JOPAA2 94(4) 771–998 (2008)(ISSN 0022-3395)

The Journal of theAmerican Society ofParasitologists

The Journal of

ANISAKID LARVAE IN ATLANTIC SALMON (SALMO SALAR L.) GRILSE AND

POST-SMOLTS: MOLECULAR IDENTIFICATION AND HISTOPATHOLOGY

T. M. Murphy, M. Berzano, S. M. O’Keeffe*, D. M. Cotter , S. E. McEvoy , K. A. Thomas , N. P. O Maoileidigh , andK. F. WhelanCentral Veterinary Research Laboratory, Backweston Campus, Young’s Cross, Celbridge, County Kildare, Ireland. e-mail: tom.murphy@agriculture.gov.ie

ANISAKID LARVAE IN ATLANTIC SALMON (SALMO SALAR L.) GRILSE AND

POST-SMOLTS: MOLECULAR IDENTIFICATION AND HISTOPATHOLOGY

T. M. Murphy, M. Berzano, S. M. O’Keeffe*, D. M. Cotter�, S. E. McEvoy�, K. A. Thomas`, N. P. O Maoileidigh�, andK. F. Whelan�Central Veterinary Research Laboratory, Backweston Campus, Young’s Cross, Celbridge, County Kildare, Ireland. e-mail: tom.murphy@agriculture.gov.ie

ABSTRACT: The molecular identification and histopathology are described for the parasitic larvae of a nematode species present in theabdominal cavity of Atlantic salmon (Salmo salar) grilse caught in fish traps on their natal river in the west of Ireland and post-smoltscollected during experimental trawls on the continental shelf edge of the northeast Atlantic Ocean. Larvae in the adult and juvenilesalmon were identified as Anisakis simplex sensu stricto by PCR amplification and RFLP and sequencing of the ITS gene and PCRamplification and sequencing of the cox2 gene. Parasitic nematode larvae in the grilse were either encapsulated in the abdominalmesentery associated with the pyloric ceca or on the serosal surface of the liver and in the vent region. In some fish, larvae were foundin the parenchyma of the liver and muscularis circularis of the intestine. In general, the larvae induced a limited cellular response apartfrom the occurrence of focal melanin macrophage aggregates and individual eosinophilic granular cells in the connective tissue capsule.Melanin macrophage aggregates were also present among the hepatocytes adjacent to encapsulated larvae in the liver. The reaction tothe parasites was more severe in the wall of the intestine. Encapsulated nematode larvae caused displacement, vacuolation, and necrosisof the circular muscle fibers. The stratum compactum was also disrupted with focal areas of degeneration. Overall, the intestinal wallhad a hypercellular appearance with extensive cellular infiltration comprising eosinophilic granular cells, macrophages, lymphocytes,and fibrocytes. The post-smolts were caught in May during the early oceanic phase of their life cycle. In these fish, A. simplex sensustricto larvae were found lying free on the serosal surface of the intestine and liver without any apparent histologic changes. This is theearliest in the marine migration of Atlantic salmon that A. simplex sensu stricto infection has been recorded.

Nematodes of the Anisakidae have a complex life cycle using

marine crustacea and fish as paratenic hosts with fish eating

vertebrates such as marine mammals and birds as definitive hosts.

Anisakis spp. are a major problem for commercial fishing

industries and a public health concern in many countries. They

can cause significant illness in humans, manifested by either

gastric or intestinal symptoms, or an allergic hypersensitivity

often induced by material from dead parasites (Chai et al., 2005;

Audicana and Kennedy, 2008). Spotted chub mackerel, squid,

herring, hake, anchovies, and sardines are considered the main

sources of human anisakiasis (Audicana et al., 2002).

The use of genetic markers such as enzyme loci has allowed

parasitic anisakid larvae to be used as biologic tags for stock

identification of demersal and pelagic fish species, including

Atlantic salmon, Salmo salar (Beverley-Burton, 1978; Moser and

Hsieh, 1992; Mattiucci, 2006). Anisakis spp. larvae also have the

potential to be used as biomarkers of marine pollution, since they

possess the capacity to accumulate heavy metals at a rate up to

300 times greater than their cephalopod, fish, and cetacean hosts

(Pascual and Abollo, 2005).

The systematic and taxonomic status of species belonging to

Anisakis has been clarified by genetic and morphological studies.

There are 9 taxa divided into 2 main clades (Mattiucci and

Nascetti, 2006; Valentini et al., 2006). The first clade includes 3

sibling species of the Anisakis simplex complex i.e., A. simplex

sensu stricto, A. pegreffii, and A. simplex C, as well as A. typica

and A. ziphidarum plus a new taxon, provisionally identified as

Anisakis sp. A (Pontes et al., 2005). The second clade encompasses

the species A. physeteris, A. brevispiculata, and A. paggiae. Each

species occupies a distinct ecological niche with a defined

geographical distribution and specific host–parasite relationship.

Anisakis simplex s.s. is found from 30uN to the Arctic Polar

Circle in both the western and eastern Atlantic and Pacific oceans

(Mattiucci et al., 1997; Umehara et al., 2006). However, the

distribution of this parasite overlaps with A. pegreffii along the

Portuguese and Spanish coasts. Adult A. simplex s.s. have been

recorded in 9 species of cetacean hosts, and L3 larvae have been

found in 4 squid and 26 fish species (Mattiucci and Nascetti,

2006). In paratenic fish hosts, the larvae become encapsulated in

the mesentery and serosal surface of the visceral organs and

sometimes encyst beneath the serosa and in somatic musculature

(Dezfuli et al., 2007).

There are many causal factors for the continued decline in the

populations of wild anadromous Atlantic salmon (Parrish et al.,

1998). It has been suggested that diseases including infections with

metazoan parasites may affect the migratory behavior, growth

rate, and fitness of these fish to withstand changing oceanic

conditions (Bakke and Harris, 1998). To date, the biology of the

Anisakis sp. in wild salmonids, especially their molecular

taxonomy, pathology, and oceanic distribution, has not been

studied in any great detail. The prevalence of anisakid larvae in

wild sockeye salmon (Oncorhynchus nerka) and Atlantic salmon

can be 100% and 64.5%, respectively (Deardorff and Kent, 1989;

Bristow and Berland, 1991). Since 2007, there have been a number

of reports of large numbers of wild Atlantic salmon returning to

natal rivers in the British Isles to spawn with swollen, inflamed,

and bleeding vents associated with A. simplex larvae (Beck et al.,

2008; FRS, 2008).

The aim of the present study was to identify by molecular

techniques the parasitic larvae present in Atlantic salmon grilse

returning to their natal river system on the west coast of Ireland.

The methods used were PCR-based restriction fragment length

polymorphism (PCR-RFLP) of the ribosomal DNA (rDNA) of

the internal transcribed spacer (ITS) and analysis and sequencing

of the PCR amplicon of the mtDNA cox2 gene. Larvae found in

the abdominal cavity of juvenile post-smolts, which had recently

Received 5 June 2009; revised 4 August 2009; accepted 7 September2009.

*Dublin Institute of Technology, Kevin Street, Dublin 2, Ireland.{Marine Institute, Furnace, Newport, County Mayo, Ireland.{Department of Zoology, Lee Maltings, University College, Cork,

Ireland.DOI: 10.1645/GE-2194.1

J. Parasitol., 96(1), 2010, pp. 77–82

F American Society of Parasitologists 2010

77

started the marine phase of their life cycle and which had been

caught in the Atlantic Ocean off the west coasts of Ireland and

Scotland, were also analyzed using the same molecular tech-

niques. There have only been a limited number of reports

describing the pathology of larval anisakids in adult Atlantic

salmon (Anon., 2008; Beck et al., 2008). An ancillary goal of the

study was to further elucidate the histopathology induced in adult

and young salmon by the migrating larvae.

MATERIALS AND METHODS

Adult salmon

The adult Atlantic salmon examined in this study were ranched grilsecaught while returning to their natal lake/river system on the west coast ofIreland. They had been hatched from eggs that had been artificiallyfertilized and reared to the smolt stage at the hatchery of the MarineInstitute in County Mayo (53u359N, 9u559W) and released into LoughFurnace (a tidal lake adjacent to the hatchery on the Burrishoole Riversystem) as S-1 smolts in spring 2006 to begin their marine migratory phase.The grilse were captured in specialized upstream traps located betweenLough Furnace and Feeagh, as they migrated to the spawning streams inthe upper reaches of Lough Feeagh during late summer and autumn 2007.Initially, the sexually maturing grilse were held in a floating cage inbrackish water on Lough Furnace; they were subsequently transferred to afreshwater broodstock pond just prior to stripping in December.

Post-smolts

Capture: The post-smolts were caught in the Atlantic Ocean, off thewest and northwest coasts of Scotland and Ireland, respectively, duringcruises of 2 research vessels, the Celtic Voyager and Celtic Explorer. Fishwere sampled during pelagic trawls at various sampling stations duringMay 2008. The trawling speed was 3 knots, and the trawls wereapproximately 62 m wide and 13 m deep.

Collection of anisakid larvae

Adult fish: In December, 5 grilse with hemorrhagic vents were randomlyselected, killed by a blow to the head, and examined for the presence ofparasitic larvae. A further 6 adults were killed and examined in January2008 at the end of the stripping period. The length and weight of each fishwas recorded. The abdomen was opened and examined for the presence ofparasitic larvae. The number of nematodes visible in the body cavity orloosely attached to the mesentery of the intestine and serosal surface of theliver was noted. A number of larvae from 10 of these 11 adult fishes wereisolated and fixed in 70% ethanol for species identification. In December,parasites were collected from only 4 adults. Parasites from 3 of these fishwere mixed in 1 container, and larvae from the remaining grilse were storedin a second specifically identified container. A separate labeled containerwas used for larvae collected from each of 6 grilse examined in January.

Post-smolts: The post-smolts were caught in 2 trawls during a researchcruise of the Celtic Voyager in May 2008 in the northeastern region of theAtlantic Ocean. The first trawl started at 55u089N, 10u009W and finishedat 55u099N, 9u599W. The second took place at 55u089N, 10u599W to54u559N, 10u189W. Immediately after each trawl, the fish were processedin the ship’s laboratory. A total of 51 fish was caught, and their length andweight were recorded before they were stored at 220C for furtherexamination on shore. The fish were taken to the Marine Institute’slaboratory at the end of the cruise. They were then thawed and their bodycavities opened and examined for presence of parasitic larvae. Any larvafound was preserved in 100% ethanol for identification at a future date.

Histological studies

Sections of heart, liver, spleen, head kidney, dorsal kidney, stomach,and pyloric ceca, including the pancreas and intestine, from the 11 adultsalmon were fixed in 10% buffered formalin. A total of 21 post-smolts wasrandomly selected from 434 fish caught during 8 and 13 trawls carried outby Celtic Voyager and Celtic Explorer, respectively, in the area 55u089N to59u359N and 6u239W to 10u189W. The abdominal wall was cut open andparted, and the whole fish was placed in buffered formalin. The heart,

liver, spleen, head kidney, dorsal kidney, stomach, and pyloric ceca weredissected from the body cavity before processing.

The fixed tissue from the adults and post-smolts was dehydrated inascending grades of ethanol, cleared in xylene, embedded in paraffin,sectioned at 5 mm, and stained with hematoxylin and eosin.

DNA extraction

One larva collected from the readily identifiable grilse sampled inDecember and from each of the 6 adults examined in January was selectedfor molecular identification studies. Additionally, molecular identificationwas also carried out using 4 larvae taken from the pooled sample of wormscollected from 3 grilse in December and a further 4 nematodes found in 4post-smolts. The nematodes were removed from the ethanol preservativesolution and snap frozen in liquid nitrogen prior to DNA extraction with acommercial kit (Wizard Genomic, Promega, Madison, Wisconsin). Themanufacturer’s instructions were modified as follows. Each larva wasplaced in a sterile mortar with 500 ml of nuclear lysis solution and crushedwith a sterile pestle. This mixture was transferred to a microcentrifugecontaining 120 ml EDTA (0.5 M) and 50 ml proteinase K (5 g/ml; Promega)and incubated for 3 hr at 55 C. During this time, the tube was mixed byhand every 10 min and by vortex every 1 hr. A total of 3 ml RNAsesolution was added to the tube and incubated at 37 C for a further 20 min.After incubation, 200 ml of a protein precipitation solution was added andthe tube was mixed vigorously for 20 sec. The tube was then placed on icefor 5 min, and a final pellet was obtained after a series of 3 ethanolprecipitation and centrifugation steps. The pellet was air dried for 15 minand eluted in 100 ml of DNA reconstitution solution. This was incubatedovernight at 4 C, and the amount of DNA extracted was quantified usinga spectrophotometer (Nanodrop, Thermo Scientific, Burlington, Ver-mont). The extracted DNA was stored at 220 C until analyzed.

PCR amplification

The ITS region was amplified using forward primer, NC5; reverseprimer, NC2; and a procedure described by Abollo et al. (2003). The PCRmix contained 2 U of Immolase polymerase in 310 Immobuffer (Bioline,London, UK), 2.5 mM MgCL2, 0.2 mM of dNTPs, 0.2 mM NC2, 0.2 mMNC5, 0.4 mg/ml BSA, and 2 ml of template DNA made up to a finalvolume 50 ml. The PCR was performed in a Peltier thermal cycler (PTC-200, MJ Research Inc., St Bruno, Quebec, Canada), and the conditionswere as follows: 7 min at 95 C (initial denaturation), 30 cycles of 30 sec at95 C (denaturation), 30 sec at 50 C (annealing), and 30 sec at 72 C(extension) with a final elongation step of 5 min at 72 C. A negativecontrol containing no DNA template was included to detect anycontamination that may have occurred.

The cox2 mitochondrial gene was amplified using primers COX210 andCOX211 and a modification of the protocol described by Iglesias et al.(2008). The PCR mix contained 2 U of Immolase polymerase in 310Immobuffer (Bioline, London, U.K.), 2.5 mM MgCl2, 0.2 mM of dNTPs,0.4 mM COX210, 0.4 mM COX211, 0.4 mg/ml BSA, and 2 ml of templateDNA (original DNA extract diluted 1/100) made up to a final volume50 ml. The reaction was carried out in a PTC-200 thermocycler using thefollowing conditions: 7 min at 95 C, 35 cycles of 30 sec at 95 C, 30 sec at49 C, and 35 sec at 72 C with a final elongation step of 5 min at 72 C. Anegative control with no DNA template was also included to detect anycontamination that may have occurred.

Following PCR, the ITS amplicon was subjected to RFLP using 3 fastdigest enzymes Hinf1, Hha1, and Taq1 (Fermentas, York, U.K.) Theenzymes were made up in a fast digest buffer and used according to themanufacturer’s recommendations. The digested products were analyzedby electrophoresis on a 2.5% agarose gel, and the restriction patterns werevisualized under UV light with Fluro Chem SP software.

Selected amplicons from the ITS and cox2 reactions were extracted andcleaned in preparation for sequencing using 2 commercial kits (PurelinkQuick, Invitrogen, Carlsbad, California; Genelute, Sigma, St. Louis,Missouri).

The purified DNA was then prepared and dispatched as per instructionsof the commercial sequencing company (BMR Genomics, Padova, Italy).The sequences were assembled using DNAstar module SeqManII(DNAstar Inc., Madison, Wisconsin) and compared using a BLASTsearch with those available on the National Center for BiotechnologyInformation database (http:/ www.ncbi.nlm.nih.gov/BLAST/).

78 THE JOURNAL OF PARASITOLOGY, VOL. 96, NO. 1, FEBRUARY 2010

RESULTS

The weight and length of the 11 grilse were 1.73 ± 0.25 (mean ±

S.D.) kg and 58.95 ± 2.13 (mean ± S.D.) cm, respectively. A total

of 4 males and 7 females was examined, and all of them had

parasitic nematode larvae. The number of larvae evident in the

peritoneal cavity of the fish ranged from 7 to over 50. The majority

of the larvae were loosely coiled and attached to the mesentery

surrounding the pyloric ceca and to the surface of the liver. In the 5

heavily infected fish, i.e., with more than 15 larvae, parasites were

also present on the peritoneum lining the abdominal wall close to

the vent. In a number of fish, larvae were observed migrating to the

serosal surface and leaving the substance of the liver and skeletal

musculature soon after the abdominal cavity was opened and the

visceral organs exposed to the air.

The length and weight of the 51 post-smolts were 16.4 ± 2.8

(mean ± SD) cm and 47.97 ± 22.96 (mean ± SD) g, respectively.

Four of these juvenile fish were found to have a single nematode

larva either loosely attached to the mesentery or free in the

abdominal cavity.

Amplification of the ITS region in the 11 larvae from the adult

fish produced a fragment of approximately 1,000 bp in length

and, when subjected to restriction enzyme digestion with Hinf1,

produced 3 fragments (620 bp, 250 bp, 80 bp), when digested with

Hha1, produced 2 fragments (550 bp, 430 bp). Digestion with

Taq1 resulted in 3 fragments, 430 bp, 400 bp, and 80 bp.

Digestion of the amplicons from the larvae found in the 4 post-

smolts with the 3 enzymes produced a similar restriction pattern.

All the larvae were identified as A. simplex s.s. on the basis of this

restriction-banding pattern and confirmed by the results of the

sequencing analysis (D’Amelio et al., 2000). Additional sequenc-

ing of the PCR product from the amplification of the cox2

mitochondrial gene substantiated this result. Eleven ITS sequenc-

es, including the sequences of the 4 larvae found in post-smolts

and 7 larvae from the 7 readily identified adults, were submitted

to GenBank with the following accession numbers GQ169362–

GQ169372. The GenBank accession numbers for the cox2

mitochondrial gene amplicon are GQ338428–338436.

One parasitic larva was observed in the abdominal cavity of 1

of the 21 post-smolts selected for histological studies. Despite this,

there were no significant changes or microscopic evidence of

Anisakis sp. infection in the tissues of these fish. Histological

examination of tissues from the grilse revealed larvae encapsulat-

ed in either a thin (2–3 cell layers) or thick (.3 cell layers) fibrous

capsule on the peritoneal surface of the intestine, pancreas, and

liver. Focal clumps of cellular debris were often present in the

space between the parasite cuticle and the capsule. Single

eosinophilic granular cells (EGCs), mononuclear inflammatory

cells, and melano-macrophage centers were interspersed randomly

throughout the fibrous tissue surrounding the parasites (Fig. 1).

Mechanical compression was evident in those areas of the liver

adjacent to the serosal surface with larvae attached (Fig. 2). There

was no evidence of hepatocyte necrosis or an inflammatory

cellular reaction associated with this compression.

A number of larvae were observed encysted in the parenchyma

of the liver. There was no obvious necrosis or cellular reaction

associated with the parasites other than the formation of a fibrous

capsule and displacement of the hepatic tissue. Numerous

melano-macrophage centers were present among the normal

hepatocytes adjacent to the encapsulated larvae (Fig. 3).

Larvae were present in the muscularis circularis of the intestinal

wall, causing displacement of the muscle fibers. The parasites

were encapsulated in a fibrous capsule of variable thickness.

Overall, the circular muscle band had a hypercellular appearance.

There were focal areas of smooth muscle vacuolation and necrosis

adjacent to the larvae with cellular infiltration comprising EGCs,

macrophages, lymphocytes, and fibrocytes. In some areas, the

necrosis extended into the stratum compactum and stratum

granulosum. The stratum compactum was disrupted with focal

degeneration and extensive cellular infiltration, extending as far as

the lamina propria (Fig. 4). The majority of cells were EGCs, with

some macrophages and lymphocytes; many of the former cells

were undergoing degranulation. The epithelial cells of the mucosa

appeared unaffected.

FIGURE 1. Larva of Anisakis simplex s.s. attached to mesentery in thebody cavity of adult Salmo salar surrounded by fibrous connective tissueinfiltrated with eosinophilic granular cells (EGC; arrowhead) andmacrophage activated centers (MAC; arrows) stained with hematoxylinand eosin (H&E), scale bar 5 100 mm.

FIGURE 2. Displacement of hepatocytes by Anisakis simplex s.s.encysted on the serosal surface of the liver of adult Salmo salar stainedwith H&E; scale bar 5 500 mm.

MURPHY ET AL.–ANISAKIS IN ATLANTIC SALMON 79

DISCUSSION

Anisakid nematodes have a worldwide distribution, and a large

number of pelagic fish species serve as paratenic hosts (Mattiucci

and Nascetti, 2006). In general, the various Anisakis species are

geographically distributed along latitudinal gradients, but sym-

patric areas have been identified where different species co-exist,

and, in some cases, hybridization between species such as A.

simplex s.s. and Anisakis pegreffii has occurred (Marques et al.,

2006; Mattiucci and Nascetti, 2006). Thus, accurate identification

and differentiation of larval and adult stages of anisakid

nematodes are very important for studying their life cycles,

transmission patterns, ecology, and pathology in fishes, as well as

for the diagnosis of anisakiasis in humans and marine mammals.

Morphological differentiation of the different L3 is difficult, but

molecular identification methods, such as PCR-RFLP analysis

and sequencing of the ITS region of the rDNA and sequencing of

a PCR amplicon from the mtDNA cox2 gene, have proved to be

reliable techniques (Kijewska et al., 2000; Umehara et al., 2006;

Abe, 2008; Iglesias et al., 2008; Quiazon et al., 2008).

All the larvae examined in this study were identified as A.

simplex s.s. The current trend among consumers of a preference

for natural foodstuffs and the heavy parasitic burden revealed in

this and other studies (Beck et al., 2008; FRS, 2008) increase the

likelihood of parasitized wild salmon becoming an important

source of human anisakiasis. This danger is exacerbated by the

fact that the worms can survive for a long period after the

paratenic host leaves the marine environment. In the present

study, active worms were observed at necropsy even though the

grilse had been maintained in either brackish water or freshwater

for at least 5 mo before being examined.

The oceanic distribution of the A. simplex complex depends on

the migration of the final cetacean hosts of the Delphinidae,

Monodontidae, Phocoenidae, and Balaenopteridae, and the

availability of appropriate copepod species as first intermediate

hosts. Previous reports have recorded the presence of larvae of the

A. simplex complex in Pleuronectiformes and other fishes off the

Portuguese and Madeiran coasts and in myctophid fish species in

the mid-Atlantic ridge of the central Atlantic (Pontes et al., 2005;

Marques et al., 2006; Klimpel et al., 2008). In the present study,

Anisakis sp. larvae were found in post-smolts in the area 55u089 to

55u099N and 9u599 to 10u189W in the northeast Atlantic circa

350 km west of Scotland and 240 km from the northwest coast of

Ireland on the continental shelf edge. This suggests that salmon

are exposed to the parasite early in the marine migratory phase of

their life cycle close to shores of the United Kingdom and Ireland.

Individual fish may have heavy burdens of anisakid larvae. In

general, this high abundance is attributable to the longevity of the

host since fish accumulate more parasites with age (Margolis,

1970). The severity of parasitism reported here in salmon grilse,

which have spent only 12–15 mo at sea, may be unusual, but as

the present study has shown, they are exposed to infection early in

their marine migration.

On account of the dearth of information on the marine

migration of salmonids, the reasons for the increased abundance

of A. simplex s.s. in adult salmon returning to their natal rivers

can only be surmised. It is possible that changing oceanic

conditions may have increased the abundance of crustacean

intermediate hosts in the feeding areas off Greenland and in the

Nordic Seas. Recently, changes associated with increasing sea

surface temperatures have been detected in the pelagic ecosystems

of the northeast Atlantic. There has been a geographic shift

northward, of at least 10u in latitude, of warm water calanoid

copepods and a decrease in the colder water species (Beaugrand

and Reid, 2003). Another likelihood is that changing sea

temperatures and currents may have affected the behavior of

the mammalian definitive hosts resulting in an increased

prevalence of infection among the zooplankton and fish species

on which post-smolts and adults prey.

The exposure of post-smolts to infection so early in their marine

migration indicates that these fish feed on marine intermediate/

paratenic hosts soon after the transition from freshwater to salt

water. Fish larvae appear to be important prey early in the marine

phase. A previous study has shown that the stomachs of post-

smolts, caught at 619N off the northwest coast of Scotland,

FIGURE 3. Macrophage activated centers (arrows) present amongsthepatocytes adjacent to the fibro-connective tissue capsules surroundinglarvae of Anisakis simplex s.s. in the liver of adult Salmo salar stained withH&E; scale bar 5 200 mm.

FIGURE 4. Anisakis simplex s.s. larvae encapsulated in the musculariscircularis of the intestinal wall of an adult Salmo salar. Extensivedegeneration of the stratum compactum (between the arrowheads) witheosinophilic granular and mononuclear cell infiltration, proximal end ofan intestinal villous (arrow), stained with H&E; scale bar 5 500 mm.

80 THE JOURNAL OF PARASITOLOGY, VOL. 96, NO. 1, FEBRUARY 2010

contained mainly fish, such as sandeel (Ammodytes spp.) and blue

whiting (Micromesistius poutassou) and that crustaceans represent-

ed only 8% of the diet of these fish (Haugland et al., 2006).

The response of vertebrates to chronic inflammation induced

by foreign bodies such as parasitic nematodes is to isolate them in

a granulomatous reaction and to sequester them in a fibrous

capsule (McGavin and Zachary, 2007). Encapsulation of anisakid

larvae attached to the abdominal mesentery and serosal surface of

the intestine and other visceral organs, accompanied with a

variable amount of lymphocytic and macrophage infiltration, has

been reported to occur in herring and plaice (Hauck and May,

1977; Dezfuli et al., 2007). In the adult salmon studied here, the

cellular infiltration associated with the fibrosis surrounding larvae

attached to mesentery and serosal surfaces was limited to

scattered individual EGCs and mononuclear cells with focal

melano-macrophage centers. There was no obvious hepatocyte

pathology or cellular infiltration associated with the encysted

larvae in the liver parenchyma. Similarly, there was no evidence of

hemorrhagic tracts or necrosis indicative of larval migration

within the liver. However, the presence of discrete focal melano-

macrophages among the liver cells adjacent to the fibrous capsule

belies the hypothesis that macrophage aggregates proliferate only

in areas of severe tissue damage caused by chronic inflammation

(Agius and Roberts, 2003). The ill effects of mechanical

compression and displacement of cells in the substance, and on

the border, of the liver on metabolic activity may be limited in

these grilse, since sexually maturing salmonids do not feed.

In contrast to the liver and mesentery, there was a more intense

response to encapsulated larvae in the pyloric cecum. The lack of

resident EGCs in the connective tissue of liver and mesentery may

be an explanation for the limited inflammatory reaction in these

organs. The recruitment of EGCs, which are present in variable

quantities in the lamina propria and submucosa, is a common

response in the intestine of salmonids to parasitic infections

(Bullock, 1963). Eosinophilic granular cells are generally consid-

ered to be analogous to mammalian mast cells (Reite and Evensen,

2006). During the process of degranulation, they release enzymes,

leucocyte chemotractants, and other factors, which influence

vascular permeability and the mobilization of inflammatory cells.

The histological changes in the cecum reported here were similar

to those described by Hauck and May (1977), Dezfuli et al. (2007),

and Beck et al. (2008) for herring, plaice, and the vent region of adult

Atlantic salmon, respectively. The vent was not included in this

study, since the presence of larvae in this area was considered a

normal manifestation of nematodes seeking a suitable site to encyst

and the swollen red vents an external sign of heavily parasitized fish.

Hauck and May (1977) suggested that the pyloric cecum was the

optimum route for larval penetration into the peritoneal cavity. In

fish with high burdens of Anisakis sp. larvae, the inflammatory

reaction induced by earlier migrating parasites may make the pyloric

cecum an inhospitable environment for further migrations. Thus,

worms ingested later may have to travel farther down the intestinal

tract to undergo their migration into the abdominal cavity.

There is very little information on the role disease plays during

the marine phase of the life cycle of Atlantic salmon (Bakke and

Harris, 1998). The effect of this high level of parasitism on sea

survival and natural spawning on an already depleted salmon stock

needs to be investigated. In the ranched broodstock studied here,

the presence of Anisakis sp. had no effect on ova/milt production

and fertility (D. Cotter, pers. comm.). However, sexually mature

salmon do not feed, and it is likely that the histologic changes

observed in the intestines and to a lesser extent the liver during this

study would have an adverse affect on digestion, absorption, and

metabolic activity in younger feeding fish. Continued surveillance

of post-smolts is required to identify the geographic locations and

dietary source of infection and elucidate the epidemiology and

pathogenesis of Anisakis sp. in salmon during their marine phase.

Finally, investigations on the genetic structure and the potential of

anisakid larvae as biological markers could be enhanced with

further sequencing of rDNA and mtDNA.

ACKNOWLEDGMENTS

The post-smolts were collected during research cruises supported by theEU Framework 7 SALSEA Merge programme (www.salmonatsea.com).

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