Veterinary Parasitology 163 (2009) 207–216

Selected parasitosis in cultured and wild fish

F.C. Guo a,*, P.T.K. Woo b

a Novartis Animal Health Canada Inc., Aqua Heath Business, 797 Victoria Road, Victoria, PE, C0A 2G0, Canadab Department of Integrative Biology, University of Guelph, Guelph, ON, Canada

A R T I C L E I N F O

Keywords:

Caligus rogercresseyi

Cryptobia salmositica

Gyrodactylus salaris

Lepeophtheirus salmonis

Loma salmonae

Aquaculture

Parasitosis

Wild and farmed fish

A B S T R A C T

While intensive aquaculture has and will continue to supply the ever growing population

with highly nutritious protein, it also comes with problems which include more frequent

outbreaks of diseases in fish farms and transmission of diseases between farmed and wild

fish. We have selected four Phyla of economically important fish parasites for our present

discussion—a haemoflagellate (Cryptobia salmositica), a microsporidian, (Loma salmonae),

a monogenean (Gyrodactylus salaries) and two copepods (Lepeophtheirus salmonis, Caligus

rogercresseyi). This review consists of two parts with a brief description of each parasite

and its biology related to transmission, followed by discussions on epizootic outbreaks in

both wild and farmed fish, interactions between wild and farmed fish, and disease

prevention and control.

� 2009 Published by Elsevier B.V.

Contents lists available at ScienceDirect

Veterinary Parasitology

journal homepage: www.elsev ier .com/ locate /vetpar

1. Introduction

The demand for animal protein will continue to escalateas the world population increases from 6 billion (estimatedin 1989) to about 8 billion within the next 15 years. Thisincrease will exert additional pressures on food productionwhich is already competing with other essential humanactivities (e.g., cultivation of crop, transportation, housing,industry) for the finite amount of usable land. Animalprotein contains essential amino acids which are impor-tant for a nutritional and well-balanced diet. Free rangingland animals are no longer a significant source of proteinand the production costs of farm-raised animals continueto increase as land becomes more expensive. Conse-quently, farms have to be large and are usually close tohuman habitations to increase efficiency, and reduceproduction and transportation costs. The large scalebreeding of warm-blooded animals close to humanpopulations presents some serious problems, includingdischarge and/or storage of animal wastes and associatedpublic health issues. There are also increased risks of

* Corresponding author. Tel.: +1 902 367 7438; fax: +1 902 658 2261.

E-mail address: fuci.guo@novartis.com (F.C. Guo).

0304-4017/$ – see front matter � 2009 Published by Elsevier B.V.

doi:10.1016/j.vetpar.2009.06.016

disease outbreaks in the animals and the subsequentinterspecies transmission of zoonotic diseases to humans(e.g., Nipah virus in pigs, avian influenza H5N1 virus inbirds, cryptosporidian parasite in cattle) (Woo, 2006a).

Fish is an excellent protein and is easily digestible.Many species of marine fish have the beneficial poly-unsaturated fatty acids (PUFA; Omega 3 fatty acid) whichinclude docosahexaenoic acid (DHA) and eicosapentaenoicacid (EPA). Both DHA and EPA are physiologically essentialnutrients for normal brain function. DHA is also animportant component of phospholipids of cell membraneand hence is necessary for proper function of the retina andbrain. PUFA has other beneficial effects and they includereducing risks of cardio-vascular diseases. The capture-fishery is on the decline or at least stagnant, and this haspartly been brought about by over and/or indiscriminatefishing, destruction of spawning grounds, and no newlydiscovered fishing grounds. Also, industrial wastes (e.g.,organochlorine pesticides, PCBs, heavy metals) dischargedinto the aquatic environment can reduce fish growth andreproduction and in some areas fish are no longerrecommended for human consumption because they haveaccumulated such high levels of pollutants. Aquaculture offish is a good option as production costs are lower overtime, especially in cage cultures. It has been the fastest

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216208

growing food production sector since 1970 – with anincrease at a compounded rate of about 9.2% per year. Also,it does not have some of the problems (e.g., public healthissues) associated with the large populations of warm-blooded animals close to human habitations (Woo, 2006a).

Intensive aquaculture of fish is not without itsproblems, and these include disease outbreaks andconsequences of introducing parasites to new hosts and/or new localities with the transportation of live fish. Severeepizootics have occurred and many of these are man-made. Selected outbreaks are discussed to better under-stand the nature and cause of the epizootics. The currentdiscussion is in two parts and is intended to highlightepizootics caused by relatively well-studied parasites fromfour Phyla (Phylum Euglenozoa – a haemoflagellate;Phylum Microspora – a microsporidian; Phylum Platyhel-minths – a flatworm; Phylum Arthropoda – sea lice) whichcause morbidity and mortality in fishes reared inhatcheries, sea cages, and in their natural habitats. Part Iof the discussion is to provide sufficient relevant back-ground information on the biology of these parasites asthey relate to the epizootics while Part II is on selectedepizootic outbreaks with control measures in bothcultured and wild fish stocks.

2. Biology of parasites

2.1. Cryptobia salmositica

Cryptobia salmositica (Fig. 1; Phylum Euglenozoa) is anelongated extracellular haemoflagellate. It has a prominentkinteoplast at the anterior end, a central nucleus and twoflagella. The parasite has been found in many fish species (allthe Pacific salmon, Oncorhynchus spp. and sculpins, Cottus

spp.) in streams and rivers along the west coast of NorthAmerica. It is normally transmitted indirectly from fish tofish by the freshwater leech, Piscicola salmositica. Briefly, theparasite is ingested during a blood meal, multiplies in thecrop of the leech, and is inoculated into a new host whenthe leech feeds again. In the absence of the leech vectordirect transmission between fish can occur under certainaquaculture conditions (e.g., during grading and weighingwhen fish are brought into direct contact with each other,crowding in tanks or in rearing cages). The prevalence of theparasite in downstream migrants (pre-smolt salmon) can be

Fig. 1. Cryptobia salmositica (Phylum Euglenozoa) with a red blood cell

from a fish with microcytic and hypochromic anaemia. Reproduced from

Woo (2006b).

quite variable (e.g., 3–21%) in streams in Oregon, UnitedStates, and fingerlings have detectable infections as early as60 days after they hatch. Sexually mature salmon becomeinfected within days of returning to fresh water from the sea.The parasite is not pathogenic to sculpins (reservoir hosts)but causes disease in salmonids. Severe outbreaks ofsalmonid cryptobiosis have occurred in fish maintainedin sea cages, in freshwater hatcheries and in streams(Woo, 2003).

2.2. Loma salmonae

Microsporidians are a diverse group of unicellularorganisms that live as obligate intracellular parasites inmany invertebrates and are reported from all classes ofvertebrate hosts. Many microsporidian species are widelydistributed in teleosts in freshwater, brackish and marinehabitats (Dykova, 2006). Once considered as the mostprimitive eukaryotes, they are now regarded as highlyspecialized fungi with simple life cycles consisting ofmerogony and sporogony stages (Keeling and Fast, 2002).Microsporidians are single walled spores with length of2–10 mm, mostly ellipsoidal or egg shaped and lackmitochondria. The spores contain extrusive apparatuscapable of injecting sporoplasm into host cell via extrud-able polar tube (Dykova, 2006).

