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Interactions between Paramoeba perurans, the causative agent of amoebic gill disease,
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and the blue mussel, Mytilus edulis
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Christine Rolin1,2*, Jennifer Graham2, Una McCarthy2, Samuel A. M. Martin1, Iveta
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Matejusova2*
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School of Biological Sciences, University of Aberdeen, Tillydrone Avenue, Aberdeen AB24 2TZ, UK
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Marine Scotland Science, Marine Laboratory, 375 Victoria Road, Aberdeen, AB11 9DB, UK
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Abstract: Amoebic gill disease (AGD) is caused by the ectoparasite Paramoeba perurans
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found free-living in seawater. In recent years outbreaks of AGD have occurred in most
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salmon farming countries causing significant economic losses. Mussels co-cultured with
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salmon in integrated multi-trophic aquaculture (IMTA) systems may change pathogen
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dynamics on sites by acting as reservoirs or biological controls. Through the use of an 18S
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rRNA gene quantitative real-time PCR we tested the interactions between P. perurans and
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blue mussels (Mytilus edulis) under experimental conditions by means of water-borne
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transmission. Quantification of DNA from water samples revealed a rapid decrease in P.
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perurans DNA over two weeks in the presence of mussels under experimental conditions. P.
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perurans was detected on swabs from mussel shells up to 48 hours post-exposure.
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Additionally, no P. perurans were detected in mussels collected from natural mussel beds
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and fish farms. These results indicate mussels are not a likely reservoir host for P. perurans
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but may in fact actively remove water-borne P. perurans.
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Keywords: Blue mussel, Paramoeba perurans, amoebic gill disease, biological control,
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Atlantic salmon, integrated multi-trophic aquaculture.
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*Corresponding
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[email protected] Present address: NAFC Marine Centre, Port Arthur, Scalloway,
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Shetland, ZE1 0UN, UK. Iveta Matejusova: E-mail address:
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[email protected] .
authors: Christine Rolin: Tel.: + 44 (0) 1595 772300; E-mail address:
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1. Introduction
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Amoebic gill disease (AGD) is an economically significant disease of Atlantic salmon
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(Salmo salar Linnaeus, 1758) caused by an amoeboid protozoan parasite Paramoeba
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perurans (synonymous to Neoparamoeba perurans, according to Feehan et al., 2013) (Shinn
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et al., 2015). In Tasmania, AGD has long been one of the main problem in salmonid
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aquaculture, with typical losses of 10-20 % of production costs (Munday et al., 2001). In
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Scotland typical mortalities on Atlantic salmon farms range between 10-20 % but have been
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reported to be as high as 70 % at some sites (Marine Scotland Science, 2012). In recent years
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a global emergence of this disease has been documented with frequent outbreaks across the
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Northern and Southern Hemispheres i.e. Ireland, Scotland, Norway, Spain, France, North
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America, Japan, Chile and South Africa (Rodger & McArdle, 1996; Palmer et al., 1997;
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Findlay & Munday, 1998; Steinum et al., 2008; Crosbie et al., 2010; Bustos et al., 2011;
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Mouton et al., 2014). Paramoeba perurans is a free-living amoeba, however, when it
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colonises Atlantic salmon gills it can cause white swollen lesions, epithelial hyperplasia, and
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increased mucus production leading to lethargy, apoxia and eventually death (Munday et al.,
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2001).
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Very little is known about the origin of P. perurans responsible for disease outbreaks.
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Fish import and subsequent spread between farms is not generally thought to be a feasible
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explanation for emergence of this disease in Europe (Steinum et al., 2008). On the other
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hand, prolonged periods of increased seawater temperature and/or high salinities combined
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with densities of fish in aquaculture sites most likely provided an advantage to P. perurans
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naturally present in marine environment (Nowak, 2007; Bridle et al., 2010; Wright et al.,
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2015). In aquaculture, infections with P. perurans were reported from a range of farmed fish
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in addition to Atlantic salmon (Crosbie et al., 2010; Karlsbakk et al., 2013; Mouton et al.,
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2014) while wild fish collected in the proximity of infected Atlantic salmon farms in
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Australia were tested negative (Douglas-Helders et al., 2002).
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Diversification of aquaculture, reduction of potential adverse environmental impacts of
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Atlantic salmon farm systems and expansion of the shellfish industry are major aspirations
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intrinsic to the successful expansion of the aquaculture sector in Scotland. Integrated multi-
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trophic aquaculture (IMTA) systems are being developed in Atlantic salmon farming
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countries to test the concept of integration of Atlantic salmon, blue mussels and/or seaweed
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production. The aim is to deliver a diverse aquaculture system with a parallel reduction in
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organic waste from fish and dissolved inorganic nutrients (Barrington et al., 2009; Chopin et
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al., 2012; Wang et al., 2012). Co-culturing different species can change disease dynamics on
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farm sites and create new risks which need to be addressed to ensure good biosecurity in the
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production systems. Mussels are filter-feeders and have the potential to digest and inactivate
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some pathogens such as sea lice (Lepeophtheirus salmonis Krøyer, 1837) (Molloy et al.,
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2011; Webb et al., 2013) or infectious salmon anaemia virus (ISAV) (Molloy et al., 2012).
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On the other hand Vibrio anguillarum (Pietrak et al., 2012), Loma salmonae (Putz, Hoffman
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& Dunbar, 1965) Morrison & Sprague, 1981 (McConnachie et al., 2013) or infectious
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pancreatic necrosis virus (IPNV) (Molloy et al., 2013) may persist and accumulate in
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shellfish posing a threat of disease transmission to farmed fish.
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The aim of the study was to assess mussels as a potential wild reservoir of P.
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perurans and investigate interactions between Mytilus species (Linnaeus, 1758) and P.
