, M.; Laurent, D. First evidence of palytoxin and 42-hydroxy-palytoxin in the marine cyanobacterium Trichodesmium. Mar. Drugs 2011, 9, 543–560. [Google Scholar] [CrossRef] [PubMed]
  157. Alexander, J.; Benford, D.; Boobis, A.; Ceccatelli, S.; Cravedi, J.P.; Di Domenico, A.; Doerge, D.; Dogliotti, E.; Edler, L.; Farmer, P.; et al. EFSA Panel on Contaminants in the Food Chain [CONTAM]; Scientific Opinion on marine biotoxins in shellfish—Palytoxin group. EFSA J. 2009, 7, 1393. [Google Scholar]
  158. Botana, L.M. Seafood and Freshwater Toxins: Pharmacology, Physiology, and Detection; CRC Press: Boca Raton, FL, USA, 2014. [Google Scholar]
  159. García-Altares, M.; Tartaglione, L.; Dell’Aversano, C.; Carnicer, O.; de la Iglesia, P.; Forino, M.; Diogène, J.; Ciminiello, P. The novel ovatoxin-g and isobaric palytoxin [so far referred to as putative palytoxin] from Ostreopsis cf. ovata [NW Mediterranean Sea]: Structural insights by LC-high resolution MSn. Anal. Bioanal. Chem. 2015, 407, 1191–1204. [Google Scholar] [CrossRef]
  160. Lenoir, S.; Ten-Hage, L.; Turquet, J.; Quod, J.P.; Hennion, M.C. Characterisation of new analogues of palytoxin isolated from an Ostreopsis mascarenensis bloom in the south-western Indian Ocean. Afr. J. Mar. Sci. 2006, 28, 389–391. [Google Scholar] [CrossRef]
  161. Habermann, E.; Chhatwal, G.S. Ouabain inhibits the increase due to palytoxin of cation permeability of erythrocytes. Naunyn-Schmiedeberg Arch. Pharmacol. 1982, 319, 101–107. [Google Scholar] [CrossRef]
  162. Miller, M.A.; Kudela, R.M.; Mekebri, A.; Crane, D.; Oates, S.C.; Tinker, M.T.; Staedler, M.; Miller, W.A.; Toy-Choutka, S.; Dominik, C.; et al. Evidence for a novel marine harmful algal bloom: Cyanotoxin [microcystin] transfer from land to sea otters. PLoS ONE 2010, 5, e12576. [Google Scholar] [CrossRef]
  163. Chorus, I.; Bartram, J. Toxic Cyanobacteria in Water: A Guide to Their Public Health Consequences, Monitoring and Management; CRC Press: Boca Raton, FL, USA, 1999. [Google Scholar]
  164. Gantar, M.; Sekar, R.; Richardson, L.L. Cyanotoxins from black band disease of corals and from other coral reef environments. Microb. Ecol. 2009, 58, 856–864. [Google Scholar] [CrossRef] [PubMed]
  165. Stanić, D.; Oehrle, S.; Gantar, M.; Richardson, L.L. Microcystin production and ecological physiology of Caribbean black band disease cyanobacteria. Environ. Microbiol. 2011, 13, 900–910. [Google Scholar] [CrossRef] [PubMed]
  166. Ramos, A.G.; Martel, A.; Codd, G.A.; Soler, E.; Coca, J.; Redondo, A.; Morrison, L.F.; Metcalf, J.S.; Ojeda, A.; Suárez, S.; et al. Bloom of the marine diazotrophic cyanobacterium Trichodesmium erythraeum in the Northwest African Upwelling. Mar. Ecol. Prog. Ser. 2005, 301, 303–305. [Google Scholar] [CrossRef] [Green Version]
  167. Vareli, K.; Zarali, E.; Zacharioudakis, G.S.A.; Vagenas, G.; Varelis, V.; Pilidis, G.; Briasoulis, E.; Sainisf, F. Microcystin producing cyanobacterial communities in Amvrakikos Gulf [Mediterranean Sea, NW Greece] and toxin accumulation in mussels [Mytilus galloprovincialis]. Harmful Algae 2012, 15, 109–118. [Google Scholar] [CrossRef]
  168. Frazão, B.; Martins, R.; Vasconcelos, V. Are known cyanotoxins involved in the toxicity of picoplanktonic and filamentous North Atlantic marine cyanobacteria? Mar. Drugs 2010, 8, 1908–1919. [Google Scholar] [CrossRef] [PubMed]
  169. Proença, L.A.O.; Tamanaha, M.S.; Fonseca, R.S. Screening the toxicity and toxin content of blooms of the cyanobacterium Trichodesmium erythraeum [Ehrenberg] in northeast Brasil. J. Venom. Anim. Toxins Incl. Trop. Dis. 2009, 15, 204–215. [Google Scholar] [CrossRef]
  170. Charpy, L.; Palinska, K.A.; Casareto, B.; Langlade, M.J.; Suzuki, Y.; Abed, R.M.M.; Golubic, S. Dinitrogen-fixing cyanobacteria in microbial mats of two shallow coral reef ecosystems. Microb. Ecol. 2010, 59, 174–186. [Google Scholar] [CrossRef]
  171. Osborne, N.J.T.; Webb, P.M.; Shaw, G.R. The toxins of Lyngbya majuscula and their human and ecological health effects. Environ. Int. 2001, 27, 381–392. [Google Scholar] [CrossRef]
  172. Nagai, H.; Yasumoto, T.; Hokama, Y. Aplysiatoxin and debromoaplysiatoxin as the causative agents of a red alga Gracilaria coronopifolia poisoning in Hawaii. Toxicon 1996, 34, 753–761. [Google Scholar] [CrossRef]
  173. Wu, M.; Okino, T.; Nogle, L.M.; Marquez, B.L.; Williamson, R.T.; Sitachitta, N.; Berman, F.W.; Murray, T.F.; McGough, K.; Jacobs, R.; et al. Structure, Synthesis, and Biological Properties of Kalkitoxin, a Novel Neurotoxin from the Marine Cyanobacterium Lyngbya majuscula. J. Am. Chem. Soc. 2000, 122, 12041–12042. [Google Scholar] [CrossRef]
  174. Fujiki, H.; Mori, M.; Nakayasu, M.; Terada, M.; Sugimura, T.; Moore, R.E. Indole alkaloids: Dihydroteleocidin B, teleocidin, and lyngbyatoxin A as members of a new class of tumor promoters. Proc. Natl. Acad. Sci. USA 1981, 78, 3872–3876. [Google Scholar] [CrossRef]
  175. Wood, S.A.; Stirling, D.J. First identification of the cylindrospermopsin-producing cyanobacterium Cylindrospermopsis raciborskii in New Zealand. N. Z. J. Mar. Freshw. Res. 2003, 37, 821–828. [Google Scholar] [CrossRef]
  176. Edwards, D.J.; Gerwick, W.H. Lyngbyatoxin biosynthesis: Sequence of biosynthetic gene cluster and identification of a novel aromatic prenyltransferase. J. Am. Chem. Soc. 2004, 126, 11432–11433. [Google Scholar] [CrossRef] [PubMed]
  177. Méjean, A.; Peyraud-Thomas, C.; Kerbrat, A.S.; Golubic, S.; Pauillac, S.; Chinain, M.; Laurent, D. First identification of the neurotoxin homoanatoxin-a from mats of Hydrocoleum lyngbyaceum [marine cyanobacterium] possibly linked to giant clam poisoning in New Caledonia. Toxicon 2010, 56, 829–835. [Google Scholar] [CrossRef] [PubMed]
  178. Roué, M.; Gugger, M.; Golubic, S.; Amzil, Z.; Araoz, R.; Turquet, J.; Chinain, M.; Laurent, D. Marine cyanotoxins potentially harmful to human health. Outst. Mar. Mol. Chem. Biol. Anal. 2014, 1–22. [Google Scholar] [CrossRef]
  179. Orjala, J.; Nagle, D.G.; Hsu, V.; Gerwick, W.H. Antillatoxin: An exceptionally ichthyotoxic cyclic lipopeptide from the tropical cyanobacterium Lyngbya majuscula. J. Am. Chem. Soc. 1995, 117, 8281–8282. [Google Scholar] [CrossRef]
  180. Fernández, M.L.; Míguez, A.; Cacho, E.; Martínez, A.; Diogéne, J.; Yasumoto, T. Bioensayos con mamíferos y ensayos bioquímicos y celulares para la detección de ficotoxinas. Floraciones algales nocivas en el Cono Sur Am. 2002, 77–120. [Google Scholar]
  181. Kat, M. Diarrhetic mussel poisoning in the Netherlands related to the dinoflagellate Dinophysis acuminata. Antonie Van Leeuwenhoek 1983, 49, 417–427. [Google Scholar]
  182. Regulation, E.C. No 854/2004 OF THE EUROPEAN PARLIAMENT AND OF THE COUNCIL of 29 April 2004 laying down specific rules for the organisation of official controls on products of animal origin intended for human consumption. Off. J. Eur. Union L. 2012, 155, 206. [Google Scholar]
  183. EFSA CONTAM Panel [EFSA Panel on Contaminants in the Food Chain]; Alexander, J.; Auðunsson, A.G.; Benford, D.C.A.; Cravedi, J.P.; Dogliotti, E.; Domenico, A.D.F.-C.M.; Fink-Gremmels, J.; Fürst, J.; Galli, C.; et al. 2017. Marine biotoxins in shellfish–okadaic acid and analogues. EFSA J. 2008, 589, 1–62. [Google Scholar]
  184. Kleivdal, H.; Kristiansen, S.-I.; Nilsen, M.V.; Goksyr, A.; Briggs, L.; Holland, P.; McNabb, P. Determination of Domoic Acid Toxins in Shellfish by Biosense ASP ELISAA Direct Competitive Enzyme-Linked Immunosorbent Assay: Collaborative Study. J. AOAC Int. 2007, 90, 1011–1027. [Google Scholar] [PubMed]
  185. Simon, J.F.; Vemoux, J. Highly sensitive assay of okadaic acid using protein phosphatase and paranitrophenyl phosphate. Nat. Toxins 1994, 2, 293–301. [Google Scholar] [CrossRef] [PubMed]
  186. Vieytes, M.R.; Fontal, O.I.; Leira, F.; de Sousa, J.M.V.B.; Botana, L.M. A fluorescent microplate assay for diarrheic shellfish toxins. Anal. Biochem. 1997, 248, 258–264. [Google Scholar] [CrossRef]
  187. Lee, J.S.; Yanagi, T.; Kenma, R.; Yasumoto, T. Fluorometric determination of diarrhetic shellfish toxins by high-performance liquid chromatography. Agric. Biol. Chem. 1987, 51, 877–881. [Google Scholar]
  188. Darius, H.T.; Ponton, D.; Revel, T.; Cruchet, P.; Ung, A.; Fouc, M.T.; Chinain, M. Ciguatera risk assessment in two toxic sites of French Polynesia using the receptor-binding assay. Toxicon 2007, 50, 612–626. [Google Scholar] [CrossRef] [PubMed]
  189. Banner, A.H.; Scheuer, P.J.; Sasaki, S.; Helfrich, P.; Alender, C.B. Observations on ciguatera-type toxin in fish. Ann. N. Y. Acad. Sci. 1960, 90, 770–787. [Google Scholar] [CrossRef] [PubMed]
  190. Lewis, R.J.; Sellin, M. Recovery of ciguatoxin from fish flesh. Toxicon 1993, 31, 1333–1336. [Google Scholar] [CrossRef]
  191. CDC [Centers for Disease Control and Prevention]. Cluster of ciguatera fish poisoning--North Carolina, 2007. Morbidity and Mortality Weekly Report [MMWR] [Internet]. North Carolina. 2009. Available online: 5811.pdf (accessed on 20 July 2018).
