Cyanobacterial (blue-green algae) toxinsBirgit Puschner and Amber Roegner
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INTRODUCTION
Freshwater cyanobacterial blooms hold the potential to significantly impact the health of both animal and human populations utilizing surface waters for drinking water, daily living, and recreation worldwide. In addi-tion to the often visually stunning nature of these prolif-erations of cyanobacterial species, the blooms can result in the production of a variety of compounds, from malo-dorous ones that affect the taste of the water to dermal and gastrointestinal irritants and severe neurotoxicants, gastrointestinal toxicants, and hepatotoxicants. Among the 2000 species identified through morphological cri-teria, more than 80 are known to be toxigenic, and as assays for detection and toxicity continue to improve, this number will continue to grow. George Francis first reported a toxigenic bloom in the journal Nature in 1878. He reported a “poisonous Australian lake” with “a thick scum like green oil paint” and vividly described acute intoxications of sheep, horses, dogs, and pigs. Analysis of archaeological evidence coupled with evolving under-standing of modern blooms have begun to implicate the role of cyanotoxin poisoning in more widespread mam-malian die-offs dating back to the Pleistocene age (i.e., approximately 150,000 years BC; Braun and Pfeiffer, 2002), and even a controversial hypothesis about the role of cyanobacteria in the various mass extinction events has begun to emerge (Castle and Rodgers, 2009).
Since Francis’ publication in 1878, numerous case reports describing animal morbidity and mortality after exposure to cyanotoxins have been published (Fitzgerald and Poppenga, 1993; Naegeli et al., 1997; Puschner et al., 1998, 2008, 2010; Gugger et al., 2005; Nasri et al., 2008;
C H A P T E R
Wood et al., 2010). The frequency of blue-green algae poisoning in animals is likely underreported due to lack of methods to confirm exposure; in addition, geographi-cal distribution of these case reports is likely biased by available resources. Diagnostic confirmation of suspect blue-green algae poisoning cases of humans and ani-mals requires extensive effort from both toxicologists and clinicians, and resources are often not readily avail-able. New algal toxins are continuously being discov-ered, and oral bioavailability and toxicity data are often unavailable. It is probable that blue-green algae poison-ings are more common in animals than in humans due to animals’ greater direct dependence and contact with surface waters.
Pursuant to several major human intoxications in Australia, Europe, and Brazil (Falconer and Humpage, 2005), in 1998 the World Health Organization (WHO) proposed a guidance value for the maximum permis-sible concentration in potable water sources for micro-cystin-LR, the most commonly reported cyanotoxin worldwide. Water sanitation agencies in many countries in Europe, North America (Canada), South America (Brazil), and Oceania (Australia and New Zealand) adopted these guidelines. In addition, as a result of a tragedy in 1996 in which more than 100 patients at a hemodialysis clinic received inadequately treated drink-ing water (Azevedo et al., 2002), most of whom devel-oped acute liver failure as a result of being exposed to cyanotoxins, Brazil has adopted more comprehensive and stringent guidelines to include other cyanotoxins (Burch, 2008). Awareness of imminent health risks for wild and domestic terrestrial vertebrates has increased during approximately the past decade, in part due to veterinary case reports; however, the extent and
heterogeneity of the impact are still far from understood. Veterinarians thus have the opportunity to substantially deepen the understanding of the impact of these cyano-toxins on animal and human populations alike.
BACKGROUND
Nutrient-rich runoff into surface waters – particularly nitrogen- and phosphorus-rich fertilizers, soaps, and waste products – has led to significant eutrophica-tion worldwide ( 40% in Europe, Asia, and America) (Bartram et al., 1999; Smith, 2003). As a major conse-quence of shifting nutrient additions, previously nutri-ent-limited photosynthetic microorganisms proliferate. Depending on the limitations of the system and the types of nutrients added, a few species (generally one or two) outcompete the others, thereby considerably reducing the heterogeneity of the phytoplankton community. In such conditions, cyanobacteria often predominate through adaptive processes, and substantial shifts in the micro-scopic and macroscopic food web may occur. Anoxic con-ditions can also result in fish kills, and falling debris from blooms can have profound impacts on the invertebrates in the sediment below (Pearl et al., 2001; Havens, 2008).
Among the oldest microorganisms, these oxygenic photosynthetic prokaryotes may be organized as individ-ual cells (e.g., Synechococcus), filaments (e.g., Planktothrix), or colonies (e.g., Microcystis). More than 2000 cyano-bacterial species belong to four orders based on mor-phological and morphometric criteria in botanical code (Anagnostidis and Komarek, 1985); however, classifica-tion based on bacterial code defines five sections through combined use of genetic data, morphological criteria, and cellular fission (Rippka et al., 1979).
