Parasite infections of rainbow trout (Oncorhynchus mykiss) from Danish mariculture

Aquaculture 434 (2014) 486–492

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Parasite infections of rainbow trout (Oncorhynchus mykiss) from Danish mariculture Jakob Skov ⁎, Foojan Mehrdana, Moonika Haahr Marana, Qusay Zuhair Mohammad Bahlool, Rzgar Mohammad Jaafar, Diana Sindberg, Hannah Malene Jensen, Per Walter Kania, Kurt Buchmann Department of Veterinary Disease Biology, Faculty of Health and Medical Sciences, University of Copenhagen, Stigbøjlen 7, DK-1870 Frederiksberg C, Denmark

a r t i c l e

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Article history: Received 16 July 2014 Received in revised form 26 August 2014 Accepted 27 August 2014 Available online 3 September 2014 Keywords: Oncorhynchus mykiss Mariculture Runts Hysterothylacium aduncum Salmon lice Stomach content

a b s t r a c t Rainbow trout (Oncorhynchus mykiss) runts (n = 5) and harvest quality fish (n = 5) were sampled from each of all the net cage mariculture facilities in Denmark at the time of slaughter during autumns of 2012 and 2013. Thus, a total of 190 trout were obtained, represented by 95 runts and 95 fish of harvest quality. Trout were examined for macroscopic ectoparasites as well as helminths of the gastrointestinal tract and body cavity by careful visual inspection, and belly flap musculature by pepsin digestion. Stomach content analysis was performed in order to assess the risk of endoparasite transmission to the net cage cultured trout. Low numbers of salmon lice (Lepeophtheirus salmonis) (1–9 lice per fish) were found on 9 trout from western localities off the eastern coast of Jutland characterized by water salinity levels of 21–24‰, whereas no lice were detected on fish from areas of lower salinity. Body cavity and musculature of all trout were free from helminths, and the absence of medically important 3rd stage larvae of Anisakidae was thus confirmed. However, transmission of endoparasites was documented by the finding of the nematode Hysterothylacium aduncum in the intestines of 9.5% of the runts (mean intensity = 2.3) and 2.1% of the harvest quality trout (mean intensity = 2.0). A few cestodes (Eubothrium sp.) located in the pyloric caeca, and one acantocephalan (Neoechinorhynchus sp.) found in the intestine, were collected from four trout. The higher prevalence of H. aduncum among runts was associated with a markedly increased intake of parasite intermediate or paratenic hosts, i.e. small fish (mainly three-spined stickleback) and crustaceans (mainly amphipods), by these fish compared to those of harvest quality. Additionally, a higher number of runts than harvest quality fish had eaten biofouling organisms, e.g. mussels and algae, further indicating a difference in feeding behavior between the two quality classes of trout. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Parasite infections may negatively affect fish welfare, cause mortality and lead to significant economic losses in aquaculture production (Barber, 2007; Costello, 2006; Heuch et al., 2005). Additionally, the presence of zoonotic parasites, e.g. nematode larvae belonging to the family Anisakidae, will compromise the food safety status of the fish product (EFSA Panel on Biological Hazards (BIOHAZ), 2010) and may thus negatively affect the market value. Depending on parasite species and the type of life cycle they follow, infection in modern aquaculture systems may be virtually absent, e.g. Anisakis sp. infection in salmonids (Angot and Brasseur, 1993; Deardorff and Kent, 1989; Inoue et al., 2000; Lunestad, 2003; Marty, 2008; Skov et al., 2009), or occur at high levels normally not observed in wild fish populations, e.g. infection with the salmon louse Lepeophtheirus salmonis (Wootten et al., 1982). The confined life of maricultured fish in net cages and the use of heat-treated ⁎ Corresponding author. Tel.: +45 35336734; fax: +45 35332755. E-mail address: [email protected] (J. Skov).

http://dx.doi.org/10.1016/j.aquaculture.2014.08.041 0044-8486/© 2014 Elsevier B.V. All rights reserved.

