Center for Disease Control website:
http://www.cdc.gov/nceh/ciguatera/
Hawaii State Department of Health website:
Ciguatera Fish Poisoning: A Synopsis from Ecology to Toxicity
P. K. Bienfang, M. L. Parsons, R. R. Bidigare, E. A. Laws and P. D. R . Moeller
In: Oceans and Human Health: Risks and Remedies from the Sea. Chapter 14. Walsh, P. J., S. L. Smith, L. E. Fleming, H. Solo-Gabriele, and W. H. Gerwick (Eds.). Elsevier Science Publishers, New York.
Introduction
Ciguatera fish poisoning (CFP) is a world-wide health problem associated with the consumption of seafood. Humans acquire ciguatoxin (CTX), the causative poison, by eating reef fish that have accumulated the toxin via the marine food web. Ciguatoxin originates in dinoflagellates Gamberdiscus spp that are found in association with various macroalgae in coral reef ecosystems. These dinoflagellates are consumed by herbivorous fish, initiating the processes of bioaccumulation, biomagnification, and biomodification in the reef ecosystem food web, as the herbivores are consumed by carnivores, and, ultimately, by humans. The ecological complexities, biochemistry, and molecular biology associated with the production of ciguatoxin remain poorly understood, as do the etiology and epidemiology of the disease as presented in fish consuming human populations. This overview addresses the ecological, biochemical, and human health aspects of this fascinating and important disease phenomenon.
CFP: The Human Health Portion
History
Ciguatera fish poisoning (CFP) has affected coastal populations for centuries. The term, first coined in the Caribbean by the Spanish in the 17th century, is derived from “cigua”, the name used by indigenous populations in the Spanish Antilles for a marine turban snail (Halstead, 1967; Banner, 1976; Bagnis 1993). In the Pacific, ships’ records from the early 1600s in the New Hebrides, and Captain James Cook in 1774 from New Caledonia (Helfrich,1964) indicated that ships’ crews displayed clinical symptoms that today are associated with CFP. Evidence exists for CFP in all tropical ocean areas of the world. Outbreaks of fish poisoning occurred throughout the Pacific before, during and after WWII, and became quite a serious problem for military troops stationed in various island locales where CFP was endemic (Hokama and Yoshikawa-Ebesu, 2001).
In part, it was those experiences that focused scientific attention on CFP by Randall (1958), Banner et al. (1960) and Cooper (1964). Randall (1958) hypothesized that the toxin, ciguatoxin (CTX), was derived from food consumed by the fish, and Helfrich (1964) and Banner (1974; 1976) showed that toxicity could be transferred to/induced in non-toxic fish via consumption of toxic fish. These contributions evolved into the “food web concept” for CFP that is held to this day. The food chain hypothesis held that fish acquired the CTX toxin in their diet, that this acquisition occurred without marked detrimental effects to the fish, and that the ingested toxin was stored in the body without degradation (Randall 1958).
For decades, there has been especially rich and active ecological research regarding CFP in Australia and French Polynesia (e.g., Yasumoto et al., 1977; Bagnis 1977; Yasumoto et al., 1979; Bagnis et al., 1985; Holmes et al., 1991; Lewis et al., 1994; Chinain et al., 1999a), and the Caribbean (e.g., Carlson 1984; Tindall et al., 1984; Ballantine et al., 1985; Bomber et al., 1988; Pottier et al., 2002). Scheuer and colleagues (Scheuer et al., 1967; Nukina et al., 1984; Tachibana, 1980; Tacnibana et al., 1987; Nukina et al., 1986) first determined the molecular structure of CTX derived from moray eels. The etiology of CFP was advanced when work in the Gambier Islands of French Polynesia by Yasumoto et al. (1977; 1979a; 1980) revealed that the guts of toxic (herbivorous) fish contained significant numbers of a dinoflagellate that came to be known as Gambierdiscus toxicus (Adachi and Fukuyo, 1979). Numerous contributions to CTX detection methods followed (Hokama et al., 1977; 1985; 1987; 1989; Pottier et al., 2002; Lewis et al., 1999; Lewis and Jones 1997. Legrand et al. (1992) chemically characterized ciguatoxins from a variety of different fish species. Following the evolution of robust chemical methodologies, significant advances were made toward synthesis of the CTX molecule (Inoue et al., 2004; 2006).
Regional Occurrence and Under-reporting
There are at least 50,000 reported cases of CFP cases per year (Quod and Turquer, 1996; Lewis, 2001), but due to the high degree of misdiagnosis and under-reporting, it is estimated that the actual frequency of CFP cases is closer to 500,000 per year (Pearn, 2001; Arena et al., 2004). Because of this inaccuracy, it is not possible to ascertain whether CFP incidence is increasing over time. Increased awareness of CFP has given rise to improved identification and increased reporting within both the general population and the medical profession in certain endemic areas.
A CDC report for a period in the 1970s indicted that reported CFP incidences accounted for 25% of all food-borne outbreaks, which was five times the reported incidence for paralytic shellfish poisoning and neurological shellfish poisoning combined (Steidinger and Baden, 1985). Various studies for the Caribbean and Pacific have indicated frequencies of reported CFP incidence ranging from 0.1 – 3% of the population per year (Vernoux, 1988). It is believed that reported values must be increased by a factor of at least 2-20X to account for under-reporting (Arena et al., 2004; Lawrence et al., 1980; Fleming et al., 1994). The CDC estimates that in the USA only 2-10% of CFP cases are actually reported. It is estimated that 3% of the population in the U.S. Virgin Islands, and French West Indies suffer CFP annually; for St Thomas and Puerto Rico, these figures were estimated to be 4.4% and 7%, respectively (Escalona de Motta, and De la Noceda, 1985). An Australian study estimated that 1.8 – 2.5% of the total population from two Pacific island communities experienced a CFP intoxication at some time in their life (Gillespie et al., 1985).
One study presented a classic case of CFP symptoms to 36 physicians in Florida, an endemic area for CFP due to local and regional coral reefs. About 68% of the physicians correctly diagnosed CFP, but only about 17% correctly recommended mannitol therapy (see below) as the acute treatment of choice. Importantly, only about 47% of the physicians knew that CFP was a reportable disease. This study illustrates that CFP is an under-diagnosed, inadequately treated, and under-reported disease especially among US-born and US-trained physicians even in endemic areas (McKee et al., 2001). In Mexico, CFP is often misdiagnosed as some form of food poisoning from traditional sanitary microbial contamination, illustrating another way that CFP is a neglected disease.
Symptoms and Effects
Recently, Arena et al. (2004) summarized the frequency of clinical symptoms of CFP at the time of diagnosis from a number of works. CFP symptomatology has been addressed in considerable detail in numerous publications (Bagnis, 1968: Bagnis; 1990, Bagnis et al., 1979. Withers, 1982; Ragelis, 1984; Yasumoto et al., 1984; Steidinger and Baden, 1985; Calvert 1990; Nicholson and Lewis, 2006). Due to similarities in symptomatology, the diagnostic differentiation of CFP from neurological shellfish poisoning or paralytic shellfish poisoning commonly relies simply on the history of fish versus shellfish consumption as the principal guide. CFP also has symptoms in common with Type E botulism, Scombroid poisoning, eosinophilic meningitis, and organophosphate pesticide poisoning.
CTX produces gastrointestinal, neurological, and cardiovascular symptoms. These normally develop within 12-24 hours of eating the contaminated fish. Gastrointestinal effects usually disappear within 1-4 days. The normal progression of symptoms is: (a) gastrointestinal symptoms (e.g., diarrhea, abdominal pain, nausea, and vomiting), followed by (b) neurological symptoms (e.g., numbness and tingling of hands and feet, dizziness, altered hot/cold perception, muscle aches, low heart rates and low blood pressure). A pathoneumonic symptom involves the neurological paresthesia of the reversal of hot/cold sensation. Symptoms may persist in some form for weeks, months, or even years (Kodama, 1990, Cameron et al., 1991; Benoit et al., 2000; Arena et al., 2004). Generally, feelings of weakness last ~1 week, and neurosensory manifestations or paresthesias (e.g., muscle and joint aches, itching, tingling extremities, and thermal reversals) commonly represent the most prolonged discomfort. Fortunately, death is rare (i.e., <0.1%) and is most commonly the result of respiratory failure due to cardiovascular shock induced by severe dehydration during the initial onset of effects (Withers, 1982; Bagnis 1993). In the Pacific, this is normally associated with eating the most toxic portions of the fish (e.g. liver, viscera, roe).
Chronic CFP is often misdiagnosed as a psychiatric disorder of general malaise, depression, headaches, and peculiar feelings in the extremities (Chan and Wang, 1993). Interestingly, it appears that acutely gastrointestinal symptoms dominate CFP cases in the Caribbean, whereas neurological symptoms seem to be more prominent acutely in CFP cases in the Pacific (Nicholson and Lewis, 2006). It has been speculated that this difference is due to different toxin composition among the Caribbean and Pacific G. toxicus strains, and/or differences in the species of Gambierdiscus contributing to the toxicology.
