Survey of pathologies in Crassostrea gasar (Adanson, 1757) oysters from cultured and wild populations in the Sao Francisco estuary, Sergipe, Northeast Brazil.
KEY WORDS: oysters, Crassostrea gasar, histopathology, Perkinsus, Bonamia
The southern region of Brazil is the biggest national producer of bivalves, with an annual production of 21,000 t of the brown mussel Perna perna and 2,500 t of the Japanese oyster Crassostrea gigas in 2012 (EPAGRI 2013). Nevertheless, the northeast region of Brazil has a great potential to develop native oyster production along approximately 3,000 km of coastline with calm waters, deep estuarine extensions, and suitable temperatures. Two oyster species with aquaculture potential inhabit those ecosystems: Crassostrea rhizophorae and Crassostrea gasar (de Melo et al. 2010). Currently in northeastern Brazil, the production of oysters is based exclusively on the collection of wild juvenile seed oysters that are then cultivated in grow-out bags. Because the culture of C. gigas is limited by excessive water temperatures in northeastern Brazil, there is an ongoing initiative to produce native oyster species on a large scale there to supply existing Brazilian markets, and new techniques are being developed to improve aquaculture of the native oyster C. gasar (Silveira et al. 2011, Lopes et al. 2013, Ramos et al. 2013).
A critical biosecurity concern for modern bivalve aquaculture operations is management of pathogens that could affect production, including those that are listed by the World Organisation for Animal Health, which include Bonamia ostreae, Bonamia exitiosa, Marteilia rejringens, Perkinsus marinus, and Perkinsus olseni. Bonamiosis is a disease caused by protozoans of the genus Bonamia, which infect hemocytes of oysters (Carnegie & Cochennec-Laureau 2004). The pathogen Bonamia ostreae causes high mortalities among adult flat oysters Ostrea edulis during their final stage of production. Perkinsosis is a disease caused by protozoan parasites of the genus Perkinsus, which are associated with mortalities among diverse molluscs worldwide (Villalba et al. 2004, Villalba et al. 2011).
Although studies on bivalve pathologies are still scarce in Brazil, they have increased during the past 15 y as a result of the expansion of commercial bivalve culture. Several recent investigations have focused on the detection of protozoan parasites listed by the World Organisation for Animal Health (OIE) (da Silva et al. 2009, Sabry et al. 2009, Sabry et al. 2011, da Silva et al. 2012, Brandao et al. 2013, da Silva et al. 2013, Sabry et al. 2013). One of the studies reported for the first time a Perkinsus species (Perkinsus beihaiensis) infecting Crassostrea rhizophorae oysters from Ceara state, on the northeastern coast of Brazil (Sabry et al. 2009, Sabry et al. 2013). Subsequently, Perkinsus marinus infections were reported at high prevalences among C. rhizophorae oysters from Paraiba state, which is located 650 km southeast of Ceara state (da Silva et al. 2013). The current study reports the results of health evaluations of wild and cultured populations of the mangrove oyster Crassostrea gasar from the estuary of the Rio Sao Francisco, Sergipe state, which is located 450 km south of Paraiba state.
MATERIALS AND METHODS
One hundred adult mangrove Crassostrea gasar oysters were collected at each time and site during January, April, July, and September 2010 from a natural population (10[degrees]32'1.20" S, 36[degrees] 31'6.20" W). and from a commercial culture operation (10[degrees] 32'0.60" S, 36[degrees]29'36.24" W) both located in the Sao Francisco River estuary, Sergipe state, northeastern Brazil (Fig. 1). The Sao Francisco River is one of the most important Brazilian water resources, draining seven states along its 2,863-km length. The area of the Sao Francisco River estuary is approximately 192 [km.sup.2], which includes extensive mangrove forests. The estuary is subject to a moderate tidal amplitude (2-4 m). The climate is tropical, with a rainy season from April to August and a dry season from September to March. The native inhabitants of the area support themselves using artisanal methods of fishing, agriculture, and aquaculture (Santos et al. 2014).
Wild sample oysters (height, 72.2 [+ or -] 9.9 mm) were removed from the rhizophores of mangrove trees Rhizophora mangle. For culture, wild juvenile oysters were obtained from natural larval settlements on collection structures (stakes and ropes) and were placed in mesh bags for grow-out. Cultured oysters (height, 73.5 [+ or -] 10.1 mm) were first sampled after 8-10 mo of cultivation. All sample oysters inhabited the intertidal zone. After quarterly sample collections, oysters from both habitats were held up to 24 h in separate tanks of aerated seawater from their sampling sites, at temperatures near 25[degrees]C, until they were processed to obtain tissue samples for diagnostic procedures. Water salinity and temperature were similar at both experimental sites in the Rio Sao Francisco estuary during the current investigation, where salinities ranged from 24-34 and water temperatures ranged from 25-30[degrees]C.