feed without any viable parasite larvae theoretically exclude them from participating in the complex life cycles of a range of parasites, including anisakids (Skov et al., 2009). However, potential intermediate hosts, e.g. small crustaceans and fish, of parasites of both veterinary and medical importance may enter net cages and suffer predation as documented by stomach content analysis of rainbow trout (González, 1998) and Atlantic salmon (Sepúlveda et al., 2004) from Chilean mariculture and rainbow trout from Denmark (Skov et al., 2009). In line with this, transmission of anisakids to maricultured salmonids have been reported by the findings of Hysterothylacium aduncum (Anisakidae) in the gastrointestinal tract of rainbow trout, coho and Atlantic salmon in Chile (Carvajal et al., 1995; González, 1998; Sepúlveda et al., 2004) and a single specimen of Anisakis sp. detected in Atlantic salmon in Canada (Marty, 2008). González (1998) found that the abundance of H. aduncum in maricultured rainbow trout was higher in trout below 500 g compared to larger fish, and both H. aduncum abundance and the intensity of infection were higher in rejected, undersized rainbow trout compared to harvested fish. A recent study by Mo et al. (2014) further documents actual transmission of

J. Skov et al. / Aquaculture 434 (2014) 486–492

anisakid nematode larvae to cultured salmonids by detection of H. aduncum and A. simplex in Atlantic salmon runts (i.e. undersized specimens showing inferior growth), whereas harvest quality fish were free from infection. The aim of the present study was to examine rainbow trout from all marine net cage cultures in Danish waters for macroscopic ectoparasites as well as helminth endoparasites of the gastrointestinal tract, body cavity and musculature. Special attention was paid to evaluate the infection status and risk of transmission of anisakid 3rd stage larvae to runts and harvest quality fish. 2. Materials and methods 2.1. Fish A total of 190 rainbow trout (Oncorhynchus mykiss, Walbaum 1792), representing 19 different mariculture facilities (localities 1–19) in Denmark (Fig. 1), were obtained during slaughter in November– December 2012 and October–December 2013. Fish were all females except for fish (n = 10) from locality 4, which were a mixture of males and females. All fish had been raised in net cages except for fish (n = 10) from locality 11, which had been raised in land based concrete basins by the sea/harbor side. As far as possible, 5 runts and 5 harvest quality fish were sampled from each facility. Fish were stored on ice and examined for parasite infections within 5 days from the time of slaughter. However, the majority of the fish (n = 152) was examined within 1–2 days. Individual fish size was measured as the standard length from the snout to the basis of the caudal fin (Lstandard) and the total weight of the ungutted fish (Mtotal) (Table 1A). The term ‘runt’ generally characterizes fish showing inferior or no growth. In this case, runts were small and lean with pale or poorly colored flesh and no or limited abdominal fat deposits. However, the definition of a runt vs. a harvest quality fish appeared to be more or less arbitrary since the total number of sampled rainbow trout represented a wide and continuous range of size classes (Fig. 2). Therefore, in order to investigate potential differences in food intake and parasite infections among runts or small rainbow trout vs. harvest quality rainbow trout, the fish were divided into two size classes, i.e. A, the smallest rainbow trout (n = 95) [mean Mtotal (range) = 0.813 (0.380–1.444) kg], and B, the largest rainbow trout (n = 95) [mean Mtotal (range) = 2.390 (1.475–3.782) kg]. Apart from runts showing poor growth, as well as slight damages to the tail fin frequently observed among all size classes, the general health

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status of the fish appeared sound with no external signs of disease or disorders. 2.2. Parasitological examination The skin was macroscopically examined for ectoparasites and any abnormalities or signs of pathological changes. Presence of helminths in the body cavity and gastrointestinal tract was investigated by scrutinizing the peritoneum and surfaces of all visceral organs using a magnifying lens (1.9× magnification). The gastrointestinal tract was opened from esophagus to anus and scrutinized in the same manner. The presence of nematode larvae in the musculature was investigated by the following procedure: the muscle tissue of both belly flaps from individual fish was gently processed into smaller pieces and exposed to a pepsin solution (1 L tap water, 6 mL concentrated HCl, 9 g NaCl, 10 g pepsin powder (2000 FIP units/g, Orthana, Denmark)) by adding 10 mL pepsin solution per 1 g fish tissue. Samples were incubated at 37 °C and continuous magnetic stirring (300 rpm) until complete digestion was achieved within approx. 6 h. Digested samples were run through a 300 μm sieve and material caught in the sieve was examined for nematode larvae. A pilot study showed that 3rd stage larvae of A. simplex (n = 10) (presenting an ITS sequence identical to A. simplex, GenBank accession no. EU624342) collected from the body cavity of Atlantic herring (Clupea harengus) were intact and alive subsequent to at least 10 h of digestion according to the procedure described above (data not shown). All recovered parasites were preserved in 96% ethanol if not otherwise stated. 2.3. Stomach content The stomach content of all fish was recorded in order to detect any ingestion of nematode paratenic or intermediate hosts, e.g. small fishes and crustaceans, entering the net cages. 2.4. Morphological analysis of parasites Prior to preservation, and when still alive, collected nematodes were morphologically examined for genus-specific characters such as the presence or absence of ventricular appendix and/or intestinal caecum, and location of nerve ring and excretory pore (Möller and Anders,

Fig. 1. A, map of Denmark (gray) including a box framing the study area enlarged in B; B, rainbow trout mariculture facilities (localities 1–19) investigated in the present study.