An interesting feature is that CFP intoxication does not confer any immunity in its victims; on the contrary, it frequently results in heightened sensitivity to CTX, and/or fish products generally. It has been suggested (Nicholson and Lewis, 2006) that, because ciguatoxins may be sequestered in adipose tissue, the re-occurrence of symptoms may be exacerbated during periods of physical stress that catabolize fats (e.g., exercise or weight loss) and/or ingestion of alcohol and high protein foods (such as fish, chicken, or nuts, and foods with caffeine).
One well-documented case illustrates the progression of CFP symptoms. Thirty French citizens who consumed portions of a 10-12 lb barracuda (muscle tissue only) in Mexico developed gastrointestinal disorders in 4-6h, and were given antidiarrheal drugs prior to their flight home. During the flight, neurosensory disorders with diffuse pain developed, and on day 5 they were admitted to a poison treatment center. To make an objective assessment of the severity of the clinical features, the center scored each victim upon admission using a quantification method applied in Tahiti that gives points for each of the following: paraesthesiae, pain, tiredness and cardiovascular and gastrointestinal signs (De Haro, 1997). This study concluded that the severity of intoxication was proportional to the amount of fish eaten.
Pharmacology
The complex suite of symptoms associated with CFP is caused by the ciguatoxin’s capability to increase Na+ permeability through the Na channels that open at normal resting membrane potentials; this enhanced excitability of the membrane in turn affects Na+ – Ca+2 exchange and mobilizes intracellular Ca+2. The primary receptor site of the CTX action is the 5th domain of the Na+ channel where it causes increased sodium ion permeability and depolarization of the resting membrane; the intense depolarization of nerve cells is believed to cause the suite of sensory discomfort symptoms associated with CFP (Cameron et al., 1991; Hokama and Yoshikawa-Ebesu, 2001; Arias, 2006).
Prevention
The heat-stability of the ciguatoxin molecule means that normal/proper food preparation (i.e., heating) will not mitigate or eliminate toxicity from fish tissue. This property, together with the fact that exceptionally small (i.e., picogram levels) amounts of CTX toxin can confer toxicity, further reduces any reasonable likelihood for reliable detection/prevention during fish handling/cooking. Therefore, there is no way to “decontaminate” a fish having CTX. There is no way to reliably distinguish a contaminated fish by smell or appearance. In addition to the amount of contaminated fish consumed, intoxication seems to be most closely associated with the size and type of fish that was consumed. Generally, people are advised to avoid the viscera of reef fish, or the consumption of large amounts of large predatory fish (see below). Also many fish are misidentified, have little/no traceability, or there is a lack of knowledge of the CFP-propensity of the particular fish at the time of their sale. Improvements in these areas are needed to aid prevention.
Tropical islanders have used numerous folk techniques in the attempt to determine if candidate fish for consumption are ciguatoxic: (a) cooking the fish with a silver object to see if discoloration results; (b) seeing if the fish repels flies or ants; (c) avoiding sliced fish that fails to reflect a rainbow when held to the sun; and (d) rubbing one’s gums with the liver to see if a tingling results (Lobel, 1974). Giving a sample of the questionable fish to a household pet or even an elderly relative as a bioassay is still practiced in some island communities.
Clinical Responses/Treatments
By far the most effective therapy for CFP has been mannitol infusions, administered at 0.5 to 1.0 g/kg body weight, and given within 48 hours of ingestion of toxic fish (Palafox et al., 1988). The mechanism of mannitol’s effect is partly understood. CTX opens sodium channels at the nodes of Ranvier in myelinated neurons, causing the cleft to swell, eliminating saltatory conductance. Mannitol relieves the swelling at the nodes, presumably by an osmotic effect. IV mannitol is the only CFP treatment evaluated by a randomized blinded trial (Bagnis, Spiegel and Boutin et al., 1992; Schnorf et al., 2002). Although one of these randomized trials (Schnorf et al., 2002) did not find mannitol to be superior to saline, that study included patients who received mannitol up to 28 days after symptom onset and not within the recommended 2-3 days from consumption, which may have prevented the identification of benefits in patients who received it within the recommended period (Blythe et al., 1992; Friedman et al., in press).
Many other treatments have been tried with variable results and without appropriate randomized trial testing (Watters 2007). Other clinical treatments, targeting various elements of the long list of CFP symptoms, have involved a variety of agents, e.g., vitamins, antihistamines, anticholinesterases, steroids, and antidepressants. Treatments frequently focus on the most demonstrative symptoms, and thus may involve injections of steroids, non respiratory depressants, antihistamines, antidiarrhetics, and vitamins. Gut emptying and decontamination with charcoal have also used, but ongoing vomiting and diarrhea often prevents this. Atropine has been used for bradycardia, and dopamine or calcium gluconate for shock.
Folk Remedies
Folk remedies involving extracts of Argusia argentea leaves or Davalliea solida rhizomes have been used traditionally in New Caledonia (Benoit, 2000). Baden et al. (1995) and others (Blythe et al., 2001; Fleming et al., 2004) reported that there are 64 different local remedies including medicinal teas that are used in both the Indo-Pacific and West Indies regions. Folk remedies frequently involve inducing vomiting; it is generally accepted that vomiting and diarrhea should not be suppressed since it aids the body in voiding the poison.
Hazard Management
CFP ranks as a very significant hazard from seafood consumption. A risk assessment tool of ten seafood hazard/product combinations using data from Australia showed CFP to be among the most hazardous incidents. Examples of hazard/product pairs with a lower ranking (i.e., less hazardous) included: mercury poisoning, Clostridium botulinum in canned or cold-smoked fish, parasites in sushi/sashimi, viruses in shellfish from uncontaminated waters, enteric bacteria in imported cooked shrimp, Vibrio parahaemolyticus or V. cholerae in cooked prawns, Listeria monocytogenes in cold-smoked seafoods, scombroid, and V. vulnificus in oysters. The only elements deemed more hazardous than ciguatera were L. monocytogenes in susceptible populations and enteric bacteria in imported cooked shrimp eaten by vulnerable consumers (Sumner and Ross, 2002).
As noted above, CTX in fisheries products is odorless, tasteless, and generally undetectable by simple chemical tests. Assurances that susceptible foods are safe to eat will likely come from the marketplace in the form of screening, isolation of high-risk products, and (where feasible) prediction of potentially hazardous harvesting areas. Effective screening methods must be easy to use/interpret, able to test large numbers of samples quickly, accurate, low-cost, readily available, and capable of identifying the causative CFP toxins. CFP incidence does not reflect on the quality of food handling, storage, preparation, or procurement; there is no known method of cooking, boiling, baking or frying that can destroy the toxin (Bagnis, 1993).
Risk Assessment
A study in French Polynesia simultaneously evaluated three indices of CFP hazard. These included CFP cases per 1000 residents (CIR), percentage of toxic fish in a group (PCI), and the abundance of G. toxicus per gram algae (GTD). During the 20-year study period 24,000 cases were reported (CI = 8%), caused by ~100 fish species. Results indicated that G. toxicus abundance data in situ and the respective ciguatoxicity of significant samples of microphagous herbivorous fishes and ichthyophagous carnivorous fish provided a reasonable estimate of the CFP potential at any given time of the year (Bagnis, 1985). In Hawaii, a general correlation is also apparent seasonally between the frequency of reported CFP incidents and the abundance of Gambierdiscus spp.; the highest values of both being observed during the warmest period at the end of the summer.
Fish to Avoid
The fish most commonly involved in CFP incidents include: barracudas (Sphyraena spp.), groupers (Epinephelus spp.), jacks (Caranx spp.), snappers (Lutjanus spp.), surgeonfish (Ctenochaetus spp), parrot fish (Scarus spp.), and moray eels, e.g., Gymnothorax spp. (Lewis et al., 1999; Ebesu, 1998; Vernoux and Lejeune, 1993; Tosteson et al.,1988). In the Pacific, the detritivorous grazer, Ctenochaetus striatus (a surgeonfish), is frequently ciguatoxic, and is thought to be a key vector sending CTX up the food chain. Surgeonfish and parrotfish are dominant families by weight on many coral reefs, and the most common prey of large piscivores. High levels of CTX and other Gambierdiscus-associated toxins in biodetritus are believed to account for frequency of ciguatoxicity in Ctenochaetus striatus, and in some areas, mullet (Mugil spp.) because both have detritivorous grazing behaviors (Steidinger and Baden, 1985). Studies of seasonality of CFP in different regions have frequently shown temporal variability, but often at different times of the year in different locations.
