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Hematodinium sp. (Bitter Crab Disease)

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Category 2 (In Canada and of Regional Concern)

Common, generally accepted names of the organism or disease agent

Bitter crab disease, Bitter crab syndrome.

Scientific name or taxonomic affiliation

Hematodinium-like dinoflagellate. Nucleotide sequence analysis of the partial small subunit (SSU) rDNA gene and ITS1 region of several isolates indicated that the Hematodinium sp. in Chionoecetes sp. is a different species from Hematodinium perezi in Callinectes sapidus and from the Hematodinium sp. in Nephrops norvegicus (Hudson and Adlard 1996). However, as indicated below, taxonomic issues pertaining to species identification and susceptible host identity continue to be unresolved by 2012 (Small 2012). Also, because transmission experiments are problematic due to the yet unknown nature of the route of infection and the inability to experimentally generate infections under laboratory conditions in many species of crustaceans know to be infected in natural settings, the ability to address specific identity of the parasite is limited (Small and Pagenkopp 2011).

Geographic distribution

Gulf of Alaska along Southeastern Alaska, USA, including Alitak Bay on the south end of Kodiak Island (Urban and Byersdorfer 2002); Bering Sea; boundary area of the Chukchi Sea and the Arctic Ocean (for details see Meyers et al. 1996 and Morado 2011); west coast of Vancouver Island, British Columbia, Canada (Bower et al. 2003); east coast of Newfoundland and Labrador, Canada including the southern Labrador shelf south to the northern Grand Bank (with highest prevalence (12.6-25%) extending from coastal bays offshore to the middle shelf) and occassionally on the southern Grand Bank (Taylor and Khan 1995; Dawe 2001, 2002; Dawe et al. 2010; Shields et al. 2005); and the southern coast of Nova Scotia, Canada (Morado et al. 2010). Eigemann et al. (2010) detected DNA of Hematodinium in 46% of Chionoecetes opilio (n = 100) from the west coast of Greenland using nested polymerase chain reaction (PCR) based technology but none of the C. opilio (n = approximately 14,000) examined by the colour diagnostic method were found infected. However, a low prevalence (less than 0.2%) of visually infected C. opilio were encountered from that region during earlier surveys (Morado et al. 2010).

Host species

Chionoecetes bairdi, Chionoecetes opilio, Chionoecetes tanneri (Bower et al. 2003) and Chionoecetes angulatus (Jensen et al. 2010). Ryazanova (2008) and Ryazanova et al. (2010) reported Hematodinium sp. in less than 1% of Lithodidae crabs, Paralithodes camtschaticus and Paralithodes platypus, from the west coast of Kamchatka, the northeastern area of the Sea of Okhotsk from August 25, 2006 to December 15, 2006. As for infected Chionoecetes spp., the meat of the Lithodidae crabs infected with Hematodinium had a bitter astringent taste (Ryazanova 2008, Ryazanova et al. 2010). A Hematodinium sp. was also detected in a Lithodes couesi specimen captured near Vancouver Island, Canada and in the majid crab, Hyas coarctatus, from the Bering Sea (Jensen et al. 2010).

Hematodinium spp. have been described from other marine crustaceans including other species of crabs from the Atlantic Ocean, in Norway lobsters, and in crabs from the vicinity of Australia and China, and from amphipods (Stentiford 2006, Small 2012).

