logy, Part C 145 (2007) 61–72www.elsevier.com/locate/cbpc

Comparative Biochemistry and Physio

The zebrafish (Danio rerio) embryo as a model system for identificationand characterization of developmental toxins from

marine and freshwater microalgae☆

John P. Berry a,c,⁎, Miroslav Gantar b, Patrick D.L. Gibbs a, Michael C. Schmale a

a Marine Biology and Fisheries, Rosenstiel School of Marine and Atmospheric Science, University of Miami,4600 Rickenbacker Causeway, Miami, FL 33149, USA

b Department of Biological Sciences, Florida International University, Miami, FL, USAc Department of Chemistry and Biochemistry, Florida International University, 11200 SW 8th Street, Miami, FL 33199, USA

Received 29 March 2006; received in revised form 8 July 2006; accepted 10 July 2006Available online 10 August 2006

Abstract

The zebrafish (Danio rerio) embryo has emerged as an important model of vertebrate development. As such, this model system is findingutility in the investigation of toxic agents that inhibit, or otherwise interfere with, developmental processes (i.e. developmental toxins), includingcompounds that have potential relevance to both human and environmental health, as well as biomedicine. Recently, this system has been appliedincreasingly to the study of microbial toxins, and more specifically, as an aquatic animal model, has been employed to investigate toxins frommarine and freshwater microalgae, including those classified among the so-called “harmful algal blooms” (HABs). We have developed this systemfor identification and characterization of toxins from cyanobacteria (i.e. “blue-green algae”) isolated from the Florida Everglades and otherfreshwater sources in South and Central Florida. Here we review the use of this system as it has been applied generally to the investigation oftoxins from marine and freshwater microalgae, and illustrate this utility as we have applied it to the detection, bioassay-guided fractionation andsubsequent characterization of developmental toxins from freshwater cyanobacteria.© 2006 Elsevier Inc. All rights reserved.

Keywords: Zebrafish; Danio rerio; Vertebrate development; Toxins; Algae; Model system; Cyanobacteria; Everglades

1. Introduction

Marine and freshwater microalgae, encompassing photosyn-thetic microbial representatives from both prokaryotic (e.g.cyanobacteria) and eukaryotic (e.g. dinoflagellates, diatoms)taxa, are known to produce a myriad of toxic or otherwisebioactive metabolites (Shimizu, 1996; Gerwick et al., 2001;Shimizu, 2003). These compounds are finding applications inbiomedicine, including use as pharmacological and toxicolog-ical research tools, as well as development of potential phar-maceuticals (Mayer and Lehmann, 2001; Mayer and Gustafson,

☆ This paper is based on a presentation given at the conference: AquaticAnimal Models of Human Disease hosted by the University of Georgia inAthens, GA, USA, October 30–November 2, 2005.⁎ Corresponding author. Tel.: +1 305 421 4217; fax: +1 305 421 4600.E-mail address: jberry@miami.edu (J.P. Berry).

1532-0456/$ - see front matter © 2006 Elsevier Inc. All rights reserved.doi:10.1016/j.cbpc.2006.07.011

2003). Moreover, as these microbes are ubiquitous in aquaticenvironments worldwide, the possible relevance of these toxicmetabolites to human and environmental health is receivinggrowing attention (Duy et al., 2000; Hitzfield et al., 2000;Carmichael, 2001; Rao et al., 2002).

In particular, many of these toxigenic algae are categorizedamong the so-called “harmful algal blooms” (HABs) owing tothe recognized impacts they have on human and environmentalhealth when they occur in high abundance. Typically, HABtoxins are most closely associated with cases of acute poisoning,specifically including toxicoses resulting from direct exposureof humans to toxins via aerosolized seawater (e.g. Florida RedTides) or contaminated drinking water, or from indirect exposureto toxins which bioaccumulate in fish and shellfish (e.g.paralytic shellfish poisoning, ciguatera). Based on such acuteeffects, toxicology of these metabolites most often focusesexclusively on those targets which interfere with immediate and

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pronounced cell-, tissue- or organ-specific function, includingtoxicoses related to neuronal action (i.e. neurotoxicity), gas-trointestinal and hepatic functions (e.g. diarrhetic shellfishpoisoning, hepatoenteritis), respiratory and pulmonary function(e.g. bronchoconstriction associated with Red Tide) and dermalresponses (i.e. dermatotoxicity).

It is becoming increasingly clear, however, that HAB toxins,including many of the same species associated with acutepoisonings, may additionally exert an array of more insidious,chronic effects. Indeed, toxic algal metabolites frequently havemultiple targets or otherwise broad mechanisms of action suchas inhibition of phosphatases and DNA damage that maypotentially affect multiple cellular processes. Associated targetsof the chronic effects of these toxins, and even some of thetoxins themselves, may consequently remain largely unrecog-nized and uncharacterized when these effects are obscured bythe more immediate acute effects.

Of particular concern are those toxins that either interfere withprocesses related to maintenance of cell-cycle control and celldifferentiation, or cause DNA damage. Such effects may leaddirectly (via genotoxicity) or indirectly (via tumor promotion) tocarcinogenesis. For example, the hepatotoxic microcystins, mostwidely associated with acute poisoning (Beasley et al., 2000;Fischer and Dietrich, 2000; discussed further below), are alsorecognized tumor promoters, and chronic exposure via contam-ination of drinking water by microcystin has been linked to highincidence of liver cancer in China (Yu, 1989).

There is also growing concern and evidence (e.g. Pilottoet al., 1999) that algal toxins, by interfering with the coordinationof rapidly dividing and differentiating cells, may perturb path-ways and processes as they relate to vertebrate development (i.e.“developmental toxins”). Such effects would have obviousdirect relevance to human, animal and environmental healthincluding possible concerns related to infertility, birth defectsand other developmental abnormalities. Indeed, review ofthe human health impacts associated with HAB toxins (e.g.Hitzfield et al., 2000) indicates that children are often at par-ticular apparent risk, suggesting perhaps that the relationshipbetween toxicity and developmental stage may not be onlylimited to embryogenesis, but more generally to continuingvertebrate development.

Fish and other aquatic animals have been shown to be usefulmodels for a variety of diseases such as cancers (Meierjohannet al., 2004; Rahn et al., 2004), anemia (Shafizadeh et al., 2004)and glaucoma (McMahon et al., 2004) as well as for toxi-cological studies including responses to environmental pollu-tants such as dioxin (Butler et al., 2004; Yamauchi et al., 2005)and heavy metals (Feng et al., 2003; Bielmyer et al., 2005). As ameans to identify and characterize toxins from marine andfreshwater microalgae, aquatic animals models have a number ofpractical advantages. In terms of investigating developmentaltoxins, embryos of various teleost fish have been specificallyexplored as potential models for vertebrate development (e.g.Berman et al., 2005; Drummond, 2005; Morris and Fadool,2005; Puffett-Lugassay and Zon, 2005; Sherwood and Wu,2005), and consequently for the investigation of toxins thatdisrupt the pathways and processes related to development. Of

these, the embryo of the zebrafish (Danio rerio) is findingconsiderably utility owing to a number of practical advantages.

Advantages of the zebrafish embryo as a toxicologicalmodel, particularly in relation to developmental toxins, includea small size, a nearly transparent embryo, ease of husbandry,high fecundity, rapid embryogenesis and a growing under-standing of its genome. The nearly transparent zebrafish em-bryos are approximately 1-mm in diameter (and surrounded bya nearly transparent chorion) making them easily observablewith standard dissecting light-microscope, and conversely smallenough to be utilized in a high-throughput (e.g. 96-well plate)format. The embryos can be reared in a wide array of media, andare reasonably tolerant to a variety of solvents and buffers usedin toxin extraction and isolation. Laboratory breeding canproduce more than sufficient numbers of embryos, typically onthe order of hundreds and even thousands of eggs with a singledaily breeding. The entire development of the embryo from afertilized oocyte to a free-swimming hatched larval fish isapproximately 5 days. Finally, the growing knowledge of thezebrafish genome, including a completed genome sequence,and an equally growing “library” of developmental mutants,enables toxicological findings to be increasingly related to thepossible genes involved.

Here we review applications of the zebrafish embryo (as wellas other freshwater fish models) as applied to the investigationof developmental toxins from marine and freshwater algae. Inaddition, we present primary data from our on-going study ofcyanobacteria isolated from the Florida Everglades and otherfreshwater sources in South and Central Florida to illustrate theutility of this aquatic animal system for identification (i.e.screening), as well as subsequent purification (i.e. bioassay-guided fractionation) and characterization of, naturally occur-ring developmental toxins.

1.1. Freshwater fish as models for developmental toxins frommarine dinoflagellates

Of the HAB toxins, perhaps the most widely recognized, andbest characterized, are those from various marine protists, par-ticularly including members of the Bacillariophyta (diatoms),Dinoflagellata (dinoflagellates) and related Raphidophytaphyla. Notable examples of these toxic HABs, and the toxinsthey produce, are given in Table 1. As discussed above, most ofthe focus on these HAB toxins has been on acute health effects,particularly including neurological symptoms (e.g. paralyticshellfish poisoning, amnesic shellfish poisoning), bronchial andpulmonary effects (e.g. aerosolized Red Tide), and gastrointes-tinal illness (e.g. diarrhetic shellfish poisoning).

However, growing evidence suggests that these same toxinsmay either havemechanisms of action that affect multiple targetsor alternative mechanisms that could potentially interfere withprocesses of cell maintenance, division and differentiation, aswell as genotoxic (i.e. DNA damaging) effects. All of thesemechanisms may have profound impacts on development. Oka-daic acid (OA), for example, is well recognized as a potentinhibitor of the serine/threonine protein phosphatases, PP1 andPP2A (Bialojan and Takai, 1988). Typically, OA is most

Table 1Marine and freshwater HABs associated with human and environmental health concerns

Organism(s) Toxin(s) “Target” Toxicoses

ProtistaDinoflagellata(“dinoflagellates”)Karenia brevis Brevetoxins (PbTx) Activates voltage-gated

Na+ channels“Florida Red Tide”, “Neurotoxic Shellfish Poisoning”

Gambierdiscus toxicus Ciguatoxin (CTx) Activates voltage-gatedNa+ channels

“Ciguatera”

Alexandrium spp. Saxitoxin (STx) Inhibits voltage-gatedNa+ channels

“Paralytic Shellfish Poisoning”

Dinophysis spp.,Prorocentrum lima

Okadaic Acid (OA), Dinophysistoxins,Pectenotoxin, Yessotoxin

Phosphatase inhibition “Diarrhetic Shellfish Poisoning”

Protoperidiniumcrassipes a

Azaspiracids (AZA) Unknown Gastrointestinal and neurological symptoms

Bacillariophyta(“diatoms”)Pseudonitzchia spp. Domoic Acid (DA) Glutamate receptor agonist “Amnesic Shellfish Poisoning”

Monera (Bacteria)CyanobacteriaMicrocystis aeruginosa(and other genera/species)

Microcystins Inhibit Ser/Thr phosphatases(PP1, PP2a)

Hepatotoxic, tumor promotion

Nodularia spumigens Nodularin Inhibit Ser/Thr phosphatases(PP1, PP2a)

Hepatotoxic, tumor promotion

Anabaena spp. Anatoxin-a; STx Mimics acetylcholine(see STx, above)

Neurotoxic

Aphanizomenonflos-aquae

Anatoxin-a(s); Anatoxin, STx Acetylcholinesterase inhibition(see anatoxin-a and STx, above)

Neurotoxic

Cylindrospermopisraciborskii

Cylindrospermopsin Inhibits protein synthesis Hepatotoxic

Species/genera and their toxins, as well as the associated targets and toxicoses, are given.a Proposed to be the algal source of azaspiracids, but not confirmed.

63J.P. Berry et al. / Comparative Biochemistry and Physiology, Part C 145 (2007) 61–72

commonly associated with diarrhogenic effects (i.e. diarrheticshellfish poisoning [DSP]), perhaps due to cell-specificcytotoxicity to intestinal cells, although the link between phos-phatase inhibition and this effect is still unclear (Leira et al.,2003). Protein phosphatases are also recognized to be involvedin wide array of signal transduction pathways and various otherrelated cellular functions including those related to maintainingcellular viability and coordinating cell division. Accordingly,recent studies have demonstrated that OA impairs cell division,particularly via interference with F-actin (Fiorentini et al., 1996;Leira et al., 2003) and inhibition of chromosomal separation,leading to aneuploidy and polyploidy (Hégarat et al., 2005), bothof which may be related to the general mechanism of phos-phatase inhibition.

In other cases, such effects are presumably unrelated to therecognizedmodes-of-action commonly associatedwith the toxins.For example, brevetoxins (PbTx) from the Red Tide dinoflagel-late,Karenia brevis, are typically associatedwith potent activationof voltage-gated sodium channels and consequent neurotoxicity,but have recently been shown to cause DNA damage in humanleucocytes at picomolar concentrations (Sayer et al., 2005). Like-wise, domoic acid (DA), most commonly associated with neuro-toxic effects, specifically including so-called amnesic shellfishpoisoning (ASP), is now recognized to exert genotoxic effects viaDNA damage at equally low concentrations (Dizer et al., 2001).

Several of the marine HAB toxins have been investigated asdevelopmental toxins, specifically including studies employing

embryos of zebrafish or other freshwater fish species (seebelow). For example, recent studies of DA by Tiedeken et al.(2005) utilized microinjection of zebrafish eggs to demonstratemorphological and behavioral dysfunction in embryos treatedwith the toxin. Specifically, dose-dependent effects were ob-served on hatching rate, and neurobehavioral parameters, in-cluding convulsions, stereotypically incessant movement ofpectoral fins and impairment of touch–swim response, as wellvarious morphological deformities (e.g. pericardial swelling,curvature of the spine) that consequently decreased survival rate(Tiedeken et al., 2005). Given the recognized neuroexcitatoryeffects of DA via glutamate receptors, the observed behavioraldysfunction is perhaps not surprising, however, specific mechan-isms for these and other effects, particularly including morpho-logical abnormalities, have not been clarified. That said, observedbehavioral effects of DA in zebrafish embryos parallel thosereported for rodent models (Xi et al., 1997; Doucette et al., 2000)that specifically indicate structural changes in the hippocampus ofDA-exposed neonatal rats (Dakshinamurti et al., 1993).

Similarly, Lefebvre et al. (2004) demonstrated, using thezebrafish embryo, apparent impairment of neurodevelopmentby neurotoxic saxitoxin (STx), the causative agent of paralyticalshellfish poisoning (PSP). Exposure of zebrafish embryos toSTx in aqueous rearing media from 0 to 6 days post fertilization(dpf) resulted in consistent morphological abnormalities, in-cluding edemas of the eye, pericardium and yolk sac, as well ascurvature of the spine, particularly at the highest concentrations

64 J.P. Berry et al. / Comparative Biochemistry and Physiology, Part C 145 (2007) 61–72

(≥372 μg STx/L). Moreover, zebrafish exposed to all con-centrations of STx showed stage-specific sensorimotor deficits,including reduced touch response and total paralysis. Interest-ingly, there was an apparent recovery from paralysis by 6 dpf atlower doses, and, furthermore, the observed developmentaleffects, including some degree of the physical abnormalities,were reversible following depuration in STx-free water(Lefebvre et al., 2004). These latter observations may provideinsight to a possible mechanism. Specifically, the authors sug-gest that more pronounced inhibition of voltage-gated sodiumchannels in the Rohon–Beard (RB) subpopulation of neurons,versus less STx-sensitive dorsal root ganglions (DRGs) whichreplace the RB neurons after ∼5 dpf in developing embryos,may explain the eventual recovery from paralysis in the lowerdoses. Furthermore, the authors suggest, based on previousrelated observations, that some amount of the physical mal-formations may be related to impaired movement and paralysis.Specifically, it is proposed that this paralysis results in fluidimbalance, and subsequent edemas that, in turn, lead to cra-niofacial abnormalities; this mechanism would consequentlyexplain the apparent reversibility of the observed edemas fol-lowing depuration and recovery from paralysis (Lefebvre et al.,2003). Interestingly, based on these proposed explanations,the observed effects could consequently be attributed largely toneurotoxicity of STx (i.e. inhibition of neuronal function),rather than cytotoxicity, genotoxicity or other mechanism, asgenerally proposed above for developmental toxins.

In addition to the zebrafish embryo, developmental toxicityof several marine HAB toxins has been shown in other fishmodels of development, and specifically the Japanese medaka(Oryzia latipes) embryo. For example, both PbTx and the che-mically related ciguatoxin (CTx) which both act as neurotoxinsvia activation of voltage-gated sodium channels, have beenshown to impair development of medaka embryos (Edmundset al., 1999; Kimm-Brinson and Ramsdell, 2001; Colman andRamsdell, 2003; Colman et al., 2004). Developmental effects,following microinjection of medaka oocytes with both PbTxand CTx, were observed at sub-nanogram and -picogram (peregg) quantities, respectively, and included, in both cases, cur-vature of the spine, hyperkinesis and cardiovascular (i.e. heartrate) effects (Edmunds et al., 1999; Kimm-Brinson andRamsdell, 2001; Colman and Ramsdell, 2003; Colman et al.,2004). Interestingly, there was an observed difference betweentwo congeners of the brevetoxins tested, namely PbTx1 andPbTx3, in terms of their cardiovascular response; specifically,PbTx3-treated (as well as CTx-treated) embryos exhibitedpersistent tachycardia (Colman and Ramsdell, 2003), asopposed to delayed bradycardia in those exposed to PbTx1(Kimm-Brinson and Ramsdell, 2001). In addition, both PbTx1and PbTx3 lead to apparent dysfunction in hatching of embryos,specifically by abnormal “head-first hatching”, as opposed tonormal, “tail-first” emergence, consequently resulting in in-creased mortality among the toxin-treated embryos (Kimm-Brinson and Ramsdell, 2001; Colman and Ramsdell, 2003).

Developmental toxicity of the recently discovered HAB toxin,azaspiracid-1 (AZA-1), which has been associated with severehuman intoxication via consumption of contaminated shellfish,

was also recently investigated utilizing themedaka embryomodel(Colman et al., 2005). Dose-dependent effects were seen in termsof reduced cardiovascular function (i.e. bradycardia), delayedonset of blood circulation and pigmentation, and decreasedhatching success, as well as general retardation of growth athigher concentrations (≥40 pg/egg). Interestingly, when extractsof AZA-contaminated mussels were evaluated in this system,additional hyperactivity of embryos was observed, perhaps due toother forms of the toxin (i.e. AZA-2 and -3) or other toxins (i.e.OA and dinophysistoxin-2) also detected in the sample.

Similar evaluations of PbTx, CTx and AZA (and othermarine dinoflagellate toxins) in the zebrafish system remain tobe conducted, though it has been suggested in standardizationstudies that data for toxicity tests in zebrafish and medakaembryos are largely comparable (Braunbeck et al., 2005).Moreover, with continuing standardization of the zebrafish as atoxicological model more generally, it is highly likely thatstudies of these, and other marine HAB toxins, in the zebrafishsystem will soon emerge.

1.2. Freshwater fish as models for developmental toxins fromcyanobacteria

In addition to these various eukaryotic marine species(discussed above), another group of HAB organisms, namelythe prokaryotic cyanobacteria, or “blue-green algae”, is re-ceiving growing attention in terms of human and environmentalhealth concerns. The cyanobacteria are recognized to produce adiverse array of toxic or otherwise biologically activemetabolites (see Gerwick et al., 2001). Although ubiquitousin their distribution, cyanobacteria as HABs are almost ex-clusively associated with freshwater environments where theyhave been linked to a range of acute and chronic illness, parti-cularly via direct exposure to the toxin in contaminated drinkingwater or similar routes (Hitzfield et al., 2000). Examples ofperhaps the best-recognized toxigenic cyanobacteria associatedwith various negative impacts as HABs are presented in Table 1.

Of these, the most widely recognized and best characterizedare the microcystins (MCs). Approximately 70 variants of theseheptapeptide toxins have been described, particularly from thefreshwater species, Microcystis aeruginosa, although they havealso been found in several other species/genera (Botes et al.,1982; Namikoshi et al., 1990, 1992). As potent inhibitors ofprotein phosphatases, and specifically the PP1- and PP2A-typecatalytic subunits, the microcystins are associated with a rangeof acute effects, including arrest of cardiac and renal functionand circulatory shock (Beasley et al., 2000; Fischer and Dietrich,2000). However, as they accumulate primarily in the liver(specifically via ATP-dependent transporters in hepatocytes)where they cause hemorrhaging and various severe effects, themicrocystins are largely considered hepatotoxins. Moreover, inaddition to these acute effects, exposure to microcystins, asrecognized tumor promoters, has also been associated specifi-cally with increased incidence of primary liver cancer (Yu, 1989;discussed above).

Given this recognized biological activity, developmentaltoxicity of the microcystins has been investigated using

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zebrafish and other freshwater fish models in several studiesspanning nearly ten years. The first reported study (Oberemm etal., 1997) investigated developmental toxicity of microcystin–LR (MC–LR), the most common of the microcystin variants,using the zebrafish embryo model. Specifically, these investi-gators utilized direct submersion of embryos in MC–LRtreatments to assess developmental toxicity. Using thisapproach, it was shown that MC–LR had no overt effect onembryonic development (0 to 6 dpf), compared to controls,though subsequent rearing of the pre-exposed larvae inmicrocystin-free water for 21 days showed that MC–LRexposure decreased survival rate and body weight by as muchas 40% and 25%, respectively, at the highest concentrations (5and 50 μg/L), as well as minimally reducing body length(Oberemm et al., 1997).

More recently, however, the investigation of the develop-mental toxicity of microcystins has been furthered considerablyby the use of microinjection of both medaka (Jaquet et al., 2004)and zebrafish (Wang et al., 2005) embryos. Unlike many of themarine HAB toxins (discussed above), microcystins are water-soluble metabolites. Recognizing that the chorion may present asignificant barrier to the water-soluble toxin, Jaquet et al. (2004)employed microinjection techniques to expose medaka embryosto MC–LR by direct injection into one-cell stage embryos, andintravitelline injection during late neurulation (1 dpf) and earlyorganogenesis (2 dpf) of embryos. At MC–LR concentrationsas low as 100 pg/mL, injection of embryos at all three stagesreduced survival (0–10 dpf) by as much as 96%, as well asreducing time-to-hatch by 1–3 days (Jaquet et al., 2004).Moreover, surviving embryos showed hepatobiliary abnormal-ities, including hypertrophy, hepatic hemorrhage and necrosis atlater stages of development (Jaquet et al., 2004). Subsequenthistological analysis of medaka embryos injected with sub-lethal concentrations of MC–LR (Huynh-Delerme et al., 2005)further revealed an array of development defects of the digestivesystem and related organs, including lack of mucus-secretingcells in the esophagus, decreased folding of the stomach andintestinal epithelium, reduced exocrine pancreas and liver, andlack of inflation of the swim-bladder, as well as inhibition ofyolk resorption and depletion of hepatic glycogen stores.

Based on these studies in medaka, Wang et al. (2004) utilizedsimilar microinjection techniques to investigate developmentaltoxicity of MC–LR in zebrafish embryos. Injection of zebrafishembryos at 2- to 4-cell stage significantly decreased survival(0–48 hpf) in a dose-dependent manner at concentrations as lowas 300 nM (Wang et al., 2004). Assessment of embryos duringcleavage indicated that MC–LR blocked the adherence andcoherence of blastomeres in dividing cells, and subsequenthistological analysis revealed that the toxin specificallyinterfered with localization of β-catenin and cadherin that areinvolved in cell–cell adhesion (Wang et al., 2004). At laterstages of development, MC–LR injection led to a range ofdevelopmental deformities including bent tails/body axes,occasional cyclopia and severe edemas of the pericardium andhatching gland (Wang et al., 2004). Taken together with resultsfrom medaka studies (Jaquet et al., 2004; Huynh-Delerme et al.,2005), and previous zebrafish studies (Oberemm et al., 1997),

these findings support a developmental toxicity of MC–LR, aswell as clarifying the apparent role of the chorion as a potentialbarrier to water-soluble toxins, such as the microcystins.

With respect to this latter point, developmental toxicity ofMC–LR has also been investigated in loach (Misguruns mizo-lepis), as an alternative freshwater fish model, by direct sub-mersion of embryos (Liu et al., 2002). Interestingly, and incontrast to results with the zebrafish, exposure by immersioninduced an array of embryonic abnormalities, including peri-cardial edema, tubular heart, bradycardia, homeostasis, pooryolk resumption, small head and curved body and tail, as well asa reduction in hatching rate. Moreover, the magnitude of theseeffects were apparently related to developmental stage ofexposure such that embryos exposed during cleavage (approx-imately 32-cell stage) showed more pronounced effects thanthose exposed immediately post-fertilization, whereas thoseexposed even later, specifically during gastrulation, showedconsiderably less profound effects than those exposed duringeither post-fertilization or cleavage stages. Exposure of post-hatch larvae continued to reduce survival (i.e. increase mortality)of embryos, however, abnormalities were limited to ultrastruc-tural effects, specifically interfering with subcellular organiza-tion of hepatocytes, including abnormalities of the nuclei andchromatin, as well as the endoplasmic reticula (Liu et al., 2000).No apparent morphological or cellular abnormalities wereobserved for exposed juveniles, although MC–LR did lead tosignificantly higher mortality compared to controls (Liu et al.,2000). Unlike studies using zebrafish, it was shown that larvaetransferred to microcystin-free water following exposure toMC–LR did not show continued effects on survival rate or bodysize of juveniles (Liu et al., 2002). Clearly these results indicatethat, although zebrafish is finding considerable utility, charac-terization of developmental toxicity may benefit from compar-ison with additional available fish models to address possiblespecies-specificity.

Finally, in addition to the discussed application of thezebrafish and other freshwater fish embryos for characterizingdevelopmental toxicity of known toxins from cyanobacteria andother HABs, it is becoming clear that this model may hold equalutility for identifying otherwise unknown, and potentially novel,toxins. Indeed, Oberemm et al. (1997) reported that crudeextracts from cultures of M. aeruginosa and other species (e.g.Anabaena), as well as several field samples from cyanobacterialblooms (particularly dominated by Aphanizomenon flos-aquae),led to various severe developmental effects (Oberemm et al.,1997). However, these effects which included apparent in-hibition of gastrulation, neural tube formation and organogen-esis more generally did not seem to correlate with the presence ofany of the recognized toxins (i.e. microcystins, anatoxin-1,aphantoxin) from these genera (Oberemm et al., 1997), in-dicating that observed developmental toxicity may potentiallybe related to otherwise unknown toxins.

Likewise, the zebrafish system was used to identify the novellactone, mueggelone, along with the known compound, lupenylacetate, from blooms of A. flos-aquae, as developmental toxins(Papendorf et al., 1997). At concentrations of 10 μg/L purifiedmueggelone, exposed zebrafish embryos showed signs of

Table 2Isolates of freshwater cyanobacteria identified to produce developmental toxinsin the zebrafish embryo assay

Isolate Order: Family Extract

Anabaena 66-2 Nostocales: Nostocaceae CHCl3Calothrix 3-27 Nostocales: Rivulariaceae CHCl3Calothrix 22-7 Nostocales: Rivulariaceae CHCl3“Coccoid” 21-11 Chroococcales: N.A. 30% EtOHFischerella 52-1 Stigonemetales:

StigonemetaceeaeCHCl3

Leptolyngbya 69-5 Oscillatoriales:Pseudanabaenaceae

CHCl3

Limnothrix 60-1 Oscillatoriales:Pseudanabaenaceae

CHCl3

Nostoc Ev-1 Nostocales: Nostocaceae 30% EtOHNostoc 23-2 Nostocales: Nostocaceae 30% EtOHNostoc 53-2 Nostocales: Nostocaceae 30% EtOHNostoc 58-2 Nostocales: Nostocaceae 30% EtOHNostoc 31-4 Nostocales: Nostocaceae CHCl3Nostoc 33-10 Nostocales: Nostocaceae CHCl3Nostoc 15-7b Nostocales: Nostocaceae CHCl3Oscillatoria 48-3 Oscillatoriales: Oscillatoriaceae CHCl3Plectonema 33-7 Oscillatoriales: Oscillatoriaceae CHCl3Pseudoanabaena 69-7 Oscillatoriales:

PseudanabaenaceaeCHCl3

Scytonema 26-1 Nostocales: Scytonemataceae 30% EtOH, CHCl3Spirulina 21-0 Oscillatoriales: Oscillatoriaceae CHCl3Synechococcus 21-10a Chroococcales: N.A. 30% EtOHTolypothrix 30-1-4 Nostocales: Nostocaceae CHCl3

All strains were isolated, by standard microbiological methods (Rippka et al.,1979), from freshwater sources in South and Central Florida. Isolates were culturedand extracted in CHCl3 and 30% EtOH, and evaluated for developmental toxicity,using 4- to 32-cell stage embryos, from 1 hpf to 6 dpf (see Materials and methods).

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impaired development of the circulatory system, as well as peri-cardial edema and thrombosis, and reduced (45%) survival.Lupenyl acetate, on the other hand, although it showed similareffects (e.g. pericardial edema), as well as unrelated effects (e.g.bent tails), was active only at higher concentrations (i.e. 100 μg/mL).

These studies clearly demonstrate that the zebrafish hasconsiderable utility for investigating both otherwise unrecog-nized developmental toxicity of known HAB toxins, as wellidentification and characterization of potentially novel toxicmetabolites from marine and freshwater algae. Accordingly, wehave employed the zebrafish embryo model as a means toscreen cyanobacteria isolates, specifically from the FloridaEverglades and other South/Central Florida freshwater sources,that are rich in microbial (and particularly cyanobacterial)diversity. This approach is used to generate “leads” to toxinsbased on observed inhibition of development, and has sub-sequently been employed to direct purification (i.e. bioassay-guided fractionation) and eventual chemical and biologicalcharacterization of compounds identified in this way. Here wepresent data on developmental toxins from freshwater cyano-bacteria identified using the zebrafish embryo model, anddiscuss the utility of this assay for isolation and characterizationof such cyanobacterial metabolites.

2. Materials and methods

2.1. Isolation and culture of freshwater cyanobacteria

All strains of cyanobacteria were isolated from samples ofwater or algal assemblages (e.g. periphyton mats) collected fromthe Florida Everglades and other freshwater sources in South andCentral Florida. Strains were isolated by standard microbiolog-ical procedures (Rippka et al., 1979), specifically employingfiltration of samples through a 0.45 μm membrane filter, sub-sequently transferred to BG11/agar (1.5%) plates which areselective for cyanobacteria and certain “green algae” (Chlor-ophyta). Plates with filters were incubated at room temperature(24 °C) under continuous light (25 μE/m2/s), and individualcolonies picked, and subsequently grown as non-axenic,unialgal cultures in BG11 medium buffered with 2-morpholi-noethanesulfonic acid (MES), pH 7.2, to obtain sufficientbiomass (see below) for extraction. All isolates were taxonom-ically identified (to genus) by microscopic observation usingclassical morphological criteria given in Komarek and Ana-gnostidis (1986, 1989).

2.2. Extraction of toxins from cyanobacterial isolates

For screening, biomass from cyanobacterial cultures wassequentially extracted in CHCl3 and 30% EtOH in order toensure sufficient representation of non-polar (i.e. lipophilic) and“semi-polar” compounds, respectively. Specifically, biomasswas collected by centrifugation, filtration or direct “harvesting”(e.g. collection with forceps), and freeze-dried. Freeze-driedbiomass (approximately 100 mg) was extracted by sonication inCHCl3 (10 mL), followed by filtration and evaporation of the

extraction solvent. Subsequently, the extracted biomass wasextracted similarly in 30% EtOH, followed by filtration andevaporation of the extraction solvent. Dried CHCl3 and 30%EtOH extracts were re-dissolved to 1 mg crude extract per mL inabsolute EtOH and 30% EtOH, respectively, for screening.

2.3. Rearing and breeding of zebrafish for embryos

Zebrafish of the D and DGL lines (Gibbs and Schmale,2000) were bred and maintained in the UM NIEHS Marine andFreshwater Biomedical Sciences (MFBS) Center’s In Vitro andIn Vivo Fish Culture Facility Core. Zebrafish were maintained in30-L tanks at 28 °C with 14 h:10 h light/dark cycle. To breed,male and female adult zebrafish (typically ca. 10–30 indivi-duals) were collected (within 15 min of the end of the darkcycle), held above 10-L tanks in mesh enclosures and allowed tobreed freely. Following successful breeding (typically 3–5 h),eggs fell through the mesh, and were subsequently collectedfrom the bottom of tanks. Following collection and washing,eggs were transferred to plates containing “embryo rearingsolution” (ERS) prepared fresh by diluting 25 mg of neomycinsulfate, 50 μL stock solution (35% w/v) of sodium thiosulfate,125 μL Amquel® and 200 μL of stock solution (0.1% w/v) ofmethylene blue in 1 L of tap water. Eggs containing dead orobviously poor quality embryos were removed. The remainingembryos were used, usually within 2 h post-fertilization (hpf),for developmental toxicity assays (see below).

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2.4. Zebrafish embryo developmental toxicity assay

Screening of zebrafish embryo developmental toxicity wastypically conducted in 24-well plates. Prior to adding embryos,cyanobacterial extracts were either diluted (typically 100, 50, 20and 10 μg/mL final concentration of crude extracts) in wellscontaining 1 mL of ERS, or alternatively added to wells, andplaced in a fume-hood to allow solvents to evaporate, prior toadding ERS. In addition, appropriate controls of solvent vehicle,along with untreated controls, were included in each assay.Subsequently, embryos at the 4- to 32-cell stage (except whereotherwise noted; see below) were collected and transferred totest wells (typically 5 embryos per well), by pipetting to mini-mize damage to eggs.

Embryo development was observed daily (for 5 dpf) using adissecting microscope, and any abnormal developmental phe-notypes (particularly morphological abnormalities), as well asmortalities, were recorded and photographically documented.Morphological abnormalities were assessed based primarily ondevelopmental features described by Kimmel et al. (1995), andgenerally categorized when possible as early-, mid- and late-stage phenotypes. “Early-stage” phenotypes included apparentinhibition of cleavage, blastulation and gastrulation (0–24 hpf);“mid-stage” phenotypes included interference with segmenta-tion, organogenesis and pharyngulation (24–48 hpf); “late-stage” phenotypes include those developmental dysfunctionsthat manifested primarily post-hatching (2–5 dpf). Assays wererepeated as necessary to confirm any observed developmentaleffects. Furthermore, to ensure that developmental toxicity wasconsistent and reproducible, these effects were specificallyrecorded as “positive”when observed in 80% (i.e. 4 of 5, or 8 of10) of embryos tested. For untreated embryos, mortality and

Fig. 1. Developmental toxicity of Oscillatoria 48-3 extracts. All embryos shown (A–chloroform extracts (20 μg/mL) of Oscillatoria 48-3 (A–C) arrest development atcompared to control embryos treated with solvent only (E). In some cases, embryos deat the same time with solvent only (E). Embryos treated at MBT (D) developed nor

spontaneous developmental dysfunction typically occurred atrelatively low frequency (approximately 5% of embryos).

3. Results

To-date, over 122 strains of cyanobacteria isolated from theFlorida Everglades and other freshwater sources in South andCentral Florida have been screened with the zebrafish deve-lopmental toxicity assay. Approximately 17% of the strainsevaluated were found to produce compounds that inhibit embryodevelopment (Table 2). Developmental toxicity was observedfor both lipophilic (CHCl3) and polar (30% EtOH) extracts,although considerably more lipophilic fractions were active (i.e.71% vs. 41%). Taxonomically, active strains represented sixfamilies from four orders of cyanobacteria. Some taxa weremorerepresented than others; in particular, 7 of the active strainsbelonged to the genus, Nostoc, and 8 belong to the familyNostocaceae (Table 2).

Cyanobacterial toxins inhibited zebrafish embryogenesis atvarious stages of development including early, middle and latephases (see Materials and methods). Chloroform extracts of Os-cillatoria 48-3, for instance, inhibited early stages of develop-ment. At 1 dpf, greater than 80% of embryos, treated at cleavage(4- to 32-cell stage) with 20μg/mLof crude extract, were halted atmid-blastula transition (MBT; Fig. 1A) or gastrulation (Fig. 1B).Approximately 20% of embryos treated at 20 μg/mLwere able toproceed beyond gastrulation at 1 dpf, however, the developmentof these embryos was notably retarded compared to controlembryos (Fig. 1E), including truncated tails and poorly formedeyes and other organs (Fig. 1C). To assess the affect of deve-lopmental stage during exposure, embryos were additionallyexposed at 1-cell, 64- to 128-cell and MBT stages. Again,

E) are 1 dpf. Within 1 dpf, zebrafish embryos exposed at 4- to 32-cell stage tomid-blastula transition (MBT; A), or are slow to progress to gastrulation (B),velop beyond gastrulation (C), but are notably slowed compared embryos treatedmally as solvent controls (E).

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approximately 80% of embryos exposed to 20 μg/mL of extractprior toMBT (i.e. 1-cell and 64- to 128-cell stages) were halted atMBT or gastrulation (data not shown). On the other hand, allembryos exposed atMBT (also at 20 μg/mL) developed normally(Fig. 1D), and appeared similar to control embryos (Fig. 1E),indicating that the toxin specifically affects developmentalpathways preceding MBT. At concentrations of 50 μg/mL orgreater, all embryos, regardless of developmental stage duringexposure, were dead within 1 dpf. Embryos treated at lowerconcentrations (i.e. 10 μg/mL), generally developed normally,though embryos were slightly smaller compared to controlembryos (data not shown).

Extracts of Plectonema 33-7 and Tolypothrix 30-1-4, on theother hand, apparently affected embryos during (or perhaps priorto) pharyngulation (1–2 dpf), and specifically appear to inhibitpathways of organogenesis and differentiation more generally(Fig. 2A, B, D and F, respectively) when compared to controls(Fig. 2C, E and G). As shown in Fig. 2, eyes failed to developbeyond a rudimentary stage in embryos treated with chloroformextracts from either of the two strains (at 50 and 100 μg/ml,

Fig. 2. Developmental toxicity of Plectonema 33-7 and Tolypothrix 30-1-4 extracts.chloroform extracts from Plectonema 33-7 (A, B and D) and 100 μg/mL of chlopharyngulation, including organogenesis (e.g. formation of the eye) and general cell-d(C and E), as solvent-only control for A, B and D, or 100 μL of CHCl3/mL of ERadditionally an apparent lack of migration of melanophores from neural crest (B)magnification.

respectively). Likewise, in both cases, embryos developed onlyrudimentary heads and truncated tails, along with a generallymalformed body axis (Fig. 2A–B, D). Embryos, however,survived up to 3–4 dpf, and were responsive to light stimulationby stereotypical twitching behavior.

In the case of embryos treated with extracts of Plectonema33-7, the phenotype was specifically characterized by anapparent failure of melanophores to migrate from their originin the neural crest, consequently aggregating in and around thehead (Fig. 2A, B and D). This phenotype was highly consistentand reproducible (Fig. 2D), indicating inhibition of a specificdevelopmental pathway(s), rather than general dysfunction.Exposures to Plectonema 33-7 extract was repeated using eggsat 1-cell, 64- to 128-cell andMBTstages, however, no differencewas observed (data not shown), indicating that the toxin targetedpathways later in the development (i.e. post-MBT).

Chloroform extracts (100 μg/mL) of Calothrix 33-2 (Fig. 3)induced various morphological deformities in embryos at laterstages that largely manifested only post-hatching (ca. 4–5 dpf).Although embryos developed otherwise normal body plans, and

Within 2 dpf, zebrafish embryos exposed at 4- to 32-cell stage to 50 μg/mL ofroform extracts from Tolypothrix 30-1-4 (F), respectively, show inhibition ofifferentiation, compared to embryos treated with either 50 μL CHCl3/mL of ERSS (G), as solvent-only control for F. In the case of Plectonema 33-7, there isrelative to the solvent-only control (C and E). Note: D and E are the same

Fig. 3. Developmental toxicity of Calothrix 33-2 extracts. Within 4 dpf,zebrafish embryos exposed at 4- to 32-cell stage to 100 μg/mL chloroformextracts of Calothrix 33-2 (B) show morphological deformities, includingpericardial and peritoneal edemas, as well as bent tails/body axes, compared tocontrol embryos treated with solvent (i.e. CHCl3) only (A).

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organs generally elaborated as normal, embryos were consis-tently and reproducibly characterized by severe edemas ofpericardial and peritoneal cavities, and considerably bent tailsand/or body axis (Fig. 3). Embryos, however, successfullyhatched and were otherwise seemingly normal behaviorally –swimming and righting themselves, for example, as well asotherwise responding to stimuli (e.g. touch, light) similarly to

Fig. 4. Developmental toxicity of crude extracts from Fischerella 52-1. Within1 dpf, zebrafish embryos exposed at 4- to 32-cell stage to chloroform extracts ofFischerella 52-1 culture medium (left) show a developmental phenotype, termed“puffy” ,characterized by swollen appearance and general lack of pigmentation,compared to control embryos treated with solvent only (right).

control embryos. However, eventual survival of the extract-exposed larvae to adults was significantly lower than controlembryos (data not shown).

One isolate identified by screening, Fischerella 52-1, wasspecifically selected for subsequent isolation and characterizationof the developmental toxin. Embryos treated with chloroformextracts of Fischerella 52-1 culture biomass, or alternativelyculture medium, consistently and reproducibly developed a char-acteristic phenotype termed “puffy”, characterized by a generallyswollen appearance and lack of pigmentation (Fig. 4) whichdeveloped within approximately 1–2 dpf (followed by 100%mortality within 2–3 dpf). This developmental phenotype wasused to guide purification (i.e. bioassay-guided fractionation) of agroup of developmental toxins from Fischerella 52-1 for sub-sequent chemical and biological characterization. Interestingly,certain aspects of the phenotypes for purified compounds (Fig. 5)were not identical to that of the crude extract, although otherfeatures, such as the lack of pigmentation, and particularlymelanophores, were observed in both cases. However, as with thecrude extracts, the observed phenotype for embryos exposed tothe purified compounds was equally consistent for a given set oftreatment conditions (Fig. 5). Details of the purification andchemical characterization are presented elsewhere (Berry et al.,submitted for publication).

Fig. 5. Developmental toxicity of a compound purified from Fischerella 52-1.Within 3 dpf, zebrafish embryos exposed at 4- to 32-cell stage to 100 μg/mL of apure compound isolated from Fischerella 52-1 (A) show a developmentalphenotype characterized, in particular, by lack of melanophores, compared tocontrol embryos (B) treated with solvent (i.e. CHCl3) only. Moreover, thisphenotype is consistent for all embryos exposed to the toxin under a given set ofconditions (e.g. toxin concentration, stage and line of fish).

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4. Discussion

The zebrafish embryo (along with other teleost fish models,e.g. medaka) is finding growing utility in the investigation ofHAB toxins, particularly with respect to the effects of thesetoxins on processes of vertebrate development (i.e. develop-mental toxicity). This model system has been effectivelyemployed to identify developmental toxicity associated with anumber of well-recognized toxins from marine and freshwaterHABs (Oberemm et al., 1997; Edmunds et al., 1999; Kimm-Brinson and Ramsdell, 2001; Liu et al., 2002; Colman andRamsdell, 2003; Colman et al., 2004, 2005; Lefebvre et al.,2004; Tiedeken et al., 2005). Equally, this system is beingincreasingly utilized to identify and characterize novel, or other-wise unrecognized, developmental toxins from algal sources(e.g. Papendorf et al., 1997).

We have specifically applied this model system to theidentification and characterization of developmental toxinsfrom freshwater cyanobacteria, including over 100 isolates fromthe Florida Everglades and other freshwater sources in Southand Central Florida. Evaluation of these isolates has demon-strated that potential developmental toxins are widespread.Indeed, extracts from approximately 17% of isolates tested,representing a considerable diversity of the cyanobacteria(Table 2), inhibited development of the zebrafish embryo.

Moreover, developmental toxins identified from cyanobac-teria in the present study appear to affect a range of deve-lopmental stages (Figs. 1–3), and it is thereby becomingincreasingly clear that stage of exposure to toxins may play animportant role in the manifestation of these toxicoses. Lipophilicextracts ofOscillatoria 48-3, for example, appear to specificallyinhibit pathways up to mid-blastula transition (MBT), as treatedembryos are largely arrested at MBT, or slowed in their progressto gastrulation and subsequent stages, and treatment of embryosat equivalent concentrations after MBT has no apparent effect ondevelopment (Fig. 1). Others, such as Plectonema 33-7 andTolypothrix 30-1-4, appear to inhibit processes of pharyngula-tion (1–2 dpf), including organogenesis and cell-differentiationmore generally (Fig. 2). In other cases, such as Calothrix 33-2(Fig. 3), developmental defects are not apparent until embryoshatch (∼4–5 dpf) and morphological malformations (e.g. bentbody axis, edema) are observable.

In addition, the zebrafish embryo has been successfully usedfor identification, purification and characterization of develop-mental toxins. As an example, utilizing the zebrafish embryomodel for bioassay-guided fractionation, we have recently puri-fied developmental toxins from the isolate, Fischerella 52-1.Using either crude extracts or pure compounds, consistentdevelopmental phenotypes were observed, though some degreeof variability was observed between the phenotype associatedwith the crude extracts compared to the purified compounds (seeFigs. 4 and 5). Details of the purification and characterization ofthese compounds is reported elsewhere (Berry et al., submittedfor publication), however, chemical characterization has iden-tified these compounds as belonging to the hapalindoles, a groupof indole alkaloids previously identified from Fischerella andrelated genera (Moore et al., 1984, 1987; Smitka et al., 1992;

Stratmann et al., 1994). Although previous work (Moore et al.,1997; Smitka et al., 1992; Doan et al., 2000; Doan et al., 2001)has characterized biological activity of these compounds (e.g.antimicrobial activity, inhibition of RNA polymerase), this is thefirst identification of apparent developmental toxicity associatedwith these compounds. This may be of particular importance asthese compounds are currently being developed as anticanceragents (Smith et al., 1995; Zhang and Smith, 1996). Moreover,through our studies it was found that these toxins are abundant inextracellular components (i.e. culture medium) of cultures(Gantar et al., submitted), and may be released by naturallyoccurring algal populations, representing a potential environ-mental and even human health concern.

Perhaps the most striking outcome of this research was theconsistent phenotype, with very little observable morphologicaldifferences between embryos, seen after exposure to either crudeextracts, such as those from Plectonema 33-7 (Fig. 2B–C) orFischerella 52-1 (Fig. 4), or a purified compound (Fig. 5).This consistency clearly suggests that these toxins are inhibitinga specific developmental pathway, or set of pathways andprocesses, rather than resulting from general dysfunction ofembryogenesis. In each case, the consistent phenotype wasfound with a specific concentration given to same stage embryosof one particular line of fish. Although these phenotypic out-comes are reproducible within these conditions, further work isneeded to determine the range of conditions that yield consistentphenotypic outcomes. Likely key variables include toxin con-centration, stage of application and line of fish.

Marine and freshwater HABs are recognized to produce avast array of toxic or otherwise biologically active metabolites.Although most research has focused on the acute toxicosesassociated with these toxins, it is becoming clear that the samecompounds may exert, via identical, related or alternative tar-gets, more insidious and often unrecognized outcomes. Sucheffects as developmental toxicity may have profound implica-tions for human and environmental health. The zebrafish embryomodel is finding a considerable role in our understanding ofthese impacts.

Acknowledgements

The authors would like to thank the National Institute ofEnvironmental Health Sciences (NIEHS) for financial supportof this research through an NIEHS ARCH grant (S11 ES11181).This research was additionally supported, in part, by an NIEHSMarine and Freshwater Biomedical Sciences Center Grant (ES05705) to the University of Miami. We would also like to thankDrs. Kathleen Rein, Minglei Wang and Robert E. Gawley whohave contributed to the on-going chemical characterization ofcyanobacterial toxins.

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