Effects of experimental infection of juvenile edible crabs Cancer pagurus with the parasitic dinoflagellate Hematodinium sp.
KEY WORDS: dinoflagellate, Hematodinium, edible crab, Cancer pagurus, hemocytes
Infections caused by the parasitic dinoflagellete, Hematodinium spp. have been described in a wide variety of crustacean hosts, mainly within the Northern Hemisphere (see reviews by Stentiford & Shields 2005, Morado 2011, Small 2012) although infected animals have also been reported in the Southern Hemisphere in Australia (Hudson & Shields 1994) and China (Xu et al. 2007) implying a global distribution of this genus. There have been extensive studies on the epizootiology of this disease in a range of commercially important crustaceans including Alaskan Tanner crabs Chionoecetes bairdi (e.g., Meyers et al. 1987), blue crabs Callinectessapidus (e.g., Messick 1994, Messick & Shields 2000), snow crabs Chionoecetes opilio (e.g., Shields et al. 2005), Norway lobsters Nephrops norvegicus (e.g., Field & Appleton 1995, Beevers et al. 2012, Stentiford & Neil 2011), and velvet swimming crabs Necora puber (Wilhelm & Mialhe 1996). These have shown that epizootics caused by Hematodinium are likely to have significant effects on the commercial viability of crustacean populations in some locations because of the high prevalence and severity of the disease (Wilhelm & Mialhe 1996, Lee & Frischer 2004, Siddeek et al. 2010, Morado et al. 2012).
Despite our detailed understanding of the ecology of this important disease, less is known about the relationship between host and parasite and its method of transmission, and the possibility of other vectors playing some role in this event. It is believed that transmission of Hematodinium occurs as a result of dinospore formation and infected animals surrounded by "clouds" of these presumed motile infective stages have been reported (Stentiford & Shields 2005). Furthermore, some experiments, albeit small scale, have described that naive blue crabs (Callinectes sapidus) rapidly die after exposure to seawater contaminated with dinospores (Frischer et al. 2006). Because of the inherent difficulties in producing dinospores for challenge experiments, a number of workers have used artificial transmission by either intrahemocoelic injection of trophonts and plasmodial stages from infected tissues (e.g., Meyers et al. 1987, Messick & Shields 2000, Shields & Squyars 2000, Shields et al. 2005, Coffey et al. 2012) or feeding of naive crabs with infected tissues (Walker et al. 2009, Li et al. 2011). The results of such approaches have been highly variable in terms of infectivity and the time taken for host death postchallenge. For example, intrahemocoelic injection of blue crabs (C. sapidus) with varying numbers of trophonts or plasmodia found that death occurred between 17 (Messick & Shields 2000) and 40 (Shields & Squyars 2000) days postchallenge. Similar approaches with snow crabs (Chionoecetes opilio) in colder water reported 50% mortality but only after 99 days (Shields et al. 2005). Other experiments, however, have observed rapid mortality postchallenge with Hematodinium in various crustaceans. For example, Frischer et al. (2006) found that two blue crabs exposed to dinospores from a naturally infected animal, died after only 4 days post-exposure although these animals were not subject to any histopathologic examination to confirm the cause of death. Overall, although there are only a limited number of studies often using small numbers of test animals, it can be concluded that the time taken from challenge to death of the host may depend on both the nature of the parasites used for challenge and the host species under study (Small 2012).