Impact on the host

Infected crabs have drooping limbs and mouthparts, milky-white haemolymph and when cooked, the muscle has a chalky texture and an astringent after-taste that makes them unmarketable. Due to the flavour this infection imparts to the flesh, this condition is known as Bitter Crab Disease (BCD) or Bitter Crab Syndrome (Meyers et al. 1987, 1990, Morado et al. 2012). Small numbers of diseased crab can render an entire batch of crabs unpalatable if batch processed (Meyers et al. 1987, Taylor and Khan 1995). In terms of economic effect, species of Hematodinium have the potential for impact at the level of the population, the fishery and the market (Stentiford and Shields 2005). Severe economic losses (estimated to be as high as $3 million US in 1987) have been attributed to this parasite on the C. bairdi fishery, primarily localized to a few impacted fjords in southeastern Alaska (Meyers et al. 1987, 1990). Evidence from the C. opilio fishery in the bays of Newfoundland shows the establishment and catastrophic outbreaks of Hematodinium. For example, in Conception Bay, prevalences increased from 3.7% in male crabs in 1992-1993 (Taylor and Khan 1995) to over 9% of males and 25% of females in an epizootic occurring in 2000 (Pestal et al. 2003, Shields et al. 2005) and highest at 23.9% in males and 18.8% in females during the outbreak in 2005 causing concern to the industry (Shields et al. 2007). In laboratory studies, mortality rates of 50 to 100% over several months have been reported for naturally infected laboratory-held C. bairdi (Meyers et al. 1987, Love et al. 1993) and C. opilio (Shields et al. 2005).

Haemolymph surveys of C. bairdi in Alaska indicated that the prevalence and intensity of infection increased through spring, peaked in summer, from June through August with peak mortalities in August and September, and then declined into fall and winter as infected crabs died resulting in a prevalence of infection near zero by mid-winter (Meyers et al. 1990, Eaton et al. 1991, Love et al. 1993). Meyers et al. (1990) and Love et al. (1993) found that sporogony of Hematodinium in C. bairdi was seasonal with sporulation occurring in the summer months. High prevalence of infection (50-80%) were common in C. opilio from Norton Sound and west of St Lawerence Island but the parasite was relatively rare in C. bairdi (about 2%) and C. opilio (about 4%) in the Eastern Bering Sea (Morado et al. 2000). Subsequent samples from the Bering and Chuckchi seas revealed higher prevalences of infection (greater that 30%) in both species of crabs from some areas with higher prevalences of infection in C. opilio from higher latitudes (Morado 2011). Morado et al. (2000) and Urban and Byersdorfer (2002) noted a higher prevalence of infection in female crabs from Alaska but of more significance was the higher prevalence of infection observed in smaller crabs when compared to the overall size distribution in the samples (Morado et al. 2012). Conversely at some locations in southeast Alaska, Bednarski et al. (2010) reported that prevalence of patent Hematodinium infections increased as shell size increased in ‘new shell’ (recently moulted) male C. bairdi. In addition, they indicated that the reproductive success of new shell females infected with Hematodinium was reduced by 11% as indicated by visual examination of clutch fullness (Bednarski et al. 2010).

In Newfoundland prior to the outbreak of 2003-2005, surveys for Hematodinium in C. opilio indicated that female crabs had a significantly higher prevalence of patent (macroscopically diagnosed) infection than males (Taylor et al. 2002, Dawe 2002, Pestal et al. 2003, Shields et al. 2005). Among the males Hematodinium was most prevalent in the intermediate size group (41 to 59 mm in carapace width) (Dawe 2002). However, during the outbreak of 2003-2005, there was an apparent shift in the dynamics of the infection with a higher prevalence of infection in males (23.9%) than females (18.8%) and bimodal prevalence values in males with peak prevalence in smaller (35 mm carapace width) and largest crabs (about 130 mm carapace width) (Shields et al. 2007). Dawe (2002) indicated that the relationships between catch rates in trawl-caught samples and prevalence of patent infections with Hematodinium in C. opilio from the continental shelf of Newfoundland and southern Labrador showed no clear evidence of either density dependence or an effect on natural mortality by the parasite. Nevertheless, Mullowney et al. (2011) determined that in these crab populations, the density of small to medium-sized, ‘new shell’ C. opilio was directly related to prevalence and distribution of Hematodinium and much of the spatio-temporal variability in disease expression was a function of variability in host productivity, growth, and movement.