  192. Manger, R.L.; Leja, L.S.; Lee, S.Y.; Hungerford, J.M.; Wekell, M.M. Tetrazolium-based cell bioassay for neurotoxins active on voltage-sensitive sodium channels: Semiautomated assay for saxitoxins, brevetoxins, and ciguatoxins. Anal. Biochem. 1993, 214, 190–194. [Google Scholar] [CrossRef]
  193. Manger, R.L.; Leja, L.S.; Lee, S.Y.; Hungerford, J.M.; Wekell, M.M. Cell bioassay for the detection of ciguatoxins, brevetoxins, and saxitoxins. Mem. Queensl. Museum. Brisb. 1994, 34, 571–575. [Google Scholar]
  194. Manger, R.L.; Leja, L.S.; Lee, S.Y.; Hungerford, J.M.; Hokama, Y.; Dickey, R.W.; Granade, H.R.; Lewis, R.; Yasumoto, T.; Wekell, M.M. Detection of sodium channel toxins: Directed cytotoxicity assays of purified ciguatoxins, brevetoxins, saxitoxins, and seafood extracts. J. AOAC Int. 1995, 78, 521–527. [Google Scholar]
  195. Empey Campora, C.; Hokama, Y.; Yabusaki, K.; Isobe, M. Development of an enzyme-linked immunosorbent assay for the detection of ciguatoxin in fish tissue using chicken immunoglobulin Y. J. Clin. Lab. Anal. 2008, 22, 239–245. [Google Scholar] [CrossRef] [PubMed]
  196. Hokama, Y.; Banner, A.H.; Boylan, D.B. A radioimmunoassay for the detection of ciguatoxin. Toxicon 1977, 15, 317–325. [Google Scholar] [CrossRef]
  197. Hokama, Y.; Honda, S.A.A.; Uyehara, K.; Shirai, L.K.; Kobayashi, M.N. Monoclonal-antibodies to low dalton natural marine toxins. J. Toxicol. Rev. 1986, 5, 194. [Google Scholar]
  198. Hokama, Y.; Kimura, L.H.; Abad, M.A.; Yokochi, L.; Scheuer, P.J.; Nukina, M.; Yasumoto, T.; Baden, D.G.; Shimizu, Y. An Enzyme Immunoassay for the Detection of Ciguatoxin: And Competitive Inhibition by Related Natural Polyether Toxins; ACS Publications: Washington, DC, USA, 1984; pp. 1947–5918. [Google Scholar]
  199. Hokama, Y.; Shirai, L.K.; Iwamoto, L.M.; Kobayashi, M.N.; Goto, C.S.; Nakagawa, L.K. Assessment of a rapid enzyme immunoassay stick test for the detection of ciguatoxin and related polyether toxins in fish tissues. Biol. Bull. 1987, 172, 144–153. [Google Scholar] [CrossRef]
  200. Lewis, R.J.; Jones, A.; Vernoux, J.-P. HPLC/tandem electrospray mass spectrometry for the determination of sub-ppb levels of Pacific and Caribbean ciguatoxins in crude extracts of fish. Anal. Chem. 1999, 71, 247–250. [Google Scholar] [CrossRef] [PubMed]
  201. Dickey, R.W.; Bencsath, F.A.; Granade, H.R.; Lewis, R.J. Liquid chromatographic mass spectrometric methods for the determination of marine polyether toxins. Bull. Soc. Pathol. Exot. 1992, 85 Pt 2, 514–515. [Google Scholar]
  202. Yasumoto, T.; Fukui, M.; Sasaki, K.; Sugiyama, K. Determinations of marine toxins in foods. J. AOAC Int. 1995, 78, 574–582. [Google Scholar]
  203. Vilariño, N.; Fonfría, E.S.; Molgó, J.; Aráoz, R.; Botana, L.M. Detection of gymnodimine-A and 13-desmethyl C spirolide phycotoxins by fluorescence polarization. Anal. Chem. 2009, 81, 2708–2714. [Google Scholar] [CrossRef]
  204. Ciminiello, P.; Dell’Aversano, C.; Fattorusso, E.; Forino, M.; Magno, G.S.; Tartaglione, L.; Grillo, C.; Melchiorre, N. The Genoa 2005 Outbreak. Determination of Putative Palytoxin in Mediterranean Ostreopsis o vata by a New Liquid Chromatography Tandem Mass Spectrometry Method. Anal. Chem. 2006, 78, 6153–6159. [Google Scholar] [CrossRef]
  205. Marrouchi, R.; Dziri, F.; Belayouni, N.; Hamza, A.; Benoit, E.; Molgó, J.; Kharrat, R. Quantitative determination of gymnodimine-A by high performance liquid chromatography in contaminated clams from Tunisia coastline. Mar. Biotechnol. 2010, 12, 579–585. [Google Scholar] [CrossRef]
  206. Association, A.P.H. Recommended procedures for the examination of sea water and shellfish. In Recommended Procedures for the Examination of Sea Water and Shellfish; APHA: Cincinnati, OH, USA, 1970. [Google Scholar]
  207. Briggs, L.R.; Garthwaite, L.L.; Miles, C.O.; Garthwaite, I.; Ross, K.M.; Towers, N.R. The newest ELISA—Pectenotoxin. In Marine Biotoxin Science Workshop; Marine Institute: Galway, Ireland, 2000; pp. 71–75. [Google Scholar]
  208. Naar, J.; Bourdelais, A.; Tomas, C.; Kubanek, J.; Whitney, P.L.; Flewelling, L.; Steidinger, K.; Lancaster, J.; Baden, D.G. A competitive ELISA to detect brevetoxins from Karenia brevis [formerly Gymnodinium breve] in seawater, shellfish, and mammalian body fluid. Environ. Health Perspect. 2002, 110, 179–185. [Google Scholar] [CrossRef] [PubMed]
  209. Wang, W.; Cole, R.B. Enhanced collision-induced decomposition efficiency and unraveling of fragmentation pathways for anionic adducts of brevetoxins in negative ion electrospray mass spectrometry. Anal. Chem. 2009, 81, 8826–8838. [Google Scholar] [CrossRef] [PubMed]
  210. Regulation, E.U. 853/2004. Regulation [EC] no. 853/2004 of the European Parliament and of the Council of 29 April 2004. Laying down specific hygiene rules for food of animal origin. Off. J. Eur. Union 2004, 226, 22–82. [Google Scholar]
  211. McNabb, P.; Selwood, A.I.; Holland, P.T. Multiresidue method for determination of algal toxins in shellfish: Single-laboratory validation and interlaboratory study. J. AOAC Int. 2005, 88, 761–772. [Google Scholar] [PubMed]
  212. Stobo, L.A.; Lacaze, J.-P.C.L.; Scott, A.C.; Gallacher, S.; Smith, E.A.; Quilliam, M.A. Liquid chromatography with mass spectrometry—detection of lipophilic shellfish toxins. J. AOAC Int. 2005, 88, 1371–1382. [Google Scholar] [PubMed]
  213. Briggs, L.R.; Miles, C.O.; Fitzgerald, J.M.; Ross, K.M.; Garthwaite, I.; Towers, N.R. Enzyme-linked immunosorbent assay for the detection of yessotoxin and its analogues. J. Agric. Food Chem. 2004, 52, 5836–5842. [Google Scholar] [CrossRef]
  214. Satake, M.; Ofuji, K.; James, K.J.; Furey, A.; Yasumoto, T. New toxic event caused by Irish mussels. Harmful Algae 1998, 468–469. [Google Scholar]
  215. [EFSA] EFSA. Marine biotoxins in shellfish–saxitoxin group. EFSA J. 2009, 7, 1019. [Google Scholar] [CrossRef]
  216. Krock, B.; Pitcher, G.C.; Ntuli, J.; Cembella, A.D. Confirmed identification of gymnodimine in oysters from the west coast of South Africa by liquid chromatography–tandem mass spectrometry. Afr. J. Mar. Sci. 2009, 31, 113–118. [Google Scholar] [CrossRef]
  217. Sommer, H.; Meyer, K.F. Paralytic Shell-Fish Poisoning. Arch. Pathol. 1937, 24, 560–598. [Google Scholar]
  218. Catterall, W.A.; Morrow, C.S. Binding to saxitoxin to electrically excitable neuroblastoma cells. Proc. Natl. Acad. Sci. USA 1978, 75, 218–222. [Google Scholar] [CrossRef] [PubMed]
  219. Jellett, J.F.; Marks, L.J.; Stewart, J.E.; Dorey, M.L.; Watson-Wright, W.; Lawrence, J.F. Paralytic shellfish poison [saxitoxin family] bioassays: Automated endpoint determination and standardization of the in vitro tissue culture bioassay, and comparison with the standard mouse bioassay. Toxicon 1992, 30, 1143–1156. [Google Scholar] [CrossRef]
  220. Campbell, K.; Stewart, L.D.; Doucette, G.J.; Fodey, T.L.; Haughey, S.A.; Vilariño, N.; Kawatsu, K.; Elliott, C.T. Assessment of specific binding proteins suitable for the detection of paralytic shellfish poisons using optical biosensor technology. Anal. Chem. 2007, 79, 5906–5914. [Google Scholar] [CrossRef] [PubMed]
  221. Carlson, R.E.; Lever, M.L.; Lee, B.W.; Guire, P.E. Development of Immunoassays for Paralytic Shellfish Poisoning: A Radioimmunoassay for Saxitoxin; ACS Publications: New York, NY, USA, 1984. [Google Scholar]
  222. Fonfría, E.S.; Vilariño, N.; Campbell, K.; Elliott, C.; Haughey, S.A.; Ben-Gigirey, B.; Vieites, J.M.; Kawatsu, K.; Botana, L.M. Paralytic shellfish poisoning detection by surface plasmon resonance-based biosensors in shellfish matrixes. Anal. Chem. 2007, 79, 6303–6311. [Google Scholar] [CrossRef] [PubMed]
  223. Jellett, J.F.; Roberts, R.L.; Laycock, M.V.; Quilliam, M.A.; Barrett, R.E. Detection of paralytic shellfish poisoning [PSP] toxins in shellfish tissue using MIST AlertTM, a new rapid test, in parallel with the regulatory AOAC® mouse bioassay. Toxicon 2002, 40, 1407–1425. [Google Scholar] [CrossRef]
  224. Usleber, E.; Schneider, E.; Terplan, G.; Laycock, M.V. Two formats of enzyme immunoassay for the detection of saxitoxin and other paralytic shellfish poisoning toxins. Food Addit. Contam. 1995, 12, 405–413. [Google Scholar] [CrossRef] [PubMed]
  225. Thibault, P.; Pleasance, S.; Laycock, M.V. Analysis of paralytic shellfish poisons by capillary electrophoresis. J. Chromatogr. A 1991, 542, 483–501. [Google Scholar] [CrossRef]