Loma salmonae (Putz et al., 1965; Fig. 2; PhylumMicrospora) causes microsporidial gill disease (MGD) inseven salmonid species of the genus Oncorhynchus (Brunoet al., 1995; Hauck, 1984; Kent et al., 1989; Magor, 1987;Morrison and Sprague, 1983; Speare et al., 1989), brooktrout, Salvelinus fontinalis (Shaw et al., 2000), and browntrout, Salmo trutta (Bader et al., 1998). The typical clinicalsigns include pale gills with petechial hemorrhages,hyperplasia, and white cysts called xenomas (Wales andWolf, 1955; Hauck, 1984; Kent, 1992). Fish with highnumbers of xenomas in their gills suffered respiratorydistress, secondary infection and high mortality rates.(Speare et al., 1998; Becker and Speare, 2007). Laboratorystudies show that L. salmonae can be transmitted directlyvia ingestion of spores by fish. Briefly, the spore enters thegut and injects sporoplasm into an epithelial cell, itmigrates to the heart, then to the gill, and forms a spore-laden xenoma within the gill filaments. The rupture ofxenoma causes proliferative branchitis and massivenumber of spores release (Shaw et al., 1998; Sanchezet al., 2001b, Becker and Speare, 2007).

2.3. Gyrodactylus salaris

Gyrodactylus salaris (Fig. 3; Phylum Platyhelminthes) isa parasitic flatworm of fishes in freshwater rivers andlakes. Numerous other species of monogeneans are knownto cause morbidity and mortality in freshwater and marinefishes. Monogeneans have a direct life cycle and are eitherviviparous (give birth to free-swimming larvae, e.g.,Gyrodactylus) or are oviparous (produce eggs which hatchafter they are laid, e.g., Pseudodactylogyrus). In either casethe free-swimming larva (oncomiracidium) migrates tospecific sites (e.g., on the gills, the fins or the body surface)where it attaches. On susceptible hosts the parasite has a

Fig. 2. Loma salmonae (Phylum Microspora) (a) Xenoma within the

primary gill filament of a Chinook salmon containing a hypertrophied

nucleus (N), meronts (arrowhead) and spores. Bar = 8 mm. (b) L. salmonae

spore with the electron-dense exospore (arrow) adjacent to the

translucent endospore and characteristic coiled polar tube

(arrowheads). Bar = 700 nm. Reproduced from Becker and Speare

(2007). (Photos courtesy of Dr. Jan Lovy).

Fig. 3. Viviparous gyrodactylid monogenean (Gyrodactylus sp., Phylum

Platyhelminthes), drawn by Beth Beyerholm. Reproduced from

Buchmann and Bresciani (2006).

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216 209

high reproductive rate which rapidly increases the intensityof infection, and this also promotes efficient transmissionbetween fish especially when fish are under crowdedconditions. Most monogeneans are ectoparasitic on specificsites on the fish (e.g., on branchial arches, the fins, the head)and they attach to the host using their opisthaptor (an organlocated at the posterior end of the worm) which is usuallyequipped with large hooks, clamps and/or suckers. Specieslike G. salaris cause morbidity and mortality in fish becauseof their high intensities, and significant damage at the pointof attachment (with considerable host reactions) throughtheir opisthaptor and by grazing on exposed structures andvulnerable integument. The parasite is mainly a pathogen ofAtlantic salmon, Salmo salar, in freshwater rivers and lakes inNorway. Artic charr, Salvelinus alpinus, and rainbow trout,Oncorhynchus mykiss, are known reservoir hosts, and theparasite is in Norway, Sweden, Finland, Denmark, Russia andGermany. The pathogen is normally on the fins, to a lesserdegree on the body surface, cornea and nostrils, and veryrarely on the gill apparatus. The parasite is viviparous, andworms with three generations (a worm with an embryowhich already has an offspring) in the uterus are sometimesseen. (Buchmann and Bresciani, 2006).

2.4. Sea lice

Sea lice are common marine ectoparasites that belongto the order of Copepoda (Phylum Arthropoda). In NorthAmerica (United States and Canada), Ireland, Scotland,

Fig. 5. Female (a) and male (b) Caligus rogercresseyi (Phylum Arthropoda).

Photos courtesy of Dr. Sandra Bravo. Reproduced from Dr. Sandra Bravo’s

presentation at World Aquaculture Conference 2007. Available at http://

www.puresalmon.org/pdfs/bravo_present_sealice_WAS.pdf (last accessed

on 21 April 2009).

Fig. 4. Lepeophtheirus salmonis (Phylum Arthropoda) adult female with

egg strings removed.

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216210

Faeroes and Norway, there are mainly three species ofsea lice that parasitize farmed salmonids, Lepeophtheirus

salmonis (Fig. 4), Caligus elongatus on the North Atlanticcoasts, and Caligus clemensi on the North Pacific coast. L.

salmonis is larger and more abundant with hosts confinedmainly to salmonids while Caligus spp. are morecosmopolitan in distribution and have broad host rangesthat include salmonids and non-salmonids (Kabata,1979; Treasurer and Grant, 1994). L. salmonis is a majorspecies of interest in northern hemisphere, while in thesouthern hemisphere, the major species of interest isCaligus rogercresseyi (Fig. 5) which parasitize farmedsalmonids and non-salmonids in Chile (Boxshall andBravo, 2000).

Adult sea lice usually show sexual dimorphism with thefemale larger than the male. The female (10–18 mm long)has a more prominent genital segment than the male (5–7 mm long) (Kabata, 1979) and a paired of egg strings.Caligus has frontal lunules which are absent inLepeophtheirus. Life cycle of L. salmonis has 10 stages,two free living naupliar stages, one infective copepodidstage, four attached chalimus stages, two mobile pre-adultstages and one adult stage (Johnson and Albright, 1991)whilst C. rogercresseyi has eight stages, two pre-adultstages were not observed (Gonzalez and Carvajal, 2003).

3. Epizootic outbreaks of parasitosis

3.1. Cryptobiosis

3.1.1. Wild fish stocks

C. salmositica is considered a lethal pathogen of salmonin semi-natural and intensive fish culture facilities in NorthAmerica (Bower and Thompson, 1987). Pre-smolt salmonretained their experimental infections when transferredfrom fresh to salt water and mortality due to cryptobiosiswas not reduced after they were transferred to seawater.As indicated earlier (Biology of parasites) the prevalence ofthe Cryptobia in downstream salmon migrants (pre-smolts) ranged from 3% to 21% in some streams in theUnited States. Consequently, it has been suggested that thedisease may be an important cause of salmon mortality in

the sea; however, to-date no field studies have beenconducted to validate this suggestion (Woo, 2006b).

The leech vector hatches from cocoons in late summerand early autumn. Newly hatched leeches pick up theinfection by feeding on infected sculpins; however theyprefer to feed on salmon when they are available later inthe season. Briefly, the parasite from the torrent sculpin,Cottus rhotheus (the main reservoir host of the pathogen) istransmitted to other fishes including the adult salmonsoon after their return from the sea to freshwater streams.The prevalence of the parasite in sculpins can be high, isseasonal, and can be quite variable in streams (e.g., about60% in the United States, and 8–95% in Canada). In general,the prevalence is higher in large sculpins than in small fish(Woo, 2003). According to Bower and Margolis (1984)returning adult salmon in some streams in the Fraser River,Canada had detectable infections within 5 days ofreturning to fresh water (in November) and their para-sitaemias increased with time. Parasitaemias were veryhigh at spawning salmon and many fish died before theyspawned. Also, the prevalence was initially relatively lowbut steadily increased to 100% by December and January –the assumption was that as the season progressed therewere increasing numbers of leech in the streams totransmit the pathogen.

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216 211

3.1.2. Freshwater hatcheries and sea cages

Serious outbreaks of salmonid cryptobiosis (highmortality) in hatcheries had been reported in the UnitedStates as early as 1955. These outbreaks were mainly injuvenile Chinook salmon, Oncorhynchus tshawytscha,sexually mature Chinook salmon and post-spawningrainbow trout, O. mykiss. Also, high fish mortalities inRussian hatcheries due to cryptobiosis had been reportedin pre-spawning pink salmon, Oncorhynchus gorbuscha,and adult Caspian salmon, S. trutta caspius. However, verylittle is known about this Cryptobia other than the parasiteis morphologically similar to C. salmositica and with similarclinical signs in infected fish. As in salmonid cryptobiosis inNorth America, and the infected fish also died with massivenumbers of parasites in the blood (Woo, 2006b).

As indicated earlier (Bower and Margolis, 1984)sexually mature salmon become infected within daysafter they return to fresh water. About 50% of sexuallymature Chinook salmon brought into hatcheries as brood-stock from streams in the United States annually die fromcryptobiosis (Woo, 2006b). Currie and Woo (2007)confirmed experimentally that sexually mature (‘sm’)rainbow trout were more susceptible than juveniles tocryptobiosis. They also showed that ‘sm’ females weremore susceptible (higher parasitaemias and mortality)than ‘sm’ males and that there was a factor(s) in the plasmaof ‘sm’ fish that promoted rapid parasite multiplicationunder in vitro conditions. Also, plasma from ‘sm’ femaleswere much more efficient (or in higher concentrations)than plasma from ‘sm’ males in enhancing in vitromultiplication of the pathogen.

Outbreaks of salmonid cryptobiosis were not merelyconfined to freshwater hatcheries. In 1997 the parasitecaused significant morbidity and mortality in smolts andpre-harvest Chinook salmon in a hatchery on VancouverIsland, Canada. It was low (about 1%) in post-smolts afterthe fish were transferred to sea cages, but the disease re-emerged later to cause higher mortality in pre-harvest fish.According to the hatchery personnel fish mortality seemedto be associated with age and major stressors such asharassment by marine mammals. In 2001 another out-break of the disease occurred in sea cages in the samehatchery. Briefly, the parasite was detected in late 1999 inthe blood of some fish while they were still in fresh water.Clinical cryptobiosis (e.g., exophthalmia, anemia, anorexia)was noticed in fish in sea cages in 2001 and the parasitewas isolated for confirmation studies. Fish mortality variedbetween cages (ranged from 3.3% to 24.9%). It wassuggested that the outbreak was initially because ofrelapse of the infection in a few infected fish and thepathogen was subsequently transmitted directly betweenfish especially when they were brought into direct contactwith each other during grading and weighing (Woo, 2003).

3.1.3. Control and prevention

3.1.3.1. Breeding Cryptobia-resistant fish. Some naıve brookcharr, S. fontinalis, cannot be infected with the parasite(Forward et al., 1995). The resistance to Cryptobia infectionin charrs is controlled by a dominant Mendelian locus andit is inherited by progeny (Forward et al., 1995). Briefly, the

pathogen is lysed in the blood via the alternative pathwayof complement activation in Cryptobia-resistant charrs(Forward and Woo, 1996). Little is known about this typeof immunity; further studies on the inheritance of naturalresistance by progeny would be rewarding as it could beexploited to protect fish against pathogens (Woo, 1998).

3.1.3.2. Vaccination. Two experimental vaccines (a liveattenuated Cryptobia vaccine and a DNA-vaccine) havebeen developed.

(a) Live vaccine: The attenuated live vaccine infects andproduces low parasitaemia in rainbow trout. It does notcause disease, circulates in the blood for at least 6 months,and protects 100% of the vaccinated fish from re-infection(Woo and Li, 1990). In trout partial protection occurs at 2weeks post vaccination and full protection is at 3–4 weekspost vaccination (Li and Woo, 1995). A single dose of thevaccine protects fish for at least 24 months (Li and Woo,1997), and it has no detectable bioenergetic cost tojuvenile rainbow trout (Beamish et al., 1996).

Fish vaccinated in fresh water and transferred toseawater were still protected (Li and Woo, 1997). Thecomplement fixing antibody titres (e.g., Li and Woo, 1995)and cell-mediated response (Mehta and Woo, 2002) invaccinated fish rose significantly soon after parasitechallenge (a classical secondary response). Protection isthrough the production of complement fixing antibodiesand enhanced cell-mediated cytotoxicity (antibody-inde-pendent and antibody-dependent). Also, macrophagesfrom vaccinated fish are much more efficient in engulfinglive parasites under in vitro conditions, especially withantibodies (Li and Woo, 1995).

(b) DNA-vaccine: A recombinant metalloprotease-plas-mid vaccine (DNA-vaccine) have been produced (Tan et al.,2008). Trout injected intramuscularly with the DNA-vaccine had a slight anaemia during the first 3–4 weekspost-vaccination (wpv), and had detectable agglutinatingantibodies against Cryptobia between 5–7 wpv. Fishinjected with the DNA-vaccine had lower parasitaemiawhen challenged, delayed peak parasitaemia and fasterrecovery.

3.1.3.3. Chemotherapy. The trypanocidal drug, isometami-dium chloride, inoculated intramuscularly into fish iseffective against C. salmositica in experimentally infectedadult rainbow trout, Atlantic salmon and juvenile Chinooksalmon. The drug also provides prophylactic protectionagainst the parasite (Ardelli and Woo, 1999, 2001) and itseffectiveness in protecting against naturally infectedChinook broodstock is encouraging (Chen, M. personalcommunication, 2007).

3.1.3.4. Vector control. As well as infected fish, the leechvector is often introduced into hatcheries via the watersupply as water is nearly always from nearby streams.Leech cocoons have been subjected to various treatmentsintended to reduce their viability. The results wereencouraging, for example chlorine was suggested as apotential chemical that could be used on cocoons inhatcheries to control the leech population (Bower andThompson, 1987; Bower et al., 1985).

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216212

3.2. Microsporidiosis (microsporidial gill disease of

salmonids, MGDS)

3.2.1. Wild fish

Although L. salmonae is common, it is not usuallyconsidered a severe pathogen in wild salmonids or in thosereared in freshwater hatcheries.

L. salmonae was detected in wild salmonids collectedfrom four rivers and one lake in British Columbia, Canada.The prevalence was 30% in pre-spawning sockeye salmon(Oncorhynchus nerka) that had recently entered fresh waterduring their return migration to spawn. L. salmonae wasalso present in migrating smolts (detected using a PCR test)(Shaw et al., 2000). The smolts were likely infected byconspecific freshwater salmonids or by spores depositedwith eggs as high prevalence of the parasite was found inthe ovaries of sexually mature Chinook salmon (Dockeret al., 1997).

A survey of fishes captured from open waters forsalmonid pathogens was conducted in the coastal watersof British Columbia. L. salmonae was found in Chinooksalmon, chum salmon (O. keta), coho salmon (O. kisutch),sockeye salmon (O. nerka), and pink salmon (O. gorbuscha),all of which were captured well away from net-pens (Kentet al., 1998).

3.2.2. Farmed fish

All cultured salmonids (Oncorhynchus), as well as brooktrout, brown trout, and Atlantic cod are susceptible toMGDS, however, Atlantic salmon (S. salar) are resistant(Kent et al., 1995). The most economically significantproblems with MGDS occur in Chinook salmon and cohosalmon reared in seawater (SW) net-pen aquaculture inBritish Columbia (Kent and Poppe, 1998; Constantine,1999). The disease most often affects salmon in theirsecond summer when the fish are near harvesting, andoutbreak can lead to a cumulative mortality rate over 30%(Kent and Speare, 2005; Speare et al., 2007).