Many investigators reported that patent infections occurred predominantly in ‘new shell’ crabs (Meyers et al. 1990; Eaton et al. 1991; Dawe 2002; Urban and Byersdorfer 2002; Shield et al. 2005, 2007; Bednarski et al. 2010, Mullowney et al. 2011). Possibly, Hematodinium transmission occurs during the crab moulting process (Meyers et al. 1990, Eaton et al. 1991, Shield et al. 2005). Because smaller (younger) crabs moult more frequently, they would be more vulnerable to infection than larger (older) crabs. However, although adult crabs do not molt as frequently, their cumulative exposure to the parasite is longer, creating more chance exposures via wounds, appendage loss, or shell erosion (Bednarski et al. 2010). Also, sporulation of Hematodinium occurs after the primary spring moulting period for these crabs and the higher prevalence in ‘new shell’ crabs was presumably not due to their acquiring recent infections because infections were thought to take at least 15 to 18 months to develop (Meyers et al. 1990). Nevertheless, infections in C. opilio appear to take 9 to 12 months to develop with moulting more clearly associated with transmission than it is in C. bairdi (Shields et al. 2005). Shields et al. (2007) attributed the 2003-2005 outbreak of Hematodinium in Conception Bay, Newfoundland to rising temperatures stimulating moulting activity which led to an increase in susceptible host in the population. Morado et al. (2000) suggested that this parasite may have an impact on crab recruitment.

Research sampling in both Alaska and Newfoundland indicated that the hydrography of bays may contribute to the epizootic as infections were centered within the deeper confines of the bay (Urban and Byersdorfer 2002, Shields et al. 2005, Dawe et al. 2010). Outbreaks in C. opilio have been associated with fjords with shallow sills or otherwise constricted areas (Meyers et al. 1987, 1990; Eaton et al. 1991; Pestal et al. 2003; Shields et al. 2005). However, outbreaks are also known from more open ocean regions (Meyers et al. 1996). Here, depth may facilitate the concentration of infectious stages since female C. opilio from greater than 250 meters had prevalences almost twice as high as those from shallower areas (Pestal et al. 2003) and infections were rare at depths less than 200 meters (Shields et al. 2005). Habitat or substrate type was important to the parasite in C. opilio as prevalence was highest in crabs from mud/sand habitats compared to other habitats (Shields et al. 2005). Such habitat restrictions hint at the presence of alternate hosts or dietary factors. In many of these cases, the enclosed areas had somewhat high-density, ‘closed’ populations, relatively high potential for entrainment of water, and stressful conditions such as seasonal hypoxia or predation pressure (Stentiford and Shields 2005). In open ocean systems, such as those for C. bairdi and C. opilio populations in the Bering Sea, the prevalences of Hematodinium were variable, with most but not all regions exhibiting low prevalences (Meyers et al. 1996).

Muscular degeneration occurs in C. opilio that are heavily infected with Hematodinium but the nature of this pathology remains to be determined (Stentiford and Shields 2005). A reduction in hemocyanin concentration and consequently a major reduction in serum proteins was noted in the plasma of infected C. bairdi (Stentiford and Shields 2005). Septicemia (haemolymph infections with bacteria) commonly associated with stress in crustaceans and unidentified ciliates generally considered facultative parasites have been reported from infected C. bairdi (Meyers et al. 1987, Love et al. 1993). The pronounced haemocytopenia (a decline in haemocytes) associated with Hematodinium infection hinders the normal immune response (e.g., clotting, phagocytosis, encapsulation of foreign material and other antibiotic factors) and may fascilitate secondary infections of opportunist pathogens (Mortensen et al. 2007). Dinospores have been shown to exit the infected host via the gills in C. bairdi which died within hours of sporulation (Love et al. 1993).