  226. Dell’Aversano, C.; Hess, P.; Quilliam, M.A. Hydrophilic interaction liquid chromatography–mass spectrometry for the analysis of paralytic shellfish poisoning [PSP] toxins. J. Chromatogr. A 2005, 1081, 190–201. [Google Scholar] [CrossRef]
  227. Franco, J.M.; Fernández-Vila, P. Separation of paralytic shellfish toxins by reversed phase high performance liquid chromatography, with postcolumn reaction and fluorimetric detection. Chromatographia 1993, 35, 613–620. [Google Scholar] [CrossRef]
  228. Lawrence, J.F.; Menard, C. Liquid chromatographic determination of paralytic shellfish poisons in shellfish after prechromatographic oxidation. J. Assoc. Off. Anal. Chem. 1991, 74, 1006–1012. [Google Scholar]
  229. Silva, M.; Rey, V.; Botana, A.; Vasconcelos, V.; Botana, L. Determination of Gonyautoxin-4 in Echinoderms and Gastropod Matrices by Conversion to Neosaxitoxin Using 2-Mercaptoethanol and Post-Column Oxidation Liquid Chromatography with Fluorescence Detection. Toxins 2015, 8, 11. [Google Scholar] [CrossRef] [PubMed]
  230. Panel, E.C. Scienti fi c opinion on marine biotoxins in shell fi sh-domoic acid. EFSA panel Contam food Chain [CONTAM]. EFSA J. 2009, 1181, 1–61. [Google Scholar]
  231. Garthwaite, I.; Ross, K.M.; Miles, C.O.; Hansen, R.P.; Foster, D.; Wilkins, A.L.; Wilkins, A.L.; Towers, N.R. Polyclonal antibodies to domoic acid, and their use in immunoassays for domoic acid in sea water and shellfish. Nat. Toxins 1998, 6, 93–104. [Google Scholar] [CrossRef]
  232. Traynor, I.M.; Plumpton, L.; Fodey, T.L.; Higgins, C.; Elliott, C.T. Immunobiosensor detection of domoic acid as a screening test in bivalve molluscs: Comparison with liquid chromatography-based analysis. J. AOAC Int. 2006, 89, 868–872. [Google Scholar] [PubMed]
  233. Quilliam, M.A.; Xie, M.; Hardstaff, W.R. Rapid extraction and cleanup for liquid chromatographic determination of domoic acid in unsalted seafood. J. AOAC Int. 1995, 78, 543–554. [Google Scholar]
  234. Pocklington, R.; Milley, J.E.; Bates, S.S.; Bird, C.J.; De Freitas, A.S.W.; Quilliam, M.A. Trace determination of domoic acid in sea water and phytoplankton by high-performance liquid chromatography of the fluorenylmethoxycarbonyl [FMOC] derivative. Int. J. Environ. Anal. Chem. 1990, 38, 351–368. [Google Scholar] [CrossRef]
  235. Van Dolah, F.M.; Leighfield, T.A.; Haynes, B.L.; Hampson, D.R.; Ramsdell, J.S. A microplate receptor assay for the amnesic shellfish poisoning toxin, domoic acid, utilizing a cloned glutamate receptor. Anal. Biochem. 1997, 245, 102–105. [Google Scholar] [CrossRef]
  236. Zhao, J.; Thibault, P.; Quilliam, M.A. Analysis of domoic acid isomers in seafood by capillary electrophoresis. Electrophoresis 1997, 18, 268–276. [Google Scholar] [CrossRef]
  237. Pineiro, N.; Leao, J.M.; Martınez, A.G.; Vázquez, J.A.R. Capillary electrophoresis with diode array detection as an alternative analytical method for paralytic and amnesic shellfish toxins. J. Chromatogr. A 1999, 847, 223–232. [Google Scholar] [CrossRef]
  238. Nguyen, A.-L.; Luong, J.H.T.; Masson, C. Capillary electrophoresis for detection and quantitation of domoic acid in mussels. Anal. Lett. 1990, 23, 1621–1634. [Google Scholar] [CrossRef]
  239. Wright, J.L.C.; Boyd, R.K.; de Freitas, A.S.W.; Falk, M.; Foxall, R.A.; Jamieson, W.D.; Laycock, M.V.; McCulloch, A.W.; McInnes, A.G.; Odense, P.; et al. Identification of domoic acid, a neuroexcitatory amino acid, in toxic mussels from eastern Prince Edward Island. Can. J. Chem. 1989, 67, 481–490. [Google Scholar] [CrossRef] [Green Version]
  240. Pardo, O.; Yusà, V.; León, N.; Pastor, A. Development of a pressurised liquid extraction and liquid chromatography with electrospray ionization-tandem mass spectrometry method for the determination of domoic acid in shellfish. J. Chromatogr. A 2007, 1154, 287–294. [Google Scholar] [CrossRef]
  241. Lawrence, J.F.; Charbonneau, C.F.; Ménard, C. Liquid chromatographic determination of domoic acid in mussels, using AOAC paralytic shellfish poison extraction procedure: Collaborative study. J. Assoc. Off. Anal. Chem. 1991, 74, 68–72. [Google Scholar] [PubMed]
  242. Quilliam, M.A.; Sim, P.G.; McCulloch, A.W.; McInnes, A.G. High-performance liquid chromatography of domoic acid, a marine neurotoxin, with application to shellfish and plankton. Int. J. Environ. Anal. Chem. 1989, 36, 139–154. [Google Scholar] [CrossRef]
  243. Quilliam, M.A.; Thomas, K.; Wright, J.L.C. Analysis of domoic acid in shellfish by thin-layer chromatography. Nat. Toxins 1998, 6, 147–152. [Google Scholar] [CrossRef]
  244. Noguchi, T.; Ebesu, J.S.M. Puffer poisoning: Epidemiology and treatment. J. Toxicol. Toxin Rev. 2001, 20, 1–10. [Google Scholar] [CrossRef]
  245. Yang, G.; Xu, J.; Liang, S.; Ren, D.; Yan, X.; Bao, B. A novel TTX-producing Aeromonas isolated from the ovary of Takifugu obscurus. Toxicon 2010, 56, 324–329. [Google Scholar] [CrossRef]
  246. Chulanetra, M.; Sookrung, N.; Srimanote, P.; Indrawattana, N.; Thanongsaksrikul, J.; Sakolvaree, Y.; Chongsa-Nguan, M.; Kurazono, H.; Chaicumpa, W. Toxic marine puffer fish in Thailand seas and tetrodotoxin they contained. Toxins 2011, 3, 1249–1262. [Google Scholar] [CrossRef]
  247. Katikou, P.; Georgantelis, D.; Sinouris, N.; Petsi, A.; Fotaras, T. First report on toxicity assessment of the Lessepsian migrant pufferfish Lagocephalus sceleratus [Gmelin, 1789] from European waters [Aegean Sea, Greece]. Toxicon 2009, 54, 50–55. [Google Scholar] [CrossRef]
  248. Hungerford, J.M. Committee on natural toxins and food allergens: Marine and freshwater toxins. J. AOAC Int. 2006, 89, 248–269. [Google Scholar]
  249. Doucette, G.J.; Powell, C.L.; Do, E.U.; Byon, C.Y.; Cleves, F.; McClain, S.G. Evaluation of 11-[3H]-tetrodotoxin use in a heterologous receptor binding assay for PSP toxins. Toxicon 2000, 38, 1465–1474. [Google Scholar] [CrossRef]
  250. Mahmud, Y.; Arakawa, O.; Ichinose, A.; Tanu, M.B.; Takatani, T.; Tsuruda, K.; Kawatsu, K.; Hamano, Y.; Noguchi, T. Intracellular visualization of tetrodotoxin [TTX] in the skin of a puffer Tetraodon nigroviridis by immunoenzymatic technique. Toxicon 2003, 41, 605–611. [Google Scholar] [CrossRef]
  251. Mahmud, Y.; Okada, K.; Takatani, T.; Kawatsu, K.; Hamano, Y.; Arakawa, O.; Noguchi, T. Intra-tissue distribution of tetrodotoxin in two marine puffers Takifugu vermicularis and Chelonodon patoca. Toxicon 2003, 41, 13–18. [Google Scholar] [CrossRef]
  252. Tsuruda, K.; Arakawa, O.; Kawatsu, K.; Hamano, Y.; Takatani, T.; Noguchi, T. Secretory glands of tetrodotoxin in the skin of the Japanese newt Cynops pyrrhogaster. Toxicon 2002, 40, 131–136. [Google Scholar] [CrossRef]
  253. Brillantes, S.; Samosorn, W.; Faknoi, S.; Oshima, Y. Toxicity of puffers landed and marketed in Thailand. Fish. Sci. 2003, 69, 1224–1230. [Google Scholar] [CrossRef]
  254. Bignami, G.S.; Raybould, T.J.G.; Sachinvala, N.D.; Grothaus, P.G.; Simpson, S.B.; Lazo, C.B.; Byrnes, J.B.; Moore, R.E.; Vann, D.C. Monoclonal antibody-based enzyme-linked immunoassays for the measurement of palytoxin in biological samples. Toxicon 1992, 30, 687–700. [Google Scholar] [CrossRef]
  255. Kawatsu, K.; Shibata, T.; Hamano, Y. Application of immunoaffinity chromatography for detection of tetrodotoxin from urine samples of poisoned patients. Toxicon 1999, 37, 325–333. [Google Scholar] [CrossRef]
  256. Tanu, M.B.; Mahmud, Y.; Takatani, T.; Kawatsu, K.; Hamano, Y.; Arakawa, O.; Noguchi, T. Localization of tetrodotoxin in the skin of a brackishwater puffer Tetraodon steindachneri on the basis of immunohistological study. Toxicon 2002, 40, 103–106. [Google Scholar] [CrossRef]
  257. Nagashima, Y.; Nishio, S.; Noguchi, T.; Arakawa, O.; Kanoh, S.; Hashimoto, K. Detection of tetrodotoxin by thin-layer chromatography/fast atom bombardment mass spectrometry. Anal. Biochem. 1988, 175, 258–262. [Google Scholar] [CrossRef]
  258. Man, C.N.; Noor, N.M.; Harn, G.L.; Lajis, R.; Mohamad, S. Screening of tetrodotoxin in puffers using gas chromatography–mass spectrometry. J. Chromatogr. A 2010, 1217, 7455–7459. [Google Scholar] [CrossRef] [Green Version]
  259. Shiu, Y.-C.; Lu, Y.-H.; Tsai, Y.-H.; Chen, S.-K.; Hwang, D.-F. Occurrence of tetrodotoxin in the causative gastropod Polinices didyma and another gastropod Natica lineata collected from western Taiwan. J. Food Drug Anal. 2003, 11, 159–163. [Google Scholar]
