Organisms living in an environment displaying a mass occurrence of toxic cyanobacteria are surrounded by water containing either toxic cells and/or dissolved toxins. Those organisms are not only threatened by the ingestion of cyanobacteria and their toxins, toxic cells and contaminated water furthermore flow through their gills and they feed, breed and develop in that water. Hence aquatic organisms can be more profoundly influenced by cyanobacterial toxins than terrestrial animals.
Regarding to this very interesting are fish. As they are located at a high trophic level within the aquatic food web, fish may be directly influenced by cyanobacterial toxins, via accumulation of cyanobacterial toxins from contaminated food (e.g. zooplankton, mussels, fish) (Engström-Öst et al., 2002; Kankaanpää et al., 2005b; Karjalainen et al., 2005; Smith & Haney, 2006), but also via indirect effects on their environment. Indeed, toxic cyanobacteria influence the zooplankton assemblage which may result in a qualitative and/or general loss of food organisms (DeMott et al., 1991; Laurén-Määttä et al., 1997; Sivonen & Jones, 1999). Moreover, cyanobacterial mass occurrences may modulate exogenous factors (e.g. pH-values, oxygen, ammonium, etc.) resulting in an insufficient sometimes even hostile environment (Wiegand & Pflugmacher, 2005 and references therein).
Whether fish are affected by cyanobacteria depends on several factors: Which cyanobacterial species are involved? Do they produce toxins and if yes how much? Are the toxins released into the water and/or accumulated by potential food organisms? Additionally important is to what extent cyanobacteria and a given fish population come into contact with one another and thus whether it is a benthic, stratified, or scum forming bloom or if cyanobacteria are evenly distributed over the entire water body. Besides the sensitivity of the fish species in question it furthermore appears crucial, if the fish recognise and subsequently may avoid an exposure to cyanobacterial toxins. The course and dimension of various cyanobacteria induced fish kills however suggest the latter occurrences to be rather low.
C
YANOBACTERIA INDUCED FISH KILLSWhilst wildlife and predominantly livestock mortalities caused by toxic cyanobacteria regularly evoke great public concern (Edwards et al., 1992; Gugger et al., 2004; Gunn et al., 1992;
Holschuh, 2001; Kuiper-Goodman et al., 1999; Mez et al., 1997; Negri et al., 1995; Wood et al., 2007, etc.), cyanobacteria induced fish mortalities mostly remain unnoticed. Nevertheless toxic cyanobacteria regularly cause considerable fish kills spanning the entire globe, involving almost all toxic cyanobacterial genera and affecting herbivorous, planktivorous, omnivorous and piscivorous fish species (Tab. 1.2).
Fish kills most likely occur with the breakdown of a cyanobacterial bloom (Albay et al., 2003;
Bürgi & Stadelmann, 2002; Nascimento & Azevedo, 1999; Rodger et al., 1994). Typically, environmental conditions change, disturbing cellular buoyancy and resulting in an accumulation of cyanobacterial cells at the water surface. Surface conditions, in particular elevated irradiance, high temperatures, wind, wave action, etc. cause bloom senescence, cyanobacteria cell lyses and toxin release. Fish remaining in those contaminated areas are then unavoidably exposed to cyanobacterial toxins.
The decomposition of the senescent bloom material often simultaneously results in substantial oxygen depletion (Jewel et al., 2003; Nascimento & Azevedo, 1999), which, depending on the dimension of the cyanobacterial bloom, may cause insufficient oxygen conditions. In consequence cyanobacteria induced fish mortalities are caused by either poisoning due to toxin release, anoxia or a combination of both (Malbrouck & Kestemont, 2006 and references therein, Jewel et al., 2003; Pollux & Pollux, 2004; Toranzo et al., 1990). Hence dead fish appear after the disappearance of cyanobacteria cells. Thus, a lot of fish kills as induced by cyanobacterial toxins remain unrealised and are rather attributed to insufficient oxygen conditions (Gaete et al., 1994;
Zambrano & Canelo, 1996). It is therefore most likely that the proportion of fish kills caused by cyanobacterial toxins is actually higher than thought.
Even so, not every cyanobacterial bloom causes a fish kill. Released toxins should usually dissolve rapidly within the water body and toxin concentrations thus mostly remain beneath lethal levels.
Moreover, fish may migrate within the water body and thus avoid an exposure to a toxic cyanobacterial bloom. It is also crucial to what extent exogenous (i.e. temperature, oxygen, alkalinity, etc.) and endogenous factors (i.e. mobility, physiological and nutritional condition, resistance to stress and diseases, etc.) modulate the consequences of a given toxin exposure (Hofer
& Lackner, 1995). Nevertheless, also non-lethal and chronic exposure to toxic cyanobacteria may cause severe organ damage and impairment also resulting in an accumulation of cyanobacterial toxins within exposed fish (Andersen et al., 1993; Carbis et al., 1997; Sipiä et al., 2001a; Sipiä et al., 2002; Xie et al., 2005, etc.).
Tab. 1.2: A summary of documented fish kills associated with cyanobacterial mass occurrence
* haptophyte, no cyanobacterium REFERENCES:
1Schwimmer & Schwimmer, 1968; 2Seydel, 1913; 3Stephens, 1948; 4Mackenthum & Herman, 1948; 5Rose, 1953; 6Davidson, 1959; 7Gorham et al., 1964; 8Devidze, 1998; 9Sawyer et al., 1968; 10Persson et al., 1984; 11Berg et al., 1986; 12Druvietis, 1998;
13Gaete et al., 1994; 14Penaloza et al., 1990; 15Bürgi & Stadelmann, 2002; 16Sevrin-Reyssac & Pleticosic, 1990; 17Toranzo et al., 1990; 18Humpage et al., 1993; 19Nascimento & Azevedo, 1999; 20Rodger et al., 1994; 21Yunes et al., 1998; 22Lindholm et al., 1999; 23Giovannardi et al., 1999; 24Albay et al., 2003; 25Fischer & Dietrich, 2000; 26Zimba et al., 2001; 27Albay et al., 2004;
CYANOBACTERIAL SP.
INVOLVED AFFECTED FISH SPECIES YEAR LOCATION
Anabaena circinalis not specified 1880 div. Polish lakes 1
Anabaena sp. carp (Cyprinus carpio), perch (Perca sp.)
& roach (Rutilus rutilus)
1913 div. German lakes 2
M. aeruginosa not specified 1913-1943 div. South-African lakes 3
Anabaena circinalis not specified 1914 div. Hungary lakes 1
Aphanizomenon sp. not specified 1931-1933 div. lakes in Iowa, USA 1
Anabaena sp. not specified 1940-1942 Lake Ymsen, Sweden 1
Aphanizomenon sp. not specified 1942 Zuiderzee, Holland 1
Aphanizomenon sp. carp, pike (Esox lucius) & perch 1946 div. rivers & lakes in Wisconsin, USA 4 Anabaena sp.,
Microcystis sp. buffalo fish (Ictiobus sp.) & carp 1948 Storm Lake, Iowa, USA 5
Nostoc sp. not specified 1956-1958 Waco, Texas, USA 6
M. aeruginosa, Anabaena sp.
carp, roach, catfish (Ictalurus punctatus)
& bream (Abramis sp.)
Aphanizomenon sp. not specified 1967 Lake Winnisquam,
New Hampshire, USA 9 Oscillatoria sp.
(red coloured)
roach 1982 Vesijarvi, Finland 10; 11
Anacystis cynaea pike & pikeperch (Stizostedion lucioperca) 1982-1987 div. lakes in Latvia 12 Microcystis sp. carp & silverside (Labidesthes sp.) 1984 Aculeo Lake, Chile 13; 14 Aphanizomenon sp. tench (Tinca tinca), bream, roach, perch, pike
& smelt (Osmerus sp.)
1984 Sempacher See, Switzerland 15
Microcystis sp. not specified 1988 Forez, France 16
Anabaena sp. rainbow trout (Oncorhynchus mykiss) 1989 fish farm, Spain 17 Anabaena sp.,
Microcystis sp.
not specified 1991 Darling river, Australia 18
Synechocystis sp. menhaden (Brevoortia tyrannus) 1991 Barra Lagoon, Brazil 19 Anabaena sp.,
Microcystis sp., Aphanizomenon sp.
brown trout (Salmo trutta) 1992 Loch Leven, Scotland 20
Microcystis sp. catfish 1996 Patos Lagoon, Brazil 21
Microcystis sp Planktothrix sp.
smelt & ruffe (Gymnocephalus cernuus) 1994-1996 Lake Ijsselmeer30
*Prymnesium sp., Oscillatoria sp.
perch, roach & bleak (Alburnus alburnus) 1997 Aland, Finland 22 Aphanizomenon sp.,
Oscillatoria sp. small fish ( not specified) 1997 Lago Varese, Italy 23
Planktothrix rubescens not specified 1997 Lake Spanca, Turkey 24
not specified bream, roach, perch, pike
& coregonids (Coregonus sp.) div. Finnish lakes 25
Microcystis sp. not specified div. ponds in southern USA 26
Microcystis sp. not specified 2000-2003 Brackish lake in Turkey 27
A. flos-aquae, M. aeruginosa
carp, tilapia (Oreochromis sp.), catla (Catla catla)
& silvercarp (Hypophthalmichthys sp.)
2002 div. fish ponds in Bangladesh 28 not specified perch, pike, tench, carp
& topmouth gudeon (P. parva)
2003 Romeinwerd, Netherlands 29
As fish is an important food source, detrimental effects such as those induced by toxic cyanobacteria may not only cause significant economic losses in aquaculture (Andersen et al., 1993; Kent, 1990; Toranzo et al., 1990; Zimba et al., 2001), but may also represent a possible route for human exposure (Deblois et al., Toxicon in press; Jewel et al., 2003; Magalhaes et al., 2003; Magalhaes et al., 2001; Soares et al., 2004; Xie et al., 2005) and result in sustained effects on the nutrition of regional populations. Thus detailed investigations on the various and multiple effects of cyanobacterial toxins on fish are essential.
T
HE ICHTHYOTOXICITY OF MICROCYSTINThe median lethal doses (LD50) for microcystins (predominantly MC-LR) have been determined for diverse fish species by different forms of toxin application (Tab. 1.3). Fish species differ in their sensitivity to microcystin, e.g. cyprinids have been shown to be up to 50-times more sensitive than trout (Andersen et al., 1993; Fischer & Dietrich, 2000; Tencalla, 1995). Differences within the same species emphasise an influence of the nutritional and physiological condition on microcystin toxicity to exposed fish (Carbis et al., 1996a; Malbrouck et al., 2004b; Råbergh et al., 1991). Although less sensitive to microcystin via the intraperitoneal route, fish are more sensitive than mice to orally applied microcystin, which suggests fish to be particularly susceptible to the effects of oral ingestion of microcystin and microcystin containing cyanobacteria.
Uptake of Microcystin
Although microcystin containing cyanobacterial blooms have unambiguously been associated with numerous fish kills, the microcystin uptake of fish has not been clarified in detail so far. Indeed, pathological alterations in moribund and dead fish resemble those of fish treated with microcystin (Andersen et al., 1993; Rodger et al., 1994; Toranzo et al., 1990; Zimba et al., 2001). However environmentally detected concentrations of microcystin and cyanobacterial crude extracts have not yet been demonstrated to cause acute fish mortality (Bury et al., 1995; Carbis et al., 1996a;
Phillips et al., 1985; Tencalla et al., 1994).
Tab. 1.3: The median lethal dose (LD50) of microcystin-LR in various fish species and mouse [µG MC-LR/KG BW] ORAL APPLICATION INTRAPERITONEAL
APPLICATION Trout
(Oncorhynchus mykiss)
1700-6600 1 400-1000 2
Carp <1700 6 300-550 3
(Cyprinus carpio) 20-50 4
Loach (Misgurnus mizolepis)
*138 5
Mouse (Mus sp.)
≥5000 7, 8 50 9
* application of MC-LRequiv. (primarily containing MC-RR) REFERENCES:
1Tencalla et al., 1994; 2Kotak et al., 1996; 3Råbergh et al., 1991; 4Carbis et al., 1996a; 5Li et al., 2005; 6Tencalla, 1995;
Principally, there are a variety of possible routes for the uptake of microcystins. Dissolved microcystins might be absorbed from water, via the gills, the intestine or the integument and fish might further be intoxicated due to the digestion of ingested cyanobacterial cells, contaminated food and subsequent microcystin release within the intestine. As deduced from various experimental microcystin exposure of trout, the most likely uptake route is the oral ingestion of either dissolved microcystin and/or microcystin containing cells with a consequent toxin uptake via the digestive system (Bury et al., 1995; Tencalla et al., 1994). Therefore, fish filtering water for food (primarily herbivorous and planktivorous species) appear to run a much higher risk of ingesting toxic cyanobacteria than species that take food selectively (e.g. most omnivorous and piscivorous species), resulting in a species-specific susceptibility to microcystin (Adamovsky et al., 2007; Chen et al., 2007). Xi et al. (2005) detected microcystins in tissue but not in gut content of fish naturally exposed to toxic M. aeruginosa, concluding, that microcystins might moreover be intoxicated via ingestion and digestion of food organisms containing accumulated toxin. However, these observations might also be attributed to far earlier ingestion of cyanobacteria, whereas cyanobacterial cells and toxins were no longer determinable in gut contents because they had already been digested and/or excreted (Soares et al., 2004).
Digestion of cyanobacterial cells in the intestine of fish varies within different cyanobacterial species. Cells of various Aphanizomenon sp. for example have been shown to be almost totally decomposed during fish digestion while others, predominantly species comprising cell walls including additional exopolysaccharides (e.g. Microcystis sp.), were barely digested (Carbis et al., 1997; Gavel et al., 2004; Kamjunke et al., 2002a; Kamjunke et al., 2002b; Lewin et al., 2003).
However, even cell walls of obviously intact cells have been shown to become permeable throughout the gut passage and may thus allow toxin release within the digestive tract (Moore &
Scott, 1985).
As microcystins are highly resistant to acidic and enzymatic degeneration, once released into the fish intestine microcystin degradation is assumed to occur slowly, resulting in high microcystin availability. It is suggested, that microcystins are absorbed across the epithelium of the ileum and thus reach the venous bloodstream of fish via bile acid membrane transporters (Boaru et al., 2006; Fischer & Dietrich, 2000; Meier-Abt et al., 2007). Bury et al. (1998b) conclude from studies on the uptake of radiolabeled microcystin in trout, that microcystins may partly pass the ileal epithelium via passive transport mechanisms. Nevertheless, the ileal epithelium provides a barrier for the transfer of microcystin into the bloodstream, for which reason a substantial proportion of microcystin appears to remain in faeces (Tencalla & Dietrich, 1997;
Xie et al., 2004).
Indeed less than 5% of microcystin orally applied to trout (Oncorhynchus mykiss) has been shown to enter the bloodstream (Bury et al., 1998b; Tencalla & Dietrich, 1997), and also salmon (Salmo salar) intraperitoneally exposed to microcystins incorporated only 38-62 % of the applied microcystin into their tissues (Williams et al., 1997a; Williams et al., 1995). The proportion of
microcystin absorbed into blood has been shown to depend on time, dose, route of exposure and on the chemical properties of the specific microcystin congeners (Bury et al., 1998b; Fischer &
Dietrich, 2000; Tencalla, 1995; Williams et al., 1997a; Williams et al., 1995; Xie et al., 2004).
Ileal absorption of microcystin is assumed to be affected by the constitution and length of the species intestine. Trout possessing a short, thick-walled gut have been shown to be much less sensitive to microcystin than cyprinids which possess a comparably long intestine (Fischer &
Dietrich, 2000; Tencalla, 1995). Differences in the microcystin sensitivity of diverse fish species can, besides the species foraging strategy, also be ascribed to differing organ morphologies, in particular the intestine.
Transport and Organotropism of Microcystins
Having traversed the epithelial barrier, microcystin accumulates primarily in the liver of exposed fish following the portal venous bloodstream and distribution of OATPs (Cazenave et al., 2005;
Fischer & Dietrich, 2000; Soares et al., 2004; Williams et al., 1997a; Williams et al., 1995).
Microcystins have been shown to accumulate in liver of carp (Cyprinus carpio) and trout orally exposed to M. aeruginosa as of 1 h post application (Fischer & Dietrich, 2000, Tencalla et al., 1997). Maximum hepatic microcystin accumulations were usually determined within 3-8 h post application subsequent to intraperitoneal and oral exposure of both, cyprinids and trout (Fischer
& Dietrich, 2000, Malbrouck et al., 2003; Tencalla & Dietrich, 1997; Williams et al., 1995, Williams et al., 1997a).
Smaller amounts have also been shown to accumulate in kidney, blood, gill, bile, intestine and brain while only minimal amounts have been detected in muscle tissue (Cazenave et al., 2005;
Fischer & Dietrich, 2000; Williams et al., 1995; Xie et al., 2005). It is yet not clear, whether microcystins reach those organs directly, via the bloodstream bypassing the hepatopancreas, or downstream of the hepatopancreas subsequent to an overload of presystemic hepatic elimination11 capacities (Carbis et al., 1996b; Fischer & Dietrich, 2000).
Biochemical Changes as Indicators of Microcystin Toxicity in Fish
Upon reaching the metabolism of an exposed organism, the toxicity of a given contaminant is principally based on molecular alterations. Only the interaction(s) of a toxin with receptors or other target molecules, their inhibition or activation initialise physiological, pathological and other functional alterations observable and measurable in exposed organisms (Hofer & Lackner, 1995; Schlenk & Di Giulio, 2002). In this respect, the ichthyotoxicity of microcystin is mainly mediated by four basic molecular effects: inhibition of protein phosphatases (PPs), an inhibition of ATPases, an elevated formation of reactive oxygen species (i.e. oxidative stress) and effects on detoxification via glutathione conjugation.
PP-INHIBITION: Also in fish, PPs appear the main target molecules of microcystin (Råbergh et al., 1991; Tencalla & Dietrich, 1997) and their inhibition causes substantial disturbance of various elementary metabolic processes (Mezhoud et al., Aquatic Toxicology in press; see also chapter 1.2). The timecourse and extent of PP-inhibition has been investigated in several studies including various fish species. Tencalla & Dietrich (1997) demonstrated that hepatic PP-activity in trout is reduced to 60% within 1 h and to 100% within 3 h post oral application of toxic M. aeruginosa (equiv. to 5.7 mg MC-LRequiv./kg bw). Also in carp, the hepatic PP-activity totally disappeared within 3 h post gavage of M. aeruginosa (equiv. to 1.8 mg MC-LRequiv./kg bw) (Tencalla, 1995). The hepatic PP-activity in catfish (Ictalurus punctatus) decreased to 66-81%
24 h post intraperitoneal application of 0.5-2 mg MC-LR/kg bw and was also shown to decrease significantly following immersion in water containing microcystin (≥1 mg MC-LR/l) (Snyder et al., 2002).
The IC50 of MC-LR induced PP-inhibition in liver and kidney homogenates of diverse cyprinid species has been determined to be 0.1-0.3 nM (Tencalla, 1995; Xu et al., 2000), those in liver homogenates of trout to 0.05-0.2 nM (Sahin et al., 1995; Tencalla & Dietrich, 1997) thus indicating low inter-species variation. However, the extent of PP-inhibition has been shown to depend on the nutritional and physiological status of exposed fish (Malbrouck et al., 2004a;
Malbrouck et al., 2004b) and also on the characteristics of specific congeners. For example, MC-RR has been identified as a weaker PP-inhibitor than MC-LR and MC-YR (Xu et al., 2000).
Xu et al. (2000) moreover demonstrated differences within the inhibition of various PPs, showing the inhibition of PP1 (IC50 = 0.9-3.6 nM) isolated from grass carp (Ctenopharyngidin idellus) to be at least three-times less effective than inhibition of isolated PP2A (IC50 = 0.3-0.6 nM). This is in accordance with data obtained in mammals where IC50s for PP-inhibition of 1.7 and 0.04 nM were reported for PP1 and PP2A, respectively (Honkanen et al., 1990). Xu et al. (2000) further suggests that PP2A might be the principal microcystin sensitive enzyme in fish tissues, as the IC50 of MC-LR induced PP-inhibition in crude tissue homogenates was similar to those determined for isolated PP2A.
ATPASE-INHIBITION: Studies investigating the effect of M. aeruginosa extracts on gill fractions derived from carp and tilapia (Oreochromis sp.) suggest cyanobacterial-induced inhibitory effects on the activity of Na+/K+ ATPases in fish gill (Bury et al., 1998a; Bury et al., 1996b; Gaete et al., 1994; Zambrano & Canelo, 1996). ATPases are important enzymes, carrying various ions across cell membranes. Thus ATPase-inhibition in gill of fish exposed to cyanobacteria is assumed to disturb the transport of ions across the gill epithelium resulting in a sustained ionic imbalance and consequently gill dysfunction. While Gaete et al. (1994) and Zambrano & Canelo (1996) attribute the observed ATPase inhibition primarily to microcystin, Bury et al. (1998a & 1996b) regard the epithelial ATPase inhibition to result from bioactive metabolites other than microcystins (e.g. fatty acids). Malbrouck et al. (2003) determined neither ionic imbalance nor detrimental effects on the activity of gill Na+/K+ ATPases in goldfish (Carassius auratus)
intraperitoneally exposed to microcystin (125 µg MC-LR/kg bw), thus corroborating the latter point of view and revealing the need for further research.
ELEVATED ROS FORMATION (OXIDATIVE STRESS): The biotransformation of contaminants in fish often results in an elevated formation of reactive oxygen species (ROS) (Schlenk & Di Giulio, 2002). This also applies to microcystin (Bláha et al., 2004; Jos et al., 2005; Li et al., 2003; Li et al., 2005; Prieto et al., 2006) and is possibly promoted by a phosphorylation imbalance as a consequence of PP-inhibition (Li et al., 2003). ROS increase may overload the ROS-complementary antioxidative systems including both enzyme (e.g. superperoxiddismutase, glutathionperoxidase, catalase, etc.) and compound-based response (e.g. glutathione, vitamin E &
C, etc.) and thus result in an ROS surplus (Hofer & Lackner, 1995; Schlenk & Di Giulio, 2002). As ROS can oxidise intracellular molecules (causing enzyme inhibition, lipid peroxidation, DNA damage, etc.), elevated ROS levels cause severe cellular damage (i.e. oxidative stress) and may finally culminate in cell death (Hofer & Lackner, 1995; Zhang et al., 2007).
Microcystin-mediated oxidative stress in fish has been confirmed by three approaches: (i) via measurement of ROS levels, (ii) via alterations in components of the antioxidative systems, and (iii) via an increase of products resulting from lipidperoxidation. In vitro experiments on isolated hepatocytes and lymphocytes demonstrated an increase of ROS concentrations after 15 min and a doubling of ROS concentrations within 2 h of exposure to 10 µg MC-LR/l (Li et al., 2003; Zhang et al., 2007). This, despite a simultaneous activation of several ROS-eliminating enzymes (Li et al., 2003). A microcystin-mediated activation of antioxidative enzymes has also been determined in vivo (Jos et al., 2005; Li et al., 2005; Prieto et al., 2006), following intraperitoneal application of microcystins (500 µg MC-LRequiv./kg bw) as well as chronic gavage of microcystins and toxic Microcystis to tilapia (equiv. to 1.2 mg MC-LRequiv./kg bw for up to 21 days) and loach (Misgurnus mizolepis; equiv. to 10 µg MC-LRequiv./kg bw for 28 days). Those studies suggest the antioxidative enzyme response to be time-, dose- and tissue-dependent. It is thus not surprising that antioxidant enzymes appear able to eliminate oxidative stress induced by low microcystin concentrations (Li et al., 2005), whereas very high microcystin concentrations (≥500 µg MC-LRequiv./kg bw) have been demonstrated to induce oxidative stress that results in tissue damage, e.g. lipid peroxidation (Jos et al., 2005; Prieto et al., 2006).
GLUTATHIONE LEVEL AND GLUTATHIONE S-TRANSFERASE AKTIVITY: In contrast to the enzymatic ROS response, the antioxidative response of glutathione is not consistent. While Li et al. (2003) observed a microcystin-induced decrease of the glutathione concentration in carp hepatocytes exposed to microcystin (10 µg MC-LR/l ≈ 10 nM MC-LR), Adamovsky et al. (2007) and Bláha et al.
(2004) noted an ambiguous response of glutathione levels in carp and silver carp
(2004) noted an ambiguous response of glutathione levels in carp and silver carp