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perurans in terms of pathogen accumulation and transmission. Previously published
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molecular tests (real-time PCR) (Fringuelli et al., 2012) were applied to (1) examine the
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persistence of P. perurans in water, on shell surfaces and in mussel tissues after experimental
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exposure and (2) test for the presence of P. perurans in mussel tissues from wild mussel
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individuals and from bio-fouling mussels collected on fish farms in Scotland.
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2. Materials and methods
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2.1. P. perurans culture maintenance and dilution curve for quantification
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Paramoeba perurans, maintained at the Marine Scotland Science (Aberdeen, UK),
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originated from gills of infected farmed Atlantic salmon (west coast of Scotland) collected in
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2012. A culture was established following the protocol of Crosbie et al., (2012). The P.
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perurans culture was maintained routinely at 15 – 18 oC on malt yeast agar (MYA) (0.1 %
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malt, 0.1 % yeast, 2 % Bacteriological agar, 35 ‰ 0.22 μm filtered seawater) overlaid with 7
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ml 0.22 μl filtered seawater (SSW, 35 ‰) collected offshore of Stonehaven (Aberdeenshire,
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UK). The culture was subcultured every two weeks by transferring the seawater overlay
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containing a free-floating population of amoebae to a new MYA plate. The old MYA plate
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containing a population of attached amoebae received a new seawater overlay. The culture of
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P. perurans was first isolated in November 2012 and used in experimental challenges
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described in the present study in May and June 2014. The pathogenicity of this culture has
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been confirmed under experimental conditions by challenging naïve Atlantic salmon at 7, 12
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and 24 months post-isolation (Marine Scotland Science, unpublished data).
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P. perurans were harvested from culture plates by centrifugation of culture
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supernatants at 6000 x g for 5 min. Pellets were pooled and resuspended in SSW. Before
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harvesting, quadruplicate log10 dilutions of were made in 96-well plates for enumeration of
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the total number of amoebae harvested under light microscope. After centrifugation, log10
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dilutions were made in duplicate from a starting cell density of P. perurans of 7 x 103 cells.
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Then, homogenised mantle-gill tissue was spiked with the prepared dilutions and total DNA
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was extracted using two different DNA extraction protocols: automated extraction using the
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QIAsymphony DNA DSP mini kit (Qiagen) or manual extraction using the MasterPure DNA
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and RNA Purification kit (Epicentre). Negative controls representing mussel tissue
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homogenate without P. perurans and blank controls were set up. P. perurans DNA was
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quantified using the quantitative real-time PCR (qPCR) (see section 2.4.) and Ct values
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recorded to create a standard dilution curve in R’ statistical package (version 3.0.2, R Core
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Team 2013).
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2.2. Experimental exposure of mussels to P. perurans culture
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2.2.1. Mussel maintenance
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Depurated market-size mussels were bought from a commercial supplier or vendor
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and kept in outdoor covered seawater tanks at 9 oC. The seawater was from Nigg Bay
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(Aberdeen, Scotland) was filtered (through a 1 mm wedge wire screen and 15 μm particle
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filter) before pumping to storage tanks. This water was also used in experimental beakers and
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tanks. Mussels were naturally feeding on nanophytoplankton present in the sea water,
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considered as one of significant dietary components for bivalves (Shumway et al., 1985) and
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no additional food was added to the holding tanks prior to the experiments.
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2.2.2. Experiment 1: Pilot experiment to establish the time frame for P. perurans ingestion by
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mussels and distribution in mussel tissues
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Forty eight beakers containing 500 ml 35 ‰ seawater (10 oC) with a small air stone
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were placed in a temperature-controlled room (18 oC). By the end of the experiment (144
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hours after exposure) the water temperature had risen to 14 oC. Five mussels were placed in
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each duplicate treatment beaker and allowed to acclimatise before adding P. perurans.
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Mussels were exposed to P. perurans (104 cells L-1 in suspension) in 500 ml and sampled at
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1, 2, 3, 4, 6, 24, 48 and 144 hours post-exposure (hpe). The mussel filtering activity was
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monitored throughout the experiment.
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At each time point, four beakers were sampled without replacement: two treatment
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beakers (containing P. perurans and mussels), one positive control beaker (P. perurans only)
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and one negative control beaker (mussels only). At each time point, all mussels were
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removed and seawater from the entire beaker was filtered through 1.0 µm cellulose nitrate
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membrane filters (25 mm diameter, GE Healthcare Life Sciences) which were stored dry in 2
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ml Safelock tubes (Eppendorf). The shell of each individual mussel was swabbed with a
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sterile cotton-tip swab on one side using four strokes lengthwise. Additionally, the bottom
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inside edge of each beaker was swabbed using two circular strokes to test adhesion and
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replication of P. perurans on plastic surfaces. All swabs were stored in RNAlater (VWR).
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The whole digestive gland and a section of mantle and gill tissue were sampled from each
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mussel and placed in separate tubes containing 100 % ethanol (Sigma). All samples were
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frozen at -20 oC immediately after sampling.
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2.2.3. Experiment. 2: Tank experiment to quantify removal rates of P. perurans from the
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water column
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Mussels were exposed to P. perurans (3 x 104 cells L-1 in suspension) in tanks
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containing 20 L of 35 ‰ seawater. These tanks were placed within a larger tank connected to
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a flow-through seawater system to maintain the experimental tanks at 12 oC throughout the
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experiment. Thirty mussels were placed in four tanks: three treatment tanks (containing P.
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perurans and mussels), one positive control tank (P. perurans only) and one negative control
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tank (mussels only). The mussels were fed with algae (Shellfish diet 1800 from Reed
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Mariculture Inc.) according to the manufacturer’s instructions, when a reduced filtering
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capacity was observed (day 7 post-exposure).