The edible or brown crab Cancer pagurus is a commercially important crustacean in northern Europe. This species is fished in many regions of the United Kingdom and Ireland and estimated landings in the United Kingdom alone reached over 24,000 t by 2010 (Bannister 2011). Edible crabs are subject to a wide range of diseases including "pink crab disease" caused by Hematodinium sp. (Latrouite et al. 1988, Stentiford et al. 2002, Stentiford 2008, Rowley et al. 2014, Smith et al. 2015). Field-based studies suggest that juvenile (prerecruit) edible crabs when living on-shore are highly susceptible to Hematodinium sp. probably due to their short-molt intervals (Shields et al. 2005), resulting in the significant levels of disease recently recorded at two sites in South Wales and that transmission of infection may take place in the late fall to winter with death occurring several months to a year later (Smith et al. 2015). Hence, unlike the artificial disease transmission studies reported for other crustaceans, such as snow crabs and blue crabs, this condition may take a different timescale to overwhelm juvenile edible crabs. Therefore, this study reports on experiments where prerecruit, C. pagurus were challenged with Hematodinium sp. by intrahemocoelic injection and the progression of resulting infection examined with regards to mortality, changes in the numbers of hemocytes and parasites in the blood, and histopathology of infected tissues.
MATERIALS AND METHODS
Collection, Initial Screening, anti Maintenance of Animals
Juvenile edible crabs (Cancer pagurus) were collected from the intertidal zone at low water (0.5 m above Chart Datum) from Mumbles Head, Swansea Bay, United Kingdom in January 2013 and immediately transported back to Swansea University. After a few days under aquarium conditions to acclimatize, crabs were sized, sexed, and were bled from the junction of the carapace and the third walking leg to check for the presence of Hematodinium or bacteria or fungi. Hemolymph was placed directly (i.e., without smearing) onto microscope slides and observed under phase-contrast microscopy. One hundred microlitres of hemolymph was also drawn into a 1 mL syringe filled with an equal volume of 8% paraformaldehyde + 3.4% NaCl solution and stored at 4[degrees]C for later hemocyte counts. Thirty apparently Hematodinium-free crabs, as based on the absence of these parasites in the hemolymph, were randomly assigned into two groups each consisting of 15 animals and for the duration of the experiment these were kept in individual containers in larger tanks in a recirculating seawater aquarium system. Crabs in the group for challenge with Hematodinium ranged in size from 41 to 66 mm carapace width (CW), with a mean size of 55 [+ or -] 1.8 mm (mean [+ or -] SD). The control animals ranged in size from 46 to 69 mm CW and had a mean CW of 55 [+ or -] 1.6 mm. Each group contained six male and nine female crabs. Animals were fed three times a week on blue mussel (Mytilus edulis) or white fish except for sampling days. All crabs were examined daily and a record taken of those that had not consumed any of the feed given. Molting frequency and subsequent increase in CW were also recorded. Aquarium temperatures were measured weekly and ranged from 10 to 18[degrees]C during the timespan of the experiment. The experiment commenced in January 2013 and finished in February 2014.
Preparation of Hematodinium for Challenge
A donor edible crab collected from the same locality as the recipients and with a heavy Hematodinium infection (Grade 4 severity--see Smith et al. 2015 for details of grading of severity) had 2.5 mL hemolymph aseptically removed into 2.5 mL sterile ice-cold 3% NaCl solution. This was then transferred into a 9 cm plastic Petri dish and the hemocytes left to attach to the surface of the dish for 10 min at room temperature (RT). After gentle mixing to dislodge any nonattached cells (mainly Hematodinium), the solution containing these parasites was decanted into a new dish and a further 5 mL of sterile 3% NaCl solution added and the attachment process repeated on two more occasions. The cell suspension is then removed, centrifuged (850 x g, 10 min at RT), and the pellet was resuspended in 10 mL 3% sterile NaCl solution. The numbers of Hematodinium were counted using an improved Neubauer hemocytometer and the concentration adjusted to 5 x [10.sup.6] [mL.sup.-1]. The parasites in the inoculum consisted of a mixture of uni- and multi-nucleate trophonts with no dinospores or vermiform plasmodia. There were a small number (<10% of total cells) of contaminating hemocytes in this inoculum. Sterile 3% NaCl solution was used for injection into the control group of crabs.