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Table 1 A. Locality, salinity data, date of sampling and size measures of O. mykiss.

B. No. of O. mykiss with stomach content presenting teleost fish (Fish), crustaceans (Crus) or molluscs (Moll).

C. No. of O. mykiss hosting parasite findings belonging to Cestoda (Ce), Nematoda (Ne), Acantocephala (Ac) or Arthropoda (Ar). Mean intensity of infection is presented in parentheses.

Locality ID

Salinitya range (‰)

Date of sampling

n

Lstandard, mean (range) (cm)

Mtotal, mean (range) (kg)

Fishb

Crusb

Mollb

Cec

Ned

Ace

Arf

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

11–13 11–13 11–13 10–12 8–10 12–14 11–13 11–13 8–10 22–24 12–14 21–23 22–24 21–23 20–22 16–18 12–14 18–20 18–20

29.11.2012 29.11.2012 29.11.2012 04.12.2012 11.12.2012 11.12.2012 11.12.2012 11.12.2012 18.12.2012 19.12.2012 16.10.2013 30.10.2013 04.11.2013 04.11.2013 15.11.2013 19.11.2013 11.12.2013 11.12.2013 12.12.2013

10 10 10 10 10 10 10 10 10 10 10 10 10 10 10 10 10 10 10

43.0 (32.5–54.5) 46.3 (36.5–55.0) 45.3 (37.0–54.5) 39.2 (31.5–44.0) 45.6 (39.0–58.5) 47.0 (36.0–54.0) 43.3 (37.0–55.0) 45.9 (37.0–55.5) 43.6 (34.5–52.0) 43.7 (32.5–53.5) 42.4 (35.0–51.5) 42.4 (41.0–49.0) 45.5 (37.0–55.5) 43.4 (37.5–50.5) 44.0 (33.0–52.5) 43.5 (35.0–51.0) 46.0 (34.0–56.5) 40.9 (30.5–53.0) 41.6 (32.5–51.5)

1.650 (0.391–3.400) 1.631 (0.600–3.576) 1.655 (0.532–3.300) 1.107 (0.738–1.665) 1.721 (0.584–3.176) 2.000 (0.794–3.782) 1.480 (0.619–3.209) 1.663 (0.769–3.020) 1.428 (0.606–2.622) 1.450 (0.383–2.884) 1.741 (0.895–3.085) 1.497 (0.861–2.327) 1.748 (0.515–3.214) 1.509 (0.617–2.710) 1.640 (0.459–2.978) 1.725 (0.698–2.802) 1.992 (1.444–3.215) 1.430 (0.380–3.028) 1.359 (0.382–2.585)

– – – – 1 1 3 1 – – – – – – – – – – –

1 – – – 1 2 2 – – – 4 1 1 – – – – – –

2 2 – – 2 2 3 1 – – 1 5 4 3 – – 2 – 1

– 1 (1.0) – – – – – 2 (3.0) – – – – – – – – – – –

1 (2.0) 2 (1.0) 1 (3.0) 3 (1.3) 2 (2.5) – 1 (3.0) 1 (6.0) – – – – – – – – – – –

– – – – – – – – – – – – – – – – – – 1 (1.0)

– – – – – – – – – 2 (1.5) – 5 (2.8) 2 (1.0) – – – – – –

a Presented as the range of mean salinities of surface water in the area from April to September 2006 (DHI, 2007), representing the time of year where water temperatures are expected to allow increased transmission (Tully and Nolan, 2002), prevalence and intensity of L. salmonis infection (Schram et al., 1998). Lstandard: Length from snout to basis of caudal fin. Mtotal: Mass of the total, ungutted fish. b See footnotes of Table 3 for more information on stomach content findings. c Eubothrium sp. d H. aduncum in all cases. e Neoechinorhynchus sp. f L. salmonis in all cases.