Analyses for Pathological Conditions
Evidence of pathological conditions was first evaluated macroscopically by examining both oyster shells and meats. Cytological hemolymph cell monolayers (30 per sample) were prepared by a modification of the methods of da Silva and Villalba (2004), which used a Giemsa stain, and were examined exhaustively microscopically for the presence of Bonamia sp. pathogens. Histological sections (5 pm) were stained with Mayer's hematoxylin-eosin (30 per sample) for examination by light microscopy (Howard et al. 2004). Prevalences of pathogens, parasites and pathological conditions were calculated as the proportions of affected oysters in each quarterly sample from both wild and cultured oyster populations.
Polymerase Chain Reaction Assays
Recently, an amoeba-like (Order Dactylopodida) ovarian and oocyte parasite was reported in Crassostrea gasar from southern Brazil based on polymerase chain reaction (PCR) and DNA sequencing results without in situ confirmation (Suhnel et al. 2014). Samples from the current study were tested retrospectively by PCR assays for detection of oocyte parasites that included both the Perkinsela amoebae-like organism of Sfihnel et al. (2014) and Marteilioides chungmuensis (Itoh et al. 2003).
DNA from ethanol-preserved ovary tissues of four selected sample oysters was extracted using a Wizard Genomic DNA Purification Kit (Promega) according to the manufacturer's instructions. To test for the presence of the ovarian parasites Marteilioides chungmuensis and Perkinsela amoebae-like organisms, the following primers were used respectively: OPF2/OPR2 (Itoh et al. 2003) and PERK-F/PERK-R (Suhnel et al. 2014).
Polymerase chain reactions of 25 [micro]L contained GoTaq Green Master Mix (Promega), 0.4 [micro]M primers, and 200-390 ng template DNA. Cycling parameters for the Marteilioides chungmuensis PCR were 95[degrees]C for 10 min, 40 cycles of 95[degrees]C for 1 min, 58[degrees]C for 1 min, and 72[degrees]C for 1 min, and were finished with a final extension at 72[degrees]C for 10 min. Cycling parameters for the Perkinsela amoebae-like PCR were 95[degrees]C for 3 min, 40 cycles of 95[degrees]C for 45 sec, 60[degrees]C for 40 sec, and 72[degrees]C for 2 min, and were finished with a final extension at 72[degrees]C for 10 min. Aliquots of each PCR product were separated by electrophoresis on 1 % agarose gels and were checked for the expected products of 672 bp and 1365 bp, respectively.
Differences among measures for commensals and parasites, and pathological conditions for wild and cultured oysters, and for the sampling months (January, April, July, and September), were analyzed by one-way analysis of variance (ANOVA). Percentages were arcsine-transformed to meet ANOVA requirements. One-way ANOVA was run to analyze for differences in oyster size in samples from cultured and wild populations. An index of the overall prevalence of pathological conditions (OPPC) was calculated for each sample type as follows: Prevalences of each condition in each month were ranked from the lowest to the highest. The OPPC of each environment was calculated by month as its mean rank (da Silva et al. 2005). Differences in OPPC between environments were analyzed using the Kruskal-Wallis test. Significance of statistical results were evaluated at P = 0.05, unless specified differently.
Analyses of histological sections from 240 oysters (30 wild and 30 cultured oysters for each seasonal sample) showed the presence of nonspecific pathological conditions and diverse parasitic and symbiotic associates (Table 1). Between cultured and wild oysters, prevalences varied significantly for Perkinsus sp. infections and Polydora sp. infestations only. Between sampling months, only the prevalence for hemocytic infiltration varied significantly. Specific differences are described next in detail.
Frequent hemocytic infiltration and less common granulocytomas (dense focal concentrations of infiltrating hemocytes) occurred in oysters from this study (Table 1), and they were not associated directly with the presence of commensals. Hemocytic infiltration was common among gill and other tissues. At low frequency (3 of 240) hemocytosis affected numerous tissues simultaneously, including gills, connective tissues of mantle viscera, and epithelia of the stomach and intestine. Occasionally (4 of 240), gill architectures were disrupted by heavy hemocyte infiltration. The prevalence of hemocytic infiltration was significantly less in January (18.3 [+ or -] 2.3) and April (31.6 [+ or -] 2.4), and greater in September (65.0 [+ or -] 7.1) and July (83.3 [+ or -] 9.4; ANOVA, P = 0.0024).
Prokaryotes, Fungi, and Cell Alterations
Hypertrophied (50-70 [micro]m) gametocytes containing basophilic intranuclear inclusions and condensed peripheral chromatin, typical of viral gametocytic hypertrophy (Fig. 2), were observed among germinal epithelia of male (n = 5) and female (n = 1) oysters (Table 1). In most cases, there was only one abnormal gametocyte per gonad follicle, and one to five infected gametocytes per section. In contrast, two oysters, one male and one female, showed 12 and 43 hypertrophied gametocytes per section, respectively. There were no host responses against infected gametocytes.
Cytoplasmic colonies of Rickettsia-like organisms (RLO) occurred among the epithelial cells of gills (3%-7%) and digestive gland tubules (7%-20%) of sample oysters (Table 1). Rickettsia-like organism colonies were basophilic and granular, with diameters of 20-25 pm (Fig. 3). The intensity of RLO lesions in both types of infected tissues was generally low (1-4 colonies per section), except in one wild and one cultured oyster that showed 10 and 18 RLO colonies, respectively.