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Water samples and shell swabs were taken at 1, 4, 7, 24, 48, 168 and 384 hpe. A 500
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ml water sample was taken from each tank and filtered using the method described above.
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The entire shell of three mussels in each tank was swabbed thoroughly; sampled mussels
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were placed in a net (3 mm diameter nylon mesh), within the tank to avoid repeated swabbing
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at different time points. The swabs were placed in a dry microcentrifuge tube (opposed to
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RNAlater as in experiment 1), immediately stored at -20 °C and processed within 48 hours.
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Finally, a section of net (1 x 1 cm) (retaining sampled mussels) from each tank was placed in
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a dry tube to test P. perurans adhesion. Mussels were checked daily and filtering activity and
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mortalities were recorded.
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2.3. Sampling of wild and biofouling mussels
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To assess presence of P. perurans in wild mussels fifty individuals were collected
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from each of three sampling points on the west coast of Scotland (Lunderston Bay, Loch
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Long (Ardgarten) and Rascarrel Bay) in 2013. To assess presence of P. perurans in mussels
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in aquaculture farm environment, a sample of 150 mussels, settled directly on the surface of
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Atlantic salmon fish cages, were collected in 2012 from a farm on the west coast of Scotland.
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This particular farm had an ongoing P. perurans infection in the time of sample collection. A
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pooled sample of gill and mantle was collected from each individual and stored in 100 %
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ethanol (Sigma) until further processing. All field samples were homogenised (see section
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2.4.) and stored at -80 oC for up to two years before processing in the current study.
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2.4. DNA extraction and qPCR
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2.4.1. Water filters
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DNA captured on water filters was extracted manually using the MasterPure
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Complete DNA and RNA Purification kit (Epicentre) following the protocol for DNA
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purification of tissue samples. Each filter was processed by adding 125 mg of glass beads
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(710 – 1180 μm, Sigma), 1 μl proteinase K (600 mAU/ml solution) and 300 μl of Tissue and
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Cell Lysis solution, then placed in the TissueLyser (Qiagen Ltd.) and homogenised at 25 Hz
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for 5 min. A negative control contained 1 μl proteinase K and 300 μl Tissue and Cell Lysis
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solution. Due to a large quantity of debris, protein precipitation was repeated once. As pellets
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were difficult to resuspend, samples were placed in the fridge for 30 min and then
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resuspended by pipetting. Genomic DNA was stored at -20 oC prior to further analysis.
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2.4.2. Shell swabs
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Swabs fixed in RNAlater or dry tubes were transferred to an empty 2 ml Safelock
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tube containing 20 μl Proteinase K and 180 μl ATL buffer (QIAsymphony DNA DSP mini
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kit, Qiagen), vortexed briefly and lysed by incubating at 56 oC for a minimum of 3 hours. A
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negative extraction control of 20 μl proteinase K and 180 μl ATL buffer was included. The
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tubes were centrifuged for 2 min at 13,500 rpm and supernatant was transferred into a clean
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1.5 ml tube. Total DNA was extracted using a QIAsymphony DNA DSP mini kit and the
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QIAsymphony SP robot (Qiagen) according to manufacturer’s instructions and resuspended
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in 100 μl of distilled water. Samples of nets at 384 hpe in experiment 2 were processed
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according to the same methods.
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2.4.3. Mussel tissues
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Mussel tissue (from experimental and field studies) was homogenised in 1 ml of ATL
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buffer (Qiagen) using a 7 mm metal bead at 25 Hz for 5 min, in a Tissuelyser homogeniser
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(Qiagen). DNA was extracted from a volume of tissue equivalent to 5 mg of tissue as
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described in the section above. All samples were stored at -20 oC until further analysis.
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2.4.4. Quantitative Real-Time PCR (qPCR)
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All DNA samples were tested for the presence of P. perurans using previously
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published qPCR assays targeting a 139-bp fragment specific to the P. perurans 18S rRNA
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gene (Fringuelli et al., 2012). As an endogenous control to confirm the quality of extracted
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DNA, ME15/16 Mytilus spp. qPCR assays were run in multiplex for simultaneous detection
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of all three Mytilus species (M. edulis, M. galloprovincialis, M. trossulus) according to Dias
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et al., (2008).
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The final qPCR reaction mixture contained 2 x Quanta Custom Toughmix (VWR),
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900 nM of each primer and 250 nM of each TaqMan® probe, 1 μl genomic DNA at a
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concentration of 50 or 100 ng ul-1 in final volume of 20 μl. The real-time PCR was performed
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on the LC480 Thermal Cycler (Roche). The reproducible limit of detection (RLOD) for the
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P. perurans assay was 36.22 Ct (Fringuelli et al., 2012) and this value was used to distinguish
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between P. perurans negative and positive samples. An increase of 3 Ct was considered a 10-
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fold decrease.
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2.5. Data analyses
All data analyses were performed in R’ statistical package (version 3.0.2, R Core
Team, 2013) and plotted with ggplot2 package (Wickham, 2009).
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Experiment 1 was designed as a pilot experiment to test the sampling methods and
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detection of P. perurans in water, swabs and tissues, as well as establish the rate at which P.
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perurans might be extracted from water by filtering mussels. Therefore, no models were
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applied and the results were analysed by calculating means and standard deviations of P.
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perurans Ct values for each treatment group.
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In experiment 2 all tanks were independent from each other and were sampled
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repeatedly over the duration of the experiment. Only the data from water extraction were
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modelled, to determine any change in P. perurans over time and between treatments (P.