Injection and Sampling
Crabs were surface sterilized with 70% ethanol and the 15 animals in the experimental group were injected with 100 [micro]l of sterile 3% NaCl solution containing Hematodinium (total 5 x [10.sup.5] parasites/crab) at the junction of the third walking leg and carapace. Control animals (15 individuals) had 100 [micro]l of sterile 3% NaCl injected into the same area. Subsequently, all crabs were bled usually every 4 wk and 100 [micro]l of hemolymph placed into 100 [micro]l of 8% paraformaldehyde + 3.4% NaCl solution for the estimation of total hemocyte and Hematodinium numbers. A further small drop of blood (ca. 20 [micro]l) was also examined using phase-contrast microscopy for the presence of Hematodinium and the relative percentage of parasites and hemocytes determined by counting a minimum of 250 cells over several fields of view. Animals that died during the experiment were routinely processed for histology as described in detail by Smith et al. (2013). Briefly, gills and hepatopancreas from crabs were fixed in Davidson's seawater fixative for ca. 24 h, dehydrated in graded series of industrial methylated spirit, cleared in Histoclear, and embedded in paraffin wax. Sections (ca. 7 pm) were stained with Cole's hematoxylin and eosin and photographed with an Olympus BX41 microscope. Crabs that were judged to be moribund when they showed little or no response to stimuli (i.e., no eye retraction, antennal movement or the ability to right themselves when turned on their dorsal surface) were also sacrificed for histology. Crabs that had clear symptoms of postmortem changes (e.g., putrefaction) could not be processed for histology. The experiment was concluded when all animals that had become Hematodinium positive had died (day 378 postchallenge) and at this time all remaining crabs were also sacrificed for histology.
Data were analyzed using Prism v. 5 (GraphPad Software, San Diego, CA) using one-way or two-way analysis of variances (ANOVA) with Bonferroni's multiple comparison posttests. Kaplan-Meier survival analyses were carried out using SPSS v. 19. Data are shown as means [+ or -] SEM.
Of the 15 animals injected with Hematodinium, only five (33%) became infected during the course of the experiment (Table 1). Four of these animals were found to have small numbers of trophont-like parasites in their blood after 56 days postchallenge with the remaining animal (#44) having Hematodinium parasites free in the hemolymph from ca. 3 mo postchallenge. Hence the time taken from challenge to the appearance of parasites in the hemolymph takes 2-3 mo. One of the control animals (#57), found to be Hematodinum negative during initial screening, also developed a natural Hematodinium infection first seen when a hemolymph preparation was examined at 28 days postchallenge. This animal died after 156 days of the experiment (Table 1).
Initially, Hematodinium appeared in the hemolymph as individual trophont-like cells (Fig. 1A) that, unlike the hemocytes, did not attach to the glass slides. As the disease progressed, these parasites appeared to form clumps containing variable numbers of cells (Fig. 1B, C). In one infected crab (#53, see Table 1), long, thin, ameboid stages of the parasite morphologically similar to the vermiform plasmodia described previously (Shields & Squyars 2000) were seen after 355 days postchallenge and these increased in frequency until the death of the host at 378 days (Fig. 1D-F). No sporoblasts or dinospores of Hematodinium were observed in any tissues examined.
Figure 2 shows representative examples of the changes in the total hemocyte numbers and Hematodinium during the experiment in an artificially infected individual (crab #53; Fig. 2A) and a saline-injected control (crab #70; Fig. 2B).There was little variation in the total hemocyte counts over the course of the experiment in both the examples shown in Figure 2A, B and in all 30 crabs in the experiment (data not shown). However, in the example of an infected animal shown, after ca. 250 days the number of parasites steadily increased with a concomitant fall in the number of circulating hemocytes (Fig. 2A).
The mean time of death for Hematodinium-infected crabs was 192 [+ or -] 115 days (n = 5). Kaplan-Meier survival analysis showed that there was no significant difference between the groups [i.e., experimental animals, which became infected, experimental animals, which did not become infected, and control (saline-injected) animals; P > 0.05; Fig. 3].