  260. Chen, X.-W.; Liu, H.-X.; Jin, Y.-B.; Li, S.-F.; Bi, X.; Chung, S.; Zhang, S.S.; Jiang, Y.Y. Separation, identification and quantification of tetrodotoxin and its analogs by LC-MS without calibration of individual analogs. Toxicon 2011, 57, 938–943. [Google Scholar] [CrossRef]
  261. Diener, M.; Christian, B.; Ahmed, M.S.; Luckas, B. Determination of tetrodotoxin and its analogs in the puffer fish Takifugu oblongus from Bangladesh by hydrophilic interaction chromatography and mass-spectrometric detection. Anal. Bioanal. Chem. 2007, 389, 1997–2002. [Google Scholar] [CrossRef]
  262. Nzoughet, J.K.; Campbell, K.; Barnes, P.; Cooper, K.M.; Chevallier, O.P.; Elliott, C.T. Comparison of sample preparation methods, validation of an UPLC-MS/MS procedure for the quantification of tetrodotoxin present in marine gastropods and analysis of pufferfish. Food Chem. 2013, 136, 1584–1589. [Google Scholar] [CrossRef] [PubMed]
  263. Rodríguez, P.; Alfonso, A.; Otero, P.; Katikou, P.; Georgantelis, D.; Botana, L.M. Liquid chromatography–mass spectrometry method to detect Tetrodotoxin and Its analogues in the puffer fish Lagocephalus sceleratus [Gmelin, 1789] from European waters. Food Chem. 2012, 132, 1103–1111. [Google Scholar] [CrossRef]
  264. Silva, M.; Azevedo, J.; Rodriguez, P.; Alfonso, A.; Botana, L.M.; Vasconcelos, V. New gastropod vectors and tetrodotoxin potential expansion in temperate waters of the Atlantic Ocean. Mar. Drugs 2012, 10, 712–726. [Google Scholar] [CrossRef] [PubMed]
  265. Yotsu-Yamashita, M.; Mebs, D.; Kwet, A.; Schneider, M. Tetrodotoxin and its analogue 6-epitetrodotoxin in newts [Triturus spp.; Urodela, Salamandridae] from southern Germany. Toxicon 2007, 50, 306–309. [Google Scholar] [CrossRef] [PubMed]
  266. Hirata, Y.; Uemura, D.; Ohizumi, Y. Chemistry and pharmacology of palytoxin. In Handbook of Natural ToxinsVolume 3. Marine Toxins and Venoms; Tu, A.T., Ed.; Marcel Dekker, Inc.: New York, NY, USA; Basel, Switzerland, 1988. [Google Scholar]
  267. Gleibs, S.; Mebs, D. Distribution and sequestration of palytoxin in coral reef animals. Toxicon 1999, 37, 1521–1527. [Google Scholar] [CrossRef]
  268. Wiles, J.S.; Vick, J.A.; Christensen, M.K. Toxicological evaluation of palytoxin in several animal species. Toxicon 1974, 12, 427–433. [Google Scholar] [CrossRef]
  269. CRLMB [Community Reference Laboratory for Marine Biotoxins]. Minutes of the 1st Meeting of Working Group on Toxicology of the National Reference Laboratories (NRLs) for Marine Biotoxins; CRLMB: Cesenatico, Italy, 2005. [Google Scholar]
  270. Aligizaki, K.; Katikou, P.; Nikolaidis, G.; Panou, A. First episode of shellfish contamination by palytoxin-like compounds from Ostreopsis species [Aegean Sea, Greece]. Toxicon 2008, 51, 418–427. [Google Scholar] [CrossRef] [PubMed]
  271. Bellocci, M.; Ronzitti, G.; Milandri, A.; Melchiorre, N.; Grillo, C.; Poletti, R.; Yasumoto, T.; Rossini, G.P. A cytolytic assay for the measurement of palytoxin based on a cultured monolayer cell line. Anal. Biochem. 2008, 374, 48–55. [Google Scholar] [CrossRef] [PubMed]
  272. Riobó, P.; Paz, B.; Franco, J.M. Analysis of palytoxin-like in Ostreopsis cultures by liquid chromatography with precolumn derivatization and fluorescence detection. Anal. Chim. Acta 2006, 566, 217–223. [Google Scholar] [CrossRef]
  273. Azevedo, S.M.F.O.; Carmichael, W.W.; Jochimsen, E.M.; Rinehart, K.L.; Lau, S.; Shaw, G.R.; Eaglesham, G.K. Human intoxication by microcystins during renal dialysis treatment in Caruaru—Brazil. Toxicology 2002, 181, 441–446. [Google Scholar] [CrossRef]
  274. Kankaanpää, H.; Leiniö, S.; Olin, M.; Sjövall, O.; Meriluoto, J.; Lehtonen, K.K. Accumulation and depuration of cyanobacterial toxin nodularin and biomarker responses in the mussel Mytilus edulis. Chemosphere 2007, 68, 1210–1217. [Google Scholar] [CrossRef] [PubMed]
  275. Sipiä, V.O.; Lahti, K.; Kankaanpää, H.T.; Vuorinen, P.J.; Meriluoto, J.A.O. Screening for cyanobacterial hepatotoxins in herring and salmon from the Baltic Sea. Aquat. Ecosyst. Health Manag. 2002, 5, 451–456. [Google Scholar] [CrossRef]
  276. Zimba, P.V.; Camus, A.; Allen, E.H.; Burkholder, J.M. Co-occurrence of white shrimp, Litopenaeus vannamei, mortalities and microcystin toxin in a southeastern USA shrimp facility. Aquaculture 2006, 261, 1048–1055. [Google Scholar] [CrossRef]
  277. Williams, D.E.; Craig, M.; Dawe, S.C.; Kent, M.L.; Holmes, C.F.B.; Andersen, R.J. Evidence for a covalently bound form of microcystin-LR in salmon liver and dungeness crab larvae. Chem. Res. Toxicol. 1997, 10, 463–469. [Google Scholar] [CrossRef]
  278. Organization, W.H. Guidelines for Safe Recreational Water Environments: Coastal and Fresh Waters; World Health Organization, 2003; Volume 1. Available online: (accessed on 10 November 2018).
  279. Cook, W.O.; Iwamoto, G.A.; Schaeffer, D.J.; Carmichael, W.W.; Beasley, V.R. Pathophysiologic Effects of Anatoxin-a [s] in Anaesthetized Rats: The Influence of Atropine and Artificial Respiration. Pharmacol. Toxicol. 1990, 67, 151–155. [Google Scholar] [CrossRef]
  280. Patockaa, J.; Stredab, L. Brief review of natural nonprotein neurotoxins. ASA Newslett. 2002, 89, 16–24. [Google Scholar]
  281. Aráoz, R.; Nghiêm, H.-O.; Rippka, R.; Palibroda, N.; de Marsac, N.T.; Herdman, M. Neurotoxins in axenic oscillatorian cyanobacteria: Coexistence of anatoxin-a and homoanatoxin-a determined by ligand-binding assay and GC/MS. Microbiology 2005, 151, 1263–1273. [Google Scholar] [CrossRef]
  282. Aráoz, R.; Vilariño, N.; Botana, L.M.; Molgó, J. Ligand-binding assays for cyanobacterial neurotoxins targeting cholinergic receptors. Anal. Bioanal. Chem. 2010, 397, 1695–1704. [Google Scholar] [CrossRef] [PubMed]
  283. Serdula, M.; Bartolini, G.; Moore, R.E.; Gooch, J.; Wiebenga, N. Seaweed itch on windward Oahu. Hawaii Med. J. 1982, 41, 200–201. [Google Scholar] [PubMed]
  284. Carmichael, W.W. Health effects of toxin-producing cyanobacteria:“The CyanoHABs”. Hum. Ecol. Risk Assess. 2001, 7, 1393–1407. [Google Scholar] [CrossRef]
  285. Ito, E.; Nagai, H. Morphological observations of diarrhea in mice caused by aplysiatoxin, the causative agent of the red alga Gracilaria coronopifolia poisoning in Hawaii. Toxicon 1998, 36, 1913–1920. [Google Scholar] [CrossRef]
  286. Capper, A.; Tibbetts, I.R.; O’Neil, J.M.; Shaw, G.R. The fate of Lyngbya majuscula toxins in three potential consumers. J. Chem. Ecol. 2005, 31, 1595–1606. [Google Scholar] [CrossRef]
  287. Nogle, L.M.; Okino, T.; Gerwick, W.H. Antillatoxin B, a Neurotoxic Lipopeptide from the Marine Cyanobacterium Lyngbya majuscula. J. Nat. Prod. 2001, 64, 983–985. [Google Scholar] [CrossRef] [PubMed]
  288. Edwards, D.J.; Marquez, B.L.; Nogle, L.M.; McPhail, K.; Goeger, D.E.; Roberts, M.A.; Gerwick, W.H. Structure and biosynthesis of the jamaicamides, new mixed polyketide-peptide neurotoxins from the marine cyanobacterium Lyngbya majuscula. Chem. Biol. 2004, 11, 817–833. [Google Scholar] [CrossRef]
  289. Griffiths, D.J.; Saker, M.L. The Palm Island mystery disease 20 years on: A review of research on the cyanotoxin cylindrospermopsin. Environ. Toxicol. 2003, 18, 78–93. [Google Scholar] [CrossRef]
  290. Carmichael, W.W.; Azevedo, S.M.; An, J.S.; Molica, R.J.; Jochimsen, E.M.; Lau, S.; Rinehart, K.L.; Shaw, G.R.; Eaglesham, G.K. Human fatalities from cyanobacteria: Chemical and biological evidence for cyanotoxins. Environ. Health Perspect. 2001, 109, 663–668. [Google Scholar] [CrossRef]
  291. Hawkins, P.R.; Chandrasena, N.R.; Jones, G.J.; Humpage, A.R.; Falconer, I.R. Isolation and toxicity of Cylindrospermopsis raciborskii from an ornamental lake. Toxicon 1997, 35, 341–346. [Google Scholar] [CrossRef]
  292. Eaglesham, G.K.; Norris, R.L.; Shaw, G.R.; Smith, M.J.; Chiswell, R.K.; Davis, B.C.; Neville, G.R.; Seawright, A.A.; Moore, M.R. Use of HPLC-MS/MS to monitor cylindrospermopsin, a blue–green algal toxin, for public health purposes. Environ. Toxicol. 1999, 14, 151–154. [Google Scholar] [CrossRef]
  293. Carson, B.; Masten, S. Cylindrospermopsin–Review of Toxicological Literature. Natl Inst Environ Heal Sci Res Triangle Park NC. 2000. Available online: (accessed on 10 November 2018).
