Hematodinium sp. of Norway Lobster
Category 3 (Host Not in Canada)
Common, generally accepted names of the organism or disease agent
Dinoflagellate blood disease of Norway lobsters, ‘post moult syndrome’.
Scientific name or taxonomic affiliation
Hematodinium sp. parasitic dinoflagellate of Norway lobsters. When Hudson and Adlard (1996) analysed the nucleotide sequence of the partial small subunit (SSU) and the first internal transcribed spacer (ITS1) region of the ribosomal RNA gene complex of several isolates including that of Hematodinium perezi from Callinectes sapidus and the Hematodinium sp. from Chionoecetes baridi and Chionoecetes opilio, they concluded that sequence differences warranted the creation of a new species of Hematodinium for the parasite in lobsters. However, as indicated below, taxonomic issues pertaining to species identification and susceptible host identity continue to be problematic and unresolved by 2012 (Small 2012).
Note: Further information on Hematodinium spp. described from marine crustaceans other than lobsters is available on the following web pages: crabs from the European and North American coasts of the Atlantic Ocean, the cause of bitter crab disease in Chionecetes spp. and other crabs from the North Pacific and North Atlantic oceans, and crabs from the vicinity of Australia and China.
West coast of Scotland, with greatest economic significance in commercially important Nephrops norvegicus stocks in the Clyde Sea Area. Also reported from the northeast Atlantic Ocean, the Irish Sea, English Channel, German Bight, the Swedish Skagerrak, Kattegat Strait between Sweden and Danmark and possibly in other species of crustacea from the west coast of Greenland and coastal waters of the province of Newfoundland and Labrador, Canada (Stentiford et al. 2001a, Small et al. 2007b, Eigemann et al. 2010).
Nephrops norvegicus and possibly the amphipod (Orchomene nanus), and various species of crabs including: Carcinus maenas, Cancer pagurus, Pagurus bernhardus, and Liocarcinus depurator from waters around the coast of mid-western Europe; Chionoecetes opilio from the east coast of Canada and west coast of Greenland; and Hyas araneus from the west coast of Greenland (Small et al. 2006, 2007b; Eigemann et al. 2010; Small and Pagenkopp 2011). However, Hamilton et al. (2010) used a combination of phylogenetic methods with analyses of secondary structures of variable ribosomal RNA genes to show that Hematodinium from the east and west North-Atlantic was comprised of distinct ribotypes or clades which corresponded to host specificity. Particularly, a Hematodinium ‘Langoustine’ clade was only found in N. norvegicus, whereas other clades were specific to crabs or seem to be generalist parasites (Hamilton et al. 2010). Nevertheless, Jensen et al. (2010) and Small (2012) concluded that Hematodinium in N. norvegicus is considered to be the same genetic clade (and probably same species) that is widely distributed in other decapods (Chionoecetes angulatus, Chionoecetes bairdi, Chionoecetes tanneri, Chionoecetes opilio, Hyas coarctatus, Lithodes couesi, Paralithodes camchaticus, Pagurus bernhardus, Pagurus prideaux, Munida rugosa, Cancer pagurus and Carcinus maenas) from the North Atlantic and North Pacific Oceans. The Hematodinium in the portunoid crabs, Callinectes sapidus, and Liocarcinus depurator from the North Atlantic Ocean and Scylla serrata from the coast of Zhejiang Provence, China were placed within another clade (Jensen et al. 2010).
Impact on the host
Stentiford and Neil (2011) claimed that Hematodinium is the most significant known pathogen of Nephrops norvegicus and was responsible for an ongoing epidemic in fished populations of N. norvegicus in Northern Europe since at least the early 1980s. In Scotland, the lucrative N. norvegicus fishery suffered significant losses to Hematodinium in the early 1990s and this parasite has been responsible for annual seasonal epidemics of varying proportions (between 20 and 70% prevalence) (Field et al. 1992, Field et al. 1998, Stentiford et al. 2001c). A long-term data series (1990 to 2008) for the Clyde Sea area population of N. norvegicus, Scotland, UK, showed that after the initial 5 year epizootic period, prevalence stabilised at 20 to 25% and possibly this high prevalence is maintained through high recrutment rates of susceptible N. norvegicus (Beevers et al. 2012).
Very little is known about the transmission and early stages of infection of Hematodinium because of the knowledge gap in the taxonomy, life cycle, host range and latency periods for this parasite group (Stentiford and Neil 2011). In N. norvegicus, the infectious stage is not known but infection appears to establish within the host via attached multinucleate syncytial networks including multinucleate plasmodial and filamentous trophonts either attached to the surface of host organs or ramifying between structures such as the abdominal musculature (Field et al. 1992, Field and Appleton 1995). Infected female N. norvegicus from the Irish Sea do not develop mature gonads (Briggs and McAliskey 2002) and the gonads of N. norvegicus are destroyed during patent infections (Stentiford and Neil 2011). There was evidence of host cellular defense reactions in some lobsters, in the form of haemocyte encapsulation in the gills and heart, and phagocytosis of dinoflagellates by the fixed phagocytes of the hepatopancrease. Most major organs and tissues appear to be invaded before attached stages give rise to multinucleate plasmodial stages and uninucleate forms circulating freely in the haemoymph in high numbers (Field and Appleton 1995). Taylor et al. (1996) indicated that the presence of such large numbers of parasites in the haemolymph may block haemal sinuses in the gills and appears to compromise oxygen delivery to the tissue of the host. The rates of oxygen consumption of the haemolymph were approximately five times greater and oxygen carrying capacity was approximately 50% less in heavily-infected lobsters compared to those of uninfected or less heavily infected lobsters. Also, the haemolymph pH was lower and the L-lactate concentration was significantly higher in infected lobsters indicating that these animals resort, in part, to anaerobic metabolism (Taylor et al. 1996). The main cause of death may be the disruption of gas transport and tissue anoxia caused by proliferation of large numbers of dinoflagellate cells in the haemolymph.
The uninucleate stages within the haemolymph likely correspond to the sporoblast stages described by Appleton and Vickerman (1998). Presumably, these give rise to the masses of bi-flagellated dinospore stages (macrospore and microspore forms) described by Field and Appleton (1995) and Appleton and Vickerman (1998) which were encountered in some lobsters (Stentiford and Neil 2011). Dinospores have been shown to exit the infected N. norvegicus via the gills and joint membranes and death occurring soon after (Appleton and Vickerman 1998). Stentiford and Shields (2005) indicated that despite its simultaneous production and active en masse exit from heavily infected hosts, the dinospore is not necessarily the infective stage but may be an intermediate stage preceding a resting cyst or some other non-parasitic stage. They also speculated that amphipods may act as alternate or reservoir hosts for this parasite (Stentiford and Shields 2005). However, in N. norvegicus, sporulation of Hematodinium occurs just prior to the primary moulting period and appears associated with transmission of the disease (Field et al. 1992, 1998). Stentiford et al. (2001c) showed that the synchrony of the moulting seasons between male and female lobsters plays an important role in the pattern of infection within a particular year. Specifically, in years when moulting was fairly synchronous, a sharp peak of patent infection was seen and in years when moulting was asynchronous, an extended lower-level 'plateau' of prevalence occurred. Also, peak patent infection prevalence of 70% were found in some trawl samples which seasonally coincided with the annual moult (Field et al. 1992). Although these studies suggest that the moult stage is an important period for obtaining infection, data is circumstantial, particularly considering the apparently extended incubation period to patent disease and the difficulty in relating an infection event to a mortality event or season (Stentiford and Shields, 2005).
Stentiford et al. (1999a, 2001c) and Briggs and McAliskey (2002) detected a seasonal pattern of patent infection with peak prevalences occurring in the spring (April and May) and prevalence levels seemed to be greatest in recently moulted N. norvegicus. Infection may be influenced by factors directly related to host age (immature lobsters) rather than size but is highest in small N. norvegicus (mean carapace length less than 30 mm) and in females (Stentiford et al. 2001c, Briggs and McAliskey 2002). However, low level infections have been found in apparently uninfected lobsters at all times of the year. Beevers et al. (2012) used a suite of assays over a nearly 2-yr period to show that a combination of assay insensitivity and variable parasite dynamics led to the erroneous conclusion of a highly seasonal occurrence of infected hosts; instead, it appears to be the intensity of infection, rather than prevalence per se, which varies among seasons (Behringer et al. 2012). Also, light Hematodinium infections were found using PCR assays when patent infections were at their most prevalent and intense, suggesting that infection develops at different rates in different N. norvegicus individuals and that only a portion of the total number of infected N. norvegicus die within a single year (Beevers et al. 2012). Using a nested PCR assay, Eigemann et al. (2010) found a high prevalence (between 45 and 87.5%) of Hematodinium in decapods with no morphological signs of disease and suggested that Hematodinium sp. may be more common than previously believed (assuming that the DNA detected originated from viable and infectious parasite cells) and that the hosts could develop fatal disease if physiologically stressed from other factors.