Table 1 summarizes the moult frequency, final size, and times and potential causes of death of animals that were in the trial. The last of the five Hematodinium positive crabs died after 378 days postchallenge and the six remaining live animals (three from the control group and three from the Hematodinium injected but uninfected cohort) were also sacrificed and subject to histologic analysis. Histologic examination of all of the crabs throughout the experiment was not possible as some had obvious postmortem changes that precluded such examination. Forty percent (eight out of 20) of the crabs examined using histology, regardless of their Hematodinium status, had a mikrocytid infection of the antennal gland, gills, and hemolymph, 15% had systemic bacterial infections and 15% had systemic fungal infections caused by Ophiocordyceps. Four of the five (80%) of the Hematodinium-infected animals were examined using histology and three of these animals were found to have coinfections (two caused by a mikrocytid parasite and one by a systemic bacterial infection) and 50% of these (crabs #44 and #53; Table 1) had apparent damage to the gills that may have impaired their respiratory activity. For example, crab #53 that died 378 days postchallenge, had extensive damage to one set of gills (Fig. 4A) where histologic examination revealed widespread necrosis and apparent loss of blood flow as evidenced by a lack of circulating hemocytes in the branchial stem and secondary lamellae (Fig. 4B). Similarly, crab #44 that died 154 days postchallenge had no evidence of any coinfections but contained clumps of Hematodinium at the junction between the branchial stem and secondary lamellae of the gills (Fig. 4C) potentially occluding blood flow.
The cases of infection caused by a mikrocytid parasite seen in both Hematodinium-infected and noninfected crabs were characterized by the presence of free plasmodia in the lumen of the antennal gland and large numbers of uninucleate forms within the antennal gland cells (Fig. 4D). In those cases, in which, this infection was thought to be the cause of death, there was extensive alteration of the antennal gland integrity resulting in disorganized and necrotic tissues probably due to the multiplication of the parasites and large numbers of uninucleate and plasmodial stages of the parasite free in the hemolymph (Fig. 4D, E). The systemic nature of these high severity infections was evidenced by the appearance of the parasites in the nephrocytes in the gills (Fig. 4F) and the fixed phagocytic cells surrounding blood vessels in the interstitial space of the hepatopancreas (Fig. 4G).
Of particular interest was the histologic examination of the six crabs that survived the experiment and were sacrificed at 378 days postchallenge. Three of these crabs had moderate to high severity mikrocytid infections characterized by variable necrosis of the antennal gland and the presence of plasmodia and uninucleate forms of these parasites in the gills (not shown). Two of these crabs (#41 and #42) also had low-grade fungal (Ophiocordyceps) infections characterized by encapsulated fungi in the hepatopancreas (Fig. 5A, B) and gills, but with few or no free fungi in circulation. The other three crabs sacrificed at day 378 displayed no histopathogical alterations of tissues associated with the presence of bacteria, fungi, or mikrocytid parasites (not shown).
During the study, the frequency of molting and the resulting size increase of crabs were noted (see Table 1). Because of the small number of infected crabs in this experiment, it was not possible to determine if there was any relationship between the frequency of molting in infected versus uninfected animals. It was, however, noted that the final Hematodinium-infected crab to die after 378 days of 83 mm CW had only molted once at 100 days, whereas the remaining six crabs (uninfected by Hematodinium) at this same time point had molted two or three times with a resulting mean CW of 99.3 [+ or -] 9.1 cm (mean [+ or -] SD, n = 6). Furthermore, Hematodinium-infected crabs #46 and #44 that died at 114 and 154 days postchallenge, respectively, failed to molt during the experiment whereas 91% of uninfected crabs with the exception of two animals (#43 and #68 neither of which molted before death) had molted after 84 days. Notably, those Hematodinium-infected animals that did molt did so before the infection became established in the hemolymph (crab #51 moulted after nine days and #53 molted after 100 days; Table 1). After the first molt the resulting CWs were not significantly different between Hematodinium-infected and uninfected crabs (two-way ANOVA, P > 0.05, data not shown).