Diagnostic techniques

Gross Observations

In crabs with patent infections, the cuticle usually has slightly different red colour in comparison to cohorts (cooked appearance) and with a pinkish white discolouration at the appendage joints. This change in morphology caused by heavy infections was called the colour diagnostic method. The haemolymph is opaque (cloudy or ‘milky'), musculature emaciated and heavily infected crabs are listless or lethargic in behaviour (Taylor and Khan 1995). A positive result from macroscopic examination is definitive, if the observer is well trained (Pestal et al. 2003). The haemolymph of heavily infected crabs also shows a distinct lack of clotting ability (Meyers et al. 1987). While this method remains useful for the detection of advanced cases in heavily infected hosts, it does not detect low-level ‘sub-patent’ nor potentially sub-patent (low-level, tissue-based) infections (Stentiford and Shields 2005). Ryazanova et al. (2010) indicated that no carapace color change was observed in heavily infected lithodid crabs, but diseased crabs possessed creamy-yellow hemolymph, which was visible through the arthrodial membranes of the abdomen and appendages.

Figure 1.Ventral surface of two small (sublegal, non commercial size) male Chionoecetes tanneri. The specimen at the top of the image was heavily infected with Hematodinium sp. and the other specimen was not infected with this parasite. Image provided by G. Meyer, DFO Pacific.

Figure 2. Ventral surface of two commercial-size male Chionoecetes tanneri with left set of appendages removed and flipped to demonstrate the whitish dicolouration of the gills (g) and opaque haemolymph (h) pooled in the carapace of a specimen heavily infected with Hematodinium sp. (top specimen) in comparison to an uninfected crab. Image provided by Greg Workman, DFO Pacific.

Wet Mounts

Numerous non-motile trophonts (vegetative stage) in the haemolymph. Although the trophonts are similar in size and shape to some crab haemolymph cells and easily confused with haemocytes by the uninitiated, their sheer abundance and highly granular appearance in heavily infected crabs are clues to diagnosis. Aggregates of multinucleate plasmodia with vesicular (frothy) cytoplasm may occur in some crabs (Urban and Byersdorfer 2002). During July through October, prespores and two types of motile dinospores occur in the haemolymph of C. bairdi from Alaska. Macrodinospores (about 13 -µm in size) and microdinospores (about 8 µm in size) were initally circular upon removal from crabs but gradually become more elongated (ovoid) over a 12 hour period in a wet mount preparation (Meyers et al. 1987, Eaton et al. 1991). Sporulation is the last stage of infection resulting in Tanner crab death. The morphologically different forms were observed in C. bairdi after injection with Hematodinium isolated from naturally infected crabs (Eaton et al. 1991).


Haemolymph samples, either drawn with a 22 guage needle on a 3 milliter syringe from the juncture of the basis and ischium of the cheliped or by detaching the dactyl from a walking leg of small crabs, were placed on acid cleaned poly-l-lysine coated glass microscope slides and fixed immediately in Bouins fixative. Alternately, between 1.2 and 1.5 milliters of haemolymph was drawn into a syringe preloaded with 1 milliter of ice-cold 10% formalin in filtered seawater and refrigerated until aliquots were placed on poly-l-lysine coated glass microscope slide, allowed to set for 45 seconds, fixed in Bouins fixative for 24 hours and transferred to 70% ethanol for holding. All smears were hydrated, stained with Jenner-Giemsa for 10 to 20 minutes, dehydrated through an acetone series, cleared in a xylene series and mounted in cytoseal. Using a microscope, Hematodinium were detected based on cell size, presence of condensed chromatin in the nucleus and the "notched" appearance of the chromosomes (Meyers et al. 1987, Love et al. 1993, Taylor and Khan 1995, Taylor et al. 2002). Air dried haemolymph smears can also be fixed in methanol and stained with other commercially available blood stains (e.g., Diff-QuiK, Baxter; Hemacolor, Merck). This technique confirmed the infection in all C. opilio that had macroscopic signs of disease (9 of 355 crabs examined) and in an additional 7 of 333 apparently uninfected crabs (Taylor et al. 2002, Pestal et al. 2003).