Shell disease (maladie du pied) was observed macroscopically in only 3 of 100 cultured oysters of a single sample (3%; Table 1). The condition was characterized by the presence of rigid blisters of shell and conchiolin materials at the myostracal surface of adductor muscle attachment to the valves, which contained brownish, viscous liquid. In one affected oyster, the adductor muscle attachment was ruptured; in another moribund oyster, the necrotic adductor muscle was detached from the shell.
The gregarine Nematopsis sp. occurred within vacuoles in the connective tissues of gill, labial palp, mantle, and viscera at variable levels of prevalence (3%-70%; Table 1). Oocysts of Nematopsis sp. (length, 10 [micro]m) were surrounded by refractile oocyst walls and contained basophilic, ellipsoid sporozoites (Fig. 4). One to four oocysts occurred per vacuole. Infection intensities ranged from 1-10 oocysts per section.
A microsporidial Steinhausia sp. infected oyster oocytes at variable levels of prevalence (0%-75%; Table 1) and at low intensities of up to 30 infected oocytes per section. Three developmental stages in the life cycle of the parasite were observed. Sporoplasm cells of 9-12 [micro]m in diameter showed a central basophilic nucleus surrounded by nonvacuolated, acidophilic cytoplasm (Fig. 5), and occurred in one to two cells per mature or immature oocyte. Sporont cells of 4-6 [micro]m in diameter showed eccentric nuclei and prominent vacuoles, with up to eight apparent proliferative cells occurring within perinuclear cytoplasmic vacuoles of 10-25 pm in diameter (Fig. 6). Sporoblast cells contained numerous basophilic immature sporozoites 2 pm in diameter (Fig. 7), but sporoblasts containing mature spores were not observed.
Retrospective PCR assays for the detection of oocyte parasites that included the Perkinsela amoebae-Yike organism of Siihnel et al. (2014) and Marteilioides chungmuensis (Itoh et al. 2003) were uniformly negative with samples from the current investigation.
Gametogenic activity and gonad development were generally similar among uninfected oysters and those infected by Steinhausia sp., except that two infected oysters sampled in July and September showed intense hemocytic infiltration in their gonad follicles. Oysters sampled in July had recently spawned, and most were reabsorbing gametes. In September samples, gonads were ripe or partially spawned, in January gonads were in advanced gametogenesis, and in April gonads were ripe.
The prevalence of Perkinsus sp. infections detected by histology ranged from 13.3% 60% (Table 1). Prevalence of Perkinsus sp. infection was significantly greater among the cultured oysters (ANOVA, P = 0.0161), with an overall annual prevalence of 50.8% (61 of 120) compared with 25.8% (31 of 120) among the wild oysters. Systemic infection by Perkinsus sp. was observed in one specimen only that also showed an intense focal infection within the gonad (Figs. 8-12). Most oysters showed low-intensity infections in the epithelia of digestive organs (stomach and intestine), and occasionally among connective tissues of the visceral mass.
Haplosporidian protozoa of Bonamia sp. were not detected by microscopic analyses of hemolymph cytological preparations (n = 240) or histological sections (n = 240), nor were any cases of disseminated (hemocytic) neoplasia detected by either method.
Platyhelminth turbellarians resembling a Urastoma sp. were associated with external surfaces of oyster gill filaments, where they sometimes occurred in close contact with gill epithelia. Turbellarians measured 100-450 [micro]m in length, had ciliated external epithelia, and showed two eye spots and a pharynx at opposite ends of their body (Figs. 13 and 14). The prevalence of the Urastoma sp. ranged from 7% 53% (Table 1), with infestation intensities always less than five turbellarians per section.
A copepod in an advanced stage of degradation was observed among the visceral connective tissues of one oyster, and its refractile and lightly acidophilic exoskeleton was densely surrounded by defensive hemocytes (Fig. 15).
Macroscopic examination of oyster valves revealed mud blisters on the inner surface of shells that reflected colonization by shell-infesting Polydora sp. spionid polychaetes. Infestation intensities were typically low, except in seven cases in which adjacent mantle tissues were affected, or adductor muscles showed abnormal coloration (n = 2) or partial rupture (n = 1). The overall prevalence of Polydora sp. infestation was significantly greater among cultured oysters (94 [+ or -] 11%) than wild oysters (40 [+ or -] 4%; ANOVA, P = 0.0008). Among samples of cultured oysters, those from the warmer months of January, April, and September had the greatest Polydora sp. prevalence (97% 100%) and the July sample had the lowest (77%). Among wild oyster samples, Polydora sp. prevalence was nearly constant during the year (38%-44%; (Table 1).
The OPPC was greater in cultured oysters (4.98) than in wild oysters from the Rio Sao Francisco estuary (4.02; Kruskal-Wallis, P = 0.08).