Hematodinium sp. effects the carbohydrate metabolism of N. norvegicus by reducing the concentrations of glucose in the haemolymph and glycogen in the hepatopancrease with a concomitant increase in the crustacean hyperglycemic hormone plasma concentrations, thereby placing a heavy metabolic load on the lobster (Stentiford et al. 2001d). Gornik et al. (2010) determined that the chronic stress induced by parasite infection, resulted in a complete depletion of muscle glycogen (and consequently the failure of post-mortem glycolytic fermentation), almost complete depletion of arginine phosphate, significantly altered the adenylate energy charge, and strongly influenced the post-mortem biochemical processes. In addition, the concentrations of several haemolymph free amino acid (FAA) in N. norvegicus, notably serine, were reduced in early infections, while at later stages of infection several FAAs were increased in concentration. The most significant change was in taurine, which increased 13-fold with a relative contribution to the total FAA of 41.6% (Stentiford et al. 1999b). Some FAAs (e.g., taurine and glutimate) are neuro-transmitters or neuro-modulator in N. norvegicus suggesting that infection could affect the behaviour and locomotion of N. norvegicus, thus significantly altering their ability to evade capture, both by predators and by trawlers (Stentiford et al. 1999b). Specifically, N. norvegicus showed a progressive decline in overall swimming performance as infection severity increased, with reductions in the number of tail-flips performed, the number of swimming bouts, average velocity over the swimming bout and the total distance travelled by swimming (Stentiford et al. 2000a). Also, infected lobsters spent more than ten-times increase in the percentage of the day out of the burrow compared to uninfected lobsters which would increase the availability of infected lobsters to trawlers and predators (Stentiford et al. 2001b). The resulting increased catchability of infected lobsters relative to their uninfected counterparts may lead to considerable overestimation of true prevalence of the disease on a particular fishing ground. However, if infected animals are more susceptible to predation through a reduced ability to escape, then the infection may not necessarily add to overall natural mortality, but rather replace a proportion of it (Stentiford et al. 2001c). Nevertheless, disease may contribute a significantly higher proportion of natural mortality than traditionally allowed for in fisheries models (Stentiford and Neil 2011).
Field et al. (1992), Field and Appleton (1995) and Stentiford et al. (2000b) described parasite-induced damage of muscle fibres in N. norvegicus, including parasites penetrating the fibre interstices in late infection and disrupting the peripheral regions of the fibres. Field et al. (1992) claimed that severe infection has an adverse effect on meat quality that has provoked comment from fishermen and processors. However, Stentiford and Neil (2011) indicated that the relative lack of effect of disease on the abdominal musculature is perhaps fortuitous for the fresh dead and frozen ‘tailed’ market for N. norvegicus because the product appeared not to be greatly altered, at least in terms of mechanical integrity. Nevertheless, in trials with trained sensory panels, considerable differences were detected between samples of tail meat from uninfected and heavily-parasitised lobsters with the latter being described as bland in flavour with an after-taste, slightly less firm and less chewy (i.e., more friable or sloppier in texture), and overall less palatable; but surprisingly no bitter taste (as for bitter crab disease in Chionecetes spp.) was reported (Stentiford and Neil 2011, Albalata et al. 2012). These results indicate that Norway lobsters heavily infected with Hematodinium are of inferior marketing quality even after the tails have been cooked (Albalata et al. 2012).
Gross Observations: Heavily-parasitised N. norvegicus can be recognised externally via pronounced hyperpigmentation (usually a vivid dull orange discolouration) of the carapace, particularly of the chelipeds and cephalothorax and an increased opacity attributed to milky-white haemolymph that is visible through the translucent cuticle of the ventral abdomen. Examination for this abnormal colouration has been called the body colour diagnostic method and provides a rapid assessment of advance stages of the disease (Stentiford et al. 2001c, Briggs and McAliskey 2002, Stentiford and Neil 2011). While this method remains useful for the detection of advanced infections, it does not detect low-level ‘sub-patent’ or potentially latent (low-level, tissue-based) infections in N. norvegicus (Stentiford et al. 2001c, Stentiford and Neil 2011). The hepatopancreas often has an obvious green colouration instead of the normal brown. Infected N. norvegicus usually have a reduction in swimming performance (Stentiford et al. 2000a), altered burrow-related behaviour patterns (Stentiford et al. 2001b), and difficulty in walking or supporting themselves. Severely affected individuals were often moribund.
Wet mount: The haemolymph contains nonmotile dinoflagellate cells similar in size to the haemocytes (5-14 µm in diameter, flagellated stages have not yet been observed in lobsters). In N. norvegicus, dense aggregations of the parasite appear as darkened areas within the pleopods that are removed from the lobster and examined under transmitted light of a compound microscope. This pleopod diagnostic method was consistent and effective in identifying patent infections and the degree of cell aggregation visible in the pleopod was used to measure the severity of infection (Field and Appleton 1995). In comparison to the body colour diagnostic method, the pleopod diagnostic method provided reliable, rapid and a relatively transferable tool for infection assessment, detects earlier stages of infection and gives a more accurate estimation of prevalence in field-caught lobsters (Stentiford et al. 2001c). Despite recent advances in antibody and molecular based diagnostic methodologies, the pleopod diagnostic method remains a useful staging technique (to mearure infection severity) for use by field and laboratory scientists (Stentiford and Neil 2011).
Histology: Tissue and organ samples from N. norvegicus (normally heart, hepatopancreas, cheliped muscle, abdominal muscle, gonad and gill) are excised and fixed in Davidson’s seawater fixative (several other fixatives are also suitable for histology of crustacean tissues) for 24 h before transfer to 70% industrial methylated alcohol for storage prior to processing. Fixed samples are then prepared for histology using standard protocols and stained using haematoxylin and eosin. Hematodinium are easily diagnosed in histological sections due to their condensed chromatin profiles that stain densely with haematoxylin (Stentiford and Neil 2011). In heavy infections, the Hematodinium occurs in the haemal spaces of all the major organs. The majority of the parasites were uninucleate with many undergoing mitosis and multinucleate plasmodial and filamentous (vermiform, ‘Gorgonlocks’) trophonts (with up to 5 nuclei) were also observed (Field et al. 1992). In N. norvegicus, the filamentous and plasmodial trophonts have an apparent predilection for the hepatopancreas with the parasites in close association with and possibly attached to the basal lamina of the hepatopancreatic tubules (Stentiford and Shields 2005). In addition, Field and Appleton (1995) described syncitial networks containing prominent condensed chromosomes attached to host tissue bounding haemal spaces and ramifying between muscle fibres in the abdominal muscle and heart. Field and Appleton (1995) state that despite the presence of network-like parasite syncytia between muscle fibres and the association of uninucleate forms with the sarcolemmal membrane, histology of the abdominal musculature remained largely normal in appearance, even in advanced stages of infection. Host cellular defence reactions include haemocyte encapsulation in the gills and heart, and phagocytosis of Hematodinium by the fixed phagocytes in the hepatopancrease (Field and Appleton 1995).
Culture: A characteristic cycle of development including the development of two types of flagellated uninucleated dinospores has been produced in vitro in 10% foetal calf serum in balanced Nephrops saline with added antibiotics at 8 °C. Both dinospore types (macrospores and microspores) germinated to produce multinucleate unattached filamentous (vermiform, ‘Gorgonlocks’) trophonts after 5 weeks in fresh medium. The trophonts (28 to 190 µm long) have characteristic flexing and longitudinal contraction movements and multiply by fragmentation (branching and fission). This form of growth has been subcultured indefinitely at 2 week intervals. If not subcultured, the filamentous trophonts gave rise to colonies of radiating filaments which became attached to the substratum to form web-like multinucleate (plasmodial) trophonts. These trophonts developed into sporoblasts which synthesized trichocysts and flagella and produced dinospores to initiate an new cycle (Appleton and Vickerman 1996, 1998). Developmental changes in the life cycle proceed above 8°C, but were retarded when culture temperatures exceed 15°C (Appleton and Vickerman 1998, Stentiford and Shields 2005).
Electron Microscopy: Small samples (about 2 mm3) of hepatopancreas should be fixed in 3% glutaraldehyde in 0.1 M sodium citrate buffer (pH 7.4) with 1.75% sodium chloride for 2 hours at room temperature followed by post-fixation in reduced 1% osmium tetroxide for 1 hour at 4 °C . Uranyl acetate staining will define irregularly shaped uninucleate and multi-nucleate stages (6 to 10 µm in diameter) of Hematodinium based upon their characteristic nuclei with condensed chromatin profiles, cytoplasmic trichocysts and a bounding alveolar membrane (Stentiford and Neil 2011). Although the nuclei with condensed chromatin were charasteristic of those in dinoflagellates, not all specimens contained trichocysts in the cytoplasm (Field and Appleton 1995). The parasite cells are bounded by characteristic amphiesma (a complex cell covering observed in some dinoflagellates) that lacked thecal plates and microtubules to reinforce the innermost membrane (Field et al. 1992). Micropores (about 110 to 200 nm in diameter), organelles for endocytosis (a cytostome function), were observed on the surface of filamentous trophonts (vermiform plasmodium) and amoeboid trophonts of Hematodinium in vitro and in vivo from the gonad of N. norvegicus (Appleton and Vickerman 1996). The abundant mitochondria have tubular cristae characteristic of dinoflagellates and chloroplasts were absent. Infection in the deep abdominal flexor muscle fibres of N. norvegicus caused alterations in sarcolemmal structure, and localized disruption of myofibrillar bundles around the periphery, but not throughout the centre of the fibres (Stentiford et al. 2000b). However, in contrast to the abdominal musculature, the structure of the cheliped muscles of infected N. norvegicus was significantly altered (similar to that seen in infected crabs (Stentiford et al., 2002)), even during early stages of infection (Stentiford and Neil 2011).
Immunological Assay: An indirect fluorescent antibody technique (IFAT) was developed to detect Hematodinium sp in the haemolymph and tissues of N. norvegicus (Field and Appleton 1996). This technique was more sensitive than gross observations (body colour diagnostic method) and wet mount examinations (pleopod diagnostic method) and was capable of detecting low level haemolymph infections as well as previously undiagnosable tissue infections. The polyclonal antibody IFAT of Field and Appleton (1996) was developed into an immunoblotting assay (Western blot) which was determined to be 10 times more sensitive than the pleopod diagnostic method of Field and Appleton (1995) (Stentiford et al. 2001a). Small et al. (2002) also used the antibody production procedures of Field and Appleton (1995) to develope an enzyime-linked immunosorbant assay (ELISA) which gave the same results as IFAT and was more sensitive than the immunoblotting assay with the added advantages of simplicity and the ability to assay multiple samples within a short period of time. Beevers et al. (2012) used a modified version of this ELISA (in addition to PCR assays) to detect subpatent infections. However, Stentiford et al. (2002) and Bushek et al. (2002) showed that the polyclonal antibodies used in these assays can cross react with epitopes found on other protozoan parasites including Hematodinium sp. from five species of crabs as well as two other species of parasitic dinoflagellates and to Perkinsus marinus in oysters. Stentiford and Neil (2011) cautioned that care must be taken when applying these techniques, particularly where the background pathogen fauna of the host is not well known.
DNA Probes: For Hematodinium spp., all published efforts have exclusively focused upon segments of the ribosomal RNA gene complex, which is present in the nuclear genome of eukaryotes as tandemly repeated clusters of highly conserved genes encoding the small subunit (SSU or 18S), 5.8S, and large subunit (LSU) genes, which are separated by highly variable spacer sequences, the first and second internal transcribed spacers (ITS1 and ITS2) (Small 2012). The development of a polymerase chain reaction (PCR) based diagnostic test was first investigated in Hematodinium by Hudson and Adlard (1994, 1996). A set of Hematodinium-specific PCR primers based on ribosomal DNA that amplify a 380 bp product from Hematodinium-infected N. norvegicus was described (Small et al. 2006). These primers detected Hematodinium in co-habiting amphipods (Orchomene nanus) from the Clyde Sea area of Scotland and Carcinus maenas from the English Channel (Small et al. 2006). Subsequently, Small et al. (2007b) used these primers to conduct a phylogenetic analysis using the sequence of ITS1 of the Hematodinium from N. norvegicus, Cancer pagurus, and Pagurus bernhardus from 4 locations in the United Kingdom and from the Hematodinium sp. infecting Chionoecetes opilio from the province of Newfoundland and Labrador, Canada and suggested that these crustaceans are infected with the same species of Hematodinium. Length variability of the ITS1 region was observed (324 to 345 bp) and attributed to 4 variable microsatellite regions within the sequenced ITS1 fragment. Small et al. (2007b) indicated that the observed variation may be due to co-infection of the host crustacean with several different strains of Hematodinium or differences among copies of ITS1 region within the genome of a single parasite cell. Using these and other previously published primers, Eigemann et al. (2010) described a nested PCR to detect Hematodinium in five species of crustacea from the east coast of Denmark and west coast of Greenland and indicated a higher detection sensitivity of the nested PCR in comparison to the standard PCR assay described by Small et al. (2006). Although Hematodinium were detected in 5.8% (n = 52) and 65% (n = 20) of the N. norvegicus using standard and nested PCR, respectively, none (n = 72) of the lobsters were identified as being infected by both the body colour and pleopod diagnostic methods (Eigemann et al. 2010). Hamilton et al. (2011) used different primers in a nested PCR and determined that Hematodinium DNA occurs in the water column and is harboured by planktonic organisms, including larval stages of the crustacean hosts, when infections are at their lowest in adult hosts. However, the presence of DNA does not indicate that the organism is viable or that it is metabolically active (Hamilton et al. 2011). Beevers et al. (2012) used these and other previously published primers to detect subpatent infections.
Biochemical Characterisation: The taurine:serine ratio in the haemolymph of N. norvegicus determined by reverse phase high performance liquid chromatography (HPLC) may provide a sensitive diagnostic measure of patent Hematodinium infections (Stentiford et al. 1999b). Also, unlike the Hematodinium sp. from the blue crab (Callinectes sapidus), the Hematodinium sp. from N. norvegicus secrets the enzyme acid phosphatase (AP) and AP activity was localized to cytoplasmic granules and on the membranes surrounding the cell nucleus (Small et al. 2007a). Small et al. (2007a) suggested that the pattern of activities of this and other enzymes may be useful in distinguishing among different species of Hematodinium.
Methods of control
No known methods of prevention. However, anecdotal evidence suggests that some fishing practices may help to spread diseases. Such practices include the culling or disassembly of the catch at sea, re-baiting with infected animals and moving animals between locations (culling while underway). Control strategies include culling on station, culling or removing dead animals to onshore fertilizer processing plants, limiting transportation of live animals, and changing baiting practices. Changes in fishing policies may also be warranted. Some evidence from the N. norvegicus fishery in Scotland suggests that the population structure of a given fishery is related to the prevalence of Hematodinium infection within that fishery, with populations of small, size-matched individuals having the highest prevalence (Stentiford et al. 2001c). As such, fisheries management regimes to promote normal size distributions within the fishery have potential for averting epizootics (Stentiford and Neil 2011). Finally, with the advent of live shipping of lobsters to distant markets, there is an increased potential for the inadvertent introduction of pathogenic agents to new regions (Stentiford and Shields 2005). However, the current lack of taxonomic clarity in the specific identity of Hematodinium from various crustacea in coastal areas of the North Atlantic makes if difficult to investigate the global epizootiology of this important pathogen and to calculate risk factors for its transfer to naïve geographical locations and hosts (Steintiford and Neil 2011).
Albalat, A., S.G. Gornik, N. Beevers, R.J.A. Atkinson, D. Miskin and D.M. Neil. 2012. Hematodinium sp. infection in Norway lobster Nephrops norvegicus and its effects on meat quality. Diseases of Aquatic Organisms 100: 105-112.
Appleton, P.L. and K. Vickerman. 1996. Presence of apicomplexan-type micropores in a parasitic dinoflagellate, Hematodinium sp. Parasitology Research 82: 279-282.
Appleton, P.L. and K. Vickerman. 1998. In vitro cultivation and developmental cycle in culture of a parasitic dinoflagellate (Hematodinium sp.) associated with mortality of the Norway lobster (Nephrops norvegicus) in British waters. Parasitology 116: 115-130.
Beevers, N.D., E. Kilbride, R.J.A. Atkinson and D.M. Neil. 2012. Hematodinium infection seasonality in the Firth of Clyde (Scotland) Nephrops norvegicus population: a re-evaluation. Diseases of Aquatic Organisms 100: 95-104.
Behringer, D.C., M.J.I.V. Butler and G.D. Stentiford. 2012. INTRODUCTION: Disease effects on lobster fisheries, ecology, and culture: overview of DAO Special 6. Diseases of Aquatic Organisms 100: 89-93.
Briggs, R.P. and M. McAliskey. 2002. The prevalence of Hematodinium in Nephrops norvegicus from the western Irish Sea. Journal of the Marine Biological Association of the United Kingdom 82: 427-433.
Bushek, D., C.F. Dungan and A.J. Lewitus. 2002a. Serological affinities of the oyster pathogen Perkinsus marinus (Apicomplexa) with some dinoflagellates (Dinophyceae). The Journal of Eukaryotic Microbiology 49: 11-16.
Eigemann, F., A. Burmeister and A. Skovgaard. 2010. Hematodinium sp. (Alveolata, Syndinea) detected in marine decapod crustaceans from waters of Denmark and Greenland. Diseases of Aquatic Organisms 92: 59-68.
Field, R.H. and P.L. Appleton. 1995. A Hematodinium-like dinoflagellate infection of the Norway lobster Nephrops norvegicus: observations on pathology and progression of infection. Diseases of Aquatic Organisms 22: 115-128.
Field, R.H. and P.L. Appleton. 1996. An indirect fluorescent antibody technique for the diagnosis of Hematodinium sp. infection of the Norway lobster Nephrops norvegicus. Diseases of Aquatic Organisms 24: 199-204.
Field, R.H., C.J. Chapman, A.C. Taylor, D.M. Neil and K. Vickerman. 1992. Infection of the Norway lobster Nephrops norvegicus by a Hematodinium-like species of dinoflagellate on the west coast of Scotland. Diseases of Aquatic Organisms 13: 1-15.
Field, R.H., J.M. Hills, R.J.A. Atkinson, S. Magill and A.M. Shanks. 1998. Distribution and seasonal prevalence of Hematodinium sp. infection of the Norway lobster (Nephrops norvegicus) around the west coast of Scotland. ICES Journal of Marine Science 55: 846-858.
Gornik, S.G., A. Albalat, R.J.A. Atkinson, G.H. Coombs and D.M. Neil. 2010. The influence of defined ante-mortem stressors on the early post-mortem biochemical processes in the abdominal muscle of the Norway lobster, Nephrops norvegicus (Linnaeus, 1758). Marine Biological Research 6: 223-238. (For electronic publication see: http://www.tandfonline.com/doi/pdf/10.1080/17451000903147468).
Hamilton, K.M., D. Morritt and P.W. Shaw. 2010. Genetic diversity of the crustacean parasite Hematodinium (Alveolata, Syndinea). European Journal of Protistology 46: 17-28.
Hamilton, K.M., I.F. Tew, R.J.A. Atkinson and E.C. Roberts. 2011. Occurrence of the parasite genus Hematodinium (Alveolata: Syndinea) in the water column. Journal of Eukaryotic Microbiology 58: 446-451.
Hudson, D.A. and R.D. Adlard. 1994. PCR techniques applied to Hematodinium spp. and Hematodinium-like dinoflagellates in decapod crustaceans. Diseases of Aquatic Organisms 20: 203-206.
Hudson, D.A. and R.D. Adlard. 1996. Nucleotide sequence determination of the partial SSU rDNA gene and ITS1 region of Hematodinium cf. perezi and Hematodinium-like dinoflagellates. Diseases of Aquatic Organisms 24: 55-60.
Jensen, P.C., K. Califf, V. Lowe, L. Hauser and J.F. Morado. 2010. Molecular detection of Hematodinium sp. in Northeast Pacific Chionoecetes spp. and evidence of two species in the Northern Hemisphere. Diseases of Aquatic Organisms 89: 155-166.
Mortensen, S., I. Arzul, L. Miossec, C. Paillard, S. Feist, G. Stentiford, T. Renault, D. Saulnier and A. Gregory. 2007. Molluscs and crustaceans, 220.127.116.11 Hematodinium infections in crustaceans. In: Raynard, R., T. Wahli, I. Vatsos, S. Mortensen (eds.) Review of disease interactions and pathogen exchange between farmed and wild finfish and shellfish in Europe. VESO on behalf of DIPNET, Oslo. pp. 426-433.
Small, H.J. 2012. Advances in our understanding of the global diversity and distribution of Hematodinium spp. – Significant pathogens of commercially exploited crustaceans. Journal of Invertebrate Pathology 110: 234-246.
Small, H.J. and K.M. Pagenkopp. 2011. Reservoirs and alternate hosts for pathogens of commercially important crustaceans: A review. Journal of Invertebrate Pathology 106: 153-164.
Small, H.J., S. Wilson, D.M. Neil, P. Hagan and G.H. Coombs. 2002. Detection of the parasitic dinoflagellate Hematodinium in the Norway lobster Nephrops norvegicus by ELISA. Diseases of Aquatic Organisms 52: 175-177.
Small, H.J., D.M. Neil, A.C. Taylor, R.J.A. Atkinson and G.H. Coombs. 2006. Molecular detection of Hematodinium spp. in Norway lobster Nephrops norvegicus and other crustaceans. Diseases of Aquatic Organisms 69: 185-195.
Small, H.J., J.D. Shields, D.M. Neil, A.C. Taylor and G.H. Coombs. 2007a. Differences in enzyme activities between two species of Hematodinium, parasitic dinoflagellates of crustaceans. Journal of Invertebrate Pathology 94: 175-183.
Small, H.J., J.D. Shields, J.A. Moss and K.S. Reece. 2007b. Conservation in the first internal transcribed spacer region (ITS1) in Hematodinium species infecting crustacean hosts found in the UK and Newfoundland. Diseases of Aquatic Organisms 75: 251-258.
Stentiford, G. 2006. Hematodinium spp. - parasitic dinoflagellates of crustaceans. In: DIPnet - Disease Interactions and Pathogen exchange between farmed and wild aquatic animal populations - a European network, p. 2.
Stentiford, G.D. and D.M. Neil. 2011. Diseases of Nephrops and Metanephrops: a review. Journal of Invertebrate Pathology 106: 92-109.
Stentiford, G.D. and J.D. Shields. 2005. A review of the parasitic dinoflagellates Hematodinium species and Hematodinium-like infections in marine crustaceans. Diseases of Aquatic Organisms 66: 47-70.
Stentiford, G.D., D.M. Neil and R.J.A. Atkinson. 1999a. Infection by the dinoflagellate Hematodinium in the Norway lobster (Nephrops norvegicus L.) on the west coast of Scotland, United Kingdom. Journal of Shellfish Research 18: 334. (Abstract).
Stentiford, G.D., D.M. Neil and G.H. Coombs. 1999b. Changes in the plasma free amino acid profile of the Norway lobster Nephrops norvegicus at different stages of infection by a parasitic dinoflagellate (genus Hematodinium). Diseases of Aquatic Organisms 38: 151-157.
Stentiford, G.D., D.M. Neil, R.J.A. Atkinson and N. Bailey. 2000a. An analysis of swimming performance in the Norway lobster, Nephrops norvegicus L. infected by a parasitic dinoflagellate of the genus Hematodinium. Journal of Experimental Marine Biology and Ecology 247: 169-181.
Stentiford, G.D., D.M. Neil and G.H. Coombs. 2000b. Alterations in the biochemistry and ultrastructure of the deep abdominal flexor muscle of the Norway lobster, Nephrops norvegicus during infection by a parasitic dinoflagellate of the genus Hematodinium. Diseases of Aquatic Organisms 42: 133-141.
Stentiford, G.D., D.M. Neil and G.H. Coombs. 2001a. Development and application of an immunoassay diagnostic technique for studying Hematodinium infections in Nephrops norvegicus populations. Diseases of Aquatic Organisms 46: 223-229.
Stentiford, G.D., D.M. Neil and R.J.A. Atkinson. 2001b. Alteration of burrow-related behaviour of the Norway lobster, Nephrops norvegicus during infection by the parasitic dinoflagellate Hematodinium. Marine and Freshwater Behaviour and Physiology 34: 139-156.
Stentiford, G.D., D.M. Neil and R.J.A. Atkinson. 2001c. The relationship of Hematodinium infection prevalence in a Scottish Nephrops norvegicus population to season, moulting and sex. ICES Journal of Marine Science 58: 814-823.
Stentiford, G.D., E.S. Chang, S.A. Chang and D.M. Neil. 2001d. Carbohydrate dynamics and the crustacean hyperglycaemic hormone (CHH): effects of parasitic infection in Norway lobsters (Nephrops norvegicus). General and Comparative Endrocrinology 121: 13-22.
Stentiford, G.D., M. Green, K. Bateman, H.J. Small, D.M. Neil and S.W. Feist. 2002. Infection by a Hematodinium-like parasitic dinoflagellate causes Pink Crab Disease (PCD) in the edible crab Cancer pagurus. Journal of Invertebrate Pathology 79: 179-191.
Taylor, A.C., R.H. Field and P.J. Parslow-Williams. 1996. The effects of Hematodinium sp.-infection on aspects of the respiratory physiology of the Norway lobster, Nephrops norvegicus (L.). Journal of Experimental Marine Biology and Ecology 207: 217-228.
Bower, S.M. (2013): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Hematodinium sp. of Norway Lobster.
Date last revised: February 2013
Comments to Susan Bower
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