Four distinct morphological forms occur in the haemal spaces of all tissues. The two most commonly occurring forms are the single cell trophonts (6 to 20 µm in diameter) and multinucleate plasmodia (with 2 to about 30 nuclei per plasmodium). Both of these forms have distinctive dinokaryon nuclei (6.3 ± 0.7 µm in diameter with condensed and darkly staining chromatin) and frothy cytoplasm. Plasmodia with less than 6 nuclei are often spheroid but occasionally vermiform in shape. Plasmodia with more than 6 nuclei are usually polymorphous with a lobular surface consisting of separating trophonts (Stentiford and Shields 2005). The other two morphological forms are two different sizes of biflagellated dinospores that occur only during the terminal stage of the infection (Eaton et al. 1991). In Alaska, dinospores were observed in naturally infected crabs during the summer (Love et al. 1993). Similar morphological forms were reported from lithodid crabs, P. camtschaticus and P. platypus (Ryazanova et al. 2010).

Figure 3. Trophonts (T) of various shapes and a plasmodium (P) of Hematodinium sp. with typical frothy cytoplasm and dinokaryon nuclei (arrow indicated in two trophonts) in the connective tissue with a haemocyte (H) of Chionoecetes tanneri. Haematoxylin and eosin stain.

Figure 4. A large pleomorphic plasmodium (P) of somewhat vermiform morphology of Hematodinium sp. in the heart sinus of a heavily infected Chionoecetes tanneri. Haematoxylin and eosin stain.

Figure 5. Hematodinium sp. in the process of binary fission (arrows) in the heart sinus of Chionoecetes tanneri. A crab heamocyte (H) occurs among the abundant parasites in this heavily infected crab. Haematoxylin and eosin stain.

Heavy infections cause dilation of haemal sinuses with massive infiltration by parasitic cells and degeneration of muscular tissues (Stentiford and Shields 2005). Trophonts and pre-spore stages remain in the circulatory system and show no signs of being able to actively penetrate or perforate basement membranes of various tissues until late in the disease (Morado 2011). Wheeler et al. (2007) indicated that in C. opilio with heavy infections of Hematodinium sp., pressure necrosis, due to the build up of high densities of parasites, was the primary histopathological alteration in most tissues. Details of histopathological alterations to the tissues include pressure necrosis in the spongy connective tissues of the hepatopancreas and the blood vessels in most organs. Damage to the gills ranged from apparent thinning of the cuticle, loss of epithelial cells, and fusion of the membranous layers of adjacent gill lamellae. Affected lamellae exhibited varying degrees of distention with a loss of trabecular cells, haemocyte infiltrations, and swelling (clubbing) along the distal margins. Damage was most severe during sporulation when large numbers of zoospores were located along the distal margins of affected lamellae suggesting that sporulation may cause a lysis or bursting of the thin lamellar cuticle, releasing spores (Wheeler et al. 2007).

Electron microscopy

For transmission electron microscopy, samples of heavily infected tissues can be fixed with 2.5% glutaraldehyde in sterile sea water, post-fixed with 1% osmium tetroxide in sea water, dehydehydrated in acetone and embedded in resin (Epon-Araldite). Ultrathin sections are usually stained with uranyl acetate and lead citrate and observed with a transmission electron microscope. The ultrastructure of Hematodinium sp. in lithodid crabs was similar to that reported for Hematodinium from other crustacean hosts (Ryazanova et al. 2010). Specifically, the nuclei of the cells were filled with a granular nucleoplasm along with the polymorphous electron-dense heterochromatin clumps and the nuclear chromatin was dense and V-shaped. Multivesicular bodies and vacuoles with homogeneous contents were located in the granular cytoplasm and the cytoplasm of the polynuclear and mononuclear cells contained electron-dense trichocysts of a rhomboid or a square shape (Ryazanova et al. 2010).

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). The small subunit ribosomal DNA (SSU rDNA) sequence of the Hematodinium sp. from C. tanneri and lithodid crabs was nearly identical to the Hematodinium spp. sequences in GenBank (Bower et al. 2003, Ryazanova et al. 2010). This segment of the genome with primer binding regions in the conserved 18S and 5.8S regions are known to be genus specific and may prove useful for genus specific probes but will not be applicable to differentiating between species. Eigemann et al. (2010) found that the nested PCR approach was more sensitive in detecting Hematodinium infections than conventional PCR in crustacea with no morphological indications of infection.