The current investigation detected nonspecific pathologies, parasites, and commensals in Crassostrea gasar oysters from the estuary of the Rio Sao Francisco, including possible viruses, bacteria, protozoa, and metazoa. None of these were reported previously for C. gasar, and most of them did not show major seasonal variations.
Morphological changes of gametocytes in both male and female gonad follicles suggested viral gametocytic hypertrophy disease. The cell alterations observed were similar to those reported by Garcia et al. (2006) in Crassostrea gigas from France, and to those previously reported among cultured C. gigas and wild Crassostrea rhizophorae from Santa Catarina Island in southern Brazil (Sabry et al. 2011, da Silva et al. 2012). Intracellular colonies of RLO reported here resemble those described by Harshbarger et al. (1977) and by Bower et al. (1994) in bivalves. Similar RLO were described previously among C. gigas and C. rhizophorae oysters from the coast of Santa Catarina in southern Brazil (Sabry et al. 2011, da Silva et al. 2012). The current investigation found three oysters with symptoms of the disease known as maladie du pied, which is also reported among C. gigas and C. rhizophorae from Santa Catarina Island, Brazil (Sabry & Magalhaes 2005). This condition was probably associated with the infection by the fungus Ostracoblabe implexa, and may include complete disarticulation of the adductor muscle from its valve, to compromise the adductive predation defenses of affected oysters (Bower et al. 1994).
The gregarine protozoa Nematopsis sp. were consistently observed infecting wild and cultured Crassostrea gasar oysters at variable levels of prevalence in the current study. This parasite commonly infects the connective tissue of various species of Brazilian bivalves without causing damage, including Crassostrea rhizophorae and Crassostrea gigas at high prevalence (Sabry & Magalhaes 2005, Sabry et al. 2011, da Silva et al. 2012).
A Steinhausia sp. microsporidian infected Crassostrea gasar oocytes. Almost all stages of development of the microsporidian were observed (Sagrista et al. 1998), including the infective, rarely observed sporoplasm stage (Anderson et al. 1995). The sporoplasm is released by spores into the cytoplasm of the oocyte through a polar tube mechanism common to microsporidia (Becker & Pauley 1968). However, sporoblasts containing rigid and refractile mature spores were not observed among infected C. gasar oocytes. Mature spores have been described previously as Steinhausia mytilovum among infected oocytes of the Brazilian mussel Mytellaguyanensis (Matos et al. 2005), and as Steinhausia sp. among the clam Anomalocardia brasiliana (da Silva et al. 2012), and the oysters Crassostrea gigas and Crassostrea rhizophorae (Sabry et al. 2011). However, only the current study reports observations of sporoplasm. Recently, a Perkinsela amoebae-like ovarian parasite (Order Dactylopodida) was reported in C. gasar from southern Brazil (Stihnel et al. 2014), but samples from the current study were negative by retrospective PCR for that parasite, indicating its probable absence among samples of the current investigation.
Among the Steinhausia sp.-infected mussel Mytilus galloprovincialis, a reduction in condition index is described (Rayyan & Chintiroglou 2003), and strong hemocytic infiltration and oocyte resorption are reported among 74%-95% of infected female Saccostrea glomerata oysters (Green et al. 2008). Similarly, in the current study, 2 of 23 oysters infected by Steinhausia sp. (8.7%) showed evidence of compromised gametogenesis. Although the general low intensity of Steinhausia sp. infections reported here suggests a low impact on oyster reproduction, expanded knowledge of the reproductive cycle of Crassostrea gasar in the estuary of the Rio Sao Francisco is crucial to evaluating the potential impacts of such infections there.
Concerning the OIE notifiable parasites, analyses of hemolymph cytological preparations and histological sections strongly suggested the absence of infections by Bonamia sp. protozoa among the oysters examined during the current investigation. Although the hemolymph cell monolayer is the most sensitive assay among microscopic methods (da Silva & Villalba 2004), both cytological and histological samples from 240 oysters were examined exhaustively with uniformly negative results. Nevertheless, a larger sample size (n = 150) is recommended by the OIE to detect prevalence as low as 2% with 95% confidence.
Analysis of Perkinsus sp. prevalence suggests the culture environment may have exacerbated infections by Perkinsus sp. (greatest mean prevalence). In contrast to results from the current investigation, Caceres-Martinez et al. (2012) found a similar prevalence for Perkinsus marinus infections among limited samples of the cultured and wild oyster Saccostrea palmula in western Mexico. Because no differences in temperature or salinity characterized the habitats for samples of wild and cultured oysters in the current investigation, two factors may be associated with the greater prevalence among cultured oysters: (1) frequent manipulation of cultured oysters and (2) their high-density culture conditions. Summarizing factors influencing the epizootiology of dermo disease among Chesapeake Bay oysters, Andrews (1967) reports highest mortality from P. marinus infections among Crassostrea virginica in higher-density populations.
A low prevalence of gill infestation by the turbellarian Urastoma cyprinae is commonly reported in the oyster Crassostrea virginica from Canada (Brun et al. 1999) and the mussels Mytilus galloprovincialis from Greece (Rayyan et al. 2004) and Spain (Crespo-Gonzalez et al. 2005), and has been reported previously without pathological consequences among mussels, clams, and oysters from southern Brazil (Suarez-Morales et al. 2010, Sabry et al. 2011. da Silva et al. 2012). The copepod Pseudomyicola spinosus is one of the most common external copepod associates of marine invertebrates worldwide (Ho & Kim 1991, Ho 2001, Caceres-Martinez et al. 2005), including the clam Anomalocardia brasiliana from Brazil (Narchi 1965). In the current study, a copepod was observed within one oyster's visceral tissues, where it had been largely destroyed by defensive hemocytes. A similar observation that was also characterized by an intense hemocytic response by a Crassostrea gigas oyster was speculated to be a consequence of an atypical copepod invasion via the oyster's gonoduct (da Silva et al. 2012).
Shell infestations by Poly dor a sp. mud worms may reduce market values. The greater prevalence of Poly dor a sp. observed among cultured oysters may reflect greater availability of shell and sediment habitats for polychaetes associated with high-density benthic populations of cultured oysters. Moreover, unlike benthic cultured oysters, wild oysters attached to mangrove rhizophores are exposed to air during low tides, which works as a natural deterrent against settlement and survival of worm larvae (Nel et al. 1996). However, based on the general low intensity of Polydora sp. infestations among the Crassostrea gasar oysters in the current study, only marginal local impacts on oyster growth and market value are anticipated.
In conclusion, results show that Crassostrea gasar oysters cultivated in the Rio Sao Francisco estuary had greater prevalences of diseases and pathogens than oysters from a wild population in the same estuary, including maladie du pied, RLO in gills, Steinhausia sp. oocyte infections, Polydora sp. valve infestations, and Perkinsus sp. infections. Despite a high prevalence of Perkinsus sp. infections, the generally low intensities of such infections suggested their mortality effects may also be low. This study represents a first histopathological inventory for C. gasar, provides a reference baseline for expanded future surveys, and indicates the general good health among both wild and cultured oysters despite their diverse commensal and parasitic symbionts.
The authors thank the Conselho Nacional de Desenvolvimento Cientifico e Tecnologico (CNPq) for financial support to projects 471822/2009-4 and CNPq/MPA no. 406170/2012-6. They are grateful to the producer of oysters, Miguel Ferreira dos Santos, who allowed access to his farm and helped with oyster sampling. They thank Dr. Ana Rosa R. Araujo for coordinating contact with the producer and Dr. Rogerio T. Vianna for map production. With regard to technical assistance, the authors are grateful to undergraduate students Andre L. L. Leal, Raoani C. Mendonca, and Ana C. S. Santana.
Anderson, T. J., P. M. Hine & R. J. G. Lester. 1995. A Steinhausia-like infection in the ovocytes of Sydney rock oysters Saccostrea commercialis. Dis. Aquat. Organ. 22:143-146.
Andrews, J. D. 1967. Interactions of two diseases of oysters in natural waters. Proc. Natl. Shellfish. Assoc. 57:38M9.
Becker, C. D. & G. B. Pauley. 1968. An ovarian parasite (Protista incertae sedis) from the Pacific oyster, Crassostrea gigas. J. Inverlebr. Pathol. 12:425-437.
Bower, S. M., S. E. McGladdery & I. M. Price. 1994. Synopsis of infection diseases and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4:1-199.
Brandao, R. P., G. Boehs, R. C. Sabry, L. O. Ceuta, M. S. A. Luz, F. R. Queiroga & P. M. da Silva. 2013. Perkinsus sp. infecting oyster Crassostrea rhizophorae (Guilding, 1828) on the coast of Bahia, Brazil. J. Invertebr. Pathol. 112:138-141.
Brun, N. T., A. D. Boghen & J. Allard. 1999. Distribution of the turbellarian Urastoma cyprinae on the gills of the eastern oyster Crassostrea virginica. J. Shellfish Res. 18:175-179.
Caceres-Martinez, J., J. Chavez-Villaiba & L. Garduno-Mendez. 2005. First record of Pseudomyicola spinosus in Argopecten ventricosus in Baja California, Mexico. J. Invertebr. Pathol. 89:95-100.
Caceres-Martinez, J., M. G. Ortega, R. Vasquez-Yeomans, T. J. P. Garcia, N. A. Stokes & R. B. Carnegie. 2012. Natural and cultured populations of the mangrove oyster Saccostrea palmula from Sinaloa, Mexico, infected by Perkinsus marinus. J. Invertebr. Pathol. 110:321-325.
Carnegie, R. B. & N. Cochennec-Laureau. 2004. Microcell parasites of oysters: recent insights and future trends. Aquat. Living Resour. 17:519-528.
Crespo-Gonzalez, C., R. M. R. Alvarez, H. R. Dominguez, M. S. Bua, R. Iglesias, C. A. Fernandez & J. M. G. Estevez. 2005. In vitro reproduction of the turbellarian Urastoma cyprinae isolated from Mytilus galloprovincialis. Mar. Biol. 147:755-760.
da Silva, P. M., F. Cremonte, R. C. Sabry, R. D. Rosa, L. Cantelli & M. A. Barracco. 2009. Presence and histopathological effects of the Parvatrema sp. (Digenea Gymnophallidae) in the stout razor clam Tagelus plebeius (Bivalvia Psammobiidae). J. Invertebr. Pathol. 102:14-20.
da Silva, P. M., J. Fuentes & A. Villalba. 2005. Growth, mortality and disease susceptibility of oyster Ostrea edulis families obtained from brood stocks of different geographical origins, through on-growing in the Ria de Arousa (Galicia, NW Spain). Mar. Biol. 147:965-977.
da Silva, P. M., A. R. M. Magalhaes & M. A. Barracco. 2012. Pathologies in commercial bivalve species from Santa Catarina state, southern Brazil. J. Mar. Biol. Assoc. U.K. 92:571-579.
da Silva, P. M., R. T. Vianna, C. Guertler, L. P. Ferreira, L. N. Santana, S. Fernandez-Boo, A. Ramilo, A. Cao & A. Villalba. 2013. First report of the protozoan parasite Perkinsus marinus in South America, infecting mangrove oysters Crassostrea rhizophorae from the Paraiba River (NE, Brazil). J. Invertebr. Pathol. 113:96-103.
da Silva, P. M. & A. Villalba. 2004. Comparison of light microscopic techniques for the diagnosis of the infection of the European flat oyster Ostrea edulis by protozoan Bonamia ostreae. J. Invertebr. Pathol. 85:97-104.
de Melo, A. G., E. S. Varela, C. R. Beasley, H. Schneider, I. Sampaio, P. M. Gaffney, K. S. Reece & C. H. Tagliaro. 2010. Molecular identification, phylogeny and geographic distribution of Brazilian mangrove oysters (Crassostrea). Genet. Mol Biol. 33:564-572.
EPAGRI. 2013. Sintese informativa da maricultura 2012. Available at <http://cedap.epagri.sc.gov.br.
Garcia, C., M. Robert, A. Arzul, B. Chollet, J.- P. Joly, L. Miossec, T. Comtet & F. Berthe. 2006. Viral gametocytic hypertrophy of Crassostrea gigas in France: from occasional records to disease emergence? Dis. Aquat. Organ. 70:193-199.
Green, T. J., B. J. Jones, R. D. Adlard & A. C. Barnes. 2008. Parasites, pathological conditions and mortality in QX-resistant and wild-caught Sydney rock oysters Saccostrea glomerata. Aquaculture 280:35-38.
Harshbarger, J. C., S. C. Chang & S. V. Otto. 1977. Chlamydiae (with phages) mycoplasmas and rickettsiae in Chesapeake Bay bivalves. Science 196:666-668.
Ho, J. S. 2001. Why do symbiotic copepods matter? Hydrobiologia 453/ 454:1-7.
Ho, J. S. & I. H. Kim. 1991. Copepod parasites of commercial bivalves in Korea: II. Copepods from cultured bivalves. Bull. Korean Fish. Soc. 24:369-396.
Howard, D. W., E. J. Lewis, B. J. Keller & C. S. Smith. 2004. Histological techniques for marine bivalve molluscs and crustaceans. NOAA technical memorandum NOS NCCOS 5, Coop. Oxford, MD: Oxford Laboratory. 218 pp.
Itoh, N., O. Tadashi, T. Yoshinaga & K. Ogawa. 2003. DNA probes for detection of Marteilioides chungmuensis from ovary of Pacific oyster Crassostrea gigas. Fish Pathol. 38:163-169.
Lopes, G. R., C. H. A. M. Gomes, C. R. Tureck & C. M. R. De Melo. 2013. Growth of Crassostrea gasar cultured in marine and estuary environments in Brazilian waters. Pesquisa Agropecu. Bras. 48:975-982.
Matos, E., P. Matos & C. Azevedo. 2005. Observations on the intracytoplasmatic microsporidian Steinhausia mytilovum, a parasite of mussel (Mytella guyanensis) oocytes from the Amazon River estuary. Braz. J. Morphol. Sci. 22:183-186.
Narchi, W. 1965. A new species of Pseudomyicola Yamaguti 1936. An. Acad. Bras. Cienc. 37:359-361.
Nel, R., P. S. Coetzee & G. Van Niekerk. 1996. The evaluation of two treatments to reduce mud worm (Polydora hoplura Claparede) infestation in commercially reared oysters (Crassostrea gigas Thunberg). Aquaculture 141:31-39.
Ramos, C. O., J. F. Ferreira & C. M. R. De Melo. 2013. Maturation of native oyster Crassostrea gasar at different diets in the laboratory. Bol. Inst. Pesca 39:107-120.
Rayyan, A. & C. C. Chintiroglou. 2003. Steinhausia mytilovum in cultured mussels Mytilus galloprovincialis in the Thermaikos Gulf (northern Aegean Sea, Greece). Dis. Aquat. Organ. 57:271-273.
Rayyan, A., G. Photis & C. C. Chintiroglou. 2004. Metazoan parasite species in cultured mussel Mytilus galloprovincialis in the Thermaikos Gulf (North Aegean Sea, Greece). Dis. Aquat. Organ. 58:55-62.
Sabry, R. C., P. M. da Silva, T. C. V. Gesteira, V. A. Pontinha & A. R. M. Magalhaes. 2011. Pathological study of oysters Crassostrea gigas from culture and C. rhizophorae from natural stock of Santa Catarina Island, SC, Brazil. Aquaculture 60:43-50.
Sabry, R. C., T. C. V. Gesteira, A. R. M. Magalhaes, M. A. Barracco, C. Guetler, L. P. Ferreira, R. T. Vianna & P. M. da Silva. 2013. Parasitological survey of mangrove oyster, Crassostrea rhizophorae, in the Pacoti River estuary, Ceara state, Brazil. J. Invertebr. Pathol. 112:24-32.
Sabry, R. C. & A. R. M. Magalhaes. 2005. Parasitas em ostras de cultivo (Crassostrea rhizophorae e Crassostrea gigas) da Ponta do Sambaqui, Florianopolis, SC. Arq. Bras. Med. Vet. Zootec. 57:194-203.
Sabry, R. C., R. D. Rosa, A. R. M. Magalhaes, M. A. Barracco, T. C. V. Gesteira & P. M. da Silva. 2009. First report of Perkinsus sp. infecting mangrove oysters Crassostrea rhizophorae from the Brazilian coast. Dis. Aquat. Organ. 88:13-23.
Sagrista, E., M. G. Bozzo, M. Bigas, M. Poquet & M. Durfort. 1998. Developmental cycle and ultrastructure of Steinhausia mytilovum, a microsporidian parasite of oocytes of the mussels, Mytilus galloprovincialis (Mollusca, Bivalvia). Eur. J. Protistol. 34:58-68.
Santos, L. C. M., H. R. Matos, Y. Schaeffer-Novelli. M. Cunha-Lignon, M. D. Bitencourt, N. Koedam & F. Dahdouh-Guebas. 2014. Anthropogenic activities on mangrove areas (Sao Francisco River estuary, Brazil northeast): a GIS-based analysis of CBERS and SPOT images to aid in local management. Ocean Coast. Manage. 89:39-50.
Silveira. R. C, F. C. Silva, C. H. M. Gomes, J. F. Ferreira & C. M. R. Melo. 2011. Larval settlement and spat recovery rates of the oyster Crassostrea brasiliana (Lamarck, 1819) using different systems to induce metamorphosis. Braz. J. Biol. 71:557-562.
Suarez-Morales, E., M. P. Scardua & P. M. da Silva. 2010. Occurrence and histopathological effects of Monstrilla sp. (Copepoda: Monstrilloida) and other parasites in the brown mussel Perna perna from Brazil. J. Mar. Biol. Assoc. U.K. 90:953-958.
Suhnel, S., C. da S. Ivachuk, A. L. C. Schaefer, V. A. Pontinha, M. L. Martins, A. Figueras, G. R. Meyer, S. R. M. Jones, J. C. Sewart, H. J. Gurney-Smith, A. R. M. Magalhaes & S. M. Bower. 2014. Detection of a parasitic amoeba (Order Dactylopodida) in the female gonads of oysters in Brazil. Dis. Aquat. Organ. 109:241-250.
Villalba, A., C. Gestal, S. M. Casas & A. Figueras. 2011. Perkinsosis en moluscos. In: A. Figueras & B. Novoa, editors. Enfermedades de moluscos bivalvos de interes en acuicultura. Madrid: Fundacion Observatorio Espanol de Acuicultura. pp 181-242.
Villalba, A., K. S. Reece, M. C. Ordas, S. M. Casas & A. Figueras. 2004. Perkinsosis in molluscs: a review. Aquat. Living Resour. 17:411-432.