1986). Subsequent to preservation, anterior and posterior ends of nematodes were mounted in glycerol gel on microscope slides for further examination by light microscopy of additional descriptive morphological characters such as tail tip morphology (Køie, 1993; Moravec and Nagasawa, 2000). The middle parts of the nematode specimens were subjected to molecular analysis (see Section 2.5.1). Other recovered parasites than nematodes were subjected to morphological examination only and identified according to the existing literature. 2.5. Molecular analysis of parasites

and elongation at 72 °C (2 min), and final post-elongation at 72 °C (5 min). PCR products were analyzed by electrophoresis in ethidium bromide stained 1.5% agarose gels, purified using the illustra™ GFX™ PCR DNA and Gel Band Purification Kit (Cat. No. 28-9034-70, GE Healthcare), and sequenced by Macrogen Inc. (Korea).

2.6. Terms used The terms ‘prevalence’, ‘intensity of infection’, ‘mean intensity’, and ‘abundance’ are used in accordance with Bush et al. (1997).

Only nematode findings were subjected to molecular analysis. 2.5.1. DNA isolation The middle part of nematodes was used for DNA isolation. Hence, the middle part of each worm was lysed in clear 1.5 mL Eppendorf tubes containing 100 μL lysis buffer (Tween 20 (0.45%), Proteinase K (60 μL mL−1) (Qiagen), 10 mM Tris and 1 mM EDTA in DNase/RNasefree H2O) at 55 °C and shaking at 800 rpm. Complete lysis of parasite material was confirmed by light microscopy. Subsequent to lysis, the proteinase was inactivated at 95 °C (10 min). 2.5.2. Polymerase chain reaction (PCR) and sequencing PCR was performed in a T3 Thermocycler (Biometra) in reaction volumes of 60 μL containing 5 μL nematode lysate as template, 1 unit of BioTaq DNA polymerase, 1 mM dNTP, 1.5 mM MgCl2, 10× NH4 reaction buffer (DNA-Technology), 1 μM forward primer (NC5), 1 μM reverse primer (NC2), and DNase/RNase free H2O. Forward primer (NC5) was 5′-GTA GGT GAA CCT GCG GAA GGA TCA TT-3′ and reverse primer (NC2) was 5′-TTA GTT TCT TTT CCT CCG CT-3′ (Zhu et al., 2007). The PCR procedure comprised pre-denaturation at 95 °C (2 min), followed by 45 cycles of denaturation at 95 °C (30 s), annealing at 53 °C (30 s)

Fig. 2. Distribution of the total number of sampled rainbow trout (n = 190) according to size measured as the total weight of the ungutted fish (Mtotal).

J. Skov et al. / Aquaculture 434 (2014) 486–492

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Table 2 Parasite findings from cultured O. mykiss (n = 190) separated in two size classes representing the smallest rainbow trout (n = 95) (A), and the largest rainbow trout (n = 95) (B). Size classes of O. mykiss, mean (range) (kg)

n

A: 0.813 (0.380–1.444) B: 2.390 (1.475–3.782)

Parasite findings, parasite identity (phylum) (prevalence) (mean intensity), on/in Skin

Pyloric caeca

Intestine

95

L. salmonis (Arthropoda) (2.1%) (1.5)

Eubothrium sp. (Cestoda) (3.2%) (2.3)

95

L. salmonis (Arthropoda) (7.4%) (2.3)

–

H. aduncum (Nematoda) (9.5%) (2.3) Neoechinorhynchus sp. (Acantocephala) (1.1%) (1.0) H. aduncum (Nematoda) (2.1%) (2.0)

3. Results 3.1. Parasitological findings The parasite findings described below are presented in relation to locality, water salinity level, date of sampling and stomach content in Table 1C, and in relation to size class in Table 2. No pathological changes were observed in association with any of the parasite infections detected. 3.1.1. Arthropoda A total of 19 salmon lice (L. salmonis) (1–9 lice per fish) were found on nine rainbow trout from localities 10, 12 and 13. L. salmonis was diagnosed based on morphology in accordance with Kabata (1979). 3.1.2. Nematoda 3.1.2.1. Nematode larvae in body cavity and musculature. All rainbow trout (n = 190) were negative for the presence of nematode larvae in the body cavity and belly flap musculature. 3.1.2.2. Nematodes in the gastrointestinal tract. A total of 25 nematodes (1–6 nematodes per fish) were collected from the intestine of 11 rainbow trout from localities 1–5 and 7–8. Morphological examination by light microscopy of all specimens in live condition prior to preservation confirmed the presence of ventricular appendix, intestinal caecum, and an excretory pore positioned in agreement with Hysterothylacium spp. (Möller and Anders, 1986). Further, mounts of nematode posterior ends on microscope slides showed that 23 of the 25 specimens presented a caudal end with the tip of the tail covered by minute spines. The remaining two specimens did not exhibit tail structures. The spiny caudal extremity, also termed “cactus tail” by some authors, was in accordance with descriptions of the 4th stage larva (Køie, 1993) and adult (Moravec and Nagasawa, 2000; Yagi et al., 1996) of H. aduncum and other Hysterothylacium spp. (Moravec et al., 1997; Shamsi et al., 2013), and distinguishes the present findings from 3rd stage larvae of Anisakis (Cannon, 1977; Hurst, 1984), Pseudoterranova (Hurst, 1984), and Contracaecum (Cannon, 1977; Martins et al., 2005). Molecular analysis of lysates from the 25 nematodes resulted in 11 equally sized (1030 bp) PCR products, which all presented a DNA