  294. Blahova, L.; Oravec, M.; Maršálek, B.; Šejnohova, L.; Šimek, Z.; Bláha, L. The first occurrence of the cyanobacterial alkaloid toxin cylindrospermopsin in the Czech Republic as determined by immunochemical and LC/MS methods. Toxicon 2009, 53, 519–524. [Google Scholar] [CrossRef] [PubMed]
  295. Dale, B. Marine dinoflagellate cysts as indicators of eutrophication and industrial pollution: A discussion. Sci. Total Environ. 2001, 264, 235–240. [Google Scholar] [CrossRef]
  296. Hallegraeff, G.M. Harmful algal blooms: A global overview. Man Harmful Mar. Microalgae 2003, 33, 1–22. [Google Scholar]
  297. Bragadeeswaran, S.; Therasa, D.; Prabhu, K.; Kathiresan, K. Biomedical and pharmacological potential of tetrodotoxin-producing bacteria isolated from marine pufferfish Arothron hispidus [Muller, 1841]. J. Venom. Anim. Toxins Incl. Trop. Dis. 2010, 16, 421–431. [Google Scholar] [CrossRef]
  298. Bubb, H.D. Vibrio parahaemolyticus--a marine pathogen detected in South African coastal waters. S. Afr. Med. J. 1975, 49, 1514–1516. [Google Scholar] [PubMed]
  299. Abd-Elghany, S.M.; Sallam, K.I. Occurrence and molecular identification of Vibrio parahaemolyticus in retail shellfish in Mansoura, Egypt. Food Control 2013, 33, 399–405. [Google Scholar] [CrossRef]
  300. Jean Turquet, J.-P.Q.; Ten-Hage, L.; Dahalani, Y.; Wendling, B. Example of a Gambierdiscus toxicus flare-up following the 1998 coral bleaching event in Mayotte Island [Comoros, south-west Indian Ocean]. In Proceedings of the 9th International Conference on Harmful Algae, Hobart, Tasmania, 7–11 February 2000. [Google Scholar]
  301. Kiteresi, L.; Mwangi, S.; Mary, M. Potentially Harmful Algae along the Kenyan Coast: A Norm or Threat. Harmful Algae 2013, 3. [Google Scholar]
  302. Silva, S.M.F.; Pienaar, R.N. Marine Cyanophytes from the Western Cape, South Africa: Chroococcales. S. Afr. J. Bot. 1999, 65, 32–49. [Google Scholar] [CrossRef] [Green Version]
  303. Shibl, A.A.; Thompson, L.R.; Ngugi, D.K.; Stingl, U. Distribution and diversity of Prochlorococcus ecotypes in the Red Sea. FEMS Microbiol. Lett. 2014, 356, 118–126. [Google Scholar] [CrossRef]
  304. Zubia, M.; Turquet, J.; Golubic, S. Benthic cyanobacterial diversity of iles eparses [Scattered islands] in the Mozambique channel. Acta Oecol. 2016, 72, 21–32. [Google Scholar] [CrossRef]
  305. Van der Molen, J.S.; Scharler, U.M.; Muir, D. Species composition, abundance and biomass of microphytoplankton in the KwaZulu-Natal Bight on the east coast of South Africa. Afr. J. Mar. Sci. 2016, 38, S139–S153. [Google Scholar] [CrossRef]
  306. Grindley, J.R.; Taylor, F.J.R.; Day, J.H. Red water and marine fauna mortality near Cape Town. Trans. R. Soc South Africa. 1964, 37, 111–130. [Google Scholar] [CrossRef]
  307. Silva, S.M.F.; Pienaar, R.N. Epipelic marine Cyanophytes of Bazaruto Island, lnhambane, Mozambique. S. Afr. J. Bot. 1997, 6, 459–464. [Google Scholar] [CrossRef]
  308. Sadally, S.B.; Taleb-Hossenkhan, N.; Bhagooli, R. Spatio-temporal variation in density of microphytoplankton genera in two tropical coral reefs of Mauritius. Afr. J. Mar. Sci. 2014, 36, 423–438. [Google Scholar] [CrossRef]
  309. Pitcher, G.C.; Cembella, A.D.; Joyce, L.B.; Larsen, J.; Probyn, T.A.; Sebastián, C.R. The dinoflagellate Alexandrium minutum in Cape Town harbour [South Africa]: Bloom characteristics, phylogenetic analysis and toxin composition. Harmful Algae 2007, 6, 823–836. [Google Scholar] [CrossRef]
  310. Ochieng, O.B.; Khakasa, M.K.; Sturcky, O.P. Harmful marine phytoplankton community in Shirazi Creek, Kenya. J. Fish. Aquat. Sci. 2015, 10, 266–275. [Google Scholar] [CrossRef]
  311. Nassar, M.Z.; El-Din, N.G.S.; Gharib, S.M. Phytoplankton variability in relation to some environmental factors in the eastern coast of Suez Gulf, Egypt. Environ. Monit. Assess. 2015, 187, 648. [Google Scholar] [CrossRef]
  312. Olofsson, M.; Karlberg, M.; Lage, S.; Ploug, H. Phytoplankton community composition and primary production in the tropical tidal ecosystem, Maputo Bay [the Indian Ocean]. J. Sea Res. 2017, 125, 18–25. [Google Scholar] [CrossRef]
  313. Sá, C.; Leal, M.C.; Silva, A.; Nordez, S.; André, E.; Paula, J.; Brotas, V. Variation of phytoplankton assemblages along the Mozambique coast as revealed by HPLC and microscopy. J. Sea Res. 2013, 79, 1–11. [Google Scholar] [CrossRef]
  314. Riaux-Gobin, C.; Compère, P. Olifantiella mascarenica gen. & sp. nov., a new genus of pennate diatom from Réunion Island, exhibiting a remarkable internal process. Phycol. Res. 2009, 57, 178–185. [Google Scholar]
  315. Quod, J.P.; Turquet, J.; Diogene, G.; Fessard, V. Screening of extracts of dinoflagellates from coral reefs [Reunion Island, SW Indian Ocean], and their biological activities. Harmful Mar. Algal Bloom. 1995, 815–820. [Google Scholar]
  316. El Semary, N. Benthic dinoflagellates from Red Sea, Egypt: Early records. Egypt. J. Aquat. Res. 2016, 42, 177–184. [Google Scholar] [CrossRef] [Green Version]
  317. Berland, B.; Grzebyk, D.; Thomassin, B.-A. Benthic dinoflagellates from the coral reef lagoon of Mayotte Island [SW Indian Ocean]; identification, toxicity and preliminary ecophysiological study. Bull. Pathol. Exot. 1992, 85, 453–456. [Google Scholar]
  318. Burckle, L.H. Distribution of diatoms in sediments of the northern Indian Ocean: Relationship to physical oceanography. Mar. Micropaleontol. 1989, 15, 53–65. [Google Scholar] [CrossRef]
  319. Carnicer, O.; Tunin-Ley, A.; Andree, K.B.; Turquet, J.; Diogène, J.; Fernández-Tejedor, M. Contribution to the genus Ostreopsis in Reunion Island [Indian Ocean]: Molecular, morphologic and toxicity characterization. Cryptogam. Algol. 2015, 36, 101–119. [Google Scholar] [CrossRef]
  320. Janse van Vuuren, S.; Taylor, J.C. Changes in the algal composition and water quality of the Sundays River, Karoo, South Africa, from source to estuary. Afr. J. Aquat. Sci. 2015, 40, 339–357. [Google Scholar] [CrossRef]
  321. Alkawri, A.; Abker, M.; Qutaei, E.; Alhag, M.; Qutaei, N.; Mahdy, S. The first recorded bloom of Pyrodinium bahamense var bahamense plate in Yemeni coastal waters off Red Sea, near Al Hodeida City. Turkish. J. Fish. Aquat. Sci. 2016, 16, 275–282. [Google Scholar]
  322. Alkawri, A. Seasonal variation in composition and abundance of harmful dinoflagellates in Yemeni waters, southern Red Sea. Mar. Pollut. Bull. 2016, 112, 225–234. [Google Scholar] [CrossRef]
  323. Ten-Hage, L.; Quod, J.-P.; Turquet, J.; Couté, A. Bysmatrum granulosum sp. nov., a new benthic dinoflagellate from the southwestern Indian Ocean. Eur.J. Phycol. 2001, 36, 129–135. [Google Scholar] [CrossRef]
  324. Africa D of AF and F of R of S. South African Molluscan Shellfish Monitoring & Control Programme; Cape Town, 2016; Available online: (accessed on 10 November 2018).
  325. Tanzania Food and Drugs Authority. Guidelines for Investigation and Control of Foodborne Diseases; Dar Es Salaam, 2011; Available online: (accessed on 10 November 2018).
  326. Munga, D.; Bosire, J.O.; Ruwa, R.K.; Jembe, T.; Abila, R.O.; Gichuki, J.W. Kenya Marine and Fisheries Research Institute Research Policy. 2010. Available online: (accessed on 20 June 2018).
  327. Cato, J.C. Seafood Safety: Economics of Hazard Analysis and critical Control Point (HACCP) Programmes; Food & Agriculture Org.: Rome, Italy, 1998. [Google Scholar]
  328. Bouaıcha, N.; Chézeau, A.; Turquet, J.; Quod, J.-P.; Puiseux-Dao, S. Morphological and toxicological variability of Prorocentrum lima clones isolated from four locations in the south-west Indian Ocean. Toxicon 2001, 39, 1195–1202. [Google Scholar] [CrossRef]
  329. El Masry, M.K.; Tawfik, H.M. 2011 Annual Report of the Poison Control Centre of Ain Shams University Hospital, Cairo, Egypt. Ain-Shams J. Forensic. Med. Clin. Toxicol. 2013, 20, 10–17. [Google Scholar] [CrossRef]
  330. Sector, N.C. Biodiversity conservation capacity building in Egypt. 2006. Available online: (accessed on 10 November 2018).