A study of toxic barracuda in the Caribbean concluded that CTX was accumulated/retained in barracuda tissue for extended periods of time. Fish flesh toxicity was shown to be inducible. Snapper fed CTX fish became toxic in six months and retained potency for 30 months (Banner et al., 1966; Helfrich and Banner, 1968). Toxin chemistry results suggest that CTX precursors from dinoflagellates may be oxidatively biotransformed to numerous congeners within herbivorous and carnivorous fish (Legrand 1998).
Diagnostic Kits
Development of simplified procedures for the assessment of CTX and related polyethers in ciguateric fish has been actively pursued. The first tested was the radioimmunoassay (RIA) reported by Hokama et al., (1977) in which the antibody was radiolabeled with 125I. Although the RIA was effective, its complexity and need for a radioactivity counter demonstrated that a simpler, more cost-effective method was needed (Hokama and Yoshikawa-Ebesu 2001). Next, Hokama’s group developed the stick-enzyme immunoassay (S-EIA) using sheep anti-CTX and MAb-CTX coupled to horseradish peroxidase (Hokama 1985). A UV monitor was used to detect when the MAb-CTX-horseradish complex bound to the putative CTX molecule. Herein lies a non-trivial diagnostic challenge for such kits, i.e., the fact that multiple CTX congeners having different molecular structures may both be present and may present similar symptomology. Adding to the diagnostic challenge and ranging degrees of efficacy from various therapies, there may also be a number of other toxins (such as okadaic acid, derived from Ostreopsis spp.) associated with ciguateric fish. While this method was used extensively through the mid-1980s and 1990s, the presence of ciguatoxin still could not be directly detected (an UV detector was needed).
The latest developments have focused on the membrane immunobead assay (MIA). The MIA utilizes a plastic stick for the solid phase and a synthetic hydrophobic membrane laminated onto one end that serves as the solid-phase receptor for the binding of methanol-extracted CTX and/or its related polyethers from fish using detection via colored polystyrene beads coated with MAb- to CTX (Hokama et al., 1998). The MIA was deemed successful enough to merit commercial production in the late 1990s through Oceanit® Test Systems as the Cigua-Check® Test Kit. The level of satisfaction with clarity, accuracy, or cost of commercially available CTX kits is far from universal, however (Dickey et al., 1994).
Societal Impacts
There are considerable negative impacts associated with CFP. These include: (a) the obvious public health impacts of the intoxications; (b) constraints on an important protein source for sub/tropical island communities globally; (c) constraints to the development of small-scale tropical fish resources for export; and (d) potentially immobilizing hazards to personnel involved in operations (e.g., military, construction, etc.) in tropical areas. In response to endemic ciguatera from the consumption of local fish, people in the Marshall Islands reportedly changed their eating preferences to include more imported and canned fish because of increased CFP incidence in locally caught fish (Tebana, 1992). Negative ecological impacts of various forms of fish and shellfish poisonings have included mass deaths of shellfish, seagulls, dolphins, and turtles (Ochoa, 1996).
CFP is believed to be the leading cause of non-bacterial food poisoning in the United States, and at an estimated incidence rate of 500,000 cases annually it is the most frequent seafood-toxin illness worldwide (Arena et al., 2004; Quod and Turquet, 1996; Lewis, 2001; Pearn, 2001). For example, a study of fish poisonings in a location in the Indian Ocean concluded that ~80% were due to CTX (Quod and Turquor, 1996). Studies have variously estimated the approximate cost to be $5000 - $9000 per CFP incident, particularly in non-endemic areas (Pohland et al 1990; Arena et al., 2004). It has also been suggested that by any latitudinal extension of warmer water (e.g., due to global warming) will similarly increase the incidence of CFP (De Sylva, 1999).
CFP: The Ocean Production Portion
It is now generally understood that the dinoflagellate, Gambierdiscus toxicus, the putative producer of the toxins associated with Gamberdiscus (also known as “gambiertoxins”) that are transformed to CTX, grows epiphytically on macroalgae together with a complex consortium of other symbiotic microflora. This microbial biomass is ingested by herbivorous fish which accumulate the toxin and in turn are grazed by carnivorous fish that bioamplify and chemically modify the toxin. The simplicity of the above schema belies a great deal of uncertainty about virtually every facet of the ecological and/or biochemical processes that lead to the manifestation of CFP in humans. Difficulties in resolving unknowns in the CFP process arise from the ephemeral character of CTX production by Gambierdiscus spp., and the need for extremely robust analytical methods due to the CTX molecular structure, its extreme toxicity, and its common association with other phytotoxins. What follows is our best understanding of the currently understood CFP paradigm.
Ciguatoxin
CTX is a polar, lipid-soluble, highly oxygenated polyether molecule. It is an oxygenated long chain fatty acid with cyclic oxoether linkages Derived from polyketide biosynthetic pathways, CTX is soluble in methanol, ethanol, acetone, and 2-propanol, but not benzene or water. Toxin isolated from different regions and/or organisms has been shown to exist in a number of forms that have different molecular weight, chemical structure, and toxicity. Recognition of these “congeners” necessitated new nomenclature, and in the more recent literature an I, P, or C prefix refers to the ocean of origin (i.e., Indian, Pacific or Caribbean), and a number following refers to a specific congener/form of CTX. Thus, P-CTX-1 would refer to CTX congener 1 isolated from the Pacific Ocean. The advent of more robust analytical methods has lead to important new details emerging regarding the chemical structure of CTX and its congeners. For example, the molecular weight of various P-CTX congeners from moray eels were found to range from 1023-1112 amu (Yasumoto et al., 1993 ), and P-CTX in fish from Hawaii were recently found to be 993 amu (P. Moeller, Pers. Comm.). Ciguatoxin specifically may also be present with another lipid-soluble toxin (scaritoxin), and the larger (3422 amu), water soluble maitotoxin, named after the Tahitian name for Ctenochaetus striatus. The molecular formulae and weights of these and other phytotoxins are summarized in Pohland et al., (1990).
Ciguatoxin and maitotoxin are two of the most lethal natural substances known. In mice CTX and MTX are lethal at injected (ip) levels of 0.45 ug/kg and 0.13 ug/kg (Anderson and Lobel 1987, Tachibana 1980, Yasumoto, 1985, Bagnis et al., 1987). Intake of picogram amounts of CTX can cause illness in human adults. Because of the prevalent use of bioassays in lieu of chemistry, the MU (i.e., mouse unit) is frequently encountered in the literature. A MU is defined as either the amount of toxin required to kill a 20-gm mouse in 15 minutes, or the lethal dose for 50% of the population (LD50) for 24Hr (Steidinger and Baden, 1985).
CTX Assay Methods
Analytical difficulties arising from the complexity of the CTX molecule cannot be overemphasized. Interest circumventing the complex analytical chemistry required has led to the development/application of a number of bioassays. None of these assays has proven to be completely satisfactory for routine laboratory testing, field samples, clinical studies or specific investigations because of individual limitations (Luckas, 2000).
The mouse bioassay is an historical method for determining algal toxins, though limited by low sensitivity and increasing societal aversion to animal testing. A number of other assay organisms have been used in CTX bioassay trials over the years; these have included: mongoose, rat, Artemia, cell bioassays, Diptera larvae, shrimp, rabbits, guinea pigs, cats, chicks, and mosquitoes (Labrousse, 1991; Miller, 1991 and refs cited within). A MAb(monocolonal antibody)-based assay has been used as an alternative assay to the mouse bioassay for routine monitoring (Hokama et al., 1998 ), and radioimmune (RIA) or enzyme linked immunosorbent (ELISA) assays are used for detecting CTX in fish (Labrousse, 1991; Hokama and Yoshikawa-Ebesu 2001). Cell-based assays for CTX using mouse neuroblastoma cells have proven to be especially functional and sensitive since development of refinements that enable the assessment of sodium-channel activity specifically (Manger et al., 1995). Currently, the best methodology for CTX assessment involves LC/MS/MS/NMR technologies. Though significantly expensive to purchase, maintain, and operate, this suite of applications is by far the most sensitive and informative. The National Science Foundation (NSF) and National Institute of Environmental Health Sciences (NIEHS) Center for Oceans and Human Health at the University of Hawaii uses a two-tiered CTX assessment strategy. The Na-channel neuroblastoma assay (tier 1) is used to screen samples to confirm sodium channel sensitivity, and positive samples are then analyzed via LC/MS/MS/NMR procedures (personal communication, authors).