sequence identical to H. aduncum (GenBank accession no. JX845137). Lysates from the remaining 14 nematodes were PCR negative. Based on the results of the morphological and molecular analysis described above, all 25 nematodes were considered to belong to H. aduncum. 3.1.3. Cestoda A total of seven adult cestodes (1–4 worms per fish) were collected from the pyloric caeca of three rainbow trout from localities 2 and 8. Specimens were identified as Eubothrium sp. based on scolex and apical disc morphology (Andersen and Kennedy, 1983; Chubb et al., 1987; Scholz et al., 2003). 3.1.4. Acantocephala One acantocephalan was found in the intestine of one rainbow trout (locality 19), hydrated in de-ionized water, fixed in hot 10% neutral buffered formalin and identified as Neoechinorhynchus sp. according to Bykhovskaya-Pavlovskaya et al. (1984) and Smales (2013). 3.2. Stomach content One hundred and fourteen rainbow trout had empty stomachs, whereas the remaining 86 rainbow trout had eaten either small fishes, crustaceans, mussels, eelgrass, algae, remnants (i.e. scales, bones and eggs) of conspecifics (O. mykiss), or non-digestible objects, or various combinations thereof. Trout that had eaten one or more of the above mentioned food items or objects were dominated by 57 trout from the smaller size class A as opposed to 19 trout from size class B. Further, for each food item category, there was a dominance of smaller trout from size class A (Table 3). Pelleted feed was found in the stomachs of a total of 11 trout at 4 different localities. 4. Discussion The marine and ectoparasitic salmon louse (L. salmonis) was detected at 3 localities with water salinity of 21–24‰ presenting mean intensities of infection between 1.0 and 2.8, whereas no lice were found on fish from areas of lower salinity. The optimal salinity level for L. salmonis appears to be around 30‰ (Bricknell et al., 2006; Johnson and Albright, 1991; Tucker et al., 2000). Thus, experimental studies

Table 3 Stomach content of cultured O. mykiss (n = 190) separated in two size classes representing the smallest rainbow trout (n = 95) (A), and the largest rainbow trout (n = 95) (B). Results are shown as numbers and proportion (%) of rainbow trout. Size classes of O. mykiss, mean (range) (kg)

n

NF

Pellets

Fisha

Crustaceansb

Molluscsc

Eelgrass

Algaed

Remnants of conspecifics (O. mykiss)f

Non-digestible objectse

A: 0.813 (0.380–1.444) B: 2.390 (1.475–3.782)

95 95

38 (40.0%) 76 (80.0%)

7 (7.4%) 4 (4.2%)

5 (5.3%) 1 (1.1%)

8 (8.4%) 5 (5.3%)

22 (23.6%) 7 (7.4%)

22 (23.6%) 0 (–)

27 (28.4%) 2 (2.1%)

4 (4.2%) 0 (–)

8 (8.4%) 3 (3.2%)

NF: Nothing found. a A total of 55 small fish, of which 54 were identified as three-spined stickleback (G. aculeatus) according to Muus et al. (1997), were eaten by 5 rainbow trout in size class A (n = 95), and 1 small fish were eaten by 1 rainbow trout in size class B. b Crustaceans were generally represented by approx. equal occurrences of barnacles and amphipods in size class A, and dominated by amphipods in size class B (Køie et al., 2000). c Molluscs were in all cases identified as mussels of the genus Mytilus (Brooks and Farmen, 2013; Christensen et al., 1978; Kijewski et al., 2011). d Algae were green and brownish multicellular algae (seaweed). e Inedible objects included stones, a bird feather, a leaf from a tree, and pieces of wood, plastic and rope. f Remnants of conspecifics included trout eggs and large fish scales and bones assumed to originate from O. mykiss.

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performed at approx. 12 °C have shown infection levels to be reduced at 24–26‰ compared to 34‰ (Bricknell et al., 2006; Tucker et al., 2000), and the survival of free-swimming copepodids and the attachment of chalimus to the host to be gradually compromised at decreasing salinities below 24‰ and 34‰, respectively (Bricknell et al., 2006). However, copepodids may distribute over a wider range of salinities (15.4–30.4‰) when provided the opportunity (Heuch, 1995), and motile stages on the host may tolerate decreased salinities between 7‰ and 28‰ for up to 7 days (Connors et al., 2008). Egg development fails at 10‰ at 10–12 °C (Johnson and Albright, 1991; Wootten et al., 1982), and the lower salinity level for development of L. salmonis has been reported to be 16‰ (Berger, 1970). Hence, regarding the present study, salinity levels of 8–14‰ characterizing the areas off the western and southern coast of Zealand probably inhibit the development of L. salmonis, whereas the life cycle may complete at suboptimal conditions along the eastern coast of Jutland as indicated by the L. salmonisinfested localities 10 and 12–13 (21–24‰). The mean intensities of infection found were generally below those of O. mykiss cultured in Japan (9.7) (Ho and Nagasawa, 2001) and wild-caught in the North Pacific and Bering Sea (3.9–8.6) (Nagasawa, 2001), sea trout (Salmo trutta) from south-eastern Norway (7.6) (Mo and Heuch, 1998), and Atlantic salmon (Salmo salar) from Ireland (11.6) (Jackson et al., 2013). However, it should be noticed that lowered salmon louse counts leading to underestimation of prevalence and intensities of infections may occur due to dislodgement of salmon lice depending on the mode of capture and killing (e.g. cervical dislocation vs. overdose by an anesthetic) (Nagasawa, 1985; Treasurer and Pope, 2000) or post mortem loss of parasites (Tingley et al., 1997). Thus, since the trout in the present study were sampled subsequent to standard slaughter procedures involving CO2 anesthesia and examined for parasites after 1–5 days storage on ice, the salmon louse prevalence and infection intensities recorded may be underestimates. Therefore further studies on salmon louse populations on rainbow trout maricultured in Danish waters are needed. Third stage larvae of anisakid nematodes were absent from body cavity and musculature of all rainbow trout cultured in marine net cages in Danish waters. Nevertheless, transmission of marine endoparasites was documented by findings of H. aduncum in the intestines of 9.5% of the smaller trout (size class A, representing the runts) and 2.1% of the larger trout (size class B, representing the harvest quality fish). This was associated with an increased intake of potential intermediate or paratenic hosts of endoparasites, i.e. small fish and crustaceans, among trout of size class A compared to trout of size class B. Further, H. aduncum was only detected in trout cultured off the coast of the islands Lolland, Falster and the southern part of Zealand and these findings were associated with trout stomach contents including crustaceans (e.g. amphipods) and small fish in particular (mainly sticklebacks (Gasterosteus aculeatus)). The marine life cycle of H. aduncum is complex and may involve two or more hosts including predatory fishes as final hosts harboring the 4th stage larva and the adult nematode in the intestine. A broad range of invertebrates as well as fishes act as intermediate and/or paratenic hosts harboring the 3rd stage larva, which has hatched from the egg voided with feces from the final host (González, 1998; Køie, 1993). Natural as well as successful experimental infections of various amphipods (Køie, 1993; Marcogliese, 1996) and the stickleback (G. aculeatus) with H. aduncum 3rd stage larvae (Køie, 1993) have been documented. Thus, it is likely that the transmission of H. aduncum to the cultured rainbow trout found in the present study was due to their intake of amphipods and sticklebacks. Transmission of H. aduncum to maricultured salmonids has previously been detected in rainbow trout and coho salmon in Chile (González, 1998) and in Atlantic salmon in Chile (Sepúlveda et al., 2004) and Norway (Mo et al., 2014). Hysterothylacium spp. are generally considered to constitute a low or hypothetical risk of infection in man or even without zoonotic potential (Deardorff and Overstreet, 1981; Möller and Anders, 1986; Sakanari and

McKerrow, 1989). However, a rare case of disease associated with the passage of an adult H. aduncum through the gastrointestinal system of a human patient has been reported from Japan (Yagi et al., 1996). Migration of H. aduncum 3rd stage larvae into the edible muscle of the fish host may occur but may be fish host species dependent (Adroher et al., 1996; Durán et al., 1989). Natural infections of farmed Atlantic salmon (Mo et al., 2014) and experimental infection of brown trout, rainbow trout and Atlantic salmon (Haarder et al., 2013) with H. aduncum showed absence of larval migration into the fish fillets. Based on the available data on the life cycle and the migration in salmonid fish hosts of H. aduncum, we suggest that the risk of human anisakidosis caused by H. aduncum from maricultured salmonid fish products is negligible. The smaller trout from size class A were characterized by no or limited abdominal fat deposits and pale or poorly colored flesh indicating a limited intake of pelleted feed during the production cycle. Overall, pelleted feed was only detected in the stomachs of a few trout (n = 11), which is in agreement with the common practice of short-term fasting of cultured fish prior to slaughter (FAWC, 1996; López-Luna et al., 2013). Hence, the majority of the trout did probably not have access to pelleted feed prior to slaughter and sampling for the present investigation. Analysis of stomach contents showed a preference among the smaller trout for predation of small fish, crustaceans and mussels as well as ingestion of algae, and objects probably drifting into the net cages (e.g. eel grass, pieces of wood and plastic), and consumption of remnants (i.e. scales, bones and eggs) of presumably dead conspecifics. This foraging behavior indicates a spatial distribution of the smaller trout along the periphery and bottom of the net cage putting them at risk of acquiring endoparasite infections. Accordingly, endoparasite infections were more often detected among the smaller trout compared to the larger trout of the present study. This is in line with a higher abundance of H. aduncum found among small and discarded rainbow trout compared to larger and harvested fish from Chilean mariculture (González, 1998) and to the restriction of A. simplex and H. aduncum to runts of farmed Atlantic salmon in Norway (Mo et al., 2014). Altogether, these and the present data suggest that runts may serve as an indicator of transmission of anisakids to fish cultured in marine net cages. Further and large-scale studies are needed in order to identify critical levels of prevalence and intensity of infection among runts, which may indicate if nematode transmission to harvest quality fish occurs within a given net cage system. Similarly, such investigations might determine whether the absence of nematode larvae in runts would be a reliable indicator of a nematode-free status of harvest quality fish. A few trout were found to be infected with a cestode (Eubothrium sp.) and in a single case an acantocephalan (Neoechinorhynchus sp.). Eubothrium sp. has previously been found in freshwater cultured rainbow trout and maricultured Atlantic salmon in Denmark and Norway, respectively (Bristow and Berland, 1991; Buchmann and Bresciani, 1997), and may even at low infection levels be associated with reduced growth of the fish host (Bristow and Berland, 1991; Saksvik et al., 2001). Since infection with Eubothrium sp. may occur in both freshwater and marine environments, it could not be settled whether the trout had acquired the infection before or after stocking in the marine net cages (Andersen and Kennedy, 1983; Buchmann, 1987; Buchmann and Bresciani, 1997). This does also apply to the finding that pertains to Neoechinorhynchus sp. (Smales, 2013). Species of Eubothrium and Neoechinorhynchus matures to adult parasites in fish, and fish and turtles, respectively (Scholz et al., 2003; Smales, 2013), and are not assigned any zoonotic potential. In conclusion, the present investigation documented the presence of salmon lice (L. salmonis) on rainbow trout cultured at some locations off the east coast of Jutland. In comparison with trout of harvest quality, stomach contents of runts showed a markedly increased ingestion of biofouling organisms, e.g. mussels and algae, as well as potential intermediate or paratenic hosts of endoparasites, i.e. small fish and

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crustaceans. This difference in feeding behavior indicated that runts were exposed to an increased risk of acquiring endoparasite infections. Accordingly, increased prevalence and intensity of H. aduncum infection were found among runts compared to harvest quality fish. Parasites found in the present study were not considered to constitute a risk of zoonosis, and medically important 3rd stage larvae of Anisakidae were absent from body cavity and flesh of both runts and harvest quality rainbow trout. Acknowledgments This project was supported by the Ministry of Agriculture, Food and Fisheries of Denmark and the EU Commission through an EFF grant 33010-12-a-0228. Ulrik Bo Pedersen (University of Copenhagen) created the background map for Fig. 1, which is highly appreciated. All Danish rainbow trout mariculture companies are sincerely thanked for their participation and kind collaboration. References Adroher, F.J., Valero, A., Ruiz-Valero, J., Iglesias, L., 1996. Larval anisakids (Nematoda: Ascaridoidea) in horse mackerel (Trachurus trachurus) from the fish market in Granada (Spain). Parasitol. Res. 82, 319–322. Andersen, K.I., Kennedy, C.R., 1983. Systematics of the genus Eubothrium Nybelin (Cestoda, Pseudophyllidea), with partial re-description of the species. Zool. Scr. 12, 95–105. Angot, V., Brasseur, P., 1993. European farmed Atlantic salmon (Salmo salar L.) are safe from anisakid larvae. Aquaculture 118, 339–344. Barber, I., 2007. Parasites, behaviour and welfare in fish. Appl. Anim. Behav. Sci. 104, 251–264. Berger, V.J., 1970. The effect of marine water of different salinity on Lepeophtheirus salmonis, ectoparasite of salmon. Parazitologiya 4, 136–138 (in Russian). Bricknell, I.R., Dalesman, S.J., O'Shea, B., Pert, C.C., Luntz, A.J.M., 2006. Effect of environmental salinity on sea lice Lepeophtheirus salmonis settlement success. Dis. Aquat. Org. 71, 201–212. Bristow, G.A., Berland, B., 1991. The effect of long term, low level Eubothrium sp. (Cestoda: Pseudophyllidea) infection on growth in farmed salmon (Salmo salar L.). Aquaculture 98, 325–330. Brooks, S.J., Farmen, E., 2013. The distribution of the mussel Mytilus species along the Norwegian coast. J. Shellfish Res. 32, 265–270. Buchmann, K., 1987. Cestodes of migratory trout (Salmo trutta L.) from the Baltic Sea. Bull. Eur. Assoc. Fish Pathol. 7, 115–117. Buchmann, K., Bresciani, J., 1997. Parasitic infections in pond-reared rainbow trout Oncorhynchus mykiss in Denmark. Dis. Aquat. Org. 28, 125–138. Bush, A.O., Lafferty, K.D., Lotz, J.M., Shostak, A.W., 1997. Parasitology meets ecology on its own terms: Margolis et al. revisited. J. Parasitol. 84, 575–583. Bykhovskaya-Pavlovskaya, I.E., Gusev, A.V., Dubinina, M.N., Izyumova, N.A., Smirnova, T.S., Sokolovskaya, I.L., Shtein, G.A., Shul'man, S.S., Epshtein, V.M., Nagibina, L.F., Raikova, E.V., Strelkov, Y.A., Bykhovskii, B.E., 1984. Key to parasites of freshwater fish of the U.S.S.R. Academy of Sciences of the U.S.S.R., Moskva-Leningrad. Cannon, L.R.G., 1977. Some larval ascaridoids from south-eastern Queensland marine fishes. Int. J. Parasitol. 7, 233–243. Carvajal, J., Gonzáles, L., Toledo, G., 1995. New record of Hysterothylacium aduncum (Rudolph, 1802) (Nematoda: Anisakidae) in salmonids cultured in sea farms from southern Chile. Res. Rev. Parasitol. 55, 195–197. Christensen, J.M., Larsen, S., Nyström, B.O., 1978. Havmuslinger. Gyldendalske Boghandel, Nordisk Forlag A/S, København (in Danish). Chubb, J.C., Pool, D.W., Veltkamp, C.J., 1987. A key to the species of cestodes (tapeworms) parasitic in British and Irish freshwater fishes. J. Fish Biol. 31, 517–543. Connors, B.M., Juarez-Colunga, E., Dill, L.M., 2008. Effects of varying salinities on Lepeophtheirus salmonis survival on juvenile pink and chum salmon. J. Fish Biol. 72, 1825–1830. Costello, M.J., 2006. Ecology of sea lice parasitic on farmed and wild fish. Trends Parasitol. 22, 475–483. Deardorff, T.L., Kent, M.L., 1989. Prevalence of larval Anisakis simplex in pen-reared and wild-caught salmon (Salmonidae) from Puget Sound, Washington. J. Wildl. Dis. 25, 416–419. Deardorff, T.L., Overstreet, R.M., 1981. Larval Hysterothylacium (=Thynnascaris) (Nematoda: Anisakidae) from fishes and invertebrates in the Gulf of Mexico. Proc. Helminthol. Soc. Wash. 48, 113–126. DHI (Dansk Hydraulisk Institut), 2007. NOVANA programmet 2004–2009. Det marine modelkompleks (MMK): 2006 resultater fra havmodellen. Rapport. (170 pp. (in Danish). Durán, M.L.S., Quinteiro, P., Ubeira, F.M., 1989. Nematode parasites of commercially important fish in NW Spain. Dis. Aquat. Org. 7, 75–77. EFSA (European Food Safety Authority) Panal on Biological Hazards (BIOHAZ), 2010. Scientific opinion on risk assessment of parasites in fishery products. EFSA J. 8, 1543 (91 pp.). FAWC (Farmed Animal Welfare Council), 1996. Report on the welfare of farmed fish, (Surbiton, Surrey).

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