  331. Joyce, L.B.; Pitcher, G.C.; Du Randt, A.; Monteiro, P.M.S. Dinoflagellate cysts from surface sediments of Saldanha Bay, South Africa: An indication of the potential risk of harmful algal blooms. Harmful Algae 2005, 4, 309–318. [Google Scholar] [CrossRef]
  332. Matthews, M.W.; Bernard, S. Eutrophication and cyanobacteria in South Africa’s standing water bodies: A view from space. S. Afr. J. Sci. 2015, 111, 1–8. [Google Scholar] [CrossRef]
  333. Kopczyńska, E.E.; Fiala, M. Surface phytoplankton composition and carbon biomass distribution in the Crozet Basin during austral summer of 1999: Variability across frontal zones. Polar Biol. 2003, 27, 17–28. [Google Scholar] [CrossRef]
  334. Sebastián, C.R.; Etheridge, S.M.; Cook, P.A.; O’ryan, C.; Pitcher, G.C. Phylogenetic analysis of toxic Alexandrium [Dinophyceae] isolates from South Africa: Implications for the global phylogeography of the Alexandrium tamarense species complex. Phycologia 2005, 44, 49–60. [Google Scholar] [CrossRef]
  335. Bauer, K.; Díez, B.; Lugomela, C.; Seppälä, S.; Borg, A.J.; Bergman, B. Variability in benthic diazotrophy and cyanobacterial diversity in a tropical intertidal lagoon. FEMS Microbiol. Ecol. 2008, 63, 205–221. [Google Scholar] [CrossRef] [PubMed]
  336. Díez, B.; Nylander, J.A.A.; Ininbergs, K.; Dupont, C.L.; Allen, A.E.; Yooseph, S.; Rusch, D.B.; Bergman, B. Metagenomic analysis of the Indian ocean picocyanobacterial community: Structure, potential function and evolution. PLoS ONE 2016, 11, e0155757. [Google Scholar] [CrossRef] [PubMed]
  337. Hamisi, M.I.; Mamboya, F.A. Nutrient and phytoplankton dynamics along the ocean road sewage discharge channel, Dar es Salaam, Tanzania. J. Ecosyst. 2014, 2014, 271456. [Google Scholar] [CrossRef]
  338. Kyewalyanga, M.; Lugomela, C. Existence of potentially harmful microalgae in coastal waters around Zanzibar: A need for a monitoring programme? 1999. Available online: (accessed on 10 June 2018).
  339. Lugomela, C.; Pratap, H.B.; Mgaya, Y.D. Cyanobacteria blooms—A possible cause of mass mortality of Lesser Flamingos in Lake Manyara and Lake Big Momela, Tanzania. Harmful Algae 2006, 5, 534–541. [Google Scholar] [CrossRef]
  340. Lugomela, C. Population dynamics of Pseudo-nitzschia species [bacillariophyceae] in the near shore waters of Dar es Salaam, Tanzania. Tanzan. J. Sci. 2013, 39, 38–48. [Google Scholar]
  341. Lundgren, P.; Bauer, K.; Lugomela, C.; Söderbäck, E.; Bergman, B. Reevaluation of the nitrogen fixation behavior in the marine non-heterocystous cyanobacterium Lyngbya majuscula. J. Phycol. 2003, 39, 310–314. [Google Scholar] [CrossRef]
  342. Kotut, K.; Ballot, A.; Krienitz, L. Toxic cyanobacteria and their toxins in standing waters of Kenya: Implications for water resource use. J. Water Health 2006, 4, 233–245. [Google Scholar] [CrossRef]
  343. Robinson, R.; Champetier de Ribes, G.; Ranaivoson, G.; Rejely, M.; Rabeson, D. KAP study [knowledge-attitude-practice] on seafood poisoning on the southwest coast of Madagascar. Bull. Soc. Pathol. Exot. 1999, 92, 46–50. [Google Scholar] [PubMed]
  344. Diogène, J.; Campàs, M. Recent Advances in the Analysis of Marine Toxins; Elsevier: Centro Rio de Janeiro, Brazil, 2017; Volume 78. [Google Scholar]
  345. Grzebyk, D.; Berland, B.; Thomassin, B.A.; Bosi, C.; Arnoux, A. Ecology of ciguateric dinoflagellates in the coral reef complex of Mayotte Island [SW Indian Ocean]. J. Exp. Mar. Biol. Ecol. 1994, 178, 51–66. [Google Scholar] [CrossRef]
  346. Glaizal, M.; Tichadou, L.; Drouet, G.; Hayek-Lanthois, M.; De Haro, L. Ciguatera contracted by French tourists in Mauritius recurs in Senegal. Clin. Toxicol. 2011, 49, 767. [Google Scholar] [CrossRef] [PubMed]
  347. ISO. IEC 17025: 2005 General Requirements for the Competence of Testing and Calibration Laboratories; ICS: Geneva, Switzerland, 2005; p. 20. [Google Scholar]
  348. Ministery of Ocean, Economy, Marine Resources, Shipping F and Annual Report on Performance Fiscal Year 2016/17. 2017. Available online: (accessed on 10 November 2018).
  349. Banguera-Hinestroza, E.; Eikrem, W.; Mansour, H.; Solberg, I.; Cúrdia, J.; Holtermann, K.; Edvardsen, B.; Kaartvedt, S. Seasonality and toxin production of Pyrodinium bahamense in a Red Sea lagoon. Harmful Algae 2016, 55, 163–171. [Google Scholar] [CrossRef]
  350. Catania, D.; Richlen, M.L.; Mak, Y.L.; Morton, S.L.; Laban, E.H.; Xu, Y.; Anderson, D.M.; Chan, L.L.; Berumen, M.L. The prevalence of benthic dinoflagellates associated with ciguatera fish poisoning in the central Red Sea. Harmful Algae 2017, 68, 206–216. [Google Scholar] [CrossRef]
  351. Sabrah, M.M.; El-Ganainy, A.A.; Zak Cembella, A.D.; Lewis, N.I.; Quilliam, M.A. The marine dinoflagellate Alexandrium ostenfeldii (Dinophyceae) as the causative organism of spirolide shellfish toxins. Phycologia 2000, 39, 67–74. [Google Scholar]
  352. Lopez, J.A.V.; Al-Lihaibi, S.S.; Alarif, W.M.; Abdel-Lateff, A.; Nogata, Y.; Washio, K.; Morikawa, M.; Okino, T. Wewakazole B, a Cytotoxic Cyanobactin from the Cyanobacterium moorea producens Collected in the Red Sea. J. Nat. Prod. 2016, 79, 1213–1218. [Google Scholar] [CrossRef]
  353. Mohamed, Z.A.; Al-Shehri, A.M. Occurrence and germination of dinoflagellate cysts in surface sediments from the Red Sea off the coasts of Saudi Arabia. Oceanologia 2011, 53, 121–136. [Google Scholar] [CrossRef] [Green Version]
  354. Abd-Elhaleem, Z.A.; Abd-Elkarim, M.A. Pattern of food poisoning in Egypt, a retrospective study. J. Pharmacol. Toxicol. 2011, 6, 505–515. [Google Scholar] [CrossRef]
Figure 1. Chemical structure of OA and main derivatives [DTX1, DTX2, and DTX3].
Figure 1. Chemical structure of OA and main derivatives [DTX1, DTX2, and DTX3].
Toxins 11 00058 g001
Figure 2. Chemical structure of major CTXs analogs from Pacific (P-CTXs) (a) and Caribbean (C-CTXs) (b) regions. The major CTXs from Indian region (I-CTXs) have a similar structure with C-CTX-1. (c) Chemical structure of maitotoxin (MTX).
Figure 2. Chemical structure of major CTXs analogs from Pacific (P-CTXs) (a) and Caribbean (C-CTXs) (b) regions. The major CTXs from Indian region (I-CTXs) have a similar structure with C-CTX-1. (c) Chemical structure of maitotoxin (MTX).
Toxins 11 00058 g002aToxins 11 00058 g002b
Figure 3. Chemical structures of CI (SPXs (a), GYMs (b), PnTXs (c), and PtTXs (c),) and Silva et al. [79,83,84,85,86].
Figure 3. Chemical structures of CI (SPXs (a), GYMs (b), PnTXs (c), and PtTXs (c),) and Silva et al. [79,83,84,85,86].
Toxins 11 00058 g003aToxins 11 00058 g003b
Figure 4. Chemical structures of the main group of PbTxs (PbTxs-A and PbTxs-B). The capital letter A in first ring indicates type A and type B (also called type 1and type 2, respectively [4]). These rings contain lactone group that is most important for the toxin activity.
Figure 4. Chemical structures of the main group of PbTxs (PbTxs-A and PbTxs-B). The capital letter A in first ring indicates type A and type B (also called type 1and type 2, respectively [4]). These rings contain lactone group that is most important for the toxin activity.
Toxins 11 00058 g004aToxins 11 00058 g004b
Figure 5. Chemical structures of main pectenotoxins.
Figure 5. Chemical structures of main pectenotoxins.
Toxins 11 00058 g005
Figure 6. Chemical structures of YTXs n corresponds to the number of methyl groups in the molecule.
Figure 6. Chemical structures of YTXs n corresponds to the number of methyl groups in the molecule.
Toxins 11 00058 g006
Figure 7. Chemical structure of AZAs.
Figure 7. Chemical structure of AZAs.
Toxins 11 00058 g007
Figure 8. Chemical structure of DA and analogs.
Figure 8. Chemical structure of DA and analogs.
Toxins 11 00058 g008
Figure 9. Chemical structures of STX group.
Figure 9. Chemical structures of STX group.
Toxins 11 00058 g009
Figure 10. Chemical structure of TTX and their main analogues.
Figure 10. Chemical structure of TTX and their main analogues.
Toxins 11 00058 g010
Figure 11. Chemical Structure of PlTXs [PTX and Ostreocin-D].
Figure 11. Chemical Structure of PlTXs [PTX and Ostreocin-D].
Toxins 11 00058 g011
Figure 12. Chemical structure of MC.
Figure 12. Chemical structure of MC.
Toxins 11 00058 g012
Figure 13. Chemical structures of Aplysiatoxin (AT) and Debromoaplysiatoxin (DAT) (a); kalkitoxins (KTX) (b); lyngbyatoxins A, B and C (LA, LB and LC) (c); cylindrospermopsins (CYN) (d); jamaicadimes (JCD) (e); anatoxin-a (ANTX) and homoanatoxin-a (HANTX) (f) and antillatoxins (ATX) (g).
Figure 13. Chemical structures of Aplysiatoxin (AT) and Debromoaplysiatoxin (DAT) (a); kalkitoxins (KTX) (b); lyngbyatoxins A, B and C (LA, LB and LC) (c); cylindrospermopsins (CYN) (d); jamaicadimes (JCD) (e); anatoxin-a (ANTX) and homoanatoxin-a (HANTX) (f) and antillatoxins (ATX) (g).
Toxins 11 00058 g013aToxins 11 00058 g013b
Figure 14. Map of the incidence of marine toxins (MT) along African countries of the Indian Ocean and the Red Sea, from EgypttoSouth Africa and nearby islands. Red circles [ Toxins 11 00058 i001]—confirmed or suspected seafood poisoning episodes caused by MT; green circles [ Toxins 11 00058 i002]—MT or Harmful Algal Blooms monitoring programmes or Centers of seafood poisonings; Toxins 11 00058 i003—Saxitoxins group; Toxins 11 00058 i004—Okadaic Acid group; Toxins 11 00058 i005—Ciguatoxin group; Toxins 11 00058 i006—Palytoxin group; Toxins 11 00058 i007—Domoic Acid group and Toxins 11 00058 i008—Tetrodotoxin group.
Figure 14. Map of the incidence of marine toxins (MT) along African countries of the Indian Ocean and the Red Sea, from EgypttoSouth Africa and nearby islands. Red circles [ Toxins 11 00058 i001]—confirmed or suspected seafood poisoning episodes caused by MT; green circles [ Toxins 11 00058 i002]—MT or Harmful Algal Blooms monitoring programmes or Centers of seafood poisonings; Toxins 11 00058 i003—Saxitoxins group; Toxins 11 00058 i004—Okadaic Acid group; Toxins 11 00058 i005—Ciguatoxin group; Toxins 11 00058 i006—Palytoxin group; Toxins 11 00058 i007—Domoic Acid group and Toxins 11 00058 i008—Tetrodotoxin group.
Toxins 11 00058 g014
Table 1. Marine toxins and their symptoms, producers, permitted limit, detection methods, limit of detection/limit of quantification [LOD/LOQ] and toxicity equivalency factors [TEF] according to the European Food Safety Authority [EFSA].
Table 1. Marine toxins and their symptoms, producers, permitted limit, detection methods, limit of detection/limit of quantification [LOD/LOQ] and toxicity equivalency factors [TEF] according to the European Food Safety Authority [EFSA].
Toxin (Syndrome)SymptomsDetectionPermitted LimitToxin (TEF)Producer
MethodsLOD, μgKg−1LOQ, μgKg−1
OA and analogs (DSP)diarrhea, nausea, vomiting, abdominal pain and tumor formation in the digestive system [50]BA [180,181]160 0.16mg OA equivalents/Kg shellfish meat in EU region [182]OA[1.0]Dinoflagellates: Prorocentrum spp. [8], Dinophysis spp. [2,6,9,10,15,53,54] and Phalacroma rotundatum [55]
EIA [183,184,185,186]10–26 3–41
DTX2 [0.6]
LC-MS [183], -UVD [187]15–301–50
DTX3 [1.0; 1; 0.6]
CTXs and analogs (CFP)vomiting, diarrhea, nausea,
tingling, itching, hypotension, bradycardia. In extreme cases, death through respiratory failure in 30 min and 48 h after fish consumption [50]
BA [188,189]0.16–0.560 P-CTX [190] 0.01 μg P-CTX-1 equivalents/kg of fish in USA [191]P-CTX-1[1.0]Dinoflagellates: Gambierdiscus toxicus, Ostreopsis siamensis and Prorocentrum lima [59]
CTA [192,193,194]~106 - 0.039 C-CTX P-CTX-2[0.3]
2,3-dihydroxy P-CTX-3C[1.0]
EIA [72,189,195,196,197,198,199]-0.032 P-CTX
[67,70,71,74,200], -UVD [62,201,202]
CIsnon-specific symptoms such as gastric distress and tachycardia in humans [82]BA5.6–77 PnTXE Not regulated 13-desmethyl SPX C[1.0]Dinoflagellates: SPXs: Alexandrium spp. [1,76], GYMs: Gymnodium spp. [77], PnTXs: Vulcanodinium rugosum [78] and PtTXs: biotransformation from PnTXs via metabolic and hydrolytic transformation in shellfish [1,5,77,78,79]
FPA [203]80–85 13-SPXC
LC-MS/MS [79,204], - UVD [205]0.8–20 13-SPXC/GYMA
PbTxs and analogs (NSP)nausea, vomiting, diarrhea, paresthesia, cramps, bronchoconstriction, paralysis, seizures in 30 min to 3 h [87]BA [206] 800 μg BTX-2 equivalents/kg shellfish in USA [98], New Zealand, and Australia [99,100]BTX-2, BTX-3, BTX2-B2 and S-deoxy-BTX-B2 [same TEF]Dinoflagellate: Karenia spp. [4,16,87]
CTA [192]250 BTX-1
RB [108]30BTX-3
EIA [207,208]1 BTXs and 25 BTXs
LC – MS/MS [209]0.2 – 2 BTXs
PTX and analogsNo specific symptomsMBA- 160 µg OA equivalents./kg shellfish meat in EU region [210]PTX [1,2,3,4,6 and 11][1.0]Dinoflagellate: Dinophysis acuta [101]
EIA [207]-
PTX [7,8,9 and 2SA] and 7-epiPTX2 SA [<<10]
LC – MS/MS [211,212]1
YTX and analogsNo specific symptomsBA 3.75 mg YTX equivalents/Kg shellfish meat in EU region [124]YTX[1.0]Dinoflagellate: Protoceratium reticuatum [4,109], Lingulodinium polyedrum [4] and Gonyaulax polyhedral [4]
EIA [213] 1a-homoYTX[1.0]
LC-MS/MS [111]0.017
AZA and analogs (AZP)nausea, vomiting, diarrhea and decreased reaction to stomach cramps, deep pain, dizziness, hallucinations, confusion, short-term memory loss, seizure [214]BA [181]0.05 0.16 mg AZA1equivalents/Kg shellfish in EU region [210]AZA1[1.0]Dinoflagellates: Azadinium spinosum [117] and Protoperidinum crassipes [118]
STX and analogs (PSP)Numbness in the face and neck; headache,
dizziness, nausea, vomiting, diarrhea, muscular paralysis; pronounced respiratory difficulty;
death through respiratory paralysis [215]
BA [216,217] 0.8 mg STX equivalent/Kg shellfish in EU region [210]STX[1.0]Dinoflagellates: Alexandrium spp. [2,3,7], Gymnodinium catenatum [3], Pyrodinium bahamense [3] and cyanobacteria Trichodesmium erythraeum [131]
SBA [218] GTX1[1.0]
CTA [192,219] GTX4[0.7]
Antibodies Assay [220,221,222,223,224] GTX[0.1]
Eletrophoresis [225] C4[0.1]
LC-MS/MS [226,227,228,229]23–42 STX de-GTX3[0.2]
DA and analogs (ASP)nausea, vomiting, diarrhea or abdominal cramps] within 24 h of consuming DA contaminated shellfish and/or neurological symptoms or signs [confusion, loss of memory or other serious signs such as seizure or coma] occurring within 48 h BA [230]40 20 mg DA equivalents/Kg shellfish in EU region [210] Diatoms: Pseudo-nitzschia spp. [126] and red algae: Chondria armata [127].
(a) ASP- EIA [184,231]0.003 0.01
SPR [232]20
RB [233,234,235]20
Capillary electrophoresis [236,237,238]0.15 -1
LC -MS/MS [211,239,240], UVD [241,242]0.015
TLC [243]10
TTX and analogsVomiting, strong headache, muscle weakness, respiratory failure, hypotension and even death in hours [244]BA [144,245,246,247]1.1 [247] 2 mg TTX equivalents/Kg shellfish in Japan [248]S/R 11-norTTX-[6]-ol[0.19/0.17]Bacteria: Serratia marcescens, Vibrio spp. [83], V. Aeromonas sp. [138], Microbacterium, arabinogalactanolyticum [139], Pseudomonas sp. [140], Shewanella putrefaciens [141], Alteromonas sp. [142], Pseudoalteromonas sp. [143], and Nocardiopsis dassonvillei [144]
RB [249]2–4.10−3TTX
EIA [245,246,247,250,251,252,253,254,255,256]0.002/mL [255], 0. 0001/mL [253]
TLC [139,257]2 [257] 4,9-anhydroTTX[0.02]
GC-MS [28,258,259]500 1000 [258]
LC-MS/MS [260,261,262,263,264] – FLD [265]0. 00009?-24.5 [260,261,262,263,264]40 [265] – 100 [265]
PlTXVasoconstriction, hemorrhage, myalgia, ataxia, muscle weakness, ventricular fibrillation, ischemia and death [266,267] and rhabdomyolysis [268]BA Not regulated toxin but proposed value is 0.25mg PlTX equivalent/Kg shellfish in EU region [269]PlTX[1.0]Zoanthids: Palythoa spp. anddinoflagellates: Ostreopsis ovata. [153,154,155] and possibly cyanobacteria: Trichodesmium sp. [156]
Hemolysis assay [270]1.6
CTA [107]50
EIA [254]1/mL
LC-MS/MS [204,271]–FLD and–UVD [272]2,5.10−5–0, 50.10−5
MCliver hemorrhage within a few hours of an acute dose and death [273]LC-MS [167,274,275,276] and EIA [277] Tolerable daily intake: 0.04 μg/kg of MC body weight/day [278] Cyanobacteriaof genus: Pseudoanabaena, Phormidium, Spirilia [164], Leptolyngbya, Oscillatoria, Geitlerinema [165], Trichodesmium [166] and Synechococcus [167]
ANTX and HANTXHypersalivation, diarrhea, shaking and nasal mucus discharge [279], respiratory arrest and death [280]RB and GC/MS [281,282] Cyanobacteria: Hydrocoleum lyngbyaceum [177]
AT and DATContact dermal: dermatitis initiating with erythema
and burning sensations, appearing a few hours after exposure,
gave way to blister formation and deep desquamation,
lasting up to several days
[283,284] and consumption of contaminated seafood; burningsensation in the mouth and throat, vomiting and diarrhea [285]
LC-MS/MS [286] Algae Gracilaria coronopifolia [172] and cyanobacteria Lyngbya majuscula [171]
LA, LB, and LC Cyanobacteria Lyngbya majuscule [174]
ATX and analogsNo specific symptomsLC [287] Cyanobacteria: Lyngbya majuscula [179]
JCD and analogsNo specific symptomsLC, TLC and [288] Cyanobacteria: Lyngbya majuscula [176]
KTX and analogsNo specific symptomsLC [173] Cyanobacteria: Lyngbya majuscula [173]
CYN and analogsGastroenteritis [289]LC-MS/MS [290],–PDAD [291]1 [292]–200 [293] Cyanobacteria: Cylindrospermopsis raciborskii [175]
EIA [294]
Toxins: DA—domoic acid, DTX, CTX -ciuatoxin, AZA—azaspiracid, CI—cyclic imines, PTX—pectenotoxin, YTX—yessotoxin, STX—saxitoxin, OA—okadaic acid, BTX—revetoxin, PlTX—palytoxin, TTX -tetrodotoxin, MC—microcystin, ANTX—anatoxin, HANTX—homoanatoxin, LA, LB and LC—lyngbyatoxins A, B and C respectively. ATX—antillatoxin, KTX—kalkitoxin, CYN—cylindrospermopsins AT—aplysiatoxin, DAT—debromoaplysiatoxin, JCD—jamaicamides, Syndrome: PSP—Paralyc Poisoning, DSP—Diarrheic Shellfish Poisoning, ASP—Amnesic Shellfish Poisoning, AZP—Azaspiracid Shellfish Poisoning, CFP—CiguateraShellfish Poisoning, NSP—Neurologic Shellfish Poisoning, Detection methods: CTA—Cytotoxicity assay, EIA—Enzyme-ImmunoAssay, SPR—Surface Plasmon Resonance, RB—Receptor-based, GC—Gas Chromatography, BA—Bioassay; UVD—Ultra Violet Detection; LC—Liquid Chromatography and MS—Mass Spectroscopy, FPA—Fluorescence Polarization Assay, TLC—Thin Layer Chromatography, SBA—Saxitoxin Binding Assay, PDAD—photo diode array detection.
Table 2. MT monitoring scenario of the African countries of the Indian Ocean and the Red Sea.
Table 2. MT monitoring scenario of the African countries of the Indian Ocean and the Red Sea.