Gambierdiscus toxicus
One of six species of Gambierdiscus, the dinoflagellate, G. toxicus, is roughly 93 x 83um (Chinain et al., 1999b). It is named after the Gambier Islands in French Polynesia (Adachi and Fukuyo, 1979) where it was found in high abundance and association with CFP (Yasumoto et al., 1977). There is considerable uncertainty about the growth rates and/or relative toxicity of clones, strains, and/or Gambierdiscus species from different locals (e.g., Holmes et al., 1991; Legrand, 1998), but several Gambierdiscus species are reported to produce gambiertoxins/ciguatoxins (Chinain et al., 1999b). Examination of the 5.8S + ITS rDNA and the LSU rDNA D8-D10 regions in eleven Polynesian isolates proved to be a reliable biogeographic identifier for the various isolates (Babinchak,1996). Isozyme analysis of intraspecific variation among nineteen isolates of G. toxicus from French Polynesia, New Caledonia, and French West Indies revealed biochemically distinct strains. Growth rates and cell sizes varied considerably among the clones; however no relationship was found between the electrophoretic profiles of these isolates and their capacity to produce CTX compounds (Sako et al., 1996). Recently, Parsons (Pers. Comm., 2006) isolated what appear to be two new Gambierdiscus species from Hawaiian waters. Other dinoflagellate genera frequently found in the epiphytic assemblages with Gambierdiscus spp. include Ostreopsis, Coolia, Amphidinium, and Prorocentrum (Steidinger 1993; Parsons and Preskitt, submitted).
In Vitro Growth
Like other dinoflagellates Gambierdiscus spp. are sensitive to excessive agitation, abrupt changes in temperature, salinity, light, and high silicate and metal levels, particularly copper (Bomber et al., 1988; Guillard and Keller, 1984; Durand-Clement, 1987). The modified K-media (Keller and Guillard 1985) and Harrison et al.’s (1980) ESNW are widely used; the latter has higher levels of chelation preventing Fe-deficiency in the medium. Culture is best at 32-35 ppt salinity, and growth is optimal within 26-30°C though possible from 19.5 – 34°C. Strong evidence points to G. toxicus as a shade-loving species (Villareal and Morton, 2002); it prefers low light (<10% surface intensity), and a 12:12 to 14:10 light to day (L:D) cycle. G. toxicus is a slow growing species (i.e., maximum growth rates ~0.3-0.5/d, Bomber et al., 1988).
G. toxicus produces mucoid sheaths that aid their epiphytic life history, but complicate in vitro culture. The degree of toxicity has frequently been shown to be variable. Toxicity has been seen to increase with increasing temperature and light, and in culture, toxicity has frequently been observed to be highest in mid to late log phase. There is some evidence for an inverse relationship between toxicity and chlorophyll content. Because of the slow growth and the fact that CTX production may well be out-of-phase from optimal growth conditions, it may take considerable time (e.g., weeks) for the actual toxin production to be normalized to a given set of environmental conditions under evaluation. Large scale culture has been performed by several investigators (Babinchak et al., 1994), and G. toxicus has continued to produce CTX in culture. It has also been regularly reported to stop producing toxin (presumably CTX) under in vitro culture conditions.
It is unclear why dinoflagellates produce potent toxins, how these secondary metabolites are synthesized, or why G. toxicus may cease production of CTX when grown in vitro. Although this may in part reflect the limitations of the chemistry employed, it may also be that the provision of optimal growth conditions in vitro eliminates the stress/austerity in the growth environment that triggers toxin production. The challenges of maintaining Gambierdiscus spp. cultures, the limited concentrations of these targeted secondary metabolites, their ephemeral production in vitro, and possibly the role of viable but non-cultivable bacteria have limited significant CTX production from culture facilities.
In Vivo Growth
Gambierdiscus spp. are found in coastal tropical and subtropical waters within the 35°N - 35°S band of all oceans. The preferred habitat is typically not high energy areas, but rather quiet, high-salinity leeward areas with minimum fresh-water runoff, and at depths where cells can chromatically adapt to low light levels (Steidinger and Baden, 1985; Taylor 1985). Application of PAM fluorometry to in vivo cells was used to show that G. toxicus exploits the three-dimensional structure of the macroalgal host to achieve shading necessary to permit these high-light intolerant organisms to thrive in shallow, high-light tropical coastal waters (Villareal and Morton, 2002). G. toxicus shows strong association with macroalgae, but may also be found in detritus and/or on dead coral surfaces. Normal cell densities range from 1-50 cells/g algae; densities >1000 cells/g algae are considered “high”, and the maximum abundance ever recorded (~450,000 cells/g algae) was off Gambier Island (Bagnis et al., 1988). It is far from clear whether increased amounts of toxic fish that may occur from time to time are due to increased G. toxicus biomass, or increased specific toxicity of the biomass (i.e., higher CTX content or composition).
Among the hypothesized causes for the observed spatial and/or temporal variations in toxicity are variations in G. toxicus abundance, specific toxicity, clonal participants, or some combination of these. There exists some direct and indirect evidence for each. It is not known whether the increased toxicity in stationary phase is due to increased toxin production, increased toxin flux to the foodweb, or merely a conversion of a toxin congener to more potent congeners. Studies in the Virgin Islands showed no seasonality in either the number of CFP cases (Morris et al.,1982) or the abundance of G. toxicus (Bomber, 1985), whereas coincident seasonality in both CFP cases and G. toxicus abundance appear evident in Hawaii (Nakata, Pers. Comm., 2006; Parsons, Pers. Comm., 2006). In the Florida Keys, seasonal minima in G. toxicus abundance correlated with minimum winter water temperatures (Bomber et al., 1988). Weekly sampling off Tahiti from 1993-1997 showed cell densities frequently >1000 cells/g, and attaining maximum abundance at the beginning and end of the hot season. A noticeable increase in peak densities (~5000 cells/g) was preceded by unusually high water temperatures severe enough to cause a serious coral-bleaching episode (Chinain et al., 1999a). No correlation was found between the toxicity of these G. toxicus blooms, and their biomass or the seasonal temperature patterns. In an impressively extensive database from 1993-2001, Chateau-Degat et al., (2005) correlated G. toxicus densities, seawater temperatures and reported CFP cases, and concluded that peak cell densities lagged peak temperatures by 13-17 months, and reported CFP cases lagged peak cell densities by 3 months, at least in French Polynesia.
Early CFP incidents in environmentally disturbed areas led to speculation that anthropogenic disturbances (e.g., shipwrecks, dredging, etc.) or natural disturbances (e.g., cyclones, hurricanes, tsunamis, coral bleaching) triggered CTX incidence (Randall, 1958; Steidinger and Baden, 1985; Bagnis, 1994). Despite a low correlation rate between such disturbances and the onset of CFP, this remains a common perception, and for that reason is addressed here (Banner, 1974; Kohler and Kohler, 1992). The rationale for this causation theory is that disturbances that create new surfaces create new substrate for colonization by algal turf that could theoretically provide additional macroalgal colonization, and thus associated epiphytes which may grow on such algae (i.e., Randall’s “new surface theory”) (Randall 1958; Quod et al., 2001). In general, available analyses of areas affected by natural or man-made perturbations have not shown casual linkages to changes in CFP incidence that could be isolated from natural, periodic or stochastic processes (Brusle, 1998).
An intriguing paradigm, recently introduced to the literature (Cruz-Rivera and Villareal, 2006) focuses attention on survival strategies of the macroalgal symbionts of Gambierdiscus spp. to explain the persistence of these slow growing species and the stochastic nature of increases in cell abundances and/or CFP incidence. Various algal hosts may tolerate/manage the high rates of herbivory that are characteristic on reef environments by growing fast, having poor nutritional quality, and/or using chemical or structural defenses. The macroalgal hosts exhibiting these various strategies likewise represent drastically different fluxes of gambiertoxins into herbivorous grazers and subsequently to the foodweb. Similarly, the relatively inedible algal species are seen to serve as refuges for the slow-growing Gambierdiscus spp., which may well attain elevated densities, but serve primarily to re-seed the algae that are more heavily grazed. In this way the CTX flux potential to the foodweb is enhanced. By requiring that the ecology of both Gambierdiscus spp. and their host algae are essential to causing patterns of CFP, this paradigm provides an explanation for the lack of correlation between Gambierdiscus spp. abundances, frequencies of CTX-positive fish, and CFP in humans.
Symbiotic Relationships
G. toxicus have been associated with foliose or filamentous red, brown, green and even blue-green algae, especially those genera with structural interstices, e.g., Ceramium, Chondria, Wrangelia, Gelidium, Amphiroa, Sargassum, Padina, Turbinaria, Caulerpa, Pseudobryopsis, Hypnea, Gracilaria, Jania, Halimeda, Colpomenia, Spyridea, Cladophora sp., benthic detritus, Heterosiphonia, and Acanthophora(Bomber et al., 1988; Gillespie et al., 1988; Carlson and Tindall, 1988; Cruz-Rivera and Villareal, 2006) . There is some evidence that various macroalgae may supply important growth factors (Carlson et al., 1984; Nakahara et al., 1996; 1987). Nonetheless, the occurrence of G. toxicus on many macroalgae species suggests opportunism in regard to its macroalgal substrate, and a low likelihood that a particular algal metabolite regulates its abundance.
Macroalgae are leaky and release a variety of substances from polysaccharides to complex enzymes, and the derivation of chelating agents from the macroalgae has been suggested as a strategy of G. toxicus to mitigate its hypersensitivity to Cu and other metals (Bomber 1987; Bomber et al., 1988). The apparent benefits to the epiphytic Gambierdiscus that are received from the association with macrobiota are location fixation, higher nutrient availability, shading from direct sunlight, protection from turbulence, and access to organic compounds within the thallisphere (Villareal and Morton, 2002). For slowly growing organisms that cannot compete via reproductive rates, and are relatively intolerant of variations in physico-chemical conditions, these benefits from the symbiosis seem to be significant. Although allelopathic functions in the thallisphere that are CTX-specific are largely unknown, the capability for extracts of epiphytic dinoflagellates to suppress growth of other dinoflagellates (Sakamoto et al., 1996; Fistarol et al., 2004; Kubanek et al., 2005) has led to the suggestion that the diversity of toxins synthesized by dinoflagellates play a role in competition among algal species of the thallisphere.
The complex chemical structure of CTX suggests that multiple biochemical steps are required for its synthesis (Murata et al., 1989; Plumley, 1997; Snyder et al., 2003). This, together with the fact that the CTX molecule resembles a polyketide, a class of secondary metabolites known to be produced by other microorganisms (e.g., bacteria, filamentous fungi), has led to speculation that other members of the symbiotic microbial consortium may participate in the production of CTX. Some species of dinoflagellates have been shown to contain bacteria in the cytoplasm and/or nucleus (Ashton et al., 2003), and it has been suggested that dinoflagellate toxins may originate from bacteria (or viral recombinant RNA). In bacteria, the polyethers often occur as a complex of closely related compounds, as is the case with dinoflagellate polyethers such as CTX. Efforts to evaluate toxin production by bacteria associated with dinoflagellate cultures have been done, but the meaning of negative data is not clear since it could simply mean that the proper environment for toxigenesis was not provided. This hypothesis has been supported by knowledge that a very potent nonproteinaceous marine toxin (palytoxin) is actually produced by an endocellular bacterium associated with zoanthid polyps (Moore et al., 1982; Steidinger and Baden, 1985; Sakami et al., 1999). The fact that only a fraction of marine bacteria are cultivable and that CTX production has been seen to cease over time in dinoflagellate cultures has been taken to be supportive of this microbial symbiont hypothesis. However, the failure of TEM analysis to show intracellular bacteria in a very toxic dinoflagellate strain, or good correlation between toxicity and presence of intracellular bacteria in multiple samples, leaves this intriguing bacteria theory lacking empirical support from well-designed experiments.
Though not prominent in the literature, it is not unreasonable to suspect endocellular viruses of participating/inducing the production of toxic secondary metabolites in toxic dinoflagellates (Plumley, 1997). Only in 1977 did studies show that pelagic bacteria were very abundant, and a decade later that viral densities 10X bacterial densities were common (Azam and Worden, 2004). Viral studies have shown species specificity, host density-dependence, and a variety of behaviors potentially influential to the CTX dynamics in the thallisphere (Brussaard et al., 2004; Tarutani et al., 2000). At present, there are only a few phytoplankton viruses in culture, and most of these infect HAB species. The primitive origin of both dinoflagellates and viruses implies the potential for a co-evolution relevant to CTX dynamics. Just as viruses aid in the mitigation of pelagic bloom “hot spots”, it has been speculated that they may play a role in controlling hot spots in the sessile, epiphytic realm. Studies where viruses were shown to reduce the density of bloom-forming phytoplankton have also shown that the surviving cells were resistant to most of the virus clones. This implies a change in the properties of the dominant cells pre- and post- hot spot in time. Thus, viruses’ capability to quantitatively and qualitatively affect pelagic phytoplankton populations has suggested that they may similarly influence temporal and/or spatial changes in toxicity of epiphytic dinoflagellates.
Issues and Questions
Due to the molecular complexity of the ciguatoxins, it seems unlikely that these secondary metabolites are merely waste products that provide no benefit to the dinoflagellates. Clearly, the biosynthesis of CTX did not evolve as a means to harm humans. The bacterial literature suggests that such compounds may be allelochemic agents directed against closely competing or pathogenic organisms. Many such algal, dinoflagellate and bacterial compounds have been shown to influence the growth of organisms in their vicinity, including inhibiting the growth of diatoms. Their role in allelopathic competition for space in the thallisphere seems a reasonable speculation, but awaits improved understanding of the eco- physiological benefits of toxin production to the dinoflagellates.
Given the extreme potency of CTX and its precursors, the negligibly low incidence of fatality in humans is notable. Although there have been reports of abnormal fish behavior associated with CFP fish (Yasumoto et al., 1987), the toxins produced by G. toxicus have shown limited evidence of ichythiyotoxicity (Capra et al., 1988; Lewis, 1992; Coleman et al., 2004). One wonders whether an ecological factor such as enhanced predation on heavily intoxicated fish constrains the accumulation of CTX in fish so that the toxin dose within a reasonably-sized portion of fish consumed by humans is below sub-lethal levels.
In the future, prevention of ciguatera fish poisoning is most likely to result from the application of new knowledge at the fish--human interface. Improved assays to expeditiously assess marine toxins are needed by clinicians, fishermen, seafood distributors, restaurateurs and food safety regulators, and appear to be tractable, based on advances of technologies now under development.
Improved understanding of how and why environmental factors control toxin production will require good data on the effects of environmental conditions on the growth and toxin production by Gambierdiscus spp., improved information on the genetics and biochemical pathways of toxin biosynthesis, and development of a molecular understanding of the genes involved in toxin synthesis.
Marine organisms are a rich, largely untapped source of a wide variety of biologically important secondary metabolites (Faulkner 2002; Snyder et al., 2003). The ability to manipulate the pathways for secondary metabolites has stimulated interest in marine organisms for novel biosynthetic functions. Extracts from cultured CTX-producing dinoflagellates are potential sources of biomedically useful compounds. These polyethers have proven antifungal and anti-neoplastic properties. As a group, the polyether class of antibiotics is known to improve food conversion in ruminant animals, and to have usefulness in molecular probes for the study of essential ligand-receptor interactions in living systems. In the biomedical research area, these dinoflagellate bio-products could be used to enhance or inhibit nerve conduction through the voltage-sensitive sodium channel, as well as other biomedical research venues.
AcknowledgementS
This publication was made possible through the Centers for Oceans and Human Health (COHH) program, of the National Institute of Environmental Health Sciences (P50ES012740), National Institutes of Health, and the National Science Foundation (OCE04-32479).
References
Adachi, R., Fukuyo, Y., 1979. The thecal structure of a marine toxic dinoflagellate Gambierdiscus toxicus gen. et sp. nov. collected in a ciguatera endemic area. Bull. Jpn. Soc. Sci. Fish. 45, 67-71.
Ashton, M., Rosado, W., Govind, N.S., Tosteson, T.R., 2003. Culturable and nonculturable bacterial symbionts in the toxic benthic dinoflagellate Ostreopsis lenticularis. Toxicon 42, 419-424.
Anderson, D. M. and P. S. Lobel., 1987. The Continuing Enigma of Ciguatera. Biol. Bull. 172, 89-107.
Arena, P., Levin, B., Fleming, L.E., Friedman, M.A., Blythe, D.G., 2004. A pilot study of the cognitive and psychological correlatyes of chronic ciguatera poisoning. Harmful Algae 3, 51-60.
Arias, H. R., 2006. Marine Toxins targeting ion channels. Mar. Drugs. 4, 37-69
Azam, F., and A. Z. Worden., 2004. Microbes, molecules and marine ecosystems. Science, pp. 1622-1624.
Babinchak, J. A., Doucette, G.J., Ball, R.M., 1996. Partial characterization of the LSU rRNA gene from the ciguatoxic dinoflagellate, Gambierdiscus toxicus. Harmful and Toxic Algal Blooms Pp. 459-462.
Baden, D., Fleming, L.E., Bean, J.A., 1995. Marine Toxins. In: Handbook of Clinical Neurology: Intoxications of the Nervous System Part II. Natural Toxins and Drugs. FA deWolff (Ed). Amsterdam: Elsevier Press, 21(65),141-175.
Bagnis, R., 1977. Modalites evolutives et biogenese de la ciguatera en Polynesie francaise. Ph.D. dissertation, Universite de Bordeaux (France), 134 pp.
Bagnis, R., 1994. Natural versus anthropogenic disturbances to coral reefs -- comparison in epidemiological patterns of ciguatera. Mem. Queensld. Mus. Brisbane 34, (3), 455-460.
Bagnis, R., Bennett, J., Prieur, C., Legrand, A.M., 1985. The dynamics of three toxic benthic dinoflagellates and the toxicity of ciguateric surgeonfish in French Polynesia. In: Anderson, D.M., White, A.W., Baden, D.G. (Eds.), Toxic Dinoflagellates. Elsevier Science Publishing Co., New York, pp. 177-182.
Bagnis, R., 1977. Modalites evolutives et biogenese de la ciguatera en Polynesie francaise. Ph.D. dissertation, Universite de Bordeaux (France), 134 pp.
Bagnis, R., 1994. Natural versus anthropogenic disturbances to coral reefs -- comparison in epidemiological patterns of ciguatera. Memoirs of the Queensland Museum. Brisbane 34, (3), 455-460.
Bagnis, R.A., Barsinas, M., Prieur, C., Pompon, A., Chungue, F., Legrand, A.M., 1987. The use of the mosquito bioassay for determining the toxicity to man of ciguateric fish. Biol. Bull. 172, 137-143.
Bagnis, R., Spiegel, A., Boutin, J.P., Burucoa, C., Nguyen, L., Cartel, J.L., Capdevielle, P., Imbert, P., Prigent, D., Gras, C., Roux, J., 1992. Evaluation de l’efficacité du mannitol dans le traitement de la ciguatera en Polynésie Française. Médecine Tropicale 52, 67-73.
Ballantine, D.L., Bardales, A.T., Tosteson, T.E., 1985. Seasonal abundance of Gambierdiscus toxicus and Ostreopsis sp. in coastal waters of southwest Puerto Rico. In: Delesalle, B., Galzin, R., Salvat, B. (Eds.), Proceedings of the Fifth International Coral Reef Congress, Tahiti, Antenne Museum-EPHE, Moorea (French Polynesia), pp. 417-422.
Banner, A.H., 1974. The biological origin and transmission of ciguatoxin. In: Bioactive Products from the Sea, ed. by H. J. Hun and C. E. Lance, pp. 15-36. Marcel Dekker, New York.
Banner, A.H., 1976. Ciguatera: a disease from coral reef fish. IN: Bioactive Products from the Sea, H. J. Humm and C. E. Lane, eds. Marcel Dekker, New York. pp 177-215.
Banner, A. H. and P. Helfrich, 1964. The distribution of ciguatera in the tropical Pacific. Tech. Rep. Hawaii Mar. Lab. 3,1-48.
Banner, A.H., Helfrich, P., Piyakarnchana, T., 1966. Retention of ciguatera toxin by the red snapper, Lutjanus bohar. Copeia 2, 297-301.
Banner, A. H., Sasaki, S., Helfrich, P., Alender. C., Scheuer. P., 1960. A new bioassay method for toxins from fish. Nature. 189, 229-230.
Bomber, J.W., Guillard, R.R.L., Nelson, W.G., 1988. Roles of temperature, salinity, and light in seasonality, growth, and toxicity of ciguatera-causing Gambierdiscus toxicus Adachi et Fukuyo (Dinophyceae). Journal of Experimental Marine Biology and Ecology 115, 53-65.
Blythe, D.G., De Sylva, D.P., Fleming, L.E., Ayyar, R.A., Baden, D.G., Shrank, K., 1992. Clinical experience with IV mannitol in the treatment of ciguatera. Bulletin de la Société de Pathologie Exotique, 85, 425-426.
Blythe, D.G., Hack, E., Washington, G., Fleming, L.E., 2001.The Medical Management of Seafood Poisoning. Hui, Kits, Stanfield. Seafood and Environmental Toxins. Marcel Dekker, pg. 311-319.
Brusle, J., Brusle-Sicard, S., Fourcault, B., 1998. Gonads of groupers (Serranidae from Mururoa and Hao (French Polynesia) used as indicators of the quality of the coral reef environment: histological and ultrastructural aspects. Cybium 22, 371-382.
Brussaard, C.P. D., Short, S.M., Frederickson, C.M., Suttle, C.A., 2004. Isolation and phylogenetic analysis of novel viruses infecting the phytoplankton Phaeocystis globosa (Prymnesiophyceae). Applied and Envir. Microbiol. 70, 3700-3705.
Cameron, J., Flowers, A.E., Capra, M., 1991. Electrophysiological studies on ciguatera fish poisoning in men (Part II). J Neurol. Sci. 101,93-97.
Capra, M.F., Cameron, J., Flowers, A.E., Coombe, I.F., Blanton, C.G., Hahn, S.T., 1988. The effects of ciguatoxin on teleosts. Proceedings of the 6th International Coral Reef Symposium, Australia 1988, vol. 3., pp. 37-41.
Carlson, R.D., 1984. Distribution, periodicity, and culture of benthic/epiphytic dinoflagellates in a ciguatera endemic region of the Caribbean. Ph.D. thesis. Southern Illinois University, Carbondale, Illinois.
Carlson, R.D., Morey-Gaines, G., Tindall, D.R., Dickey, R.W., 1984. Ecology of toxic dinoflagellates from the Caribbean Sea: Effects of macroalgal extracts on growth in culture, p. 272-287. In: Ragelis, E.P. (Ed.). Seafood Toxins, Amer. Chem. Soc. Washington, D.C.
Carlson, R. D., Tindall, D.R., 1988. Distribution and Periodicity of Toxic Dinoflagellates in the Virgin Islands, In Toxic Dinoflagellates , D. B. Anderson Editor. 1985, Proc. 3rd Intl Conf. Toxic Dinoflagellates: New Brunswick.
Cooper, M.J., 1964. Ciguatera and other marine poisoning in the Gilbert Islands. Pacific Science 18, 411-440.
Colman, J.R., Dechraoui, M.-Y. B., Dickey, R.W., Ramsdell, J.S., 2004. Characterization of the developmental toxicity of Caribbean ciguatoxins in finfish embryos. Toxicon 44, 59-66.
Chan, T. Y. K., Wang, A.Y.M., 1993. Life-threatening bradycardia and hypotension in a patient with ciguatera fish poisoning. Trans. R. Soc Trop. Med. Hyg. 87, 71-7.
Chateau-Degat, M.L., Chinain, M., Cerf, N., Gingras, S., Hubert, B., Dewailly, E., 2005. Seawater temperature, Gambierdiscus spp. variability and incidence of ciguatera poisoning in French Polynesia. Harmful Algae 4, 1053-1062.
Chinain, M., Germain, M., Deparis, X., Pauillac, S., Legrand, A.M., 1999a. Seasonal abundance and toxicity of the dinoflagellate Gambierdiscus spp. (Dinophyceae), the causative agent of ciguatera in Tahiti, French Polynesia. Marine Biology 135, 259-267.
Chinain M., Faust M.A., Pauillac S., 1999b. Morphology and molecular analyses of three toxic species of Gambierdiscus (Dinophyceae): G. pacificus, sp. nov., G. australes, sp. nov., and G. polynesiensis, sp. nov. J. Phycol. 35, 1282-1296.
Cruz-Rivera, E.,Villareal, T.A., 2006. Macroalgal palatability and the flux of ciguatera toxins through marine food webs. Harmful Algae 5, 497-525.
De Haro, L., Hayek-Lanthois, M., Jooson, F., Castanier, L., Jouglard, J., 1997. Mediacal management in the Marseilles Poison Centre of a ciguatera poisoning collective case after a meal of one barracuda in Mexico. Toxicon. 35, 810.
Dickey, R. W., H. R. Granade, McClure, F.D., 1994. Evaluation of a solid-phase immunobead assay for detection of ciguatera-related biotoxins in Caribbean finfish. Memoirs of the Queensland Museum.Brisbane 34, 481-488.
Dickey, R., Jester, E., Granade, R., Mowdy, D., Moncreiff, C., Rebarchik, D., Robi, R., Musser, S., Poli, M., 1999. 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. Natural Toxins 7, 157-165.
Durand-Clement, M., 1986. A study of toxin production by Gambierdiscus toxicus in culture. Toxicon 24, 1153-1157.
Ebesu, J. S. M., 1998. Isolation and Characterization of Novel Ciguateric Compounds from Acanthurus triostegus (Manini). Ph.D. Dissertation, University of Hawaii, Honolulu, Hawaii.
Escalona de Motta, G., De la Noceda, G.G., 1985. Epidemiological, clinical and experimental aspects of ciguatera in Puerto Rico. Proc. 5th Intl Coral Reef Congress, Tahiti (French Polynesia). Vol. 4, Pp. 415-416.
Fistarol, G. O., Legrand, C., Selander, E., Hummert, C., Stolte, W., Graneli, E., 2004. Allelopathy in Alexandrium spp.: Effect on a natural plankton community and on algal monocultures. Aquatic Microbial Ecology 35, 45-56.
Fleming, L., E. Dewailly, Baden, D.G., 2000. The epidemiologic of marine harmful algal blooms, Epidemiology. 11, 143.
Friedman, M.A., Arena, P., Levin, B., Fleming, L.E,, Weisman, R., Bernstein, J., Schrank, K., Blythe, D., Backer, L., Reich, A, In Press. Neuropsychological Study of ciguatera fish poisoning: A longitudinal case-control study. Archives Clinical Neurology.
Gillespie, N. C., M. J. Holmes, J. Burke, Doley, J., 1985. Distribution and periodicity of Gambierdiscus toxicus in Queensland, Australia. In: D. M. Anderson, A. White and D. G. Baden, eds. Toxic Dinoflagellate, Elsevier, London. Pp. 183-188.
Guillard, R. R .L., Keller, M.D., 1984. Culturing Dinoflagellates. In: Dinoflagellates. Academic Press. Inc., New York. Pp. 391-441.
Halstead, B. W., 1967. Poisonous and Venomous Marine Animals of the World. Vol. II. U. S. Government Printing Office. 1070pp.
HARRNESS, 2005. Harmful Algal Research and Response: A National Environmental Science Strategy 2005–2015. Ramsdell, J.S., Anderson, D.M., Glibert, P.M. (Eds.), Ecological Society of America, Washington, DC.
Harrison, P.J., Waters, S.E., Taylor, F.J.R., A broad spectrum artificial seawater medium for coastal and open ocean phytoplankton. Journal of Phycology 16, 28-35.
Helfrich, P., 1964. Fish Poisoning in Hawaii. Haw. Med J. 22,361-372.
Helfrich, P., Banner, A.H., 1963. Experimental induction of ciguatera toxicity in fish through diet. Nature 197, 1025-1026.
Helfrich, P., Banner, A.H., 1968. Ciguatera fish poisoning. 2. General patterns of development in the Pacific. Occasional Papers of the Bernice P. Bishop Museum 23, 371-382.
Hokama, Y., Banner, A. H., Boylan, D.B., 1977. A radioimmunoassay for the detection of ciguatoxin. Toxicon 15, 317-325.
Hokama, Y., Abad, M.A., Kimura, L.H., 1983. A rapid enzyme-immunoassay for the detection of ciguatoxin in contaminated fish tissues. Toxicon 21, 817-824.
Hokama, Y., Shirai, L.K., Kurosawa, M.N., Iwamoto, L.M., Goto, C.S., Osugi, A.M., 1987. Assessment of a rapid enzyme immunoassay sitck test for the detection of ciguatoxin and related polyether toxins in fish tissues. Biol. Bull. 172, 144-153.
Hokama, Y., Nishimura, K., Takenaka, W., Ebesu, J.M., 1998. Simplified solid-phase membrane immunobead assay (MIA) with monoclonal anti-ciguatoxin antibody (MAb-CTX) for detection of ciguatoxin and related polyether toxins. Journal of Natural Toxins 7, 1-22.
Hokama, Y., Wachi, K.M., Shiraki, A., Goo, C., Ebesu, J.S.M., 2001. The biological assessment of flora and fauna as standards for changes in the nearshore ocean environment: a study of Barbers Point Harbor. Journal of Natural Toxins 10, 57-68.
Hokama, Y., Yoshikawa-Ebesu, J.S.M., 2001 Ciguatera Fish Poisoning: a Foodborne Disease J. Toxicol.—Toxin Reviews 20(2), 85-139.
Holmes, M. J., 1998. Gambierdiscus yasumotoi sp. nov. (Dinophyceae), a toxic benthic dinoflagellate from Southeastern Asia. Journal of Phycology 34, 661-668.
Holmes, M.J., Lewis, R.J., 1994. The origin of ciguatera. Mem. Queensld. Mus. 34, 497-504.
Holmes, M. J., Lewis, R. J., Poli, M. A., Gillespie, N.C., 1991. Strain dependent production of ciguatoxin precursors (gambiertoxins) by Gambierdiscus toxicus (Dinophycean) in culture. Toxicon 29, 761-775.
Inoue, M., 2004. Convergent Synthesis of Polycyclic Ethers. Illustrations of the utility of acetal-linked intermediates. Org. Biomol. Chem. Vol. 2, 1811-1817.
Inoue, M., 2005. Convergent Strategies for Synthesis of trans-funsed Polycyclic Ethers. Chem. Rev. 105, 4379-4405.
Inoue, M ., Ishihara, Y., Tamashita, S., Masahiro, M., 2006. Convergent Assembly of Polycyclic Ethers via Acyl Radical Addition to Unactivated Enol Ether. Organic Letters 8, 5801-5804.
Kodama, M., 1990. Production of paralytic shellfish toxins by a bacterium Moraxellasp. isolated from Protogonyaulax tamerensis. Toxicon. 28,707-714.
Keller, M. D., Guillard, R.L.L., 1985. Factors significant to marine dinoflagellate culture. In: Toxic Dinoflagellates, ed. D. M. Anderson, A. W. White, and D. G. Baden, pp. 113-116. New York: Elsevier.
Kohler, S.T., Kohler, C.C., 1992. Dead bleached coral provides new surface for dinoflagellates implicated in ciguatera fish poisonings. Environ. Biol. Fish. 35,413-416.
Kubanek, J., Hicks, M.K., Naar, J., Villareal, T.A., 2005. Does the red tide dinoflagellate Karenia brevis use allelopathy to outcompete other phytoplankton? Limnology and Oceanography 50, 883-895.
Lawrence, D. N., Enriquez, M.B., Lumish, R.M., 1980. Ciguatera fish poisoning in Miami. J. Am. Med. Assoc. 244, 254-258.
Labrousse, H., Pauillac, S., Jehl-Marinex, C., Legrand, A.M., Avrameas, L., 1991. In vivo and in vitro techniques for the detection of ciguatoxin. Oceanic 18, 182-245
Legrand, A.-M., 1998. Ciguatera toxins: Origin, transfer through the food chain and toxicity to humans. In: Harmful Algae, ed. B. Reguera, J. Blanco, M. L. Fernández, and T. Wyatt., pp. 39-43. Vigo, Spain: Xunta de Galicia and IOC.
Lewis, R.J., 1992. Ciguatoxins are potent ichthyotoxins. Toxicon 30, 207-211.
Lewis, R.J., 2001. The changing face of ciguatera. Toxicon 39, 97-106.
Lewis, R. J., Jones, A., Vernoux, J.P., 1999. HPLC/Mass spectrometry and HPLC/MS/MS methods for the detection of multiple ciguatoxins. Proceedings of the 5th Indo-Pacific Fish conference, Noumea – New Caledonia Pp. 739-744.
Lewis, R.J., Holmes, M.J., Sellin, M., 1994. Invertebrates implicated in the transfer of gambiertoxins to the benthic carnivore Pomadasys maculates. Memoirs of the Queensland Museum 34, 561-564.
Lewis, R.J., Jones, A., Vernoux, J. P., 1999. Societe Francaise d' Ichtyologie, Paris (France), and Institut de Recherche pour le Developpement, Paris (France). HPLC/Mass spectrometry and HPLC/MS/MS methods for the detection of multiple ciguatoxins. 5. Indo - Pacific fish conference, Noumea (New Caledonia), 3-8 nov 1997., edited by B. Seret, J. Y. Sire. Paris (France): Societe Francaise d' Ichtyologie.
Lewis, R. J., Jones, A., 1997. Characterization of ciguatoxins and ciguatoxin congeners present in ciguateric fish by gradient reverse-phase high-performance liquid chromatography/mass spectrometry. Toxicon 35, 159-168.
Lobel, P. S., 1979. Folk remedies for tropical fish poisoning in the Pacific. Sea Frontiers 25, 239-245.
Luckas, B., 2000. Phycotoxins in the marine environment: control of marine organisms for contamination with algal toxins. Int. J. Environ. Pollut. 13, 148-172.
Manger, R.L., Leja, L.S., Lee, S.Y., Hungerford, J.M., Hokama, Y., Dickey, R.W., Granade, R.H., Lewis, R., Yasumoto, T., Wekell, M., 1995. Detection of sodium channel toxins: directed cytotoxicity assays of purified ciguatoxins, brevetoxins, saxitoxins, and seafood extracts. Journal of AOAC International 78, 521-527.
McKee, D. B., Fleming, L.E., Tamer, R., Weisman, R., Blythe, D., 2001. Physician diagnosis and reporting of ciguatera fish poisoning in an endemic area. Harmful Algal Blooms 2000. 451-453.
Murata, M., Legrand, A.M., Ishibashi, Y., Yasumoto, T., 1989. Excretion of ciguatoxin from moray eels (Muraenidae) of structure of ciguatoxin and its congener. J. Am. Chem. Soc. 111, 8929-8931.
Nakahara, H., Sakami, T., Chinain, M., Ishida, Y., 1996. The role of macroalgae in epiphytism of the toxic dinoflagellate Gambierdiscus toxicus (Dinophyceae). Phycological Research 44, 113-117.