Both pelagic (suspended in the water column) and benthic (along the bottom) cyanobacteria can proliferate into blooms. Pelagic blooms, which are easier to visually detect because of the evident scum formation at the sur-face, usually occur in mesotrophic and eutrophic ecosys-tems (concentrations in phosphorus 30 µg/L), during the summer, in water temperatures greater than 20°C, and in low turbulence. Proliferations of benthic species generally occur during the summer on the surfaces of sediments, stones, or macrophytes in small oligotrophic rivers or in oligotrophic lakes (Mez et al., 1997).
In the past, most cases were diagnosed by positive identification of the algae in the suspect water source along with the occurrence of consistent clinical signs and pathological findings. However, new analytical methods can now be applied to detect toxins in biological speci-mens of animals or humans with suspect exposure to toxic algal blooms Figure 72.1 illustrates the wide variety
in chemical structures of cyanotoxins and the need for specific detection methods. (Yuan et al., 2006; Humbert, 2010). These capabilities will allow for in-depth diag-nostic investigations and a better estimate of the true frequency of blue-green algae poisonings in livestock, pets, and wildlife. Table 72.1 provides an overview of cyanobacterial species known to produce a large number of toxins. Some species can produce a variety of cyano-toxins and thus it is difficult to predict the nature and the level of the toxin production during a bloom event. This chapter focuses on the several types of cyanotoxins known to have the greatest impact on veterinary species and presents the current understanding of their toxic mechanisms, toxicokinetics, and diagnostic and thera-peutic approaches with a focus on veterinary medicine.
MICROCYSTINS
Produced by multiple cyanobacteria, including spe-cies within the genera Microcystis, Anabaena, Planktothrix, Nostoc, Oscillatoria, and Anabaenopsis, microcystins have been detected worldwide (Fromme et al., 2000; Hitzfeld et al., 2000; Ballot et al., 2004; Briand et al., 2005; Karlsson et al., 2005a; Ndetei and Muhandiki, 2005; Agrawal et al., 2006). Not all strains are capable of producing microcystins. In recent years, a useful diagnostic tool to test for the pres-ence of toxin-producing genes has emerged (Hisbergues et al., 2003). Although the reason for production is not understood, environmental factors, such as pH, nutrient concentrations, and water temperature, clearly trigger pro-duction, increasing with water temperature, elevated con-centrations of phosphorus and nitrogen, iron limitation, and globally with the growth rate (Briand et al., 2005; Downing et al., 2005; Sevilla et al., 2008). Microcystin concentrations may be highest when the growth of the cyanobacteria is high, but toxin concentrations do not necessarily correlate with cell count, and toxins may occur any time of the year. Although predominantly found in freshwater, microcystin-producing blooms have also been described in saline eco-systems (Atkins et al., 2001; Carmichael and Li, 2006).
Potent cyclic heptapeptides causing acute hepatotoxi-cosis in mammals, microcystins have also been demon-strated to be toxic to reptiles, amphibians, and aquatic species, as well as invertebrates and even plant species (McElhiney et al., 2001; Malbrouck and Kestemont, 2006; Nasri et al., 2008; Amado and Monserrat, 2010). In fresh-water, the toxins are retained inside the cyanobacteria and only released upon cell damage, lysis, and death; destruction of algal mats (either naturally or through the application of herbicides) may result in a pulse of micro-cystin release following destruction of the individual cell walls. After oral exposure to microcystin-containing
MiCRoCysTins 955
algae, the acidic environment of the stomach can result in the release of microcystins. Commercially available blue-green algae food supplements also present a poten-tial route of oral exposure (Schaeffer et al., 1999; Dietrich and Hoeger, 2005).
More than 80 different structural variants of micro-cystins have been identified from various genera of cyanobacteria (Luukkainen et al., 1994; Lawton et al., 1995; Welker and von Döhren, 2006). The shared structure involves an amino acid called ADDA
Microcystins
Homoanatoxin-a
Nodularins
Cylindrospermopsin Anatoxin-a
Anatoxin-a(s)
PSPs
O3SO
HO
O
+
OH
NHN
Me
NH
NH
HN
H
H
HN
NH
R2
R1
CH2
CH3
OCH3
H3CH3C
H3C
HN
COOH O
O
RHOO
O
OS
SZ
H HH
H
H
H H H
H COOH
NX
NHH
SSS
67
1
2
3
4
5
CH3
NH2
O
+
1 2
9
10
3
4
56
7
8
1 2
910 11
3
4
56
8
7
NH2
CH2CH3O
+
CH3
Mdhb
D–GluCH3
R1
CH3
H3C
CH3
OR2 O
O
O
O
O
N
NH
D-MeAspD-Asp
NH
COOH
COOH
HN
N
Z
Adda(DMAdda)
H2
345
1
67891011
12
13
14
1516
17
1819
20
3
4
5
1
2
HHN
N
OHN H
NH2
H2N
R3R2
R1N
R5
+
+
O
R4
1716
65
7
8
9 15
10
13
11
1214
342
1 +
NH2CH3
CH3
CH3
N
+
+
N
O
O
P
O
O
HN
54
3
21
6
+
FIGURE 72.1 Structural formulas of cyanobacterial toxins.
72. CyAnobACTERiAl (bluE-gREEn AlgAE) Toxins956
(3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca; 4,6-dienoic acid) and six other amino acids, includ-ing two variable positions that designate the congener. The most extensively studied, microcystin-LR, has been found worldwide and has caused acute, lethal hepato-toxicoses in farm animals (Carbis et al., 1994; Mez et al., 1997; Puschner et al., 1998) and hepatic injury in humans (Azevedo et al., 2002; Rao et al., 2002).
Pharmacokinetics/toxicokinetics
Despite the abundant literature on microcystins, understanding of pharmacokinetics remains limited,
particularly with regard to potential species variations. Most studies have been conducted in mice after intra-venous (i.v.) and intraperitoneal (i.p.) administration of cyanobacteria, their filtrates, and, in some instances, purified microcystins. After i.v. and i.p. administration in mice and rats, microcystins are rapidly distributed to the liver (Falconer et al., 1986; Robinson et al., 1991). Plasma half-lives of microcystin-LR in mice after i.v. administra-tion were 0.8 and 6.9 min for the alpha and beta phases of elimination (Robinson et al., 1991). Interestingly, the hepatic concentration of 3H-microcystin-LR remained constant throughout the 6-day study, indicating accu-mulation in this target organ. This study also demon-strated that approximately 9 and 14% of the dose was
TABLE 72.1 Potential toxins produced by the different cyanobacterial species
MiCRoCysTins 957
excreted in urine and feces, respectively, after 12 h, with 60% of it being excreted unchanged. Additional studies in swine have also indicated that the majority is excreted unchanged, with only two metabolites detected; biliary excretion is also noted after less than 1 h of i.v. adminis-tration (Stotts et al., 1997). The exact route of metabolism is yet to be defined, but glutathione and cysteine con-jugation have been identified and may represent major detoxification pathways (Kondo et al., 1996; Pflugmacher et al., 1998). Other metabolites have been identified in vivo and in vitro, but further work is needed to define their roles.
Data on bioavailability for microcystins are needed to better evaluate risk from oral ingestion. Absorption occurs in the small intestine (Ito et al., 2000); thus, the integrity of the intestinal mucosa can significantly impact the degree of absorption (Zeller et al., 2011). Altered cell permeability of the small intestine in aged mice lends them more susceptible than young ani-mals (Ito et al., 1997). Once absorbed, microcystins are rapidly distributed to the liver (Runnegar et al., 1981; Fischer et al., 2000), but they can also reach lung, heart, and capillaries (Ito et al., 2000). Based on radiolabeled experiments in which microcystins were administered i.v. in Wistar rats, uptake into the kidney appears to be important for excretion. Results of uptake experi-ments with radiolabeled dihydro-MC-LR demonstrated that OATP1B1 (organic anion transporter protein) and OATP1B3 are involved in uptake of MC-LR (Seithel et al., 2007). Studies in fish (i.v.) have demonstrated that the liver is the primary target, followed by kidney and gonads, although uptake does occur into muscle tissue and cardiovascular effects are observed (Malbrouck and Kestemont, 2006). Absorption of microcystin via the res-piratory route (Ito et al., 2001) has been demonstrated to lead to lethality in mice and also induce damage to nasal epithelium at lower exposures (Benson et al., 2005), thus posing a threat from aerosolization of the compound.
Mechanism of action
Specifically toxic to liver, microcystins cause severe hepatomegaly macroscopically and progressive centri-lobular hepatocyte rounding, dissociation, and necrosis microscopically. Breakdown of the sinusoidal endothe-lium and intrahepatic hemorrhage ultimately result in death (Hooser et al., 1991a; Falconer and Yeung, 1992). Unable to permeate cell membranes, microcystins enter hepatocytes via the bile acid transporter mechanism (Hooser et al., 1991b). Once inside the hepatocytes, microcystins are potent inhibitors of protein phos-phatases 1 and 2A (Falconer and Yeung, 1992; Runnegar et al., 1993). The disruption of the cytoskeletal compo-nents and the associated rearrangement of filamentous
actin within hepatocytes account for the morphologi-cal changes, although other mechanisms play a role in the development of liver lesions. Microcystins induce apoptosis of hepatocytes via induction of free radi-cal formation and mitochondrial alterations (Ding and Ong, 2003). A single-dose i.v. in rats demonstrated an increase in liver sphingolipid levels at higher doses (implicating ceramide-mediated apoptosis), a dose-dependent decreased PP2A expression, and ultimately a dose-dependent decreased expression of Bcl2 family pro-teins, involved in cell cycle/apoptosis regulation (Billam et al., 2008). The role of oxidative stress has become increasingly apparent, and the ultimate toxic effect may depend on the ability of antioxidant pathways to coun-ter the stressors (Ding and Ong, 2003; Jayaraj et al., 2006; Xiong et al., 2010). In addition, microcystins are classi-fied as tumor-promoting compounds (Humpage and Falconer, 1999). Investigations have indicated the role of protooncogenes in this tumorigenesis, hypothesized to be a sequelae of dysregulation of phosphorylation (Li et al., 2009). Several studies have demonstrated the abil-ity of microcystins to induce DNA damage in liver cells (Zegura et al., 2011).
Clinical signs of microcystin poisoning have been described in a number of reports in livestock, humans, and wildlife in the United States (DeVries et al., 1993; Fitzgerald and Poppenga, 1993; Galey et al., 1987; Puschner et al., 1998) and other countries (Done and Bain, 1993; Van Halderen et al., 1995; Mez et al., 1997; Naegeli et al., 1997; Azevedo et al., 2002; Ballot et al., 2004; Ndetei and Muhandiki, 2005; Handeland and Østensvik, 2010; Wood et al., 2010). Interestingly, laboratory ani-mals select water with microcystin-producing strains of cyanobacteria over a water source with nontoxic strains (Lopez Rodas and Costas, 1999), suggesting an increased risk for toxicosis in animals due to behavioral prefer-ences. Microcystin intoxication should be suspected in cases of acute hepatotoxicosis with clinical signs of diarrhea, vomiting, weakness, pale mucous membranes, and shock. Although most animals die within a few hours of exposure, some animals may live for several hours and develop hyperkalemia, hypoglycemia, nerv-ousness, recumbency, and convulsions. Animals that survive the acute intoxication may develop hepatog-enous photosensitization. Nephrotoxic effects have been described in laboratory animals after chronic microcys-tin exposure (Milutinovic et al., 2003). Evidence suggests potential suppression of immune function at sublethal exposures (Shi et al., 2004). In humans, primary liver can-cer as well as colorectal cancer have been associated with microcystin-contaminated drinking water (Ueno et al., 1996; Zhou et al., 2002). In mice, subchronic exposure i.p. of microcystin-LR (20 µg/kg) causes the appearance of hepatic nodules, a characteristic not observed after oral subchronic administration (Ito et al., 1997).
72. CyAnobACTERiAl (bluE-gREEn AlgAE) Toxins958
Toxicity
The lethal doses 50 (LD50s) for microcystins vary between 50 µg/kg and 11 mg/kg, depending on the microcystin analog, the species affected, and the route of administration. In mice, the oral LD50 value for micro-cystin-LR is 10.9 mg/kg, whereas the i.p. LD50 is 50 µg/kg. Because most blooms contain a number of structural variants of microcystins, it is difficult to estimate the toxicity potential of a bloom. The no-observed-adverse-effect level for orally administered microcystin LR to mice is 40 µg/kg/day (Fawell et al., 1994). In pigs, the lowest-observed-adverse-effect level for microcystin-LR is 100 µg/kg/day (Falconer et al., 1994), and in rat it is 50 µg/kg/day (Heinze, 1999). WHO set the tolerable daily intake (TDI) for human ingestion of microcystin-LR at 0.04 µg/kg/day (Kuiper-Goodman et al., 1999). The potential risk to humans by ingesting food prod-ucts derived from animals exposed to microcystins was evaluated in beef (Orr et al., 2003) and dairy cattle (Orr et al., 2001). Based on these studies, it is unlikely that consumption of milk, meat, or liver poses a significant health risk to humans. It might be prudent to establish specific guidelines for nonlethal, chronic microcystin exposure in livestock.
Treatment
No specific antidote for microcystins exists. The rapid onset of acute hepatotoxicosis renders therapeutic inter-vention quite difficult, and mortality rates are very high. In addition, despite the evaluation of numerous treatment options, no specific therapy has been proven to be effective. The most promising strategy appears to be prevention of uptake into hepatocytes through the administration of compounds that may compete for the specific transporters associated with microcystin uptake; administration of the antibiotic rifampin (i.p.) in mice effectively reduced mortality after exposure (i.p.) to microcystin-LR (Hermansky et al., 1991). By contrast, other compounds, such as glutathione, silymarin, and cyclosporine A, were only beneficial if administered as a prophylactic (Hermansky et al., 1991; Rao et al., 2004). These compounds may help reduce microcystin toxicity in chronic exposure scenarios. Due to the role of oxida-tive stress, antioxidants such as vitamin E, selenium, and green tea polyphenols also appear to be beneficial pro-phylactically (Gehringer et al., 2003a,b; Jayaraj et al., 2007; Xu et al., 2007). Although the adsorption of microcystins by activated charcoal was used successfully to decon-taminate drinking water (Warhurst et al., 1997), this decontamination procedure was not protective in mice dosed with microcystins (Mereish and Solow, 1989); no data are available for other species.
Diagnosis of microcystin toxicosis is corroborated by identification of microcystin-containing water in the environment of the animal. Identification of algae mate-rial in water and gastric contents is an important com-ponent of the diagnostic workup but does not confirm intoxication. As described previously, the toxicity of the cyanobacteria is strain specific, and morphological observations alone cannot predict the hazard level, but polymerase chain reaction detection of microcystin- producing genes can help identify a potentially culpable species. Detection of microcystins in gastric contents is confirmatory, but these tests are not routinely avail-able at diagnostic laboratories and are limited to a few structural variants. In the past, the mouse bioassay was used to determine the toxicity of crude algal biomass in suspicious blue-green algae poisonings. Although many assays are available to analyze water samples for microcystins (Maizels and Budde, 2004; McElhiney and Lawton, 2005; Frias et al., 2006), there are only limited methods available to reliably and accurately detect microcystins in biological specimens collected from animals suspected to have died from microcystin intoxication (Bogialli et al., 2005; Karlsson et al., 2005b; Chen et al., 2009). An electrospray ionization liquid chromatography–mass spectrometry method has been developed to determine the bound microcystin concen-trations in animal tissues, which provides an estimate of the total microcystin burden in exposed animals (Ott and Carmichael, 2006).
Differential diagnoses in animals with a clinical pres-entation of liver failure include other toxic ingestions, such as amanitins, cocklebur, cycad palm, aflatoxin, xyli-tol, certain heavy metals, and acetaminophen overdose. Careful evaluation of the history, feed, and environment of the animal can help eliminate most of the toxicant dif-ferentials on the list.
ANATOXINS
Anatoxins are mainly produced by cyanobacteria in the Anabaena genus (Beltran and Neilan, 2000) but also by other genera, such as Plantkothrix, Oscillatoria, Microcystis, Aphanizomenon, Cylindorspermum, and Phormidium. Unfortunately, specific data on factors resulting in anatoxin-a production are lacking, which makes toxin production unpredictable. Although anatoxin-a is con-sidered unstable in the environment, certain environ-mental conditions are known to result in continuous toxin production. Reports of anatoxin poisoning are less frequent than those of microcystin toxicosis; how-ever, poisoning has occurred worldwide (Edwards et al., 1992; Gunn et al., 1992; Beltran and Neilan, 2000;
AnAToxins 959
Fromme et al., 2000; Gugger et al., 2005; Yang and Boyer, 2005; Wood et al., 2007; Puschner et al., 2008, 2010). Anatoxins are neurotoxins and can generally be divided into anatoxin-a, homoanatoxin-a, and anatoxin-a(s). Anatoxin-a is a secondary amine and has been detected in blooms worldwide. Homoanatoxin-a is a methyl derivative of anatoxin-a and has been identified in blooms in Japan (Namikoshi et al., 2004), Ireland (Furey et al., 2003), Sweden (Skulberg et al., 1992), and New Zealand (Wood et al., 2007). Anatoxin-a(s) is a unique N-hydroxyguanidine methyl phosphate ester that has been detected in the Americas (Monserrat et al., 2001) and Europe (Henriksen et al., 1997). Commercially avail-able blue-green algae dietary supplements also represent a potential source (Rellán et al., 2009).
Pharmacokinetics/toxicokinetics
Definite data on the toxicokinetics of anatoxin-a, homo-anatoxin-a, and anatoxin-a(s) have not been established. Based on the rapid onset of clinical signs after oral exposure, rapid absorption of the toxins is suspected. Anatoxin-a has been detected in the urine and bile of a poisoned dog, confirming that anatoxin-a is, at least in part, excreted unchanged in urine and bile (Puschner et al., 2010).
Mechanism of action
Anatoxin-a is a potent cholinergic agonist at nicotinic acetylcholine receptors in neurons and at neuromuscu-lar junctions (Thomas et al., 1993). Anatoxin-a has two enantiomers, with ()anatoxin-a having a higher bind-ing affinity than the (–) form (Spivak et al., 1980; Zhang and Nordberg, 1993). Compared to nicotine, anatoxin-a is approximately 20 times more potent than acetylcho-line. After continuous electrical stimulation at the neu-romuscular junctions, a nerve block may follow and result in death due to respiratory paralysis. Furthermore, anatoxin-a has modulatory action at presynaptic neuro-nal nicotinic acetylcholine receptors, which can lead to dopamine as well as noradrenaline release (Barik and Wonnacott, 2006; Campos et al., 2010). Clinical signs of anatoxin-a poisoning include a rapid onset of rigidity and muscle tremors followed by convulsions, paralysis, respiratory failure, cyanosis, and death. Death usually occurs within minutes to a few hours. Anatoxin-a poi-sonings have been reported in dogs in Europe (Edwards et al., 1992; Gunn et al., 1992; James et al., 1997; Gugger et al., 2005) and the United States (Puschner et al., 2008, 2010). Anatoxin-a is also considered a contributing fac-tor in the deaths of Lesser Flamingos in Kenya (Krienitz et al., 2003). Homoanatoxin-a is a methyl derivate of
anatoxin-a with similar pharmacological and toxicologi-cal properties (Wonnacott et al., 1992), and it has been implicated in dog deaths in New Zealand (Wood et al., 2007). In addition to being a nicotinic agonist, homo-anatoxin-a can increase the release of acetylcholine from peripheral cholinergic nerves through opening of endog-enous voltage-dependent neuronal L-type Ca2 channels (Aas et al., 1996).
Anatoxin-a(s) is different from anatoxin-a and homo-anatoxin-a. This neurotoxin has a unique chemical structure and is a naturally occurring irreversible acetyl-cholinesterase inhibitor. The increased concentrations of acetylcholine in the synapse lead to persistent stimulation, followed by a neuronal muscular block (Cook et al., 1990; Hyde et al., 1991). The mechanism of toxic action is similar to that of organophosphorus and carbamate insecticides, as well as some chemical warfare nerve agents (Patocka et al., 2011). However, one of the main differences is that anatoxin-a(s) acts only in the periphery, whereas the insec-ticides inhibit acetylcholinesterase in the brain and retina (Cook et al., 1989). Animals poisoned with anatoxin-a(s) show a rapid onset of excessive salivation (“s” stands for salivation), lacrimation, diarrhea, and urination. Clinical signs of nicotinic receptor overstimulation including trem-ors, incoordination, convulsions, recumbency, and respira-tory arrest are most commonly observed in cases with a lethal outcome. Animals often die within 30 min of expo-sure. Animals that die from anatoxin-a, homoanatoxin-a, or anatoxin-a(s) toxicosis do not show specific gross or microscopic lesions. Anatoxin-a(s) poisoning has been reported in pigs, birds, dogs, and calves in the United States and Europe (Mahmood et al., 1988; Cook et al., 1989; Onodera et al., 1997). Because of the lack of specific detec-tion methods for anatoxin-a(s), the natural occurrence of this neurotoxin has not been fully evaluated.
Toxicity
In mice, the i.p. LD50 of anatoxin-a is 200 µg/kg (Stevens and Krieger, 1991), whereas the i.v. LD50 is estimated to be less than 100 µg/kg. The oral toxicity of anatoxin-a is much higher, with an oral LD50 in mice reported to be greater than 5 mg/kg. Several studies have shown that there are significant species differences with regard to anatoxin-a toxicity. Whereas an anatoxin-a containing Aphanizomenon flos-aquae bloom was toxic to sheep after i.p. administration, oral administration failed to induce toxicity (Runnegar et al., 1988). In contrast, calves devel-oped toxicity after oral administration of an anatoxin-a containing A. flos-aquae bloom (Carmichael et al., 1977). The i.p. LD50 of homoanatoxin-a in mice is 250 µg/kg (Skulberg et al., 1992). Anatoxin-a(s) is much more toxic than anatoxin-a or homoanatoxin-a, with an i.p. LD50 in mice of 20 µg/kg (Briand et al., 2003).
72. CyAnobACTERiAl (bluE-gREEn AlgAE) Toxins960
Treatment
There is no specific antidote for anatoxin-a. Because of the rapid onset of clinical signs, emesis is not likely to be useful. Although no studies have evaluated the efficacy of specific decontamination procedures, administration of activated charcoal has been recommended. In addi-tion, artificial respiration may be of benefit along with general supportive care. Specific measures to control sei-zures include benzodiazepines, phenobarbital, or pento-barbital. If given, they may cause central nervous system and respiratory depression, and careful monitoring of the animal is necessary. In any seizuring animal, control of body temperature is an important part of the sympto-matic care.
Treatment of animals poisoned with anatoxin-a(s) is primarily symptomatic and supportive. Decontamination procedures can be considered but have not been evaluated. It has been shown that 2-PAM is not able to reactivate the inhibited acetylcholinesterase and is therefore not recommended (Hyde and Carmichael, 1991). Atropine should be given at a test dose to deter-mine its efficacy in animals with life-threatening clinical signs. After the test dose, atropine can be given repeat-edly until cessation of salivation. It is important to care-fully monitor the animal for anticholinergic effects and to reduce or discontinue atropine if adverse effects develop.
As with other cyanobacteria toxins, toxicity is strain-specific, and identification of the cyanobacteria alone cannot predict the toxicity level. Therefore, detection of anatoxin-a in biological specimens is confirmatory, but these tests are not routinely available (James et al., 1998; Puschner et al., 2010). Anatoxin-a was confirmed in stom-ach content, liver, urine, and bile of dogs (Gugger et al., 2005; Puschner et al., 2010). In suspect cases, environ-mental and biological samples should be saved for toxi-cological and phylogenetic analysis.
Diagnosis of anatoxin-a(s) toxicosis is aided by the determination of blood acetylcholinesterase activity. However, organophosphorus and carbamate insecticides can also inhibit acetylcholinesterase, and additional diag-nostic workup is needed to establish a firm diagnosis. This includes the determination of brain acetylcholinesterase postmortem (unchanged in cases of anatoxin-a(s) poison-ing), screening of gastrointestinal contents for insecticides, examination of stomach contents (possible identification of cyanobacteria), and a careful evaluation of the envi-ronment (access to freshwater and access to insecticides). Detection methods for anatoxin-a(s) are rare. A biosensor method has been developed that allows the quantitation of anatoxin-a(s) in environmental samples (Devic et al., 2002). New analytical methods for anatoxin-a(s) are necessary to better document the distribution of this neurotoxin in freshwater worldwide. Phylogenetic analysis of 16S rRNA gene sequences will help in the species identification.
MISCELLANEOUS FRESHWATER CYANOBACTERIAL TOXINS
Although microcystin and anatoxin poisonings comprise the majority of cases reported in animals, other cyanoto-xins are of concern. Saxitoxins and derived forms belong to the group of paralytic shellfish poisoning (PSP) tox-ins and have been produced by a number of freshwater cyanobacteria, including A. flos-aquae, Cylindrospermopsis raciborskii, Anabaena circinalis, Lyngbya wollei, Planktothrix sp., and Aphanizomenon gracile (Carmichael et al., 1997; Kaas and Henriksen, 2000; Molica et al., 2005; Ballot et al., 2010). All saxitoxin analogs have high toxicity in mam-mals by blocking voltage-gated sodium channels, lead-ing to respiratory arrest, neuromuscular weakness, and cardiovascular shock. Whereas intoxications of birds and cats have been associated with ingestion of PSP contami-nated fish and clams (Landsberg, 2002), there is only one confirmed report of PSP toxin-associated mortality from exposure to contaminated freshwater (Negri et al., 1995). Fourteen sheep showed signs of trembling, recumbency, and death in Australia after exposure to toxic concentra-tions of PSPs produced by Anabaena circinalis. Because PSP toxins are produced by several species of freshwater cyanobacteria, terrestrial animals and humans are at risk of being exposed to these toxins. Thus, it is important to reliably evaluate animals with neurologic signs after access to freshwater for possible PSP exposure because the risk certainly exists.
Another cyanotoxin, the alkaloid cylindrospermopsin, has caused deaths in cattle (Saker et al., 1999) and severe gastrointestinal disease in humans. Cylindrospermopsin is a potent inhibitor of protein synthesis and can lead to various degrees of injury to the liver, kidneys, adrenal gland, intestine, lung, thymus, and heart (Griffiths and Saker, 2003). Furthermore, this cyanotoxin is of particu-lar concern because of its mutagenic and possibly carci-nogenic activities. Cylindrospermopsin has been found in Europe, Australia, New Zealand, and Asia (Hawkins et al., 1997; Saker and Griffiths, 2001; Fastner et al., 2003), but it should be considered a potential worldwide prob-lem. Cylindrospermopsin and deoxycylindrospermopsin have been produced by C. raciborskii (Ohtani et al., 1992), and 7-epicylindrospermopsin has been produced by Aphanizomenon ovalisporum (Banker et al., 1997). After oral exposure, the LD50 of cylindrospermopsin obtained with culture extracts of C. raciborskii ranged from 4.4 to 6.9 mg/kg in equivalent cylindropsermopsin (Seawright et al., 1999; Shaw et al., 2000).
Nodularins are cyclic pentapeptides that lead to severe hepatotoxicosis in the same way as microcystins (Harding et al., 1995). In addition, nodularin is a more potent tumor promoter than microcystin (Sueoka et al., 1997). The only cyanobacterium species known to
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produce nodularin is Nodularia spumigena. This cyano-bacterium can form extensive blooms in the Baltic Sea and in brackish waters in the summer (Francis, 1878; Sivonen et al., 1989). The risk of nodularin intoxication is twofold because toxin exposure can occur not only through recreational or drinking water but also via con-tamination of seafood (Van Buynder et al., 2001). In cases of acute hepatotoxicity, exposure to microcystins as well as nodularins must be considered.β-N-methylamino-l-alanine (BMAA), a neurotoxic
amino acid, has been confirmed to be produced by a newly discovered cyanobacterial species associated with avian vacuolar myelinopathy (Bidigare et al., 2009). The algal species is in the order Stigonematales and is found on the surface of Hydrilla verticillata, aquatic vegetation commonly found in wetlands in the southeastern United States (Wiley et al., 2007). BMAA is a neurotoxic amino acid that has been associated with the pathogenesis of human amyotrophic lateral sclerosis–parkinsonism–dementia complex of Guam (Guam ALS-PD) and lathy-rism. Birds with avian vacuolar myelinopathy develop ataxia, tilting, weakness, and death, and they have char-acteristic postmortem lesions of bilateral symmetrical vacuolation of the white matter of the brain and spinal cord (Thomas et al., 1998). Characterization of BMAA provides a critical tool to study the impact of this cyano-bacterial toxin on animals (and humans) and to develop control strategies.
CONCLUDING REMARKS AND FUTURE DIRECTIONS
The frequency and extent of harmful cyanobacterial blooms appears to increase with the addition of nutrients to surface freshwater ecosystems throughout the world. As demonstrated in Table 72.1, numerous cyanobacterial species have the potential to produce a number of toxins with a range of target organs, but for each cyanobacte-rial species, it is very difficult to predict the nature and the level of the toxin production for a specific bloom. Although newer detection methods allow for better monitoring of potentially harmful blooms, there is still a need to apply these existing methods spatially and temporally and to develop lower cost, field-ready alter-natives accessible to even remote areas. It is also impor-tant to develop more sophisticated methods that allow testing for a wider range of cyanotoxins and matrices in order to reliably confirm intoxications and improve overall risk assessment. Some recently developed meth-ods have been useful in analyzing biological specimens in order to confirm a diagnosis of poisoning, but due to lack of availability and high cost, they are rarely pursued
in suspect cases. The lack of methods to confirm expo-sure is most likely responsible for the low number of reported cases in the veterinary literature during the past 20–30 years. The incorporation of new analytical meth-ods into diagnosis should provide insight into the true frequency of cyanotoxin poisoning in animals. For exam-ple, in the United States, several dog intoxications due to anatoxin-a have recently been identified in different regions in relation with the development of an analyti-cal method using high-performance liquid chromato-graph–mass spectrometry analysis (Puschner et al., 2008, 2010). In addition, information is needed on the efficacy of therapeutic measures. Similar to advisories for human populations, it is advisable to take preventative meas-ures to avoid contact with surface waters containing vis-ible blooms for all veterinary species, particularly when the water is utilized for a drinking source or for bathing.
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