Based on phylogenetic analysis of the sequences of the first internal transcribed spacer (ITS1) region of the SSU rDNA complex, Small et al. (2007) suggested that the Hematodinium sp. infecting C. opilio from Conception Bay, Newfoundland, Canada and the Hematodinium from Nephrops norvegicus, Cancer pagurus, and Pagurus bernhardus from the United Kingdom were the same species of Hematodinium. Small et al. (2007) indicated that the observed variability in length within the sequenced ITS1 fragments may be due to co-infection of the host crustacean with several different strains of Hematodinium or differences among copies of ITS1 regions within the genome of a single parasite cell. Jensen et al. (2010) concluded that Hematodinium in C. angulatus, C. bairdi, C. tanneri, C. opilio, L. couesi and P. camchaticus are considered to be the same genetic clade (and probably same species) that is widely distributed in other decapods (such as Nephrops norvegicus, Hyas coarctatus, Pagurus bernhardus, Pagurus prideaux, Munida rugosa, Cancer pagurus and Carcinus maenas) from the North Atlantic and North Pacific Oceans but in a clade separate from the Hematodinium sp. infecting the portunoid crabs, Callinectes sapidus from the east coast of the United States, Liocarcinus depurator from the English Channel and Scylla serrata from China. Following analysis of the secondary structures of the ITS1 and ITS2 sequences, Hamilton et al. (2010) determined that Hematodinium from the east and west North-Atlantic is comprised of distinct ribotypes or clades with the Hematodinium from C. opilio being a generalist parasite that occurs in Scottish crustaceans (various species of crabs and N. norvegicus) and is not specific to a particular geographic region. However, Hamilton et al. (2010) also suggested that the presence of a distinct ‘C. opilio’ clade would not be surprising, as C. opilio not only occurs in a different geographic region to the investigated Scottish crustaceans but also at a much greater depth and in colder waters. In support of this suggestion, Eigemann et al. (2010) reported differences in the ITS1 sequence of Hematodinium from C. opilio from the west coast of Greenland and suggested that two of the isolates with aberrant sequences (outliers to all other equivalent Hematodinium sequences from various host) were likely to belong to a unrecognised species.

Methods of control

Management of bitter crab syndrome may be possible by harvesting Chionoecetes spp. in the winter when fewer crabs are severely parasitized and meats are more marketable. This strategy should also reduce the necessity of culling heavily infected crabs which are more prevalent later in the season (Stentiford and Shields 2005). Proper disposal of infected crabs is essential in controlling dissemination of the parasite. Control strategies include culling on station or within a watershed, culling or removing dead animals to onshore fertilizer processing plants, limiting transportation of live animals, and not using potentially infected crabs for bait (Stentiford and Shields 2005). Shields et al. (2005) recommended that fishery management programs for Chionoecetes fisheries employ non-selective gear to monitor for Hematodinium infections in female and juvenile crabs because these under-sampled members of the population may forewarn of impending recruitment declines that might otherwise remain unexplained. Furthermore, the prevalence of Hematodinium in small and medium sized crabs may reflect the relative strength of a recruitment pulse and perhaps could be used as a predictor of medium- to long-term recruitment to a fishery (Mullowney et al. 2011, Morado et al. 2012). As indicated by Siddeek et al. (2010), it is important to include the potential effect of additional mortality that bitter crab disease could have on rebuilding performance of lightly and heavily infected Chionoecetes stocks. Specifically, under the assumed recruitment scenario, the new control rules were adequate to rebuild the depleted lightly infected Eastern Bering Sea C. bairdi stocks, but not the heavily infected southeast Alaska stock (Morado et al. 2012).


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Citation Information

Bower, S.M. (2013): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Hematodinium sp. (Bitter Crab Disease).

Date last revised: February 2013
Comments to Susan Bower

Date modified: