why won’t they grow? – inhibitory substances and mollusc hatcheries

Why won’t they grow? – Inhibitory substances and mollusc

hatcheries

J. B. JONES*Department of Fisheries, Government of Western Australia, P.O. Box 20, North Beach, WA 6920,

Australia; *Author for correspondence (e-mail: [email protected]; phone: +61-8-93683649;

fax: +61-8-94741881)

Received 8 February 2005; accepted in revised form 20 December 2005

Key words: Biological effects, Contamination, Metals, Seawater storage, Toxins

Abstract. Molluscs are known to be seriously affected by trace amounts of environmental pollu-

tants such as tributyltin at concentrations in seawater that are below the level of detection by all but

the most sensitive chemical analytical techniques. This extreme sensitivity by molluscs has led to use

of both adults and larvae as biomonitors for environmental pollution. Mollusc aquaculture has led

to an increasing demand for commercial hatcheries to supply seed stock, including selected genetic

lines of spat and juveniles. It is becoming apparent that many of the unexplained ‘‘crashes’’, ill thrift

or failures of larvae to metamorphose in such hatcheries are primarily due to their being com-

promised for a range of reasons including traces of inhibitory or toxic substances in the water

supply. Because dead and dying larvae are ideal substrates for bacterial and ciliate growth, such

invaders are often assumed to be the primary cause of the problem and this hinders finding a

solution. In addition, many of the toxins which may be implicated in crashes are sporadic in

occurrence and are both difficult to detect and hard to remove from the water supply. This paper

provides evidence for these toxic effects and suggests ways of reducing the problems.

Introduction

Mollusc hatcheries are an established source of larvae for commercial andresearch purposes and the principles behind good hatchery production are welldocumented. It is recognised that success is strongly dependent on quality ofthe broodstock and quality of fertilised eggs that are obtained (Utting andMillican 1997). Bacteria are also a universal threat and are kept at bay by stricthygiene and/or the use of antibiotics or probiotics (Gomez-Leon et al. 2005).However, no matter how long a hatchery has been operating or how skilled themanager, crashes or unexpected mortalities do occur. These problems areusually indicated by slow larval growth, larvae ceasing to swim and droppingout of the water column, or simply as mortalities. When examined, the deadlarvae may show heavy bacterial and/or ciliate contamination and it is oftendifficult to decide if these are the primary cause of the problem or secondaryinvaders of compromised larvae (Walne 1958; Estes et al. 2004). The usualpractice is to discard the batch, clean or sterilise the equipment, select newbroodstock and start again. However, in some cases, the problem does notresolve itself by such a direct approach and batch after batch can fail.

Aquaculture International (2006) 14:395–403 � Springer 2006

DOI 10.1007/s10499-005-9040-z

The reason for these multiple failures is often the presence of inhibitors andtoxins in the water supply or equipment. Molluscs, particularly larval molluscs,are known to be affected by trace amounts of environmental pollutants such astributyltin that are often below the level of detection by all but the mostsensitive chemical analytical techniques (Ruiz et al. 1995; Alzieu 2000). Thisextreme susceptibility of molluscs to environmental pollutants has led to use ofmolluscs and their larvae as biomonitors for environmental pollution (Rain-bow 1995; His et al. 1999; Nascimento et al. 2000; Sarver et al. 2003). Tocompound the problem, levels of toxic pollutants and inhibitors in marinewaters are seldom static but fluctuate with environmental perturbationsincluding storms, changes in temperature and salinity as well as with short termpulses due to human activity in the hatchery environment. These fluctuationscan cause hatchery failures to come and go with a frustrating lack of pattern.

The first step with any batch failure is to evaluate the hatchery managementprotocols, which should be well documented. Antibiotic use may induce apredominance of virulent antibiotic-resistant strains of bacteria, or may simplybe directly toxic to molluscs (Walne 1958; D’Agostino 1975); likewise, hypo-chlorite and ozone, commonly used to disinfect hatcheries, can inhibit normallarval development either directly or through toxic compounds they can pro-duce (Richardson et al. 1982). If chemicals used in the hatchery have beeneliminated from contention, the possibility of metal contamination should beconsidered.

Known toxic and inhibitory metals

The toxicity of metals such as copper to molluscs is well known since they havebeen used as antifouling agents for centuries. Molluscs also have a requirementfor metals and a complete lack of metals in the environment will affect growthand survival. However, an excess of metal ions can be toxic. For example,copper levels above 6 lg/l in the water supply causes a significant reduction indevelopment of Crassostrea gigas embryos, while at 10 lg/l only 50% devel-oped normally and at over 12.0 lg/l there was a decreasing percentage oflarvae showing normal embryonic development (Coglianese and Martin 1981).Reduction in salinity (for example, after washing down equipment with freshwater) can also have a marked effect on toxicity of copper, particularly below22.7 ppt (Coglianese 1982). Sources of copper in hatcheries include brass fit-tings and pump bearings, both of which can start to ‘‘leak’’ copper ions ascorrosion sets in. However, copper is not the only problem metal. Toxicity ofmercury to Argopecten irradians juveniles varies in a complex relationship withboth salinity and temperature (Nelson et al. 1977). Other metals, and indicativelevels at which they are known to cause toxic effects in the marine environ-ment, are shown in Table 1. It should be noted that toxicity could be furtheraffected if metal ions occur in combination or where metals forming organo-metal complexes combine with organic matter in the hatchery environment.

396

Table 1. Metals known to cause problems in marine mollusc culture, concentrations in seawater at

which effects on larvae have been noted and the published reference.

Metal Species affected Concentration

mg/l

Effect on larvae Reference

Arsenic Crassostrea gigas 0.326±0.0699 Abnormal development Martin et al. 1981

Mytilus edulis >3.0 Abnormal development Martin et al. 1981

Cadmium Argopecten irradians >0.06 Reduced growth, Pesch & Stewart

1980

Argopecten irradians 0.25 10% mortality Pesch & Stewart

1980

Crassostrea gigas 0.611±0.195 Abnormal development Martin et al. 1981

Mytilus edulis >1.2 Abnormal development Martin et al. 1981

M. galloprovincialis 0.424 EC50 Beiras & Albentosa

2004

Ruditapes decussatus 1.925 EC50 Beiras & Albentosa

2004

Chromium Crassostrea gigas 4.538±0.724 Abnormal development Martin et al. 1981

Mytilus edulis 4.469±0.739 Abnormal development Martin et al. 1981

Copper Crassostrea gigas 0.0053±0.0005 Abnormal development Martin et al. 1981;

Coglianese 1982

Crassostrea gigas 0.022 50% mortality Mandelli 1975



2004

Pteria colymbus 0.007 Abnormal development Rumbold &

Snedaker 1997


2004

Lead Crassostrea gigas 0.758±0.02 Abnormal development Martin et al. 1981



2004

Ruditapes decussatus 0.156–0.312 EC50 Beiras & Albentosa

2004

Mercury Crassostrea gigas 0.0067±0.0015 Abnormal development Martin et al. 1981



2004


2004

Nickel Crassostrea gigas 0.349±0.049 Abnormal development Martin et al. 1981


Mercenaria

mercenaria

0.310 LC50 Calabrese et al.

1977

Zinc Crassostrea gigas 0.119±0.012 Abnormal development Martin et al. 1981

Mytilus edulis 0.175 Abnormal development Martin et al. 1981

M. galloprovincialis 0.165–0.320 EC50 Beiras & Albentosa

2004


2004

This table provides indicative levels; it is not a review of the literature on the subject, for which see His

et al. (1999). (EC50 = median effective concentration; LC50 = lethal concentration).

397

In freshwater the effects are more complex since toxicity is affected by waterhardness and cannot be generalised across metals and species affected (Mance1987). Likewise, salinity and pH are also important. For example, the fresh-water bivalve Anodonta cygnea has an avoidance response to aluminium withavoidance reactions being dose dependent at neutral pH. At a concentration of250lg/l there was no effect while 500 lg/l reduced mean duration of shellopening by 50%, and the effect was irreversible over 15 days observationperiod (Kadar et al. 2001).

The best method to combat the effects of metal contamination in hatcheriesis to have a rigorous maintenance schedule to remove sources of corrosion andwear that can contribute to background metal ion levels. Care should also betaken to ensure that the water supply has not become contaminated. Activatedcharcoal filters are effective at removing metals and other pollutants fromincoming water (Kobya et al. 2005).

Other toxic compounds

Apart from metals, hydrocarbons from diesel oil at concentrations of 28 lg/lare cytotoxic and cause atresia in affected adult Mytilus (Lowe and Pipe 1986).30 lg/l of naphthalene in seawater causes ultrastructural changes to the renalepithelium of the gastropod Littorina littorea (Cajaraville et al. 1990).Hydrocarbons can be removed by use of activated charcoal filters (which is aquick way to isolate the problem). Sources of hydrocarbons and oils inhatcheries include the air supply – any oil-filled air pump can start to leakhydrocarbons and should be avoided. In addition, many non-food gradeplastics can ‘‘leak’’ hydrocarbons and parting compounds associated with theirmanufacture (personal observation) and can be toxic despite apparent highrates of dilution.

The presence of surfactants, pesticides and other chemicals in either thewater or air supply is also an issue. Estrogenic chemicals at concentrations aslow as 25 ng/l in treated sewerage water effluent can cause reproductive effectsin the snail Potamopyrgus antipodarum (Jobling et al. 2004). Development of‘‘D’’ larvae of the pearl oyster Pteria colymbus and veliger larvae of the musselPerna perna was affected by the presence of 0.8 mg/l and 0.68 mg/l respectivelyof sodium dodecyl sulphate (a common household detergent) in the water(Rumbold and Snedaker 1997; Jorge and Moreira 2005). Handcreams, insectrepellants and sunblock products can also cause problems if hands contami-nated with such products are subsequently immersed in tanks containing lar-vae. Owen et al. (2002) used the scallop Euvola (Pecten) ziczac in in-vivoexperiments to show that the organophosphate insecticide chlorpyrifos inhib-ited haemolymph acetylcholinesterase activity at concentrations as low as0.1 ng/l. Chemical spray drift can enter both the hatchery building and the airsupply or be transferred into tanks on the hands, feet and clothing of staff.Filtration of the air supply is strongly recommended, as is an enclosed hatcherywith the provision of footbaths and hand-washing facilities at entrances.

398

The location of the hatchery water inlet pipe can be critical, particularly ifnear the surface or the substrate. Sediments can be a potent source of heavymetals if disturbed, for example by wave or tidal action. This problem wasillustrated by analysis of sediment taken from a trap on the inlet pipe of amarine shellfish hatchery in Western Australia. The sediment had relativelyhigh levels of heavy metals (Cu 0.78 mg/l; Pb 0.67 mg/l; Ni 0.71 mg/l; Zn4.10 mg/l). The sea-surface layer can cause problems too. Many toxic sub-stances including hydrocarbons and heavy metals, lipids and proteins areknown to accumulate at the air-sea interface at concentrations 100 to1000 times greater than in the water column (Hardy 1982; Wurl and Obbard2004). This effect was studied by Rumbold and Snedaker (1999) who showedthat the sea surface microlayer was toxic to a variety of marine life, includingsea urchin larvae (Lytechinus variegates) and presumably would be equallylethal to many species of mollusc larvae. Subsurface water was not affected. Inabalone hatcheries the practice of floating fertilised eggs out of spawning tankstogether with the surface layer (often visibly contaminated with protein films)should be avoided and not just because of the bacteria associated with suchlayers.

‘‘Stored seawater’’

Obviously, stagnant water trapped in supply lines or in dead space in pipevalves and fittings can be extremely toxic to shellfish, particularly if hydrogensulphide is present, but there are more subtle effects that can occur. Manyhatcheries store seawater for varying periods in tanks or in supply lines.Stored seawater is not the same as fresh seawater and has been shown tohave corrosive properties different from those of open ocean water fromwhich it was taken (Craig, 1989). In addition, when raw seawater is stored,while the overall bacterial numbers do not change, the bacterial flora in theseawater does change (Yamamoto et al. 1983). Many bacteria and algalspecies when they are stressed or die release potent toxins that may survive inthe water after the death of the producer organism. Vibrio bacteria, includingV. alginolyticus, V. tubiashii and V. anguillarum may release heat stable cil-iostatic toxins and proteinases that degrade gill connective tissue (DiSalvoet al. 1978; Nottage et al. 1989). Early signs of such toxins include reducedfeeding activity and reduction of swimming in bivalve larvae (Estes et al.2004). Nottage et al. (1989) found that artificial seawater innoculated with2 · 10)3 cells/ml of V. alginolyticus could produce deleterious levels of toxinwithin 48 h. Such bacterial loads are well within the levels consideredacceptable in hatcheries. Very little work has been done on this effect butclearly it will be dependent on species present, the initial population size ofthe producer organisms and such things as temperature, organic load andsalinity of the storage container and the time the water is stored (Kaspar andTamplin 1993).

399

Toxic algae and hatcheries

Adult shellfish show a reduction in filtration rate, feeding behaviour andgrowth rate when exposed to toxic algae such as Gymnodinium and Hetero-capsa (Smolowitz and Shumway 1997), however, adult Mytilus edulis, Mytilusgalloprovincialis and Crassostrea gigas exposed to filtrates of toxic algae wereunaffected, showing that the effect is associated with the particulate algae, notwater soluble components (Gainey and Shumway 1988; Neilsen and Strømgren1991; Matsuyama et al. 1999). Provided incoming water is adequately filteredto remove algae, toxic algal blooms outside the hatchery should not be aproblem. Algae such as Trichodesmium erythraeum, commonly used in hatch-eries as a food source, may also cause problems for larval bivalves undercertain conditions (Negri et al. 2004).

Other chemical contamination

Finally, it should be noted that mollusc larvae are able to recognise the pres-ence of other larvae and appear to have the ability to attract or exclude settlinglarvae (Keen and Neill 1980). Chemical cues, particularly low molecular weightpolypeptides, are used to induce settlement in Crassostrea, Mytilus andPinctada (Zimmer-Faust and Tamburri 1994; De Vooys 2003; Zhao et al.2003). These chemicals occur naturally and are secreted by adult shellfish.However, other cues exist and substances associated with crustose corallinealgae are known to induce metamorphosis in Haliotis spp. larvae. These maybe structurally related to c-amino-butyric acid (GABA), which is sometimesused as an inducer in hatcheries (Doroudi and Southgate 2002; Suenaga et al.2004). Also of significance are the observations by Zhao et al. (2003) thatbiofilms can initiate settlement in P. maxima and by Suenaga et al. (2004) thatpotential inducer chemicals are secreted by anaerobic bacteria. Molluscs canalso secrete substances to prevent further settlement. The gastropod snailBulinus truncates uses urea as a population inhibitory substance (Leopard andIsseroff 1994; Smith et al. 1994). Clearly this confers a competitive advantagein fostering either clumping or thinning of shellfish, depending on the species –but may not be appropriate behaviour in a tank! While density dependentgrowth has been well documented in molluscs including abalone (Capinpinet al. 1999) and pearl oysters (Wada and Komaru 1994), this has usually beenattributed to micro-competition rather than the density controlling ability ofmolluscs and has not been investigated in a hatchery or farm setting.

Summary

Though hatchery technology is widely used to produce commercial quantitiesof bivalve larvae, unexpected failures of batches still occur. In many cases,

400

these are due to contaminants and toxins at very low concentration(s) and froma variety of sources. Inability to grow larvae, or sudden larval mortalities maybe due to subliminal concentrations of organic or inorganic toxins, sometimesin combination and often at specific times of the year. This requires constantvigilance on the part of the hatchery manager. However, some effects, like thepotential toxins associated with seawater storage, and density dependent effectsof larvae on each other have not yet been adequately researched.

Acknowledgements

I would like to acknowledge the cooperation of the large number of shellfishhatchery managers in Western Australia and New Zealand who, over manyyears, have freely collaborated with me to solve their problems. Drs JohnCreeper, Fran Stephens and two anonymous referees made valued suggestionsand comments on the manuscript. The editorial input from Dr Greg Maguire isalso acknowledged.

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why won’t they grow? – inhibitory substances and mollusc hatcheries

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