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perurans with and without mussels). Models were tested for effects and interactions between
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tanks before pooling data in the respective treatment groups. Data transformations were
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performed and the best model was chosen from linear models, including interactions and
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fixed factors, and from mixed effect models with tank as a random factor. The best fit model
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was chosen based on an Akaike Information Criterion (AIC) comparison, summary plots and
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residual plots. If the difference in AIC was ≤ 2 between two models, the model with less
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variables was considered the better fit. Analysis of Variance (ANOVA) was used to test any
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significant differences over time and between treatments.
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The shell swabs were analysed as binomial data based on proportion of swabs positive
and tested with a Chi test to detect significant changes over time.
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3. Results
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3.1. Detection of P. perurans in spiked mussel tissue
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The dilution curves generated from P. perurans-spiked mussel tissue showed that the
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MasterPure extraction protocol has a greater extraction efficiency than the DNA DSP
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protocol and confirm that P. perurans can be detected down to 70 cells for the DNA DSP and
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7 cells for the MasterPure kit (Table 1). A linear relationship between the concentration of P.
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perurans-spiked mussel tissue and the Ct values was observed for both the DNA DSP mini
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kit (y = -4.28x + 43.5, R2 = 0.99) and MasterPure kit (y = -3.38x + 37.91, R2 = 0.99) (Fig. 1).
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However, DNA yield from the DNA DSP kit might be underestimated (by up to 1 Ct) as only
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200 μl of lysed solution was extracted using the QIAsymphony compared to 300 μl used for
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manual MasterPure kit extraction.
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3.2. Experimental exposure of mussels to P. perurans culture
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Observed mortality in experiment 1 and 2 reached 6 % and 22 %, respectively, by the
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end of the experiment. In experiment 2, all mortalities occurred in one out of three individual
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treatment tanks, 3 mussels died before day 7 and 15 mussels by the end of the experiment.
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Causes of mussel mortality were not investigated.
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3.2.1. Detection of P. perurans in water samples
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In experiment 1, a clear reduction in quantity of P. perurans (103-fold) was detected
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in water after 1 hour and it remained low until day 16 (P. perurans Ct = 33.4 ± 2.0 0 with
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mussels and Ct = 25.7 ± 0.9 without mussels) (Fig. 2).
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In experiment 2, only data from 1 to 168 hpe (i.e. 7 days) were modelled as P.
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perurans Ct values were below RLOD (i.e. positive) in only one of three with mussels on 384
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hpe. Due to a non-linear relationship between Ct values and time, the values for hpe were
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log10 transformed. Despite a significant variation in the rate of the reduction of P. perurans
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between tanks (F = 3.69, df = 2, p < 0.05) data from all tanks were pooled in the final model
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as the model excluding the tanks provided the same or better fit based on summary plots,
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residual plots and the AIC values (< 2 AIC difference). The final model included an
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interaction between treatment tank and hpe as explanatory variables.
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A rapid and significant decrease in detectable P. perurans was recorded in the
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presence of mussels, after 1 hour the difference between the tanks with and without mussels
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was > 10-fold (Table 2). P. perurans were reduced a further 102-fold between 1 and 24 hours
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and a 104-fold between 1 hour and 7 days (F = 86.7, df = 1, p < 0.001; y = 3.73x + 25.29, R2
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= 0.83) (Fig. 3). Some reduction in P. perurans with time was also observed in the positive
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control tank (in the absence of mussels) with a 10-fold change (range 21.6 – 23.7 Ct) across
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the 164 hpe (F = 15.25, df = 1, p < 0.05; y = 0.96x + 21.47, R2 = 0.74) (Fig. 3). The rate of
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reduction in P. perurans was significantly greater in the mussel treatment group compared to
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the positive control group (F = 169.79, df = 1, p < 0.001). No P. perurans DNA was detected
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in water samples from negative control beakers or tanks. Endogenous Mytilus spp. multiplex
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assays indicated that Mytilus DNA levels in the water of all tanks were stable throughout the
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experiment (range: 26.5-32.5 Ct).
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3.2.2. Shell swabs
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Mussel shells were swabbed to determine if P. perurans attaches to shells and uses
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them as a surface for attachment, survival or replication. The proportion of shells testing
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positive in both experiments are given in Table 3. All shell swabs from negative control
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beakers and tanks tested negative for P. perurans DNA. Additionally, all swabs from the
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positive control beakers tested positive.
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In experiment 1, 1/10 shell swabs tested positive between 1 and 4 hpe with most Ct
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values below the RLOD (mean: 36.01 + 1.46, range: 33.70-38.41 Ct). In experiment 2, where
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a modified sampling protocol was used 9 out of 10 swabs tested positive between 1 and 4 hpe
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and lower Ct values were obtained for P. perurans than in experiment 1 (mean: 33.98 + 2.14
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Ct, range: 28.94 – 38.71 Ct).
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In both experiments P. perurans was detected on swabs only during the first 48 hpe.
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In experiment 2, all shells tested positive 1 hpe but only 50 % tested positive after 48 hpe. As
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there was no significant tank effect on the proportion of P. perurans shell swabs testing
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positive, data were pooled. The reduction in the proportion of positive shell swabs was
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significant over time (Chi test: p < 0.001). Additionally, swabs taken from the bottom of the
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beakers in experiment 1 all tested negative when mussels were present (range ≥ 36.58 Ct),
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when mussels were absent 3 out of 8 beakers tested positive (range ≥ 32.24 Ct). Samples of
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nets taken at 385 hpe in experiment 2 tested negative when mussels were present and positive
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when mussels were absent (range: 26.0 – 27.27 Ct). Nets were also positive for Mytilus spp.
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with lower Ct values (range: 34.73 – 36.83 Ct) than for P. perurans.