3.2.3. Interaction between cultured and wild fish

MGDS is transmitted directly to fish by either ingestionof infected tissue or by free spores in the water column, orby co-habitation with infected fish, or by horizontaltransmission from wild marine fishes (Shaw et al.,1998). Infections can persist after fish are transferred toseawater, and the associated lesions in the gills canbecome severe in the pen-reared salmon (Speare et al.,1989). Several Loma species have been described frommarine fishes, but transmission studies (Shaw et al., 1997)and comparisons of rDNA sequences from marine fishLoma spp. (Brown et al., 1998) strongly suggest that L.

salmonae only infects salmonid fishes. A marine non-salmonid reservoir has not been identified and L. salmonae

has been found in ocean-caught wild salmon (Kent et al.,1998). Thus a source of the infection for fish in marine net-pens may be wild marine-phase salmonids (Kent, 2000).

3.2.4. Control and prevention

Currently there are no approved drugs for the control ofL. salmonae infections in the aquaculture industry inCanada (Sanchez et al., 2001b). Several drugs have been

tested in rainbow trout using a Loma challenge model.Dietary monensin at 1000 ppm significantly reduces thesize of xenomas (Becker et al., 2002); whilst fumagillin andalbendazole have some potential value in controlling L.

salmonae infection in trout while pyrimethamine + sul-phaquinoxaline, amprolium and metronidazole wereineffective (Speare et al., 1999). Intraperitoneal injectionof b-1,3/1,6-glucan to rainbow trout at 100 mg per fish wasfound to produce significantly fewer xenomas (Guselleet al., 2006). A live vaccine containing a low-virulencestrain of L. salmonae, and inactivated spore-based sporevaccine are both effective in reducing xenoma counts(Sanchez et al., 2001a; Rodrıguez-Tovar et al., 2006; Speareet al., 2007).

3.3. Gyrodactylosis

3.3.1. Wild fish

Hansen et al. (2003) examined the variations inmitochondrial DNA between Swedish and Norwegianisolates of the parasite and they concluded that G. salaris

most likely was introduced to Norwegian rivers in the 1970sduring the transfer of Baltic-Atlantic salmon from Sweden toNorway. The parasite is highly pathogenic to the East-Atlantic salmon in Norway and it has devastated thepopulations of salmon fry in 46 Norwegian rivers. Since thepathogen is viviparous it spreads rapidly because it canreproduce excessively on Norwegian salmon. Also, there areobvious concerns the pathogen may spread to otherEuropean countries that have the highly susceptible East-Atlantic salmon stock (Von Gersdorff Jorgensen et al., 2008).It has been suggested the parasite does not adversely affectthe Baltic-Atlantic salmon in Norway and Scotland basicallybecause the parasite and the Baltic salmon have coexistedfor thousands of years. However, in Norway there has beenno natural selection for resistance to G. salaris in East-Atlantic salmon prior to the 1970s, consequently thedevastating outbreaks of gyrodactylosis in the rivers(Buchmann and Bresciani, 2006).

3.3.2. Control and prevention

Drastic control programs have been instituted for over adecade to eradicate the pathogen from affected rivers.Rotenone added to rivers kills both the fish and parasitewith subsequent re-stocking of the treated rivers with thesame fish stock. This control measure has met with somesuccess although a number of treated rivers have been re-infected. Another option is the addition of aluminiumcompounds to the rivers because aluminium is quite toxicto gyrodactylids but is tolerated by fish at concentrationsused (Soleng et al., 1999).

3.4. Caligidosis

Sea lice are among the most notorious and damagingparasite to the salmonid farming industry in both Europeand the Americas (Costello, 2006; Lester and Hayward,2006). Estimated costs of sea lice control based on 2006salmonid production statistics (FAO, 2008) are s305million for the seven salmon cage farming countries(Costello, 2009). The damages are not just on the fish and

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216 213

the environment, but also the public perceptions towardsaquaculture (Pike and Wadsworth, 1999; Costello et al.,2001). Because of the importance of these parasites, aseries of international conferences on sea lice, and selectedarticles has been published in special issues of interna-tional journals (Pest Management Science, 58, 2002, pp.622–629; Aquaculture Research, 35, 2004, pp 711–805;and Journal of Fish Diseases, 32, 2009, pp. 1–113). Despitemajor research efforts over three decades, as evident fromover 800 research publications, sea lice remain a persistentproblem.

3.4.1. Sea lice in wild fish and cultured fish, and their

interactions

Sea lice infestations are endemic in most countrieswhere salmonid is cultured. Declines in wild salmonidpopulations in recent years have led to the widespreadbelief that there could be a link between sea lice in farmedfish and this decline.

Norway is the world largest producer of farmed Atlanticsalmon, wild salmon consists only about 1% of farmedAtlantic salmon (Heuch et al., 2005). Consequently thepotential number of L. salmonis produced by farmedsalmonids could be large (Heuch and Mo, 2001), and cross-infestation of L. salmonis could occur between farmed andwild hosts. By examining the carbon and nitrogen stableisotopes levels in the sea louse, Butterworth et al. (2004)differentiated L. salmonis from farmed Atlantic salmon andthose from wild coho salmon. Generally sea lice are largeron wild fish than on farmed fish (Nordhagen et al., 2000),however, size is not a reliable indicator on the origin of lice(Boxaspen, 2006). The average prevalence of natural C.

elongatus infections of wild coastal fishes on the Norwe-gian south east coast is 15%, and great differences are foundbetween fish species and seasons. Lumpfish Cyclopterus

lumpus L. spawners are the most infected fish, with gadidsat the lower end, while sea trout (S. trutta L.) and herringClupea harengus L. are in between. Relatively high numbersof chalimii on found on North Sea lumpfish suggest thatoffshore fish sustain an oceanic population of C. elongatus

(Heuch et al., 2007).Scotland, the second largest salmon producing region in

Europe has marked population declines of wild sea trout (S.

trutta), particularly in the north-west where salmonculture is concentrated. On their first visit to sea in thespring of the year following hatching, sea trout may beconfronted with very high concentrations of infective sealice larval stages and quickly become infested with lice. Aburden of only 10 adult lice is thought to be sufficient tocause mortality, especially in immature fish already understress (Anon., 2002). Shinn et al. (2000) successfullydifferentiated sea lice on cultured salmonids from thoseon wild salmonids by analyzing the elements magnesium,vanadium, and uranium on the lice.

Canada is among the top five farmed salmon producingcountry. Salmon farming on the west coast of Canada hasbeen linked with declining wild pink salmon (O. gorbuscha)by the environmental groups with sea lice as the culprit. Byusing mathematical models, Krkosek et al. (2007) reportedthat sea lice spread from salmon farms in the BroughtonArchipelago, British Columbia, have placed wild pink

salmon populations ‘‘. . . on a trajectory toward rapid localextinction. . .’’ and that ‘‘. . . a 99% collapse in pink salmonpopulation abundance is expected in four salmon genera-tions.’’ However, Riddell et al. (2008) counter-argued thatthe risks were overstated as the dataset used for theprediction were selected (excluding 2004 high return data,and data from Glendale River which produces the largestpopulation of pink salmon), and the predictions wereinconsistent with recent observations of pink salmonreturns to the Broughton Archipelago. It is interested toknow that besides salmonids, wild sticklebacks (Gaster-

osteus aculeatus) appear to serve as temporary hosts for L.

salmonis, suggesting a role of this fish in the epizootiologyof the parasite (Jones et al., 2006). There is generalagreement between scientists and environmental groupsthat sea lice pass freely to and from wild salmon; however,the debate is whether the transfer of sea lice from thefarmed salmon to the wild salmon is on a sufficient scale tohave an impact (Anon., 2006). In assessing and managingwild pink salmon in BC, all potential impacts on the wildpopulations, including sea lice, should be acknowledged indeveloping an effective management strategy (Riddellet al., 2008). Brooks (2009) advocated to develop andimplement an area management plan coordinating sea licemanagement efforts by all producers and close monitoringof therapeutants efficacy and conducting specific fieldbioassays as suggested by SEARCH (2004).

3.4.2. Prevention and control

Currently the salmon farming industry adopts theintegrated pest management (IPM) approach to deal withsea lice which includes farm fallowing, regular monitoringof lice burden, chemical control (use of parasiticides),biological control (use of cleaner fish), monitoring andreporting of drug resistance, strategic rotation of medi-cines (Lees et al., 2008). With a limited range of approvedchemicals to choose and increasing speed of resistancedevelopment, sea lice control remains the biggest problemfaced by the industry.

Parasiticides can be topical (bath) treatment byenclosing the cage in canvas skirts or tarpaulins or theycan be used as oral (in-feed) treatment where themedication is mixed with feed pellets and fed to the fish.Bath treatments usually have quicker action than oraltreatments, however, it is very difficult to have a uniformdistribution of active compound within the salmon cage.‘Hot’ spots (areas with over dose) are common which maybe toxic to fish (Roth et al., 1993), while cold spots (underdose) lead to lice not being removed, and causing re-infection. Bath treatment is labour-intensive and it is verydifficult to treat all cages in one site within the same day, solice from untreated fish can transfer to re-infect previouslytreated fish. However, oral treatment takes a few days tobuild up tissue concentration, and fish that are not eatingwell will not get adequate amount of medicine, furthermore there is a withdrawal period that is regulated strictlyby the authorities (Costello, 2006), but it offers sustainedperiods of louse clearance (Stone et al., 2000a,b,c).

Organophosphates (OPs, trichlorfon and dichlorvos)were the first chemotherapeutants used to control sea liceas bath treatment. They were replaced by azamethiphos

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216214

(also an organophosphate), hydrogen peroxide, andsynthetic pyrethroids (cypermethrin and deltamethrin).The oral treatments are the chitin synthesis inhibitorsdiflubenzuron and teflubenzuron, and the widely usedemamectin benzoate (Grant, 2002; Costello, 2006).

OPs kill 100% of post chalimus stage lice, but have littleefficacy against chalimi (Roth et al., 1996). Repeatedexposure to organophosphates can lead to resistancedevelopment (Jones et al., 1992; Fallang et al., 2004) andthis has prompted for higher dosage and reduction ofsafety margins in fish (Roth et al., 1993).

Hydrogen peroxide can dislodge 85–100% of sea lice at1.5 g/L for a 20 min bath (Thomassen, 1993), however it istoxic to fish when the temperature is over 14 8C (Alder-man, 2002). Hydrogen peroxide can delay chalimusdevelopment, but some pre-adult and adult lice recoveredand re-attached to fish (Johnson et al., 1993; McAndrewet al., 1998; Pozo et al., 2008). Prolonged exposure alsocaused gill damage (Thomassen, 1993) and resistance toHydrogen peroxide was also reported (Treasurer et al.,2000).

Synthetic pyrethroids are currently the most popularcompounds used for bath treatment. These interventionsare applied more frequently toward the end of theproduction cycle when salmon are larger, and oraltreatments are consequently more costly (Lees et al.,2008). Furthermore, bath treatment requires almost nonewithdrawal period. Resistance or reduced sensitivity tothese compounds was reported in Norway (Sevatdal andHorsberg, 2003; Sevatdal et al., 2005).

Most recently, insect growth regulators (IGRs) havebeen developed as in-feed sea lice treatment, they inhibitchitin synthesis and thus not effective against adult lice.Diflubenzuron was effective at an oral dose of 75 mg/kgBW/day for 14 days, while teflubenzuron was effective at10 mg/kg BW/day for 7 days (Branson et al., 2000; Ritchieet al., 2002). The major disadvantage of IGRs is theirtoxicity to marine crustaceans (Burridge et al., 2004;Waddy et al., 2007). Emamectin benzoate (EB) waseffective at an oral dose of 50 mg/kg BW/day for 7consecutive days. There have been anecdotal reports ofreduced sensitivity and potential resistance of sea lice toEB, particularly in Chile. Most recently, Lees et al. (2008)reported that not all treatments were effective and therewas a reduction in efficacy over a period of 5 years inScottish salmon farms.

4. Conclusions

It is obvious from this brief discussion that there istransfer of (either man-made or natural) parasites fromfarmed to wild fish stocks and vice versa. When this occursthe parasite can cause severe epizootics (with high fishmortalities) in either hatcheries or cage cultures or innatural habitats (e.g., rivers).

Conflict of interest statement

FCG is an employee of Novartis Animal Health CanadaInc., Victoria, Prince Edward Island, Canada and PTKW hasno financial nor personal relationships with other people

or organizations that could inappropriately influence(bias) the contents of this publication.

References

Alderman, 2002. Trends in therapy and prophylaxis. Bull. Euro Assoc. FishPath. 22, 117–125.

Anon., 2002. Review and synthesis of the environmental impacts ofaquaculture. The Scottish Association for Marine Science and NapierUniversity. Scottish Executive Central Research Unit, UK, pp. 24–27.Available at: http://www.scotland.gov.uk/Resource/Doc/46951/0030621.pdf (last accessed on 29 April 2009).

Anon., 2006. Aquaculture Association of Canada. Available at: http://www.aquacultureassociation.ca/news/Sea_Lice_Fact_Sheet.pdf (lastaccessed on 9 March 2009).

Ardelli, B.F., Woo, P.T.K., 1999. The therapeutic use of isometamidiumchloride against Cryptobia salmositica in rainbow trout (Oncorhynchusmykiss). Dis. Aqua. Org. 37, 195–203.

Ardelli, B.F., Woo, P.T.K., 2001. Therapeutic and prophylactic effects ofisometamidium chloride (Samorin) against the haemoflagellate Cryp-tobia salmositica in chinook salmon (Oncorhynchus tshawytscha).Parasitol. Res. 87, 18–26.

Bader, J.A., Shotts Jr., E.B., Steffens, W.L., Lom, J., 1998. Occurrence of Lomacf. salmonae in brook, brown and rainbow trout from Buford trouthatchery, Georgia, USA. Dis. Aquat. Organ. 34, 211–216.

Beamish, F.W.H., Sitja-Bobadilla, A., Jebbink, J.A., Woo, P.T.K., 1996. Bioe-nergetic cost of cryptobiosis in fish: rainbow trout (Oncorhynchusmykiss) infected with Cryptobia salmositica and with an attenuatedlive vaccine. Dis. Aquat. Organ. 25, 1–8.

Becker, J.A., Speare, D.J., Daley, J., Dick, P., 2002. Effects of monensin doseand treatment time on xenoma reduction in microsporidial gill dis-ease in rainbow trout, Oncorhynchus mykiss (Walbaum). J. Fish Dis. 25,673–680.

Becker, J.A., Speare, D.J., 2007. Transmission of the microsporidian gillparasite, Loma salmonae. Anim. Health Res. Rev. 8, 59–68.

Bower, S.M., Margolis, L., 1984. Distribution of Cryptobia salmositica, ahaemoflegellate of fishes in British Columbia and the seasonal patternof infection in a coastal river. Can. J. Zool. 62, 2512–2518.

Bower, S.M., Thompson, A.B., 1987. Hatching of the Pacific salmon leech(Piscicola salmositica) from cocoons exposed to various treatments.Aquaculture 66, 1–8.

Bower, S.M., Margolis, L., MacKay, R.J., 1985. Potential usefulness ofchlorine for controlling Pacific salmon leeches, Piscicola salmositicain hatcheries. Can. J. Fish. Aquat. Sci. 42, 1986–1993.

Boxshall, G.A., Bravo, S., 2000. On the identity of the common Caligus(Copepoda: Siphonostomatoida: Caligidae) from salmonid netpensystems in southern Chile. Contrib. Zool. 69, 137–146.

Branson, E.J., Rønsberg, S.S., Ritchie, G., 2000. Efficacy of teflubenzuron(Calicide1) for the treatment of sea lice, Lepeophtheirus salmonis(Krøyer 1838), infestations of farmed Atlantic salmon (Salmo salarL.). Aqua. Res. 31, 861–867.

Brooks, K.M., 2009. Considerations in developing an integrated pestmanagement programme for control of sea lice on farmed salmonin Pacific Canada. J. Fish Dis. 32, 59–73.

Brown, A.M.V., Kent, M.L., Adamson, M.L., 1998. Phylogeny of microspor-idian parasites of fishes reveals dispersed ribosomal RNA genes inrelatives of Loma salmonae. In: Program guide and abstracts – 73rdAnnual Meeting of the American Society of Parasitologists, 16–20August. American Society of Parasitologists, Kona, Hawaii, p. 80 pp..

Bruno, D.W., Collins, R.O., Morrison, C.M., 1995. The occurrence of Lomasalmonae (Protozoa: Microspora) in farmed rainbow trout, Oncor-hynchus mykiss Walbaum, in Scotland. Aquaculture 133, 341–344.

Boxaspen, K., 2006. A review of the biology and genetics of sea lice. ICES J.Mar. Sci. 63, 1304–1316.

Buchmann, K., Bresciani, J., 2006. Monogenea (Phylum Platyhelminthes),.In: Woo, P.T.K. (Ed.), Fish Diseases and Disorders. Protozoan andMetazoan Infections, 2nd ed., vol. 1. CABI Publishing, Wallingford,U.K., pp. 297–344.

Burridge, L.E., Hamilton, N., Waddy, S.L., Haya, K., Mercer, S.M., Green-halgh, R., Tauber, R., Radecki, L.S., Wislocki, P.G., Endris, R.G., 2004.Acute toxicity of emamectin benzoate (SLICETM) in fish feed toAmerican lobster, Homarus americanus. Aqua. Res. 35, 713–722.

Butterworth, K.G., Li, W.M., McKinley, R.S., 2004. Carbon and nitrogenstable isotopes: a tool to differentiate between Lepeophtheirussalmonis and different salmonid host species? Aquaculture 241,529–538.

Constantine, J., 1999. Fact Sheet: Estimating the Cost of Loma salmonae toB.C. Aquaculture. Animal Health Branch, Aquaculture Program, BC

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216 215

Ministry of Agriculture and Food, Abbotsford Agriculture Centre,Abbotsford.

Costello, M.J., Grant, A., Davies, I.M., Cecchini, S., Papoutsoglou, S., Quigley,D., Saroglia, M., 2001. The control of chemicals used in aquaculture inEurope. J. Appl. Ichthyol. 17, 173–180.

Costello, M.J., 2006. Ecology of sea lice parasitic on farmed and wild fish.Trends Parasitol. 22, 475–483.

Costello, M.J., 2009. The global economic cost of sea lice to the salmonidfarming industry. J. Fish Dis. 32, 115–118.

Currie, J.L.M., Woo, P.T.K., 2007. Susceptibility of sexually mature rainbowtrout, Oncorhynchus mykiss to experimental cryptobiosis caused byCryptobia salmositica. Parasitol. Res. 101, 1057–1067.

Docker, M.F., Devlin, R.H., Richard, J., Kent, M.L., 1997. Sensitive andspecific polymerase chain reaction assay for detection of Loma sal-monae (Microsporea). Dis. Aquat. Organ. 29, 41–48.

Dykova, I., 2006. Phylum microspora. In: Woo, P.T.K. (Ed.), Fish Diseasesand Disorders. Protozoan and Metazoan Infections, 2nd ed., vol. 1.CABI Publishing, Wallingford, U.K, pp. 205–229.

Fallang, A., Ramsay, J.M., Sevatdal, S., Burka, J.F., Jewess, P., Hammell,K.L., Horsbergs, T.E., 2004. Evidence for occurrence of an organopho-sphate-resistant type of acetylcholinesterase in strains of sealice (Lepeophtheirus salmonis Krøyer). Pest Manage. Sci. 60, 1163–1170.

FAO Fisheries and Aquaculture Information and Statistics Service, 2008.Aquaculture Production 1950-2006. FISHSTAT Plus-Universal Soft-ware for Fishery Statistical Time Series [Online or CD-ROM]. Food andAgriculture Organization of the United Nations. Available at: http://www.fao.org/fi/statist/FISOFT/FISHPLUS.asp (last accessed on 1March 2009).

Forward, G.M., Woo, P.T.K., 1996. An in vitro study on the mechanism ofinnate immunity in Cryptobia-resistant brook charr (Salvelinus fonti-nalis) against Cryptobia salmositica. Parasitol. Res. 82, 238–241.

Forward, G.M., Ferguson, M.M., Woo, P.T.K., 1995. Susceptibility of brookcharr, Salvelinus fontinalis to the pathogenic haemoflagellate, Crypto-bia salmositica, and the inheritance of innate resistance by progeniesof resistant fish. Parasitology 111, 337–345.

Gonzalez, L., Carvajal, J., 2003. Life cycle of Caligus rogercresseyi, (Cope-poda: Caligidae) parasite of Chilean reared salmonids. Aquaculture220, 101–117.

Grant, A.N., 2002. Medicines for sea lice. Pest Manage. Sci. 58, 521–527.Guselle, N.J., Markham, R.J., Speare, D.J., 2006. Intraperitoneal adminis-

tration of beta-1,3/1,6-glucan to rainbow trout, Oncorhynchus mykiss(Walbaum), protects against Loma salmonae. J. Fish Dis. 29, 375–381.

Hansen, H., Buchmann, K., Bakke, T.A., 2003. Mitochondrial DNA variationof Gyrodactylus spp. (Monogenea, Gyrodactylidae) populations infect-ing Atlantic salmon, grayling and rainbow trout in Norway andSweden. Int. J. Parasitol. 33, 1471–1478.

Hauck, A.K., 1984. A mortality and associated tissue reactions of chinooksalmon, Oncorhynchus tshawytscha (Walbaum), caused by the micro-sporidian Loma sp. J. Fish Dis. 7, 217–229.

Heuch, P.A., Bjørn, P.A., Finstad, B., Holst, J.C., Asplin, L., Nilsen, F., 2005. Areview of the Norwegian ‘‘National action plan against salmon lice onsalmonids’’: the effect on wild salmonids. Aquaculture 246, 79–92.

Heuch, P.A., Mo, T.A., 2001. A model of salmon louse production inNorway: effects of increasing salmon production and public manage-ment measures. Dis. Aquat. Organ. 45, 145–152.

Heuch, P.A., Øines, Ø., Knutsen, J.A., Schram, T.A., 2007. Infection of wildfishes by the parasitic copepod Caligus elongatus on the south eastcoast of Norway. Dis. Aquat. Organ. 77, 149–158.

Jones, M.W., Sommerville, C., Wootten, R., 1992. Reduced sensitivity of thesalmon louse, Lepeophtheirus salmonis, to the organophosphatedichlorvos. J. Fish Dis. 15, 197–202.

Jones, S.R., Prosperi-Porta, G., Kim, E., Callow, P., Hargreaves, N.B., 2006.The occurrence of Lepeophtheirus salmonis and Caligus clemensi (Cope-poda: Caligidae) on three-spine stickleback Gasterosteus aculeatus incoastal British Columbia. J. Parasitol. 92, 473–480.

Johnson, S.C., Albright, L.J., 1991. The developmental stages ofLepeophtheirus salmonis (Kroyer, 1837) (Copepoda, Caligidae). Can.J. Zool. 69, 929–950.

Johnson, S.C., Constible, J.M., Richard, J., 1993. Laboratory investigations ofthe toxicological and histopathological effects of hydrogen peroxideto salmon and its efficacy against the salmon louse Lepeophtheirussalmonis. Dis. Aquat. Organ. 17, 197–204.

Kabata, Z., 1979. Parasitic Copepoda of British Fishes. The Ray Society,London, 179 pp.

Keeling, P.J., Fast, N.M., 2002. Microsporidia: biology and evolution ofhighly reduced intracellular parasites. Annu. Rev. Microbiol. 56, 93–116.

Kent, M.L., 1992. Diseases of seawater netpen-reared salmonid fishes inthe Pacific Northwest. Can. Spec. Publ. Fish Aquat. Sci. 116, 39–42.

Kent, M.L., Elliott, D.G., Groff, J.M., Hedrick, R.P., 1989. Loma salmonae(Protozoa: Microspora) infections in seawater reared coho salmonOncorhynchus kisutch. Aquaculture 80, 211–222.

Kent, M.L., Dawe, S.C., Speare, D.J., 1995. Transmission of Loma salmonae(Microsporea) to chinook salmon in sea water. Can. Vet. J. 36, 98–102.

Kent, M.L., Poppe, T.T., 1998. Diseases of Seawater Netpen-reared Salmo-nids. Pacific Biological Station Press, Nanaimo, British Columbia,Canada, 138 pp.

Kent, M.L., Traxler, G.S., Kieser, D., Richard, J., Dawe, S.C., Shaw, R.W.,Prosperi-Porta, G., Ketcheson, J., Evelyn, T.P.T., 1998. Survey of sal-monid pathogens in ocean-caught fishes in British Columbia, Canada.J. Aquat. Anim. Health 10, 211–219.

Kent, M.L., 2000. Marine netpen farming leads to infections with someunusual parasites. Int. J. Parasitol. 30, 321–326.

Kent, M.L., Speare, D.J., 2005. Review of the sequential development ofLoma salmonae (Microsporidia) based on experimental infections ofrainbow trout (Oncorhynchus mykiss) and Chinook salmon (O. tsha-wytscha). Folia Parasitol. (Praha) 52, 63–68.

Krkosek, M., Ford, J.S., Morton, A., Lele, S., Myers, R.A., Lewis, M.A., 2007.Declining wild salmon populations in relation to parasites from farmsalmon. Science 318, 1772–1775.

Lees, F., Baillie, M., Gettinby, G., Revie, C.W., 2008. The efficacy of ema-mectin benzoate against infestations of Lepeophtheirus salmonis onfarmed Atlantic salmon (Salmo salar L) in Scotland, 2002–2006. PLoSONE 3, e1549.

Lester, R.J.G., Hayward, C.J., 2006. Phylum Arthropoda. In: Woo, P.T.K.(Ed.), Fish Diseases and Disorders. Protozoan and Metazoan Infec-tions., 2nd ed., vol.1. CABI Publishing, Wallingford, U.K, pp. 466–565.

Li, S., Woo, P.T.K., 1995. Efficacy of a live Cryptobia salmositica vaccine, andthe mechanism of protection in vaccinated Oncorhynchus mykiss(Walbaum) against cryptobiosis. Vet. Immunol. Immunopathol. 48,343–353.

Li, S., Woo, P.T.K., 1997. Vaccination of rainbow trout, Oncorhynchusmykiss (Walbaum) against cryptobiosis: efficacy of the vaccine infresh and sea water. J. Fish Dis. 20, 369–374.

Magor, B.G., 1987. First report of Loma sp. (Microsporida) in juvenile cohosalmon (Oncorhynchus kisutch) from Vancouver Island, British Colum-bia. Can. J. Zool. 65, 751–752.

McAndrew, K., Sommerville, C., Wootten, R., Bron, J.e., 1998. The effects ofhydrogen peroxide treatment on different life-cycle stages of thesalmon louse, Lepeophtheirus salmonis (Krøyer 1837). J. Fish Dis. 21,221–228.

Mehta, M., Woo, P.T.K., 2002. Acquired cell-mediated protection in rain-bow trout, Oncorhynchus mykiss against the haemoflagellate, Crypto-bia salmositica. Parasitol. Res. 88, 956–962.

Morrison, C.M., Sprague, V., 1983. Loma salmonae (Putz, Hoffman andDunbar, 1965) in the rainbow trout, Salmo gairdneri Richardson, and L.fontinalis sp. nov. (Microsporida) in the brook trout, Salvelinus fonti-nalis (Mitchill). J. Fish Dis. 6, 345–353.

Nordhagen, J., Heuch, P., Schram, T., 2000. Size as indicator of origin ofsalmon lice, Lepeophtheirus salmonis (Copepoda: Caligidae). Contrib.Zool. 69, 99–108.

Pozo, V., Bravo, S., Erranz, F., Bassaletti, P., 2008. Monitoring the effec-tiveness of hydrogen peroxide in the control of sea lice Caligusrogercresseyi in Chile. 7th International Sea Lice Conference, PuertoVaras, Chile (Abstract).

Pike, A.W., Wadsworth, S.L., 1999. Sea lice on salmonids: their biology andcontrol. Adv. Parasit. 44, 233–337.

Putz, R.E., Hoffman, G.L., Dunbar, C.E., 1965. Two new species of Pleisto-phora from North American fish with a synopsis of Microsporidia offreshwater and euryhaline species. J. Protozool. 12, 228–236.

Riddell, B.E., Beamish, R.J., Richards, L.J., Candy, J.R., 2008. Comment on‘‘Declining Wild Salmon Populations in Relation to Parasites fromFarm Salmon’’. Science 322, 1790–11790.

Ritchie, G., Rønsberg, S.S., Hoff, K.A., Branson, E.J., 2002. Clinical efficacy ofteflubenzuron (Calicide) for the treatment of Lepeophtheirus salmonisinfestations of farmed Atlantic salmon Salmo salar at low watertemperatures. Dis. Aquat. Organ. 51, 101–106.

Rodrıguez-Tovar, L.E., Becker, J.A., Markham, R.J., Speare, D.J., 2006. Induc-tion time for resistance to microsporidial gill disease caused by Lomasalmonae following vaccination of rainbow trout (Oncorhynchusmykiss) with a spore-based vaccine. Fish Shellfish Immunol. 21,170–175.

Roth, M., Richards, R.H., Sommerville, C., 1993. Current practices in thechemotherapeutic control of sea lice infestations in aquaculture: areview. J. Fish Dis. 16, 1–26.

Roth, M., Richards, R.H., Dobson, D.P., Rae, G.H., 1996. Field trial on theefficacy of the organophosphate compound Azimethiphos for thecontrol of sea lice (Copepoda, Caligidae) infestations of farmed Atlan-tic salmon (Salmo salar). Aquaculture 140, 197–217.

F.C. Guo, P.T.K. Woo / Veterinary Parasitology 163 (2009) 207–216216

Sanchez, J.G., Speare, D.J., Markham, R.J.F., Jones, S.R.M., 2001a. Experi-mental vaccination of rainbow trout against Loma salmonae using alive low-virulence variant of L. salmonae. J. Fish Biol. 59, 442–448.

Sanchez, J.G., Speare, D.J., Markham, R.J.F., Wright, G.M., Kibenge, F.S.B.,2001b. Localization of the initial development stages of Loma salmo-nae in rainbow trout (Oncorhynchus mykiss). Vet. Pathol. 38, 540–546.

SEARCH, 2004. Sea lice resistance to chemotherapeutants: a handbook inresistance management. Prepared in association with the EU-fundedproject SEARCH (QLK2-CT-00809). Available at http://www.rotham-sted.bbsrc.ac.uk/pie/search-EU/Handbook.pdf (last accessed on 18April 2009).

Sevatdal, S., Horsberg, T.E., 2003. Determination of reduced sensitivity insea lice (Lepeophtheirus salmonis) against the pyrethroid deltamethrinusing bioassays and probit modelling. Aquaculture 218, 21–31.

Sevatdal, S., Fallang, A., Ingebrigtsen, K., Horsberg, T.E., 2005. Monoox-ygenase mediated pyrethroid detoxification in sea lice(Lepeophtheirus salmonis). Pest. Manage. Sci. 61, 772–778.

Shaw, R.W., Kent, M.L., Docker, M.F., Devlin, R.H., Brown, A., Adamson,M.L., 1997. A new Loma species (Microsporea) in shiner perch (Cym-atogaster aggregata). J. Parasitol. 83, 296–301.

Shaw, R.W., Kent, M.L., Adamson, M.L., 1998. Modes of transmission ofLoma salmonae (Microsporidia). Dis. Aquat. Organ. 33, 151–156.

Shaw, R.W., Kent, M.L., Brown, A.M.V., Whipps, C.M., Adamson, M.L., 2000.Experimental and natural host specificity of Loma salmonae (Micro-sporidia). Dis. Aquat. Organ. 40, 131–136.

Shinn, A., Bron, J., Gray, D., Sommerville, C., 2000. Elemental analysis ofScottish populations of the ectoparasitic copepod Lepeophtheirussalmonis. Contrib. Zool. 69, 79–87.

Soleng, A., Poleo, A.B.S., Alstad, N.E.W., Bakke, T.A., 1999. Aqueous alu-minium eliminates Gyrodactylus salaris (Platyhelminthes, Monoge-nea) infections in Atlantic salmon. Parasitology 119, 19–25.

Speare, D.J., Brackett, J., Ferguson, H.W., 1989. Sequential pathology of thegills of coho salmon with a combined diatom and microsporidian gillinfection. Can. Vet. J. 30, 571–575.

Speare, D.J., Arsenault, G.J., Buote, M.A., 1998. Evaluation of rainbow troutas a model for use in studies on pathogenesis of the branchialmicrosporidian Loma salmonae. Contemp. Top. Lab. Anim. Sci. 37,68–71.

Speare, D.J., Athanassopoulou, F., Daley, J., Sanchez, J.G., 1999. A preli-minary investigation of alternatives to fumagillin for the treatment ofLoma salmonae infection in rainbow trout. J. Comp. Pathol. 121, 241–248.

Speare, D.J., Markham, R.J.F., Guselle, N.J., 2007. Development of aneffective whole-spore vaccine to protect against microsporidial gilldisease in rainbow trout (Oncorhynchus mykiss) by using a low-virulence strain of Loma salmonae. Clin. Vaccine Immunol. 14,1652–1654.

Stone, J., Sutherland, I.H., Sommerville, C., Richards, R.H., Endris, R.G.,2000a. The duration of efficacy following oral treatment with

emamectin benzoate against infestations of sea lice, Lepeophtheirussalmonis (Krøyer), in Atlantic salmon Salmo salar L. J. Fish Dis. 23,185–192.

Stone, J., Sutherland, I.H., Sommerville, C., Richards, R.H., Varma, K.J.,2000b. Commercial trials using emamectin benzoate to control sealice Lepeophtheirus salmonis infestations in Atlantic salmon Salmosalar. Dis. Aquat. Organ. 41, 141–149.

Stone, J., Sutherland, I.H., Sommerville, C., Richards, R.H., Varma, K.J.,2000c. Field trials to evaluate the efficacy of emamectin benzoatein the control of sea lice, Lepeophtheirus salmonis (Krøyer) and Caliguselongatus Nordmann, infestations in Atlantic salmon Salmo salar L.Aquaculture 186, 205–219.

Tan, C.W., Jesudhasan, R.R.R., Woo, P.T.K., 2008. Towards a DNA vaccineagainst salmonid cryptobiosis caused by Cryptobia salmositica. Para-sitol. Res. 102, 265–275.

Thomassen, J.M., 1993. Hydrogen peroxide as a delousing agent forAtlantic salmon. In: Boxshall, G.A., Defaye, D. (Eds.), Pathogensof Wild and Farmed Fish. Sea Lice, Ellis Horwood, New York, pp.290–295.

Treasurer, J., Grant, A., 1994. ‘Second’ louse species must not be ignored.Fish Farmer 17, 46–47.

Treasurer, J.W., Wadsworth, S., Grant, A., 2000. Resistance of sea liceLepeophtheirus salmonis (Krøyer), to hydrogen peroxide on farmedAtlantic salmon, Salmo salar. Aqua. Res. 31, 855–860.

Von Gersdorff Jorgensen, L., Heinecke, R.D., Kania, P., Buchmann, K., 2008.Occurrence of gyrodactylids on wild Atlantic salmon, Salmo salar L., inDanish rivers. J. Fish Dis. 31, 127–134.

Waddy, S.L., Merritta, V.A., Hamilton-Gibsona, M.N., Aikena, D.E., Burrid-gea, L.E., 2007. Relationship between dose of emamectin benzoateand molting response of ovigerous American lobsters (Homarus amer-icanus). Ecotox. Environ. Safe. 67, 95–99.

Wales, J.M., Wolf, H., 1955. Three protozoan diseases of trout in California.Calif. Fish Game 41, 183–187.

Woo, P.T.K., 1998. Protection against Cryptobia (Trypanoplasma) salmosi-tica and salmonid cryptobiosis. Parasitol. Today 14, 272–277.

Woo, P.T.K., 2003. Cryptobia (Trypanoplasma) salmositica and salmonidcryptobiosis. J. Fish Dis. 26, 627–646.

Woo, P.T.K., 2006a. Strategies against piscine parasitosis: the Cryptobiamodel. In: Phang, S.M., Aisyah, S., Chong, V.C., George, M., Siti Aisyah,M., Ho, S.C. (Eds.), Innovations and Technologies in Oceanography forSustainable Development. Maritime Research Centre. University ofMalaya, Kuala Lumpur, Malaysia, pp. 17–28.

Woo, P.T.K., 2006b. Diplomonadida (Phylum Parabasalia) and Kinetoplas-tea (Phylum Euglenozoa). In: Woo, P.T.K. (Ed.), Fish Diseases andDisorders. Protozoan and Metazoan Infections., 2nd ed., vol. 1. CABIPublishing, Wallingford, U.K, pp. 46–115.

Woo, P.T.K., Li, S., 1990. In vitro attenuation of Cryptobia salmositica and itsuse as a live vaccine against cryptobiosis in Oncorhynchus mykiss. J.Parasitol. 76, 752–755.