PATRICIA MIRELLA DA SILVA, (1) * ([dagger]) MARCOS PAIVA SCARDUA, (2,3) CAIRE BARRETO VIEIRA, (4) ANALEE CRUZ ALVES (1) AND CHRISTOPHER F. DUNGAN (5)
(1) Nucleo de Engenharia de Pesca, Universidade Federal de Sergipe, CEP 49100-000, Sao Cristovao, SE, Brazil; (2) Embrapa Tabuleiros Costeiros, Bairro Jardins, CEP 49025-040, Aracaju, SE, Brazil; (3) Instituto Federal de Educacao Ciencia e Tecnologia do Ceara, CEP 62800-000, Aracati, CE, Brazil; (4) Universidade Federal da Paraiba, CEP 58051-900, Joao Pessoa, PB, Brazil; (5) Maryland Department of Natural Resources, Cooperative Oxford Laboratory, 904 S. Morris Street, Oxford, MD 21654
* Corresponding author. E-mail: email@example.com Current address: Universidade Federal da Paraiba, Centro de Ciencias Exatas e da Natureza, Departamento de Biologia Molecular, Jardim Universitario s/n. Bairro Castelo Branco, CEP 58051-900, Joao Pessoa, PB, Brasil
TABLE 1. Number of affected oysters and quarterly (2010) prevalence (measured as a percentage) of pathological conditions, parasites (P), and commensals (C) among mangrove oysters (Crassostrea gasar) from cultured and wild populations in the estuary of the Rio Sao Francisco, Sergipe. January Cultured * Wild Shell height 77.7 [+ or -] 6.7 71.1 [+ or -] 9.4 (mean [+ or -] SD) N = 127 N = 121 n = 30 n = 30 NF = 12 NF = 20 NM = 18 NM = 8 Condition Hemocytic infiltration 6 (20%) 5 (16.7%) Granulocytoma 1 (3.3%) 0 VGH (P) 0 0 RLO (P) DG 2 (6.7%) 6 (20%) Gill 2 (3.3%) 0 Maladie du pied (P) ([double dagger]) 0 0 Nematopsis sp. (P) 10 (33.3%) 21 (70%) Steinhausia sp. (P) ([dagger]) 1 (8.3%) 1 (5.0%) Perkinsus sp. (p) 18 (60%) 12 (40%) Turbellarians (c) 4 (13.3%) 6 (20%) Copepod unknown (c) 0 0 Polydora sp. (c) 126 (99.2%) 43 (35.5%) April Cultured * Wild Shell height 76.8 [+ or -] 12.8 70.5 [+ or -] 9.9 (mean [+ or -] SD) N = 100 N = 100 n = 30 n = 30 NF = 11 NF = 15 NM = 19 NM = 13 HPM = 2 Condition Hemocytic infiltration 9 (29.9%) 10 (33.3%) Granulocytoma 1 (3.3%) 0 VGH (P) 0 1 (3.3%), M RLO (P) DG 4 (13.3%) 2 (6.7%) Gill 0 0 Maladie du pied (P) ([double dagger]) 0 0 Nematopsis sp. (P) 2 (6.7%) 12 (40%) Steinhausia sp. (P) ([dagger]) 0 4 (26.7%) Perkinsus sp. (p) 13 (43.3%) 9 (30%) Turbellarians (c) 2 (6.7%) 10 (33.3%) Copepod unknown (c) 1 (3.3%) 0 Polydora sp. (c) 100 (100%) 42 (42%) July Cultured Wild Shell height 73.9 [+ or -] 6.7 76.6 [+ or -] 11.0 (mean [+ or -] SD) N = 100 N = 100 n = 30 n = 30 NF = 15 NF = 16 NM = 15 NM = 13 HPF = 1 Condition Hemocytic infiltration 27 (90%) 23 (76.7%) Granulocytoma 0 3 (10%) VGH (P) 1 (3.3%). M 0 RLO (P) DG 5 (16.7%) 2 (2.6%) Gill 2 (6.7%) 0 Maladie du pied (P) ([double dagger]) 0 0 Nematopsis sp. (P) 3 (10%) 6 (20%) Steinhausia sp. (P) ([dagger]) 1 (6.7%) 3 (33.3%) Perkinsus sp. (p) 17 (56.7%) 6 (20%) Turbellarians (c) 16 (53.3%) 7 (23.3%) Copepod unknown (c) 0 0 Polydora sp. (c) 77 (77%) 38 (38%) September Cultured * Wild Shell height 65.4 [+ or -] 9.0 70.4 [+ or -] 9.9 (mean [+ or -] SD) N = 100 N = 99 n = 30 " = 30 NF = 16 NF = 20 NM = 13 NM = 7 HPF = 1 HPF = 3 Condition Hemocytic infiltration 21 (70%) 18 (60%) Granulocytoma 3 (10%) 0 VGH (P) 2 (6.7%), M and F 2 (6.7%), M RLO (P) DG 0 3 (10%) Gill 2 (6.7%) 1 (3.3%) Maladie du pied (P) ([double dagger]) 3 (3%) 0 Nematopsis sp. (P) 3 (10%) 1 (3.3%) Steinhausia sp. (P) ([dagger]) 12 (75%) 1 (5%) Perkinsus sp. (p) 13 (43.3%) 4 (13.3%) Turbellarians (c) 5 (16.7%) 4 (13.3%) Copepod unknown (c) 0 0 Polydora sp. (c) 97 (97%) 44 (44.4%) DG, digestive gland; F, female; HPF, hermaphrodite predominantly female; HPM, hermaphrodite predominantly male; M, male; Maladie du pied, shell disease;", number of oysters analyzed histologically; N, number of oysters analyzed macroscopically; NF, number of females; NM, number of males; RLO, RickettsiaAike organisms; VGH, viral gametocytic hypertrophy. * Statistical difference (P < 0.05) in shell height of cultured and wild oysters, ([dagger]) Prevalence among female oysters. ([double dagger]) Prevalence among oysters analyzed macroscopically (TV).
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|Author:||Da Silva, Patricia Mirella; Scardua, Marcos Paiva; Vieira, Caire Barreto; Alves, Analee Cruz; Dungan|
|Publication:||Journal of Shellfish Research|
|Date:||Aug 1, 2015|
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