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3.2.3. Detection of P. perurans in mussel tissues
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To determine a potential of P. perurans accumulation in mussel tissues and to
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establish a time frame for accumulation and/or release of P. perurans, the gill, mantle and
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digestive gland were tested for presence of the parasite by qPCR. No P. perurans DNA was
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detected in any tissues in either experiment. All mussels from the negative control group
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were negative for P. perurans DNA. Endogenous Mytilus spp. multiplex assay indicated that
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all DNA extracted from mussels and tissues was of good quality (range: 20.6 – 28.0 Ct). No
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P. perurans or Mytilus DNA was detected in their respective negative controls.
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3.3. Detection of P. perurans in naturally settled and biofouling mussels
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The combined mantle-gill tissue samples collected from natural mussel beds (Lunderston,
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Long, Rascarrel Bay) and fish farm cage (west coast of Scotland) tested negative for P.
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perurans. Endogenous Mytilus spp. multiplex assay indicated that all DNA extracted from
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mussels tissues was of good quality (all values < 30 Ct). No P. perurans or Mytilus DNA was
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detected in their respective negative controls.
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4. Discussion
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Paramoeba spp. have previously been reported in shellfish such as lobsters (Mullen et
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al., 2005), sea urchins (Feehan et al., 2013) and oysters (Sühnel et al., 2014) but to date P.
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perurans has not been detected in any shellfish and the only known environmental reservoir
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is free-living in sea water (Bridle et al., 2010, Wright et al., 2015). Similarly, the present
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study did not detect any P. perurans in mussels collected from natural beds nor selected fish
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farms on the west coast of Scotland.
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Mussel shell surfaces have not previously been considered as an environmental
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reservoir for fish diseases. To assess the mussel shell’s potential as a substrate for P.
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perurans attachment and replication, the shells from mussels experimentally exposed to high
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numbers of P. perurans were swabbed according to the protocol developed for fish gill swabs
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(Bridle et al., 2010). The method yielded Ct values below RLOD in experiment 1. A change
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in the protocol (increased sample area and storage in dry tubes) was more effective, with
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more positive detections in experiment 2. The improvement in detection of P. perurans may
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be attributed to increasing the quantity of DNA on each swab or storage in dry tubes may
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have prevented loss of DNA into the fixative solution.
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The present study suggested that mussel shells only provide a suitable surface for P.
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perurans up to 48 hours post exposure when the parasite is present in high numbers. Shell
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swabs had Ct values for P. perurans comparable to those from gill swabs from AGD-infected
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fish with gill scores of 1 or 2 (Bridle et al., 2010), which is considered a light infection
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(Powell et al., 2001). Using the calibration curve from the present study, the Ct values
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detected on the surface of mussel shells at 24 hpe correspond to approximately 100 amoebae,
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and 10 amoebae after 48 days and none at the end of the experiment (144 hpe). Therefore, it
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can be concluded that mussel shells most likely do not act as a long-term environmental
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reservoir for P. perurans. In comparison, swabs of the inside of the beaker surface
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(experiment 1) and net samples taken at 384 hpe (experiment 2) tested positive for P.
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perurans when mussels were absent, but not when mussels were present, indicating P.
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perurans do attach to surfaces in the absence of mussels. When the ambient water had a
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concentration of approximately 10,000 amoebae L-1, the Ct values of the net were equivalent
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to as much as 1,000 amoebae which is more than expected considering the small size of the
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net sample (1 x 1 cm). Materials such as plastic and net surfaces are abundant on salmon
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farms and should be considered as a potential substrate for P. perurans attachment and/or
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replication.
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Development of IMTA has a clear economic and environmental benefit as it widens
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the aquaculture product portfolio and removes organic waste associated with unused fish feed
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(Stirling & Okumus, 1995; Whitmarsh et al., 2006; Macdonald et al., 2011; Handå et al.,
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2012). While species interactions in polyculture may be exploited for bio-mitigation of fish
355
farm waste or as biological controls of pathogens, they may also pose new potential pathways
356
for disease transmission. Uptake and release of variety of a finfish pathogens by mussels and
357
scallops have been tested under experimental conditions. Some pathogens appear to be taken
358
up and inactivated by mussels e.g. sea lice copepodids (Molloy et al., 2011; Bartsch et al.,
359
2013; Webb et al., 2013) and ISAV (Skår & Mortensen 2007; Molloy et al., 2012) while
360
others such as Vibrio anguillarum (Pietrak et al., 2012), IPNV (Mortensen et al., 1992;
361
Molloy et al., 2013) and L. salmonae (McConnachie et al., 2013) can be accumulated, remain
362
viable in tissues and can be shed in faeces long after exposure. For example, up to 70 %
363
mortality were observed in Atlantic cod (Gadus morhua) exposed to V. anguillarum-infected
364
mussel faeces under experimental conditions (Pietrak et al., 2012). On the other hand, no
365
mortality of Atlantic salmon cohabited with IPNV-positive mussels was observed, although
366
the rate of transmission (determined by qRT-PCR and virus culture) between mussels and
367
Atlantic salmon, and between fish infected with IPNV by intraperitoneal injection and naïve
368
fish was comparable, suggesting a risk of disease transmission at potential IMTA sites
369
(Molloy et al., 2013). However, caution must be used when interpreting and utilising results
370
of experimental challenges in estimating risk of disease transmission under the dynamic
371
conditions of open sea cage farming. For example, in a direct immersion experiment mussels
372
were able to take up ISAV but no mussels were found positive when cohabited with
373
artificially infected Atlantic salmon exhibiting mortality and clinical signs of the ISA disease
374
or during ISA outbreaks at farm sites (Skår & Mortensen, 2007).