CountryMonitored MTPermitted Limit, mgKg−1 ShellfishDetectionLaboratories for Toxin AnalysisReference
South AfricaPST0.8 STX Research centers and Universities[324]
OA, DTX1-2, PTX1-20.16 mg OALC-MS/MS
YTX, 45 OH YTX, homo YTX, and 45 OH homo YTX8 mg YTXLC-MS/MS
AST20 mg DA
AZA1-30.16 mg OALC-MS/MS
TanzaniaCTX, TTX, ASTN.D.Symptomology and vectorsN.D.[325]
KenyaMT producers [HAB]N.D.N.D.Mombasa Research Center[326]
MadagascarN.D.N.D.Educational programmesResearches centers and Universities[327]
French IslandsN.D.N.D.N.D.Researches centers[35,328]
MauritiusN.D.N.D.N.D. [324]
Somalia and SeychellesN.D.N.D.N.D.N.D.
EygptN.D.N.D.N.D.Poison Control Center, Ain Shams University [329,330]
N.D.—No Data.
Table 3. Geographic occurrence MT per country, MT producer, and MT vector along African countries of the Indian ocean and red sea coasts. TX - toxin.
Table 3. Geographic occurrence MT per country, MT producer, and MT vector along African countries of the Indian ocean and red sea coasts. TX - toxin.
ToxinDateLocationToxin ProducerDetermination MethodToxin VectorTX Concentration, (mg TX Equivalents per Kg Shellfish Meat)Cell/Extract ToxicityReference
PSTs1999South AfricaAlexandrium catenellaAOAC mouse bioassayHaliotis midae0. 01609 STX [22]
1998–2002South Africa: Yzerfontein, Alexandrium catenellaHPLC-FLD--4.8 pg STX eq cell−1[334]
Alexandrium tamiyavanichi0.14 pg STX eq cell−1
2003–2004South Africa: Cape Town Alexandrium minutumLC-FD and HILIC-MS/MS--0.65 pg GTX cell−1[309]
Red Sea
Pyrodinium bahamense, Ceratium sp., Alexandrium sp. and Protoperidinium spp.ELISA -->> 0.4 ng mL1 [349]
DSTs2000Europa Island Mozambic channel, France]Prorocentrum arenariumFR3T3 fibroblast --IC50 = 0,1 µg OA ml−1 and 50 µg extract ml−1[11]
HPLC-MS22 ng OA/mg of extract
2001Lagoons of La Reunion Mayotte and Mauritius IslandsProrocentrum
PPIA- -IC50 1.3–25 mg/mL onon fibroblast;6261.3 ± 156.5 − 128.3±17.2 ng eq OA/mg crudeextract[328]
2002–2018South Africa:Abalgold--Haliotis asinina--[324]
2008South Africa: Saldanha Bay and
Lambert’s Bay
Dinophysis acuminataLC-MS/MSCrassostrea gigas0.267 OA
Choromytilus meridionalis0.012 OA
CTXs2001Mauritius: Nazareth, Saya de Malha and Soudan-HPLC-MS/RLB, Mongoose feeding test, and MBALutjanus sebae and
Qualitative analysis-[71]
2002North of the Republic
of Mauritius, Banks fishery
-HPLC-MS/RLBLutjanus sebae -[70]
2012–2013Central Red SeaGambierdiscus belizeanus and
Ostreopsis spp.
Mouse neuroblastoma cell-based assay--6,50–1,14.10 −5 pg P-CTX−1 eq. cell−1[350]
2013Madagascar: district of Fenoarivo Atsinanana Gambierdiscus spp.CBA Carcharhinus leucas0.083
MBA 0. 09272 P-CTX-1
LC-ESI-HRMS0. 01628 P-CTX-1
MBA752 MU/g
Antalaha District
Ostreopsis siamensisMBAHerklotsichthys quadrimaculatus0. 00045 PTXs/fish [head and esophagus] [18]
Hemolysis assays0. 00002 PTXs/fish [head and esophagus]
Cytotoxicity tests0. 00000005/fish [head and esophagus]
1996Mauritius: Rodrigues IslandOstreopsis mascarenensisHPLC-diode array detector, Nanoelectrospray ionization quadrupole time-of-flight and HPLC-ESI-MS/MS analysis -- [14,160]
Hemolysis assays 8.00 ± 0.01 ng PTX mL−1
Cytotoxicity AssayIC50 = 10 μM against human H460 lung cancer cells
2008South Africa: Saldanha Bay and
Lambert’s Bay
Dinophysis acuminataLC-MS/MSCrassostrea gigas0.267 OA
Choromytilus meridionalis0.012 OA
DA cultures 2012South Africa: Algoa BayPseudo-nitzschia multiseriesELISA--0.076 pg DA cell−1–0.098 pg DA cell–1[12]
LC/MS–MS0.086 pg DA cell–1–0.086 pg DA cell–1
TTXs 1990–1991Egypt: Suez City, in the northwestern
part of the Red Sea
TLC, electrophoresis, UV, GC–MS Pleuranacanthus
752 MU/g
1998Madagascar: Nosy Be Island --MBA 16 MU/g [41]
2002–2003Egypt: Gulf of Suez MBALagocephalus sceleratus3950
2013Reunion Island MBA and LC-MS/MSLagocephalus sceleratus17 TTX-[35]
Table 4. Seafood poisoning episodes caused by MTs, observed effects/Symptoms, fish or shellfish consumed and victim number affected along African countries of the Indian Ocean and Red sea coasts. TX - Toxin
Table 4. Seafood poisoning episodes caused by MTs, observed effects/Symptoms, fish or shellfish consumed and victim number affected along African countries of the Indian Ocean and Red sea coasts. TX - Toxin
LocalDateSeafoodObserved Effects/SymptomsTXDetection MethodTX Concentration, (mg TX Equivalents/Kg Shellfish Meat)Victim NumberReference
Comoros islands:
24 December 2012Eretmochelys imbricata
Itching, Asthenia, Vomiting, Abdominal pain, Rash Myalgia
Shortness of breath, Nausea
Itching of the mouth/throat, Fever, Diarrhea Vertigo, Paresthesia, Dysphagia
Mouth burn Sore throat, Erectile dysfunction
---49 suspected cases and 8 probable cases, age range [0–40 years], 1 death[26]
North-eastern coast of MadagascarDecember 1994TurtleNausea, vomiting, dysphagia, acute stomatitis---60 persons with poisoning attack rate were 48% with a lethality of 7.7%[47],
Madagascar: district of Fenoarivo AtsinananaNovember 2013Carcharhinus leucas (shark)Paresthesia of the extremities, dysesthesia, and reversing sensitivity of hot and cold accompanied by a headache, dizziness, and arthralgia between 2 and 12h after ingestion CTXsMBA0.083
124 people, 9% deaths[20]
CBA0. 09272 P-CTX-1
Madagascar: Antalaha DistrictJanuary 1994Herklotsichthys quadrimaculatus (Fish)Malaise, uncontrollable vomiting, diarrhea, tinglings of extremities,
delirium and death
PlTXsMBA0. 00045 PTXs/fish [head and esophagus]Death of one adult[18]
Hemolysis assays0. 00002 PTXs/fish (head and esophagus)
Cytotoxicity tests0. 00000005/fish (head and esophagus)
Mass spectroscopy-
Madagascar: Nosy Be IslandJuly 1998--TTXsMBA16 MU/g (no data to covert to mg/Kg)4 people, one death[41]
Madagascar: Manakara district November 1993Carcharhinus amboinensis [shark] Deep coma and death,
body rigidity due to loss of cerebral function,
myosis, mydriasis,
convulsions, Respiratory distress due to acute pulmonary edema, cardiovascular collapse, bradycardia, gengivorrhagia
Dehydration, paresthesia on fingertips and toes, dizziness,
pruritus, narcosis, faintness, hyperthermia, ataxia asthenia, dehydration, cephalalgia, diarrhea, epigastralgia, laryngeal distress
CTXsCiguatera poisoning Symptomology-500 people, 20% deaths[21]
South Africa: Cape TownMay 1978Choromytilus meridionlis [Mussel]Paraesthesia of en
fingers/hands, Circumoral paresthesia, paranesthesia of toes/feet, Vertigo, Floating sensation, Ataxia, Weakness of upper, Weakness of lower limbs and Dysarthria
A headache
PSTsMBA72.83 STX17 people, no deaths[39]
South Africa: Natal coastDecember 1957Mytilus
meridionalis [Mussel]
lightness of the body, with a tingling around mouth, finger, and toes; no moving; feeble inarticulate noise;
PSTsMBA0.04 STX5 people and one cat[40]
South Africa: Table and False Bays1888Donax serra [Mussel]-----[37]
South Africa: Cape TownApril 1948Donax serra [Mussel] and Chloromytilus
meridionalis [Mussel]
----One death
South Africa: Natal coastDecember 1957Perna perna [Mussel]----5 people, one death
South Africa: Cape Town aMay 1958Chloromytilus meridionalis [Mussel]-- -One death
Reunion IslandSeptember 10th, 2013Lagocephalus sceleratus [fish]peri-oral paresthesia, weakness of both lower limbs, paresthesia all over the body, headache, dyspnea,
nausea and vomiting, blurring of vision, and vertigo
Flesh: 5 TTX
10 people[35]
Table 5. Recommended marine toxins to be monitored and suggestion of permitted limit to be used.
Table 5. Recommended marine toxins to be monitored and suggestion of permitted limit to be used.
ToxinSyndromePermitted Limit, mgKg−1To be adopted from
STXPSP0.8 STXeqEU region
CTXCFP0.00001 P-CTX-1eqUSA
YTX-3.75 YTXeqEU region
PTX-0.16 OAeqEU region
TTX-2 TTeqJapan
DAASP20 DAeqEU region
OADSP0.16 OAeqEU region
AZAAZP0.16 AZAeqEU region
PlTX-0.25 PlTXeq *EU region
PbTxNSP0.8 TX-2 eqUSA, New Zealand, and Australia
* This toxin is not monitored and 0.25 PlTXeq was proposed in the first meeting (Cesenatico, Italy, 24–25 October 2005) of the working group on Toxicology of the national reference laboratories [NRLs] for Marine Biotoxins.

Share and Cite

MDPI and ACS Style

Tamele, I.J.; Silva, M.; Vasconcelos, V. The Incidence of Marine Toxins and the Associated Seafood Poisoning Episodes in the African Countries of the Indian Ocean and the Red Sea. Toxins 2019, 11, 58.

AMA Style

Tamele IJ, Silva M, Vasconcelos V. The Incidence of Marine Toxins and the Associated Seafood Poisoning Episodes in the African Countries of the Indian Ocean and the Red Sea. Toxins. 2019; 11(1):58.

Chicago/Turabian Style

Tamele, Isidro José, Marisa Silva, and Vitor Vasconcelos. 2019. "The Incidence of Marine Toxins and the Associated Seafood Poisoning Episodes in the African Countries of the Indian Ocean and the Red Sea" Toxins 11, no. 1: 58.

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop