Algal Toxins in Pond Aquaculture John H

SRAC Publication No. 4605

VI PR November 2008

Algal Toxins in Pond Aquaculture John H. Rodgers, Jr.1

Algal toxins can cause problems in the world is increasing (Shumway, • the decaying biomass of a bloom the freshwater aquaculture of both 1990; Sunda et al., 2006), especially depletes dissolved oxygen (espe- vertebrates (fish) and invertebrates in the U.S. where almost every coastal cially critical in aquaculture); or (shellfish). Such problems include: state is now threatened, in some cases • blooms kill other algae impor- • off-flavor (Tucker, 2000; SRAC by more than one species of harmful tant in the food web (Codd et al., Publication No. 192), algae. Scientists are unsure why this 2005b; Landsburg, 2002). • indirect toxicity through changes trend is occurring. The causes may be HABs can cause serious economic in water quality (SRAC Publica- natural (species dispersal) or human- losses in aquaculture if they kill cul- tion No. 466), or related (nutrient enrichment, climate tured organisms or cause consumers • direct toxicity. change, and/or transport of algae in concern about food safety. Prelimi- ship ballast water) (Johnk, et al. 2008; Algal toxins are organic molecules pro- nary estimates show that the effect of Sunda et al., 2006). duced by a variety of algae in marine, HAB outbreaks on the U.S. economy brackish and fresh waters, as well as The effects of algal blooms vary wide- is more than $40 million per year, on wet soils (Falconer, 1993). They are ly. Some algae are toxic only at very or $1 billion per decade (Landsburg, a problem in aquaculture when they high densities, while others can be 2002; Hudnell, 2008). are produced in sufficient quantities, toxic at very low densities (a few cells Toxin-producing algae may become with sufficient potency, to kill cul- per liter). Some blooms discolor the more prevalent in the future (Sunda et tured organisms, decrease feeding and water (thus the terms “red tide” and al., 2006; Johnk et al., 2008), especial- growth rates, cause food safety issues, “brown tide”), while others are almost ly in eutrophic freshwater systems. or adversely affect the quality of the undetectable with casual observation This publication focuses on algal product (Shumway, 1990). (Shumway, 1990). toxins in freshwater pond aquaculture HABs can affect public health and in the South and southeastern United Algal blooms ecosystems when: States. The most common toxin-pro- The production of algal toxins is nor- • filter-feeding shellfish (clams, ducing algae in this region are cy- mally associated with algal blooms, mussels, oysters, scallops) feed anobacteria, golden algae (Prymnesium or the rapid growth and exception- on toxic phytoplankton and ac- parvum) and euglenoids. ally dense accumulation of algae. The cumulate harmful toxins that are term Harmful Algal Bloom (HAB) is passed up the food chain; Cyanobacteria: • fish, shellfish, birds and even used to describe a proliferation of al- the blue-green algae gae, or phytoplankton. Severe blooms mammals are killed by eating of even non-toxic algae can spell organisms that have consumed Cyanobacteria (blue-green algae) in- disaster for cultured animals, because algal toxins; habit fresh, brackish, marine and hy- blooms deplete the oxygen in the • light cannot penetrate the water, persaline waters, as well as terrestrial shallow waters of many aquaculture thus changing the function and environments. Cyanobacteria grow in systems. The number of HABs around structure of the aquatic ecosystem; many habitats from thermal springs • discoloration makes water aesthet- to the arctic. They play important 1 Clemson University ically unpleasant; roles in the biogeochemical cycling of elements and in the structure, In aquaculture situations, eukaryotic toxic Microcystis cells, they show no function and biodiversity of aquatic algae (green, diatoms, etc.) can often selection between ingesting toxic or communities (from microbes through grow faster than cyanobacteria. How- nontoxic cells, which indicates that vertebrates). Some cyanobacteria can ever, cyanobacteria can out-compete microcystins are not responsible for reduce both N2 and CO2. Some can algae for nutrients, thrive with low feeding inhibition (Berry et al., 2008). convert N2 into NH3 and, ultimately, dissolved oxygen, and photosynthe- into amino acids and proteins. size more efficiently at low light lev- Problems with cyanobacteria els. Cyanobacteria are less affected in aquaculture ponds Cyanobacteria have a relatively by turbidity, high concentrations of Cyanobacteria can rapidly overtake simple prokaryotic structure and lack ammonia and warm temperatures. an aquaculture pond and contribute membrane-bound organelles (nucleus, They can seize the advantage in to unstable conditions. Cyanobacteria mitochondria and chloroplasts). With eutrophic aquaculture situations. Cy- blooms can decrease fish production murien in the cell wall and reproduc- anobacteria can affect the production and kill fish because of oxygen depletion by binary fission, cyanobacteria of zooplankton and consequently the tion. Cyanobacteria can also cause are structurally and physiologically production of fish. They also produce off-flavor and objectionable odor in like other gram-negative bacteria, allelochemicals that can inhibit com- fish. but they conduct photosynthesis like peting algae and invertebrate grazers plants in aquatic systems. Cyanobac- However, the role of cyanobacteria (Gross, 2003; Berry et al., 2008). teria are much larger than other bac- and cyanotoxins in fish kills and oth- teria and make a major contribution There is compelling evidence that er problems is not clear at this time. to world photosynthesis and nitrogen cyanobacteria and their toxins (both There are more than 1 million fish fixation (Codd et al., 2005a; Huisman neurotoxins and hepatotoxins) af- ponds in the Southeast and many of et al., 2005; Hudnell, 2008). fect zooplankton (cladocerans and them have relatively frequent blooms rotifers) population structure, and of cyanobacteria that may produce Cyanobacteria occur in unicellular, that this may influence the ecological toxins (e.g., Microcystis, Anabaena, colonial and filamentous forms and processes responsible for cyanobac- etc.). Yet there are only a few reports most are enclosed in a mucilagenous terial success (Berry et al., 2008). of fish kills that are directly related sheath, either individually or in colo- Zooplankton generally avoid cy- to algal toxin production (Zimba et nies. As single cells, large colonies anobacteria as a food source (Gross, al., 2001). So the mere presence of and filaments (trichomes), blue-green 2003), which means that zooplank- toxin-producing algae does not neces- algae can become the dominant algae ton feed on algae that compete with sarily mean that enough toxin will be in nutrient-rich waters. They can cyanobacteria. In the process, they produced to harm fish in culture. form blooms so thick it appears that release essential nutrients, further blue-green paint covers the surface of Cyanobacterial toxins fertilizing cyanobacterial growth. the water. Cyanobacterial toxins can be clas- During cyanobacterial blooms, when sified several ways. They may be Several species found in the South alternative food sources for zooplank- classified according to their chemical and Southeast produce substances ton have been exhausted, Daphnia structures as cyclic peptides (mi- that cause taste and odor problems in populations may decline. Some zoo- crocystin and nodularin), alkaloids water supplies and aquacultural prod- plankton (Daphnia pulicaria, Daphnia (anatoxin-a, anatoxin-a(s), saxitoxin, ucts (Tucker, 2000). Some blue-green pulex) species have adapted to survive cylindrospermopsin, aplysiatoxins, algae, particularly Anabaena and Mi- in the presence of certain toxic cells lyngbyatoxin-a) and lipopolysaccha- crocystis, produce toxins poisonous to (Sunda et al., 2006; Gross, 2003). rides. However, cyanotoxins are more fish and to wildlife and livestock that This changes the zooplankton popula- commonly discussed in terms of their drink contaminated water. There are tion dynamics. Feeding pressure by toxicity to animals. While there are also documented cases of blue-green adapted zooplankton on cyanobac- several dermatotoxins (e.g., lyng- algal toxins harming people in other teria is reduced by fish predation, byatoxin and aplysiatoxins), which parts of the world who drank poorly which again releases nutrients that are produced primarily by benthic treated water. fuel cyanobacterial growth. It may be cyanobacteria, most cyanotoxins are premature to propose that cyanobac- Cyanobacterial ecology either neurotoxins or hepatotoxins terial dominance is guided by cyano- (Codd et al., 2005a). in aquaculture ponds toxin production. However, feeding Cyanobacteria can colonize and rap- deterrence is one of the roles sug- Neurotoxins. Neurotoxins are or- idly grow to great masses in aquacul- gested for these metabolites (Berry et ganic molecules that can attack the ture ponds. Factors that affect their al., 2008). Whether the compounds nervous systems of vertebrates and growth are nutrient status, salinity causing toxicity and deterrence are invertebrates. Three primary types or ionic strength, light conditions, one and the same has recently been of neurotoxins have been identified: turbulence and mixing, temperature questioned (Berry et al., 2008). While 1) anatoxin-a, an alkaloid, inhibits and herbivory (Sunda et al., 2006). daphnids are killed when feeding on transmissions at the neuromuscular junction by molecular mimicry of the neurotransmitter acetylcholine (blocks post-synaptic depolarization); 2) anatoxin-a(s) blocks acetylcholin- esterase (similar to organophosphate pesticides); 3) saxitoxins are carbamate alkaloids that act like carbamate pesticides by blocking sodium channels. Neurotoxins are produced by several genera of cyanobacteria including Anabaena, Aphanizomenon, Microcystis, Microcystis aeruginosa Planktothrix, Raphidiopsis, Arthrospira, Figure 1. (photos by John H. Rodgers, Jr.). Cylindrospermum, Phormidium and Oscillatoria. Neurotoxins produced by Ictalurus punctatus, can become intox- if belonging to the same species, is Anabaena spp., Oscillatoria spp. and icated at ~50 to 75 µg microcystin/L a common explanation for this oc- Aphanizomenon flos-aquae blooms have (Zimba et al., 2001). All fish may be currence. However, some species are been responsible for animal poison- killed within 24 hours of exposure. known to produce high or low levels ings around the world (Carmichael, At necropsy, severe lesions may be of toxicity under different laboratory 1997; Briand et al., 2003). observed in liver tissues. conditions. The stimulus for toxin production in such species is not Neurotoxins usually have acute One potent hepatotoxin, cylindrosper- known. Environmental parameters effects in vertebrates, with rapid pa- mopsin, is produced by Cylindrosper- such as light intensity, temperature, ralysis of the peripheral skeletal and mopsis raciborskii, a relatively small nutrients and trace metals have been respiratory muscles. Other symptoms cyanobacterium. Cylindrospermopsin mimicked under laboratory conditions include loss of coordination, twitch- is an alkaloid that suppresses glu- and their effect on cyanotoxin pro- ing, irregular gill movement, tremors, tathione and protein synthesis. C. duction investigated. Studies on light altered swimming, and convulsions raciborskii has been in the South and intensity are not definitive, but it is before death by respiratory arrest. Southeast for decades and is becom- known that intense light increases the ing more widespread. Mammals (such Hepatotoxins. Hepatotoxins are cellular uptake of iron, which may be as humans) are relatively sensitive produced by many genera of cy- responsible for more toxin production. to cylindrospermopsin and may be anobacteria and have been impli- However, low concentrations of iron affected when they eat fish that have cated in the deaths of fish, birds, lead to higher microcystin concentra- been exposed to the toxin. A study wild animals, livestock and humans tions (Huisman et al., 2005). Nutrients reporting the bioaccumulation of around the world (Briand et al., 2003; such as nitrogen and phosphorus are cylindrospermopsin in muscle tis- Carmichael, 1997). The cyclic hep- essential for cyanobacterial growth. sue of the redclaw crayfish (Cherax tapeptides, or microcystins, inhibit Phosphorus is usually the limiting fac- quadricarinatus) and visceral tissues eukaryotic protein phosphatases type tor in ponds, so small increases in this of rainbow fish (Oncorhynchus mykiss) 1 and type 2A, resulting in excessive nutrient may influence toxin produc- shows that exposure could occur from phosphorylation of cytoskeletal ele- tion simply as a result of increasing farm-raised freshwater aquatic foods. ments and ultimately leading to liver algal growth. Generally, decreased Fish are generally more tolerant of failure (Codd, 2005b). These toxins amounts of microcystin (produced by algal toxins than mammals and tend target the liver by binding the organic Anabaena, Microcystis and Oscillatoria) to accumulate them over time (Carson, anion transport system in hepato- and anatoxin-a (produced by Aphani- 2000). Although C. raciborskii has not cyte cell membranes. Microcystins zomenon) have been reported under yet been a problem in aquaculture, it are the largest group of cyanotoxins, the lowest phosphorus concentrations could become a problem in the future. with more than 70 structural variants tested (Watanabe et al., 1995). (Malbrouk and Kestemont, 2006). Environmental effects Managing toxic cyanobacteria Microcystin is the only cyanotoxin on toxin production Monitoring and diagnosing the for which the biosynthetic pathway The effects of environmental factors problem. While not all blooms of and gene cluster have been identified on toxin production are much studied toxin-producing cyanobacteria result (Huisman et al., 2005). Microcys- and widely disputed (Codd, 2000; in toxin production, most do. Once tins are produced in fresh waters by Codd et al., 2005a). Blooms in the a bloom is observed, the onset of species of Microcystis, Anabaena and same body of water can be toxic or toxicity will be rapid (hours to a day Planktothrix. Symptoms of poisoning non-toxic from one year to the next. or two). To confirm the problem, a in fish include flared gills because of A different strain composition (i.e., diagnostician will need fresh samples difficulty breathing and weakness or toxic versus non-toxic), which can- (unpreserved) of the water contain- inability to swim. Channel catfish, not be distinguished microscopically ing the suspected cyanobacteria 9.1 µm (Rottmann et al., 1992). A sample of both sick and dead fish will also Haptonema be needed, along with informa- saddle-shape chlaroplasts tion on fish behavior and any other symptoms observed. Young fish are generally more sensitive than older fish. The diagnostician may look for lesions on fish livers, although this is inconclusive as the sole method of 2 flagella diagnosis (Zimba et al., 2001). Figure 2. Prymnesium parvum (photos by Dave Buzan and Greg Southard). Treatment. Most of the time, managing a pond specifically to prevent toxic brackish waters and inland waters Signs of intoxication blue-green algae blooms is not justi- with relatively high mineral content Dense growths of Prymnesium par- fied, and the treatments themselves on five continents (Otterstrom and vum may color the water yellow to are risky. An algicide should not be Steemann-Nielsen, 1940; Holdway et copper-brown or rust. The water may applied without considering the size al., 1978; James and de la Cruz, 1989; foam if aerated or agitated. Affected of the affected pond, the number and Kaartvedt et al., 1991; Guo et al., fish behave erratically. They may type of fish at risk, the age and condi- 1996; Lindholm et al., 1999). accumulate in the shallows and be tion of the fish, the sensitivity of the P. parvum cells contain chlorophylls observed trying to leap from the wa- cyanobacterium to treatment, and the a and c as well as yellow-brown ac- ter to escape the toxins (Sarig, 1971). cost of the treatment. Non-chemical cessory pigments and are capable of Affected fish may have evident blood treatments include 1) physical mixing photosynthesis. However, the organ- in gills, fins and scales and may be and aeration, 2) increasing flow rate or ism is thought to be a mixotroph that covered with mucus. Young fish are flushing to decrease hydraulic reten- can feed on bacteria and protists often most sensitive to the toxin. If tion time, and 3) decreasing or alter- (Skovgaard et al., 2003) and phagotro- fish are removed to uncontaminated ing nutrient content and composition. phy has been observed in cultures. P. water in the early stages of intoxica- Some of these options may not be parvum has a vitamin requirement in tion, their gills can recover within viable at all sites and in all situations. laboratory culture (Droop, 1954). hours (Shilo, 1967). However, feeding and growth will usually be reduced if Prymnesium Prymnesiophytes: The toxins fish survive (Barkoh and Fries, 2005). Prymnesium parvum produces at least Aquatic insects, birds and mammals the golden-brown algae three toxins—an ichthyotoxin (Ulitzer are not affected by P. parvum toxins. The haptophyte genus Prymnesium is and Shilo, 1966), a cytotoxin (Ulitzer The golden alga is not known to harm comprised mainly of toxin-producing and Shilo, 1970) and a hemolysin (a humans either, although dead or dy- species that form harmful blooms protein that lyses red blood cells) ing fish should not be eaten. usually in brackish water (West et al., (Ulitzer, 1973). These toxins are col- 2006). Blooms of P. parvum have been lectively known as prymnesins and Managing prymnesiophytes responsible for fish kills and signifi- all can alter cell membrane perme- Monitoring and diagnosing the cant economic losses in Europe, North ability (Shilo, 1971). Bloom density problem. Identifying Prymnesium America and other continents. Texas does not correlate strongly with toxic- parvum requires examining un- has been hit with recurrent blooms ity (Shilo, 1981), probably because preserved, discrete water samples. in several reservoirs and rivers and toxicity may be enhanced by tem- P. parvum can pass through many Texas Parks and Wildlife has offered perature lower than 30 ºC (Shilo and plankton nets and their structure can some detailed advice regarding man- Aschner, 1953), pH greater than 7.0 be altered and distorted by preserva- agement options (Sager et al., 2007). and phosphate concentration (Shilo, tives or fixatives. P. parvum cells can 1971). be detected at low concentrations Prymnesiophyte ecology 2 Prymnesium parvum is commonly The P. parvum ichthyotoxin affects (~10 cells /mL) using fluorescent mi- called the “golden” alga. It is con- gill-breathing aquatic animals such croscopy on live or reasonably fresh sidered to be a haptophyte protist as fish, brachiated tadpoles and mol- samples. P. parvum cell densities can (Green and Leadbetter, 1994). It is lusks (Shilo, 1967). It causes gill cells be determined using a hemacytom- a relative small (~10 µm), generally to lose their selective permeability eter (Barkoh and Fries, 2005). halophilic organism that intermit- and, thus, makes them vulnerable to Prymnesium parvum N. Carter are tently produces an ichthyotoxin. any toxin in the water (Ulitzer and small, subspherical, swimming cells This organism has been implicated Shilo, 1966; Shilo, 1967). about 8 to 15 µm long with two equal in numerous extensive fish kills in or subequal heterodynamic flagella 12 to 20 µm long and a short (3 to 5 affect some fish (Barkoh et al., 2004). Relatively recent research has con- µm), flexible, non-coiling haptonema If not carefully used, the copper firmed that Euglena species produce (Green et al., 1982). Cells have two sulfate may kill desirable algae along an ichthyotoxin in freshwater aqua- chloroplasts that may be “c-shaped” with the P. parvum and decrease food culture (Zimba et al., 2004). The hy- and olive green in color. They swim resources for the zooplankton, thereby brid striped bass mortalities in North with a smooth forward movement disrupting fish feeding. Neither barley Carolina were caused by E. san- while the cell spins on the longitudi- straw nor a commercial bacterial guinea, a widely distributed species nal axis and flagellar pole. P. parvum bioaugmentation product were effec- in many shallow, relatively calm, cells have diagnostic calcareous scales tive for controlling P. parvum in Texas eutrophic freshwater systems. This that can be observed with electron ponds (Barkoh et al., 2008). Barley species also killed laboratory-reared microscopy (Green et al., 1982). Expe- straw extract was also ineffective in channel catfish, tilapia (Oreochromis rience is required to identify this alga. laboratory tests (Grover et al., 2007). niloticus) and striped bass. In confir- West et al. (2006) have developed a Treatments with high concentrations matory studies, Zimba et al. (2004) specific monoclonal antibody used in of ammonium (0.72 mg NH4-N/L) noted that cultures of Euglena granu- conjunction with solid-phase cytom- were successful, though they pose a lata (UTEX LB2345) caused similar etry to rapidly quantify P. parvum. risk to non-target species. Repeated mortalities and symptoms in chan- treatments of ammonium chloride nel catfish and sheepshead minnows A bioassay can be used to estimate and phosphoric acid were somewhat (Cyprinodon variegatus). ichthyotoxin production by Prymne- successful for controlling P. parvum sium parvum (Sager et al., 2007). This growth in limnocorrals in aquaculture Diagnosis assay is useful in deciding whether to ponds (Kurten et al., 2007), but they Microscopic examination of a water apply an algicide. The assay involves are also hazardous to non-target spe- sample can confirm the presence exposing fish such as larvalPimephales cies. of Euglena since E. sanguinea is promelas to the pond water in ques- morphologically distinct. The typi- tion, and to dilutions of the suspect In the Chinese aquaculture of carp cal progression of symptoms from water amended with a cofactor or species, suspended solids (mud), exposure to Euglena toxin begins promoter of P. parvum toxin. Ulitzer organic fertilizer (manure) and with the fish going off feed for no and Shilo (1966) discovered that the decreased salinity have been used to apparent reason. Within 24 hours of potency of the P. parvum ichthyotoxin control P. parvum (Guo et al., 1996), cessation of feeding, the fish swim was augmented by the cation DADPA with the best results from decreased at or near the surface in an agitated (3,3-diaminodipropylamine) in labora- salinity and ammonium sulfate. or disorientated state, often with the tory tests because it increased the When using an algicide, obtain all dorsal fin extending out of the water sensitivity of Gambusia to the toxicant. regulatory approvals and permits or swimming on their sides and even This bioassay can identify waters that and follow label instructions and re- upside down. If steps are not taken have sufficient toxin (or are develop- strictions to comply with federal law. immediately after observing this ing sufficient toxin concentrations) to state, within 24 hours the fish will be pose a risk to fish in culture. dead. Euglenoids For diagnostic analysis, 10-liter water Field observation of dead fish indi- Since 1991, several outbreaks of samples should be collected at least 6 cated rapid onset of mortality with toxic Euglena (Fig. 3) have occurred inches below the surface because P. reddening of gill tissue. Mortality in North Carolina hybrid striped parvum is sensitive to UV radiation. of laboratory-reared channel catfish bass (Morone saxatilis x M. chrysops) Both cell counts (microscopy) and the occurred within 6.5 minutes to 2 production ponds, causing the loss toxin bioassay are needed to confirm hours after exposure. No water qual- P. parvum, so unpreserved water of more than 20,000 pounds of fish. samples must be shipped expedi- tiously to the lab. Treatment. Texas Parks and Wild- life has detailed advice for managing Prymnesium parvum (Sager et al., 2007). One method used in isolated pond culture is applying ammonium sulfate and copper sulfate (Barkoh et al., 2003). The ammonium sulfate concentrations required to control P. 25 µm parvum (~0.17 mg/L of un-ionized ammonia) may produce un-ionized ammonia concentrations that adversely Figure 3. Euglena sanguinea. ity characteristics (ammonia, nitrate, References Advances in Botanical Research, pH, temperature or dissolved oxygen) Vol. 27. London: Academic Press. Baker, J.W., J.P. Grover, B.W. Brooks, were abnormal or exceptional during pp. 211-256. F. Urena-Boeck, D.L. Roelke, the fish mortalities. Fish exposed to Carson, B. 2000. Cylindrospermop- R.M. Errara and R.L. Kiesling. E. sanguinea in culture or to filtrate sin – Review of Toxicological 2007. Growth and toxicity of from cultures were disoriented Literature. National Institute of Prymnesium parvum (Haptophyta) rapidly. Their respiration was ac- Environmental Health Sciences: as a function of salinity, light and celerated and they lost the ability to Research Triangle Park, NC. 49 temperature. Journal of Phycology maintain equilibrium. Although no pp. 43:219-227. distinct hemorrhaging was observed, Codd, G.A. 2000. Cyanobacterial Barkoh, A., D.G. Smith and J.W. gill tissue was reddened (Zimba et toxins, the perception of water Schlecte. 2003. An effective mini- al., 2004). quality, and prioritization of mum concentration of un-ionized eutrophication control. Ecological Euglena ammonia nitrogen for controlling toxin Engineering 16:51-60. Prymnesium parvum. North Ameri- Based on behavioral changes in Codd, G.A., J. Lindsay, F.M. Young, can Journal of Aquaculture 65:220- exposed fish, Zimba et al. (2004) L.F. Morrison and J.S. Metcalf. 225. suggested that the euglenoid toxin 2005. Harmful cyanobacteria: Barkoh, A., D.G. Smith, J.W. functions as a neurotoxin. The toxin from mass mortalities to manage- Schlechte and J.M. Paret. 2004. is water soluble, non-protein, heat ment measures, in J. Huisman, Ammonia tolerance by sunshine stable (30 °C for 10 minutes), freez- H.C.P. Matthijs and P.M. 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The work reported in this publication was supported in part by the Southern Regional Aquaculture Center through Grant No. 2006- 38500-16977 from the United States Department of Agriculture, Cooperative State Research, Education, and Extension Service.