This study has shown that artificially induced Hematodinium infections in juvenile edible crabs (Cancer pagurus) manifest themselves in the hemolymph after ca. 2 mo postchallenge and that diseased animals may survive for over a year after showing initial signs of infection. Both the length of the latent phase and the time taken for mortality to occur is in close agreement with the model for the development of Hematodinium sp. in juvenile edible crabs from the field-based studies (Smith et al. 2015). Although the numbers of infected animals in the experiment were small (five individuals, 33% of challenged crabs) Hematodinium did not appear to be the sole cause of death in any of these crabs and there was no evidence of dinospore production sometimes characteristic of terminal infections (Stentiford & Shields 2005, Coffey et al. 2012). Instead, secondary infections, such as that caused by a mikrocytid parasite (probably Paramikrocytos canceri) similar to that recently reported in edible crabs from a variety of locations in the United Kingdom (Hartikainen et al. 2014) and bacterial and fungal septicemia (sepsis), were judged to be the primary cause of death in some individuals. Others, although showing no obvious sign of other infections, appeared to die as a result of either respiratory impairment because of obstruction of blood flow in the gills or a failure to molt successfully. Meyers et al. (1987) also suggested that organ or respiratory dysfunctions are primary factors contributing to host mortality in Hematodinium-infected infected animals. Similarly, Shields et al. (2003) reported that blue crabs (Callinectes sapidus) naturally infected with Hematodinium showed "metabolic exhaustion" as evidenced by reductions in concentrations of hemolymph-based hemocyanin and glycogen reserves in the hepatopancreas. Stentiford et al. (2001) also observed marked reductions in plasma glucose and hepatopancreatic glycogen with increasing severity of Hematodinium infections in the Norway lobster Nephrops norvegicus. Hence, in juvenile edible crabs Hematodinium appears to be a chronic infection that causes a limited reduction in the numbers of circulating hemocytes resulting in reduced clotting and defense abilities that probably leaves these crabs susceptible to other infections (Rowley et al. 2015).
One of the main observations of our study was that the survival rates for crabs either infected or uninfected with Hematodinium did not differ. It must be stressed, however, that our experiments were based on small numbers of animals (n = 15) with a low level of infectivity (33%) and under these conditions statistical analysis of the data becomes difficult. Table 2 summarizes the results of several other aquarium-based studies designed to determine the mortality of various species of crustaceans postchallenge with Hematodinium spp. Whereas some of these showed significantly higher rates of mortality in the infected animals in comparison with the controls (e.g., Shields & Squyars 2000, Shields et al. 2005) others were unable to make such conclusions as they either made no use of uninfected controls (Messick & Shields 2000) or were based on very small numbers of test animals (e.g., Meyers et al. 1987, Frischer et al. 2006). Finally, it should be stressed that aquarium-based experimental challenge with various life history stages of the parasite other than with the naturally infectious dinospores, may not adequately reflect the outcome of natural infections in the wild.
The time taken from challenge to the appearance of Hematodinium in the hemolymph was significantly greater in our study than many reports on other species of crabs (see Table 2). For instance, in some experiments, various stages of Hematodinium have been reported to be present in the hemolymph as early as only two (Walker et al. 2009) or three (Coffey et al. 2012) days postchallenge whereas we found a latent phase of infection in edible crabs of at least 2 mo. The only similar result to the present study comes from the experiments of Meyers et al. (1987) in Tanner crabs (Chionoecetes bairdi) where infection by Hematodinium was first seen as late as 55-91 days postchallenge. Furthermore, the latter study reported that these infected crabs were still alive after 148 days postchallenge suggesting a protracted development of parasites that was probably linked to the low water temperature (4-9[degrees]C) that this experiment was conducted under. Further analysis of the previously published experiments reviewed in Table 2 shows a surprising lack of data on mortality rates in both Hematodinium positive and negative animals such that only a few studies (e.g., Shields & Squyars 2000, Shields et al. 2005) have provided compelling evidence of mortality caused by the direct presence of these parasites. Furthermore, to date ours is the only study to examine the histopathology of such animals at the time of death to ensure that the cause of this is actually infection by Hematodinium and not secondary infections caused by other pathogens or parasites.
Crabs that were subject to histologic examination revealed that ca. 45% of these animals were affected by a mikrocytid parasite that initially infects the epithelial cells in the antennal gland and in high severity infections spreads to others tissue via the hemolymph. Because this infection could not be identified by any changes in external features or behavior of live crabs, and no polymerase chain reaction (PCR)-based methods for their detection in hemolymph were available when this experiment commenced (Thrupp et al. 2013), we were unable to screen for its presence. As it has been observed in juvenile edible crabs collected at Mumbles Head (Smith & Rowley unpublished observations), it is probable that the mikrocytid parasites were present in some crabs at the onset of the experiment. This infection was first reported by Bateman et al. (2011) in edible crabs collected from the English Channel and at that time it was thought to be caused by a "haplosporidian-like" parasite. More recently, Thrupp et al. (2013) found that between 15% and 70% of juvenile edible crabs from two sites in Pembrokeshire (South West Wales, United Kingdom) close to our current collection site, were affected by the same condition. They found that heavily infected crabs showed evidence of infiltration of the parasites into other tissues such as the gills but they were unable to prove that this condition could cause the death of the host. The antennal gland produces waste products from the hemolymph that are voided through openings close to the antennae (Friere et al. 2008). Thrupp et al. (2013) suggested that infections caused by mikrocytid parasites may damage the antennal gland and because it also regulates fluid volumes within crabs, this could cause mortality especially during ecdysis when a large volume of water is taken up by crabs to help remove the old carapace (Chung et al. 1999). Although it may be premature to propose that this infection can cause host mortality, one of the crabs (#68; see Table 1) that failed to molt had large numbers of uninucleate and plasmodial forms of mikrocytid parasites in the hemolymph with evidence of necrosis in the epithelial cells of the antennal gland.
A further infection that we observed in crabs in our experiment was caused by a fungus believed to belong to the genus, Ophiocordyceps (Smith et al. 2013). This fungal infection was first reported by Stentiford et al. (2003) as a coinfection in Hematodinium-infected crabs, perhaps suggesting that the presence of the latter agent is necessary for fungal infection. However, a more recent survey of disease status of edible crabs in South Wales found examples of Ophiocordyceps infections in Hematodinium-negative animals (Smith et al. 2015). Furthermore, all three crabs with this fungal infection in our current experiment were Hematodinium negative showing that this fungus is a primary pathogen in its own right. One of these crabs (#67; see Table 1) was thought to be killed by this infection because hemolymph preparations made from this animal a few days before its death showed large numbers of fungal elements in the hemolymph with only small numbers of circulating hemocytes. The potential interaction between Ophiocordyceps and Hematodinium infections is not well understood. Although initially it was suggested that the presence of Hematodinium facilitated the development of the fungus in the host (Stentiford et al. 2003), experiments where edible crabs with or without low severity Hematodinium infections were challenged with Ophiocordyceps, revealed that fungal growth was actually lower in those animals with the dinoflagellate parasites (Smith et al. 2013). As high-severity infections with Hematodinium result in a marked reduction in hemocyte numbers, it would be expected that this would leave crabs more vulnerable to septicemia caused by microbial agents such as fungi and bacteria (Rowley et al. 2015).
Crabs in the experiment molted up to three times over the course of the experiment. The crabs with Hematodinium infections appeared to stop molting or only molted before the infection became pronounced in the blood. Meyers et al. (1987) have also suggested that Tanner crabs infected with Hematodinium are unlikely to undergo successful ecdysis. In our experiment one of the Hematodinium-infected crabs apparently died as a result of incomplete molting but unfortunately because of the postmortem state of this animal at the time of examination, we were unable to examine it histologically to determine if other infectious agents were present. If indeed the presence of Hematodinium in crabs inhibits molting, then this merits investigation to find a mechanistic explanation. The "metabolic exhaustion" described by some authors (e.g., Shields et al. 2003) as symptomatic of such infections may be one answer, whereas hormonal changes, such as those described by Stentiford et al. (2001), offer a further potential explanation for the putative interference of molting induced by Hematodinium.
In conclusion, our studies, albeit carried out on small cohorts of edible crabs, imply that Hematodinium is a long-term chronic infection that on its own is not a significant driver of host mortality at least under our aquarium conditions. As suggested by other authors (e.g., Stentiford & Shields 2005) this infection leaves it host immunocompromised resulting from the metabolic stress induced in the host leaving it susceptible to other secondary infections.
This work was supported in part by the ERDF Interreg 4A Ireland Wales project, SUSFISH, awarded to AFR. ALS was supported by the College of Science. Swansea University under a postgraduate bursary. We gratefully acknowledge the help of Dr. Emma Wootton, Dr. Ed Pope, and Dr. Claire Vogan with crab collection in the field.
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AMANDA L. SMITH AND ANDREW F. ROWLEY *
Department of Biosciences, College of Science, Swansea University, Swansea SA2 8PP, Wales, United Kingdom
* Corresponding author. E-mail: firstname.lastname@example.org
TABLE 1. Summary table showing of infectivity, moulting frequency, and times and potential causes of mortality in edible crabs. Hematodinium Time of Animal # Sex Group infection status death (days) 61 Male Control No 30 64 Female Control No 82 45 Male Experimental Yes 93 68 Male Control No 111 46 Female Experimental Yes 114 60 Male Control No 131 43 Male Experimental No 146 47 Female Experimental No 146 59 Male Control No 149 44 Female Experimental Yes 154 57 Female Control Yes 156 48 Male Experimental No 191 49 Male Experimental No 196 51 Female Experimental Yes 219 56 Female Control No 219 65 Female Control No 247 62 Male Control No 252 54 Male Experimental No 257 55 Female Experimental No 257 63 Female Control No 280 67 Female Control No 285 52 Female Experimental No 303 41 Female Experimental No 378 42 Male Experimental No 378 50 Female Experimental No 378 53 Female Experimental Yes 378 58 Female Control No 378 69 Female Control No 378 70 Male Control No 378 Potential cause of death (ascertained by histology Animal # and gross structure) 61 Cause unknown 64 Cause unknown * 45 Moulting defect * 68 Moulting defect with large numbers of mikrocytid plasmodia in hemolymph 46 Bacterial septicaemia 60 Cause unknown * 43 Cause unknown * 47 Cause unknown * 59 Cause unknown * 44 Respiratory impairment due to Hematodinium infection? 