375
In the present study, no evidence was found to suggest that P. perurans is capable of
376
persisting or accumulating in mussels under experimental conditions. Despite exposing
377
mussels to high numbers of viable P. perurans (experiments 1 and 2: 2 x 104 amoebae L-1 and
378
3 x 104 amoebae L-1 respectively) no values above the RLOD were detected in gill, mantle or
379
digestive gland using a qPCR assay (Fringuelli et al., 2012). Sensitivity of the qPCR assay
380
was tested in the present study and it was confirmed that P. perurans can be detected down to
381
70 amoeba in spiked mussel tissue spiked material. Similar studies considered an adverse
382
effect of PCR inhibitors on the ability of qPCR assays to detect accumulated pathogens in
383
mussels, particularly in the digestive gland (Schrader et al., 2012; Molloy et al., 2013).
384
However, the endogenous Mytilus spp. assay used in the current study did not indicate that
385
inhibition was occuring, with the range of Ct values in the digestive gland (22.6 ± 1.3 Ct)
386
comparable to other tissues such as mantel and gill (23.3 ± 1.6 Ct).
387
Preservation and storage of samples can affect the quality of DNA and the final result
388
of the analysis (Ørpetveit et al., 2010). Gill swabs are commonly stored in ethanol before
389
AGD diagnosis (Young et al., 2008; Crosbie et al., 2010; Karlsbakk et al., 2013). The storage
390
of tissue samples in ethanol with the addition of a freezing step may have led to lysis of P.
391
perurans cells and loss of material into the fixative solution. Water filters and shell swabs
392
were stored in dry tubes instead of ethanol and consistently tested positive in the experiments.
393
Other studies have successfully detected sea lice, ISAV and IPNV from digestive gland
394
stored directly in lysis buffer after sampling (Molloy et al., 2011, 2012, 2013) and P.
395
perurans on water filters stored at room temperature (Bridle et al., 2010; Wright et al., 2015).
396
Despite the absence of P. perurans DNA in mussel tissues, in both experiments a
397
significant decrease of P. perurans DNA signal over time was observed in water from the
398
beakers/tanks where mussels were kept compared to the positive control tank. It should be
399
emphasised that the P. perurans Ct values in the positive control beakers/tanks did not rapidly
400
change, and remained stable increasing by approximately 10-fold across 16 days (Table 2)
401
and therefore reduction of P. perurans DNA signal in treatment tank cannot be explained by
402
DNA degradation over time. When compared to the positive control, after only 1 hour P.
403
perurans decreased by 103-fold in experiment 1 (0.1 L of water per mussel) and 10-fold (2
404
litres of water per mussel) in experiment 2 (overall 104-fold across 7 days), indicating a very
405
rapid removal that increases with mussel density. Furthermore, to support the hypothesis that
406
higher density increases removal rate it can be mentioned anecdotally that the tank with the
407
highest mussel mortalities (i.e. lowest filtering capability) experienced a lower removal rate
408
than the other two tanks. A rapid uptake and re-release of P. perurans could explain the lack
409
of accumulation/detection of P. perurans in any analysed mussel tissues, however, as the
410
water samples would contain faeces and pseudofaeces, it is unlikely any accumulation in the
411
waste products occurred as there was no secondary spike in detection of P. perurans target
412
DNA. Changes to mussel feeding behaviour observed towards the end of tank experiment
413
could have affected results of the study as filtering activity was reduced at this point.
414
However, no P. perurans was detected in the mussel tissues even in the very early time points
415
where active feeding was observed, in both beaker and tank experiments.
416
Prevention is the key to controlling outbreaks of diseases that are difficult to treat
417
and/or manage. An understanding of the source and potential spread of pathogen in natural
418
ecosystems as well as systems such as IMTA is therefore crucial for aquaculture expansion
419
and sustainability. The present study indicates that the presence of mussels on Atlantic
420
salmon farms do not pose an increased risk in terms of accumulation of P. perurans and
421
release of viable parasites. This is an agreement with observations made in Shetland where
422
mussels on rafts appeared unaffected during widespread AGD outbreaks (personal
423
communication, Margaret Nugent, SSQC Ltd). However, P. perurans may be able to settle
424
on hard surfaces such as plastic or net structures on fish farm cages, these should therefore be
425
considered as potential environmental reservoirs for P. perurans. Further work is needed to
426
understand the mechanism leading to reduction of P. perurans in the presence of blue
427
mussels in order to determine if mussels can be considered as biological control against
428
AGD.
429
430
Acknowledgements: The authors would like to thank staff at Marine Scotland Science
431
Patricia White, Rebecca McIntosh, Julia Black and Mark Fordyce for their technical
432
assistance and invaluable feedback on the project. Thanks also go to Alex Douglas at the
433
University of Aberdeen for his advice on data analysis and statistics. For feedback on the
434
manuscript thanks to Lesley McEvoy and Rhiannon Inkster at the NAFC Marine Centre. The
435
study was supported by the Marine Collaborations Forum (MarCRF) which aims to develop
436
cross-disciplinary research between the University of Aberdeen and Marine Scotland
437
Science. Finally, thanks are also due to Scottish Fishermen’s Trust for a student support
438
bursary.
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References
453
Barrington, K., Chopin, T., Robinson, S., 2009. Integrated multi-trophic aquaculture (IMTA)
454
in marine temperate waters. In: Soto D (ed) Integrated Mariculture: A Global Review, p 7–
455
46. FAO Fisheries and Aquaculture Technical Paper No. 529 FAO Rome.
456
Bartsch, A., Robinson, S.M.C., Liutkus, M., Ang, K.P., Webb, J., Pearce, C.M., 2013.
457
Filtration of sea louse, Lepeophtheirus salmonis, copepodids by the blue mussel, Mytilus
458
edulis, and the Atlantic sea scallop, Placopecten magellanicus, under different flow, light and
459
copepodid-density regimes. J. Fish Dis. 36, 361–370.
460
Bridle, A.R., Crosbie, P.B.B., Cadoret, K., Nowak, B.F., 2010. Rapid detection and
461
quantification of Neoparamoeba perurans in the marine environment. Aquaculture 309, 56–
462
61.
463
Bustos, P.A., Young, N.D., Rozas, M.A., Bohle, H.M., Ildefonso, R.S., Morrison, R.N.,
464
Nowak, B.F., 2011. Amoebic gill disease (AGD) in Atlantic salmon (Salmo salar) farmed in
465
Chile. Aquaculture 310, 281–288.
466
Chopin, T., Cooper, J.A., Reid, G., Cross, S., Moore, C., 2012. Open-water integrated multi-
467
trophic aquaculture: environmental biomitigation and economic diversification of fed
468
aquaculture by extractive aquaculture. Rev. Aquaculture 4, 209–220.
469
Crosbie, P.B.B., Ogawa, K., Nakano, D., Nowak, B.F., 2010. Amoebic gill disease in
470
hatchery-reared ayu, Plecoglossus altivelis (Temminck and Schlegel), in Japan is caused by
471
Neoparamoeba perurans. J. Fish Dis. 33, 455–458.
472
Crosbie, P.B.B., Bridle, A.R., Cadoret, K., Nowak, B.F., 2012. In vitro cultured
473
Neoparamoeba perurans causes amoebic gill disease in Atlantic salmon and fulfils Koch’s
474
postulates. Int. J. Parasitol. 42, 511–515.
475
Dias, P.J., Sollelis, L., Cook, E.J., Piertney, S.B., Davies, I.M., Snow, M., 2008.
476
Development of a real-time PCR assay for detection of Mytilus species specific alleles:
477
Application to a sampling survey in Scotland. J. Exp. Mar. Biol. Ecol. 367, 253–258.
478
Douglas-Helders, G.M., Dawson, D.R., Carson, J., Nowak, B.F., 2002. Wild fish are not a
479
significant reservoir for Neoparamoeba pemaquidensis (Page, 1987). J. Fish Dis. 25, 569–
480
574.
481
Feehan, C.J., Johnson-Mackinnon, J., Scheilbling, R.E., Lauzon-Guay, J.-S., Simpson,
482
A.G.B., 2013. Validating the identity of Paramoeba invadens, the causative agent of
483
recurrent mass mortality of sea urchins in Nova Scotia, Canada. Dis. Aquat. Org. 103, 209–
484
227.
485
Findlay, V.L. & Munday, B.L., 1998. Further studies acquired resistance to amoebic gill
486
disease (AGD) in Atlantic salmon, Salmo salar L. J. Fish Dis. 23, 193–198.
487
Fringuelli, E., Gordon, A.W., Rodger, H., Welsh, M.D., Graham, D.A., 2012. Detection of
488
Neoparamoeba perurans by duplex quantitative Taqman real-time PCR in formalin-fixed,
489
paraffin-embedded Atlantic salmonid gill tissues. J. Fish Dis. 35, 711–724.
490
Handå, A., Min, H., Wang, X., Broch, O.J., Reitan, K.J., Reinertsen, H., Olsen, Y., 2012.
491
Incorporation of fish feed and growth of blue mussels (Mytilus edulis) in close proximity to
492
salmon (Salmo salar) aquaculture: Implications for integrated multi-trophic aquaculture in
493
Norwegian coastal waters. Aquaculture 356–357, 328–341.
494
Karlsbakk, E., Olsen, A.E., Einen, A.-C.B., Mo, T.A., Fiksdal, I.U., Aase, H., Kalgraff, C.,
495
Skår, S.-J., Hansen, H., 2013. Amoebic gill disease due to Paramoeba perurans in ballan
496
wrasse (Labrus bergylta). Aquaculture 412–413, 41–44.
497
Marine Scotland Science, 2012. Amoebic gill disease. Topic sheet no. 96 V1.
498
http://www.scotland.gov.uk/Resource/ 0039/00393037.pdf on the 15/7/2015.
499
McConnachie, S.H., Guselle, N.J., Speare, D.J., 2013. Retention of viable microsporidial
500
(Loma salmonae) spores within the blue mussel (Mytilus edulis): Use of an experimental
501
laboratory model probing pathogen transfer within a multi-trophic aquaculture setting.
502
Aquaculture 376–379, 1–5.
503
Molloy, S.D., Pietrak, M.R., Bouchard, D.A., Bricknell, I., 2011. Ingestion of Lepeophtheirus
504
salmonis by the blue mussel Mytilus edulis. Aquaculture 311, 61–64.
505
Molloy, S.D., Pietrak, M.R., Bouchard, D.A., Bricknell, I., 2012. The interaction of infectious
506
salmon anaemia virus (ISAV) with the blue mussel, Mytilus edulis. Aquacult. Res. 1–10.
507
Molloy, S.D., Pietrak, M.R., Bricknell, I., Bouchard, D.A., 2013. Experimental transmission
508
of infectious pancreatic necrosis virus (IPNV) from the blue mussel, Mytilus edulis, to
509
cohabiting Atlantic salmon (Salmo salar) smolts. Appl. Environ. Microbiol. 79, 5882–5890.
510
Mortensen, S.H., Bachere, E., Le Gall, G., Mialhe, E., 1992. Persistence of infectious
511
pancreatic necrosis virus (IPNV) in scallops Pecten maximus. Dis. Aquat. Org. 12, 221–227.
512
Mouton, A., Crosbie, P., Cadoret, K., Nowak, B., 2014. First record of amoebic gill disease
513
caused by Neoparamoeba perurans in South Africa. J. Fish Dis. 37, 407–409.
514
Mullen, T.E., Nevis, K.R., O'Kelly, C., Gast, R.J., Frasca, Jr. S., 2005. Nuclear small-subunit
515
ribosomal RNA gene-based characterization, molecular phylogeny and PCR detection of the
516
Neoparamoeba from Western Long Island Sound lobster. J. Shellfish Res. 24, 719–731.
517
Munday, B.L., Zilberg, D., Findlay, V., 2001. Gill disease of marine fish caused by infection
518
with Neoparamoeba pemaquidensis. J. Fish Dis. 24, 497–507.
519
Nowak, B., 2007. Parasitic diseases in marine cage culture – An example of experimental
520
evolution of parasites? Int. J. Parasitol. 37, 581–588.
521
Ørpetveit, I., Mikalsen, A.B., Sindre, H., Evensen, Ø., Dannevig, B.H., Midtlyng, P.J., 2010.
522
Detection of Infectious Pancreatic Necrosis Virus in subclinically infected Atlantic Salmon
523
by virus isolation in cell culture or Real-Time reverse transcription polymerase chain
524
reaction: Influence of sample preservation and storage. J. Vet. Diagn. Invest. 22, 886–895.
525
Palmer, R., Carson, J., Ruttledge, M., Drinan, E., Wagner, T., 1997. Gill disease associated
526
with Paramoeba, in sea reared Atlantic salmon in Ireland. Bull. Eur. Assn. Fish P. 17, 112–
527
114.
528
Pietrak, M.R., Molloy, S.D., Bouchard, D.A., Singer, J.T., Bricknell, I., 2012. Potential role
529
of Mytilus edulis in modulating the infectious pressure of Vibrio anguillarum 02β on an
530
integrated multi-trophic aquaculture farm. Aquaculture 326-329, 36–39.
531
Powell, M.D., Parsons, H.J., Nowak, B.F., 2001. Physiological effects of freshwater bathing
532
of Atlantic salmon (Salmo salar) as a treatment for amoebic gill disease. Aquaculture 199,
533
259–266.
534
R Core Team, 2013. R: A language and environment for statistical computing. R Foundation
535
for Statistical Computing, Vienna, Austria. ISBN 3-900051-07-0, URL http://www.R-
536
project.org/.
537
Rodger, H.D. & McArdle, J.F., 1996. An outbreak of amoebic gill disease in Ireland. Vet.
538
Rec. 139, 348–349.
539
Schrader, C., Schielke, A., Ellerbroek, L., Johne, R., 2012. PCR inhibitors – occurrence,
540
properties and removal. J. Appl. Microbiol. 113, 1014–1026.
541
Shumway, S.E., Cucci, T.L., Newell, R.C., Yentsch, C.M., 1985. Particle selection, ingestion,
542
and absorption in filter-feeding bivalves. J. Exp. Mar. Biol. Ecol. 91, 77–92.
543
Shinn, A.P., Pratoomyot, J., Bron, J.E., Paladini, G., Brooker, E.E., Brooker, A.J., 2015.
544
Economic costs of protistan and metazoan parasites to global mariculture. Parasitology 142,
545
196–270.
546
Skår, C.K. & Mortensen, S., 2007. Fate of infectious salmon anaemia virus (ISAV) in
547
experimentally challenged blue mussels Mytilus edulis. Dis. Aquat. Org. 74, 1–6.
548
Steinum, T., Kvellestad, A., Rønneberg, L.B., Nilsen, H., Asheim, A., Fjell, K., Nygård,
549
S.M.R., Olsen, A.B., Dale, O.B., 2008. First cases of amoebic gill disease (AGD) in
550
Norwegian seawater farmed Atlantic salmon, Salmo salar L., and phylogeny of the causative
551
amoeba using 18S cDNA sequences. J. Fish Dis. 31, 205–214.
552
Stirling, H.P. & Okumus, I., 1995. Growth and production of mussels (Mytilus edulis L.)
553
suspended at salmon marine cages and mussel farms in two sea lochs on the West Coast of
554
Scotland. Aquaculture 134, 193–210.
555
Sühnel, S., Ivachuk Cda, S., Schaefer, A.L., Pontinha, V.A., Martins, M.L., Figueras, A.,
556
Meyer, G.R., Jones, S.R., Stewart, J.C., Gurney-Smith, H.J., Magalhães, A.R., Bower, S.M.,
557
2014. Detection of the parasitic amoeba (Order Dactylopodida) in the female gonads of
558
oysters in Brazil. Dis. Aquat. Org. 109, 241–250.
559
Young, N.D., Dyková, I., Nowak, B.F., Morrison, R.N., 2008. Development of a diagnostic
560
PCR to detect Neoparamoeba perurans, agent of amoebic gill disease. J. Fish Dis. 31, 285–
561
295.
562
Wang, X., Olsen, L.M., Reitan, K.I., Olsen, Y., 2012. Discharge of nutrient wastes from
563
salmon farms: Environmental effects, and potential for integrated multi-trophic aquaculture.
564
Aquacult. Environ. Interac. 2, 267–283.
565
Webb, J.L., Vandenbor, J., Pirie, B., Robinson, S.M.C., Cross, S.F., Jones, S.R.M., Pearce,
566
C.M., 2013. Effects of temperature, diet, and bivalve size on the ingestion of sea lice
567
(Lepeophtheirus salmonis) larvae by various filter-feeding shellfish. Aquaculture 406–407,
568
9–17.
569
Whitmarsh, D.J., Cook, E.J., Black, K.D., 2006. Searching for sustainability in aquaculture:
570
An investigation into the economic prospects for an integrated salmon-mussel production
571
system. Mar. Policy 30, 293–298.
572
Wickham, H., 2009. ggplot2: elegant graphics for data analysis. Springer New York.
573
Wright, D.W., Nowak, B., Oppedal, F., Bridle, A., Dempster, T., 2015. Depth distribution of
574
the amoebic gill disease agent, Neoparamoeba perurans, in salmon cages. Aquacult. Environ.
575
Interact. 7, 67–74.