Nicholson, G. M., Lewis, R.J.., 2006. Ciguatoxins: Cyclic Polyether Modulators of Voltage-gated Iion Channel Function. Marine Drugs 4, 82-118.
Nukina, M., Koyanagi, I.M., Scheur, P.J., 1984. Two interchangeable forms of ciguatoxin. Toxicon. 22 ,169-176
Nukina, M., Tachibana, K., Scheurer, P.J., 1986. Recent developments in ciguatera research. In: Natural Toxins, Animal, Plant and Microbial. J.B. White. ed. Clarendon Press, Oxford. 45-55
Ochoa, J., Sanchez-Paz, L.A., Crus-Villacorta, A., Nunez-Vazquez, E., 1996. Toxic events in the Northwest Pacific coastline of Mexico during 1992-1995. Hydrobiologia 352, 195-200.
Palafox, N. A., Jain, L.G., Piezano, A.Z., 1988. Successful treatment of ciguatera fish poisoning with intravenous mannitol. JAMA 259, 2740-2742.
Parsons, M.L., Preskitt, L.B., in press. A survey of epiphytic dinoflagellates from the coastal waters of the island of Hawai‘i. October.
Pearn, J., 2001. Neurology of ciguatera. J. Neurol. 70, 4-8.
Plumley, F. G., 1997. Marine Algal Toxins: Biochemistry, Genetics, and Molecular Biology. Limnol. Oceanogr., 42, 1252-1264.
Pohland, A. E., Dowell, V.R., Richard, J.L., (eds.), 1990. Microbial Toxins in Foods and Feeds: Cellular and Molecular Modes of Action. Plenum Press, New York. 566pp.
Pottier, I., Vernoux, J.P., Jones, A., Lewis, R.J., 2002. Characterization of multiple Caribbean ciguatoxins and congeners in individual specimens of horse-eye jack (Caranx latus) by high-performance liquid chromatography/mass spectrometry. Toxicon 40, 929-939.
Quod, J-P., Turquer, J., Conejero, S., Ralijaona, C., 2001. Ciguatera risk assessment in the Indian Ocean following the 1998 coral bleaching event. Pp. 166-168. CORDIO. SARREC. Mar. Sci. Prgm. Status Report. D. Souter, D. Obura, and O. Linden (eds).
Quod, J. P., Turquer, J., 1996. Ciguatera in Reunion Island (SW Indian Ocean): Epidemiology and clinical patterns. Toxicon. 34, 779-785.
Randall, J. E., 1958. A review of ciguatera tropical fish poisoning with a tentative explanation of its cause. Bull. Mar. Sci. Gulf Carib. 8, 236-267.
Sakamoto, B. H., 1998. Studies on Gambierdiscus toxicus. PhD dissertation, School of Medicine, University of Hawaii, Honolulu.
Sakami, H., Nakahara, M., Chinain, M., Ishida, Y., 1999. Effects of epiphytic bacteria on the growth of the toxic dinoflagellate Gambierdiscus toxicus (Dinophyceae). L. Exp Mar. Biol. Ecol. 233, 231-246.
Sakamoto, B., Nagai, H., Hokama, Y., 1996. Stimulators of Gambierdiscus toxicus (Dinophycease) growth: the possible role of gambieric acid-A as an endogenous growth enhancer. Phycologia. 35, 350-353.
Sako, Y., Chinain, M., Legreand, A., Nakahara, H., Kuruawa, T.S., Uchida, A., Ishida, Y., Inouye, A., 1996. Phylogentic relationship of ciguatera-causing dinoflagellate Gambierdiscus toxicus with 18s ribosomal DNA sequence comparison. Harmful and Toxic Algal Blooms Pp. 443-446.
Scheuer, P.J., Takahashi, W., Tsutsumi, J., Yoshida, T., 1967. Ciguatoxin, isolation, and chemical nature. Science 155, 1267-1268.
Schnorf, H., Taurarii, M. Cundy, T., 2002. Ciguatera fish poisoning: A double-blind randomized trial of mannitol therapy. Neurology 58, 873-880.
Steidinger, K. A.. 1993. Some taxonomic and biologic aspects of toxic dinoflagellates. Algal Toxins in Seafood and Drinking Water. Academic Press, Ltd., p. 1-28.
Steidinger, K. A., Baden, D.G., 1985. Toxic marine dinoflagellates. Pp. 210-261. In Dinoflagellates. D. L. Spector (ed.), Academic Press, Orlando.
Sumner, J., Ross. T., 2002. A semi-quantitative seafood risk assessment. Int. J. Food Microbiol. 77, 55-59.
Tachibana, K., 1980. Structural Studies in Marine Toxins. Ph.D. Dissertation, University of Hawaii, Honolulu, Hawaii.
Tachibana, K., Nukina, M., Joh, Y-G., Scheuer, P.J., 1987. Recent developments in the molecular structure of ciguatoxin. Biol. Bull. Mar. Biol. Lab. Woods Hole. 172, 122-127.
Tarutani, K., Nagasaki, K., Yamaguchi, M., 2000. Viral impacts on total abundance and clonal composition of the harmful bloom-forming phytoplankton Heterosigma akashiwo. Applied and Envir. Microbiol. 66 ,4916-4920.
Taylor, F.J.R., 1985. The distribution of the dinoflagellate Gambierdiscus toxicus in the eastern Caribbean. In: Proceedings of the fifth international coral reef congress, ed. by B. Delesalle, B. Galzin, Repsilon, D., and B. Salvat, vol. 4, pp. 423-428. Antenne Museum-EPHE, Moorea (French Polynesia).
Tebana, T., 1992. Ciguatera fish poisoning and reef disturbance in South Tarawa, Kiribati. Ciguatera Info. Bull. No. 2, pp 7.
Tindall, D.R., Dickey, R.W., Carlson, R.D., Morey-Gaines, G., 1984. Ciguatoxic dinoflagellates from the Caribbean Sea. In: Ragelis, E.P. (Ed.), Seafood Toxins, American Chemical Society Symposium Series, Washington, DC, pp. 225-240.
Tosteson, T.R., Ballantine, D.L., Winter, A., 1998. Sea surface temperature, benthic dinoflagellate toxicity and toxin transmission in the ciguatera food web. In: Reguera, B., Blanco, J., Fernández, M.L., Wyatt, T. (Eds.), Harmful Algae, Xunta de Galicia and IOC, pp. 48-49.
Vernoux, J.P., 1988. Ciguatera fish poisoning: Epidemiology, toxicology, and prevention of the illness on Saint-Barthelemy Island, French West Indies. Oceanologica acta. 11, 37-46.
Vernoux, J-P., Lejeune, J., 1993. Ciguatera in the French West Indies. Mem. Queensl. Mus. 34, 631-638.
Villareal, T. A., Morton, S.L., 2002. Use of cell-specific PAM-fluorometry to characterize host shading in the epiphytic dinoflagellate Gambierdiscus toxicus. Marine Ecology 23, 127-140.
Watters, M. R., 2007. Marine Neurotoxins as a Starting Point to Drugs. In: Seafood and Freshwater Toxins. Pharmacology, Physiology and Detection, 2nd Edition. L. M. Botana (ed). Taylor & Francis (CRC Press) publishers
Yasumoto, T., Nakajima, I., Bagnis, R., Adachi, R., 1977. Finding of a dinoflagellate as a likely culprit of ciguatera. Bull. Jpn. Soc. Sci. Fish. 43, 1021-1026.
Yasumoto, T., Inoue, A., Bagnis, R., 1979. Ecological survey of a toxic dinoflagellate associated with ciguatera. In: Taylor, D.L., Seliger, H. (Eds.), Toxic Dinoflagellate Blooms, Elsevier, New York, pp. 221-224.
Yasumoto, T., Nakajima, I., Oshima, Y., Bagnis, R., 1979b. a new toxic dinoflagellate found in association with ciguatera. Pp. 65-70 ub Toxic Dinoflagellate Blooms. D. L. Taylor and H.H. Seliger, eds. Elsevier, New York.
Yasumoto, T., Oshima, Y., Murakami, Y., Nakajima, I., Bagnis, R., Fukuyo, Y., 1980. Toxicity of benthic dinoflagellates. Bull. Jap. Soc. Fish. 46, 327-331.
Yasumoto, T., Raj, U., Bagnis, R., 1984. Seafood Poisoning in Tropical Regions. Lab. Food Sci. Hyg., Faculty of Agriculture, Tohoku, Japan, 1-74.
Yasumoto, T., Murata, M., 1993. Marine toxins. 93, 1897-1907.
Yasumoto, T., 1985. Recent progress in the chemistry of dinoflagellate toxins Pp. 259-270 in Toxic Dinoflagellates. D. M. Anderson, A. W. White, and D. G. Baden, eds. Elsevier, New York.