57 Antennal gland necrosis due to mikrocytid infection and Hematodinium 48 Antennal gland necrosis caused by mikrocytid infection 49 Infection by mikrocytid parasites 51 Combination of extensive Hematodinium and systemic mikrocytid infection 56 Cause unknown 65 Cause unknown 62 Mikrocytid infection of antennal gland and other tissues 54 Cause unknown * 55 Gill necrosis with bacterial infection 63 Cause unknown * 67 Fungal infection 52 Cause unknown * 41 Sacrificed--mikrocytid infection and low grade fungal infection 42 Sacrificed--mikrocytid infection and low grade fungal infection 50 Sacrificed--mikrocytid infection 53 Extensive gill necrosis and secondary bacterial infection 58 Sacrificed--apparently healthy 69 Sacrificed--apparently healthy 70 Sacrificed--apparently healthy Number of moults Starting Size at death Animal # during experiment size (CW; mm) (CW; mm) 61 0 51 51 64 1 61 80 45 0 (1 at death) 51 51 68 0 (1 at death) 59 59 46 0 48 48 60 1 69 85 43 0 61 61 47 1 51 65 59 1 46 65 44 0 60 60 57 0 49 49 48 1 61 85 49 1 41 56 51 1 59 74 56 2 50 75 65 1 62 85 62 2 51 83 54 2 54 80 55 1 55 65 63 1 51 69 67 1 57 71 52 1 50 62 41 2 59 96 42 2 60 110 50 2 52 90 53 1 66 83 58 2 57 90 69 3 49 100 70 3 53 110 * Animal not subject to histologic analysis. TABLE 2. Summary of results from infectivity trials of crustaceans with Hematodinium spp. Crab (stage, sex) Challenge route Infective dose (per crab) Callinectes sapidus Intrahemocoelic 10 plasmodia C. sapidus (adult, Intrahemocoelic 1 x [10.sup.3] and 1 x female) [10.sup.5] mainly plasmodia C. sapidus Intrahemocoelic 1 x [10.sup.5] ameboid trophonts C. sapidus Oral Infected tissue, 5 g/crab C. sapidus (juvenile Oral Infected crab tissue, and adults) [10.sup.4] 1.5 x [10.sup.6] parasites/ crab C. sapidus Exposure to water Not reported containing dinospores Chionoeeetes bairdi Intrahemocoelic 1 x [10.sup.5] to 2 x [10.sup.6] dinospores from culture C. bairdi Intrahemocoelic Infected hemolymph Chionoeeetes opilio Intrahemocoelic 1 x [10.sup.5] Cancer pagurus Intrahemocoelic 1 x [10.sup.5] mainly (juvenile, male trophonts and female) Mortality (%) and Crab (stage, sex) timescale Observations Callinectes sapidus 100% mortality after No uninfected 55 days controls in experiment C. sapidus (adult, 86% after 40 days; Death not associated female) median time of with sporulation of death 30 days parasite; some crabs may be immune to parasite C. sapidus 33%-39% crabs became Small number of crabs infected in trial showed rapid depending on proliferation of salinity of water parasites in blood crabs held in after only 3 days C. sapidus 33% by 48 h post 73% of crabs became challenge infected; no uninfected controls C. sapidus (juvenile Low (1.7% overall) no Infections found and adults) different to thought to be control crabs fed spontaneous uninfected tissue resulting from natural "silent" infections; no evidence of infection as a result of challenge C. sapidus 100% (total of 2 Putative infected animals) after 4 crabs were PCR days positive for Hematodinium Chionoeeetes bairdi Mortality not Crabs held at recorded but crabs 4-9[degrees]C became infected 55-69 days postchallenge. Crabs still alive after 148 days postchallenge C. bairdi Mortality not Crabs held at recorded but 100% 4-9[degrees]C crabs infected, first seen in blood after 83-91 days postchallenge Chionoeeetes opilio 50% mortality after -- 99 days. Median time of death 91 days, infection first seen in hemolymph after 2 wk Cancer pagurus 100% of infected No significant (juvenile, male animals died after difference between and female) 378 days, mean time mortality in of death 191.6 infected and days; only 33% of uninfected crabs crabs became infected postchallenge Crab (stage, sex) References Callinectes sapidus Messick and Shields (2000) C. sapidus (adult, Shields and Squyars (2000) female) C. sapidus Coffey et al. (2012) C. sapidus Walker et al. (2009) C. sapidus (juvenile Li et al. (2011) and adults) C. sapidus Frischer et al. (2006) Chionoeeetes bairdi Meyers et al. (1987) C. bairdi Meyers et al. (1987) Chionoeeetes opilio Shields et al. (2005) Cancer pagurus This study (juvenile, male and female)
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|Author:||Smith, Amanda L.; Rowley, Andrew F.|
|Publication:||Journal of Shellfish Research|
|Date:||Aug 1, 2015|
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