Printer Friendly

Reproduction and pathology of blue mussels, Mytilus edulis (L.) in an experimental longline in Long Island Sound, Connecticut.

ABSTRACT An experimental longline was deployed in Long Island Sound, Connecticut to study the biologic feasibility of commercial culture of blue mussels, Mytilus edulis, in the area. Mussels were sampled monthly for 1 year starting 6 months after the seed settled. Samples were processed for histology and studied for gonad development to predict spawning peaks for future seed line deployments. There were 2 spawning peaks; a major one in May and a minor one in August. Gametogenic activity was detected throughout the year with developing gonads present in February and spawning specimens present from April to January. Samples were also examined for pathology to determine if there were diseases or parasites that could affect the operation. Mussels acquired heavy infections of the trematode parasite Proctoeces maculatus (over 60%) in September that lasted to the end of the year. Large areas of the mantle and visceral tissues were replaced with larval and adult parasites. Abscesses were formed with adult trematodes surrounded by massive aggregations of host hemocytes. Mussels reached sexual maturity and commercial size (60 mm) during their first 6 months after seed set. Two opposing forces drove the feasibility of the operation in this atypically southern location for mussel culture; exceptionally rapid growth, but also heavy infections by a trematode parasite enzootic to tropical and temperate waters. We conclude that the biologic potential for a commercial operation exists for a seasonal product that would use the window of opportunity for parasite-free full-grown mussels from the first winter after settlement until the next midsummer.

KEY WORDS: reproduction, pathology, longline, blue mussel, Mytilus edulis, Proctoeces maculatus, Long Island Sound


Culture of blue mussels, Mytilus edulis (Linnaeus 1758) is a fast growing sector of shellfish aquaculture. Global aquaculture production of mussels has increased 20-fold (from less than 100 kilotons to over 1.3 million tons) during the past 50 y (McLeod 2002). Mussel production has been dominated by Europe (mainly Spain, Italy, the Netherlands, and France) and China with combined output exceeding 90% of world production (McLeod 2002). Mussel culture, still a diminutive player on a global scale, in North America has been growing constantly. The market leader is Prince Edward Island (PEI), Canada with an estimated 50% of the North American mussel market. Submersible longline technology is used in PEI (McDonald et al. 2002).

Three mussel species are cultured in the USA: Mytilus edulis in New England, M. trossulus (Gould 1850) in Alaska, both M. galloprovincialis (Lamark 1819) and M. trossulus in Washington State, and M. galloprovincialis in California. In the Northeast USA about half of the current production is still largely based on wild fishery (bottom harvest) and the other half coming from cultured product with a total of 7.3 million kg annually. Raft culture is used widely in Maine, the Northeast USA market leader in mussel production (King & Cortes-Monroy 2002, The Island Institute 1999). A pilot-scale mussel longline culture was established in 1999 at a site eight kilometers from the shore in the Gulf of Maine, New Hampshire, to study the feasibility of deep-water culture (Langan 2000, Langan & Horton 2002). A mussel longline was also established in Narragansett Bay, Rhode Island (Corayer 2003). Growing interest toward blue mussel culture is due to the easy collection of natural seed, relatively low operation costs and short production cycles. Unlike most bivalves mussels have a short shelf life, a problem that could be overcome with local production.

Mussel culture is usually based on collection of natural seed. Hatchery production of mussel seed is used only in Washington State, Australia, and China (King & Cortes-Monroy 2002). Because of the importance of accurate timing for deployment of seed collecting lines to catch mussel spat and avoid setting of fouling organisms, the reproductive cycle of Mytilus has been studied extensively. Studies from different European estuaries reported that spawning times differed between areas from March until July as well as annually. In general, spawning occurred later, was shorter, and had only one peak in the more northern latitudes compared with those areas further south. A second, less pronounced, spawning peak usually manifested itself later during the summer or fall in more southern areas (e.g., Bayne 1964, Chipperfield 1953, Kautsky 1982, Seed 1975, Sunila 1981, Thorarinsdottir 1996).

A similar pattern was observed on the East Coast of North America (Thompson 1984). Mussels in Newfoundland spawned in late July. The reproductive cycle resembled that of Baltic mussels rather than mussels in central European locations. The reproductive cycle of M. edulis in Long Island Sound (LIS) has been studied as follows: Hrs-Brenko (1971) sampled in Milford, Connecticut (CT), LIS from March until July; Brousseau (1983) in Fairfield, CT, LIS for a period of 2 years; and Fell & Balsamo (1985) in the Thames River and at Branford, CT, LIS from May until June. These reports studying wild mussel populations concluded that spawning occurred in May to June. Newell et al. (1982) sampled several sites between Maine and Delaware Bay and reported that at one of the sites, Stony Brook, NY, LIS, spawning occurred from April to May.

When predicting accurate timing for setting seed collecting lines, the length of the pelagic phase of mussel larvae must be taken into consideration. Peak setting of mussel larvae in Milford Harbor generally occurred between June 29 to July 6 (Engle & Loosanoff 1944) but in late fall in the Thames River (Fell & Balsamo 1985). According to their observations, spawning and setting occurred about 4 months earlier in the central shores of the Sound. Larval period is estimated as being 3-5 wk after which the peak settlement occurs (Seed 1969). However, unfavorable conditions could prolong it for up to 6 months (Nelson 1928, Lane et al. 1985). Postlarvae are capable of secreting a long drifting thread to remain pelagic, which enables the mussels to extend their distribution.

Massive mortalities of cultured M. edulis are generally not associated with disease outbreaks. Virus-like disease has been reported to cause mortalities (50% to 100%) of cultured green-lip mussels Perna canaliculus in New Zealand (Jones et al. 1996), but virus-associated granulocytomas in M. edulis in Denmark were not associated with mortalities (Rasmussen 1986). Marteiliasis (Marteilia refringens, usually a pathogen of Ostrea edulis) has been associated with mortalities of M. galloprovincialis in Spain (Villalba et al. 1993), but not in M. edulis in Brittany, France (Robledo et al. 1994a). Disseminated sarcoma has caused significant mortalities of cultured M. trossulus in Puget Sound, Washington but doesn't reach epizootic prevalences in M. edulis at different sites in the Atlantic (Elston et al. 1992). Figueras et al. (1991) sampled wild mussels from the coasts of New Jersey and Maine to provide preliminary baseline pathology data. They reported the presence of microsporidia Steinhausia mytilovum, an intracellular ciliate inside digestive cells, and trematodes Proctoeces maculatus. They also observed Rickettsiae-like prokaryotic inclusions in the digestive cells, haplosporidia-like protozoan and microcell-like organisms that resembled Bonamia. The presence of these parasites was not reported to be associated with mortalities.

Even though Mytilus is a dominant species of the northern hemisphere, a key aquaculture species and part of the natural fauna in LIS (Weiss 1995), no efforts have been made to commercially grow the mussel in LIS. Several papers were published concerning the reproduction of Mytilus in LIS, but did not provide information about the actual spawning times. Fell and Balsamo's (1985) observations from Branford were based on sampling during 2 months, whereas Hrs-Brenko' s (1971) studies covered 4 months. Newell et al. (1982) and Brousseau (1983) used stereology to assess reproductive status and expressed it as the "gamete volume fraction" which is the proportion of the mantle tissue that is composed of follicles containing developing or ripe gametes. This method doesn't include visual observation of gamete release, and pre and post spawning status can show similar values. There is a paucity of information concerning the pathologic status of mussels in LIS. Thus, an experimental mussel longline was deployed in LIS in the summer of 2001 to determine if there are diseases or parasites that could affect mussel culture in the area and to define spawning peaks for future seedline deployments.


A longline for mussel seed collection was deployed in summer of 2001 on a shellfish lease in Milford, Connecticut. The longline is simply a long piece of rope that is fixed horizontally in the water column (Fig. 1). A horizontal head rope (30 m, 9.5 mm sinking rope) was submerged to 1.8 m below the surface at mean low tide and anchored at each end with concrete blocks (68 kg, 0.2 m x 0.3 m x 0.2 m). Water depth at the lease was 10 m. Vertical seed collecting lines were hung below the head rope at 1 m intervals and their weight was supported by a series of buoys. Three-meter long collecting lines were composed of alternating 6.4 mm Polyplus rope and 12.7 mm sinking rope. Two marker buoys were tied to the headline at the quarter- and three-quarter- distance marks. Mussels were left on the seed lines for the grow-out period.


A random sample of 30 mussels was taken monthly from February 2002 to January 2003 to study the developmental stage of the gonads and pathology (a total of 360 mussels). Temperature was recorded at a distance of 300 m from the longline site in Milford Harbor (Table 1). In the laboratory, the mussels were measured with calipers (maximum length) and examined grossmacroscopically. Fouling organisms and the presence of pea crabs (Pinnotheres maculatus) were recorded and the pea crabs were removed at this point. Samples were fixed in Davidson's fixative and processed for pathology using standard histologic techniques. A 5-[micro]m thick paraffin section was stained with hematoxylin-eosin. The developmental stage was expressed by a gonad index (Chipperfield 1953). The developmental stage of the gonads was classified according to Seed (1969). Developing specimens were described by numerical scores 1 to 5 and spawning by values 4 to 1. A resting or immature gonad was defined with value 0. Gonad index varied from zero, when no sexual activity was detected, to five, when all the individuals were ripe. The number of mussels at each stage was multiplied by the numerical score of the stage and the sum was divided by the number of individuals in the sample to obtain a weighted gonad index.

Classification of the Gonad Developmental Stages

Developmental Stage

0. Immature or already spawned, resting gonads. No germinal follicles are observed in the mantle and the sexes cannot be distinguished.

1. Ducts lined with germinal epithelium start to appear in the middle of the connective tissue in the mantles. Ripe gametes cannot be observed, but early stages of gametogenesis are present.

2. Gonoducts expand displacing the connective tissue. Ripe sperm and ova appear in the middle of the follicles, but the early stages such as spermatogonia, spermatocytes, and ovocytes fastened to the germinal epithelium dominate.

3. The follicles are about half-filled with mature gametes, the remaining area consisting of early stages of gametogenesis. The ducts and follicles have expanded to fill about half of the mantle matrix.

4. Follicles have invaded almost the entire mantle. Cells undergoing gametogenesis can still be found in the margins of the ducts, but ripe gametes dominate.

5. Follicles are full of ripe gametes. Some gametocytes are still in the margins of the ducts (gametes de reserves, Lubet 1957). Tightly compacted ova have an angular configuration.

Spawning Stage

4. The release of gametes from the mussel has begun. Follicles are still full of gametes, but their numbers in the lumina of the follicles have decreased and the ova have a spherical shape.

3. Follicles are half-filled with gametes. This can be distinguished from the previous developmental stage by the scarcity of developing cells.

2. Gametes fill less than half of the follicles. The ducts continue to shrink, whereas the connective tissue is expanding.

1. The follicles have disappeared almost completely. Some sperm and ova remain in the follicles. There may be hemocytes in the follicles phagocytizing residual gametes.

All pathologic changes were recorded for characterizing the health of the mussels. Additional pea crabs were observed in histologic sections and their number was added to the count obtained from gross-macroscopic examination to determine the actual prevalence.


Mussel seed set on the longline in July of 2001. Sampling for histologic examination started in February 2002 and ended in January 2003. The mussels were followed from 6 mo of age for a full year during the experiment. The size of the mussels did not increase significantly during the experiment (Table 1) indicating that the mussels had reached approximately their maximum size during their first 6 months after setting. During the first 5 months of sampling no fouling organisms were present, but after that mussels were covered with slipper shells, Crepidula fornicata (Table 1). Barnacles and bryozoa also attached to shells, but slipper shells covered these organisms to such an extent that the prevalences of the latter were not recorded. Concurrently with the invasion of slipper shells the lines were covered with hydroids (Tubularia ssp.) that were removed prior to sampling of the mussels. The mean prevalence of pea crabs (Pinnotheres maculatus) was 9.4% during the year. Their presence had a seasonal pattern with absence during the spring and summer months and a peak from November through January (see Table 1). Nine out of the total of 34 pea crabs (26%) were observed histologically only and were not noted during gross-macroscopical examination due to their small size. Most of the infestations were single pea crab/mussel associations, but one mussel was harboring 2 pea crabs, 1 female and 1 male.


Sex distribution was 43.3% females, 39.7% males, and 15.0% sterile or indeterminate. Seven (1.9%) were hermaphrodites with 5 having ova and sperm in separate follicles. In one of the specimens, half of the body was producing female gametes and the other half male gametes. Two specimens had compound follicles with both ova and sperm present.

In February and March genital tissue started to develop in the mantle (Fig. 2A, B), the outer surface of the digestive gland and the pericardium. Female gametes were fastened with a peduncle to the germinal epithelium during development. In ripe female follicles, ova filled the lumina (Fig. 2C) and were angular in shape until spawning at which time they became more rounded. In ripe male follicles, different stages of spermatogenesis could be observed. Germinal epithelium lining the follicles was overlaid by spermatogonia and spermatocytes with numerous meiotic and mitotic figures present. Mature sperm with pink staining flagella filled the middle of follicles (Fig. 2D). At the end of spawning, follicles, with some ripe gametes present, began to shrink and were replaced by connective tissue (Fig. 2E, F).


Gonad indices of the mussels are illustrated in Figure 3. Mussels had already begun gametogenesis in February when sampling started. The major spawning peak was in May followed by a minor one in August. Percentages of developing, spawning, and spent mussels are shown in Figure 4. Spawning specimens were present from April until January with a sharp decline in July, when half of the mussels were again developing and preparing for the fall spawning. Most of the mussels showed some gonad development throughout the entire year. However, in December 50% of specimens examined were resting with no developing gametes present in their mantles, and sexes could not be distinguished (see Fig. 4).


Pathologic Observations

Pathologic observations on histologic sections are listed in Table 2. Four different types of infectious organisms were observed. Intracellular prokaryotes (Chlamydia, Rickettsia or Mycoplasma) were present in digestive epithelial cells of one specimen. These organisms formed basophilic, finely granular inclusions. Ciliates (Ancistrum mytili) were observed on the gills of some mussels. Ciliates were pear shaped with a macro- and micronucleus and were anchored between gill epithelial cells. A small percentage (5.8%) of the mussels had microsporidian infections, Steinhausii mytilovum (Mussel Egg Disease). Parasites formed vacuoles inside the cytoplasm or nuclei of ova where large numbers of microsporia were observed with their characteristic polar vacuoles (Fig. 5). Also, immature cysts with less defined, developing parasites were present. There was an accumulation of granular hemocytes in the follicles containing infected ova. The presence of S. mytilovum was first detected in June (23.3%), after which they were present at prevalences ranging from 0% to 13.3% (Fig. 6). Although S. mytilovum occurred at high prevalence, the proportion of infected ova versus uninfected ova was generally low. In some specimens, several infected ova were present in the same follicle, and several vacuoles were present in a single ovum (Fig. 5).


The October mussel samples exhibited infections by the digenetic trematode, Proctoeces maculatus. The high prevalence of infection (67%) remained until the end of sampling (Fig. 6). Trematodes invaded the gonadal follicles and different stages of maturation, from sporocysts containing germ balls and cercaria to metacercaria, could be observed (Fig. 7A). More mature forms were present in the foot musculature and the pericardial chamber. Occasionally, large abscesses were formed, harboring adult forms of the parasites. These stages of the trematodes had oral and ventral suckers (Fig. 7B) and occasionally the digestive tract appeared to be filled with host hemocytes (Fig. 7C). Sperm and ovaries, as well as ova with developing miracidia within them, were present in several trematodes. Massive hemocytic response surrounded some of the worms. Parasites in different stages of degradation, due to the response of host hemocytes, were seen towards the end of the sampling period. However, in most cases the parasites did not evoke any host response and appeared to have completed their life cycle inside the mussels. Trematodes invaded the follicles, foot, and pericardial cavity, as well as the digestive gland and kidneys; some of the infested mussels were merely a mass of trematodes in different stages of development. Surprisingly, heavily infested mussels appeared healthy even when the tissues were almost entirely displaced by the parasite mass.


The pathologic responses in the mussels included multifocal aggregations of hemocytes in the byssus gland in one specimen, focal aggregation of hemocytes in the digestive diverticulum in one specimen, and abscesses in the digestive diverticulum or the kidney in four specimens (Table 2). Three specimens had pearls in the mantle epithelium. One specimen had degeneration of digestive tubules with deposition of collagen in the surrounding connective tissue. This lesion was surrounded by hemocyte infiltration. Abnormal byssus formation with threads forming concentric structures inside the septal area of the byssus gland and not in the foot groove, as usually occurs, was present in one specimen. Kidney concretions, located inside kidney tubules with varied sizes and shapes, were observed in 12 specimens. There were also hemocyte aggregations in the stomachs of three specimens.

No significant mortalities were observed during the experiment. The meat quality remained high until the end of the summer, after which the mussels were thin and watery with visible trematode abscesses in the mantles. Concurrently, the mussels started to slide down the ropes due to their collective weight on the dropper lines.


Blue mussels occur on the East Coast of North America from the Canadian Maritimes to Cape Hatteras, North Carolina (Gosner 1978, Koehn et al. 1984). Mussels are successfully cultured on the East Coast of the US in Maine and Prince Edward Island (PEI) in Canada. There is no commercial mussel culture south of Long Island Sound. High temperatures may have concurrent negative and positive effects on the culture, therefore, evaluating the biologic potential of mussel longline culture in LIS should be cognizant of these factors.

On the East Coast of North America there are 2 species of blue mussels, Mytilus edulis and M. trossulus. These species do not have reliable morphologic characteristics and confusion about their taxonomy prevails throughout blue mussel research. Blue mussels in northern Canada and Alaska are mainly M. trossulus, but in Newfoundland and Nova Scotia, Canada M. trossulus overlaps with M. edulis (Penney & Hart 1999). In Maine there is a transition zone where M. trossulus dominates at some sites in the northeast whereas M. edulis is the prominent species in central Maine (Rawson et al. 2001). Several commercial mussel farms operate successfully with a mixed seed stock of M. edulis and M. trossulus (Mallet & Carver 1995). M. trossulus is considered to have an economic disadvantage compared with M. edulis in regard to tissue weight and shell height. Mallet and Carver (1995) estimated that the economic value of M. edulis was 1.7 times higher than that of M. trossulus. The only mussel genotype present in LIS is M. edulis (McDonald et al. 1991). This is an obvious benefit for the potential of a LIS mussel longline culture.

Mussels had an exceptional growth rate on the longline reaching market size (60 mm) during the first 6 months after seed set (Table 1). There was no significant additional growth during their second summer (Fig. 8) thus demonstrating that the population had reached their full size during the first growing season. Growth rate is an important criterion in assessing the potential for mussel culture in a given area. Depending on geographic latitudes and environmental factors (temperature, light, salinity, primary productivity, currents, and tides), growth of mussels varies. In Iceland, mussels reach market size (50 mm) in 24 mo after settlement (Thorarinsdottir 1996). In Galicia, Spain, mussels (Mytilus galloprovincialis) generally reach commercial size (80-90 mm) in 1218 mo. In France, M. edulis reach commercial size (40-50 mm) in 24 too, and in the Netherlands commercial size (72 mm, bottom culture) is reached in 36 mo (Chew & King 2000). Langan and Horton (2002) reported a 13-mo production cycle to 55 mm in their New Hampshire deep-water longline. In PEI the production cycle varies from 17-28 mo (McDonald et al. 2002).


No significant mortality was detected in the mussels during the experiment. However, massive mussel beachings on the shores of LIS were observed during the past summers (Sunila 2001). Mussel beaching occurred in the area during August and September of this experiment. This phenomenon affects intertidal mussels during unusually hot summers in LIS. During emersion mussels gape slightly for air breathing, which exposes the animals to desiccation and possible death (De Zwaan 1977). Mussels react to environmental stress, such as high temperature, by cutting their byssi off and trying to sink. During hot summer days emersed mussels jettison their byssi, but because of gas bubbles trapped between their shells they are unable to sink and instead float. Animals are finally washed on shore in piles that may reach a meter high (Sunila 2001). Beached mussels are live and based on histologic observations in good health, but without byssi (Sunila, pers. obs.).

Summer mortality is a well-published and frequent event in cultured mussels (e.g., Myrand & Gaudreault 1995, Incze et al. 1980, Emmett et al. 1987). According to Myrand and Gaudreault (1995) as much as 80% of the population can be affected in PEI. The phenomenon usually affects mussels during their second growing season. There can be significant differences in the temperature tolerance of mussels originating only kilometers apart. Susceptibility to summer mortality appears to be due more to the genetic origin of the mussels than to culture site. Incze et al. (1980) reported decreased growth rate of mussels in an experimental raft culture in Maine when water temperature exceeded 20[degrees]C. Thompson and Newell (1985) compared mussels from Newfoundland, Canada and mussels from LIS (Stony Brook, NY) and demonstrated more severe stress (measured as depression of the clearance rate) at 25[degrees]C in the Newfoundland mussels than the LIS mussels. The Newfoundland mussels also had a considerably higher mortality rate (46%) than the LIS mussels (10%) at 25[degrees]C. They concluded that the mortality patterns and physiologic responses suggested that mussels from southern, warmer waters are better adapted to high temperatures than mussels from northern, cooler waters.

Along with rising seawater temperature, mussels acquired Proctoeces maculatus (Looss 1901), Fellodistomidae, infection (Table 2 and Fig. 6). At the same time mussels started to slide down the ropes, their meats became watery and thin and parasite abscesses were observed on the mantles by the naked eye. In the past, this condition has been referred to as "orange sickness" due to the characteristic orange color of the parasite. The first to observe P. maculatus in LIS mussels was Uzmann (1953), who named them Cercaria milfordensis. He reported a prevalence of 4.3% from Mill Neck, Long Island, New York and prevalences of 6.6% and 7.7% in subsequent years in a pooled sample from Milford and Bridgeport, CT. Later, Stunkard and Uzmann (1959) sampled Mytilus edulis from Connecticut and Massachusetts and observed several life stages of these trematodes, from unencysted metacercaria to gravid adults, in the mussels. They concluded that C. milfordensis is the larval stage of P. maculatus. Cercaria tenuans and C. brachidontis also were conspecific with C. milfordensis and the cercarial stage of P. maculatus. This species may be able to complete its entire life cycle in mussels and a vertebrate final host is not needed. However, in more southern locations tropical and subtropical shallow water bottom feeding marine fish belonging to several different families are reported to harbor adult Proctoeces in their hindguts (Wardle 1980). The ability to accomplish their life cycle within the mussel host may represent a mechanism for P. maculatus to extend its limits of distribution into temperate waters without dependence on its usual tropical fish hosts (Lang & Dennis 1976).

Data about the economic importance of this cosmopolitan parasite to the mussel industry are controversial. Robledo et al. (1994b) reported a low prevalence (less than 1/1000) of P. maculatus in M. galloprovincialis in Spain and concluded that although the parasite caused severe lesions, the low prevalence did not make it a real threat to the mussel industry. This view was supported by Villalba et al. (1997) who reported average prevalences of 0.7% from different locations in Spain, and Canzonier (1972) who reported prevalences of 4% from Ria de Arosa, Spain and 4% in Laguna Veneta, Italy. The parasite was later associated with extensive mortalities of cultured mussels in Laguna Veneta (Munford et al. 1981). However, other etiologic agents as putative causes of the observed mortality were not excluded in the study. In this study, there was no mortality associated with the infection, and the presence of disintegrating parasites surrounded by a strong host response at the end of the year suggested that the mussels were able to suppress the parasites and modulate their life cycle. However, the exceptionally high prevalences observed in this study and Uzmann's (1953) early observations suggest that the mussel longline was deployed right in the midst of an area with high infection potential.

P. maculatus elicited a strong hemocyte response in the host mussel (see Fig. 7). This differs from most mollusk-trematode infections when the host usually doesn't react to healthy parasites. Tripp and Turner (1978) described adult P. maculatus in the pericardial cavity of mussels from New Jersey. They claimed that the trematodes actively ingested hemocytes and actually "grazed" the mussel tissues. Besides eliciting hemocyte response, P. maculatus affects reproduction of the mussels. Feng (1988) studied infected mussels from Ram Island Reef, CT. He reported 33% mean prevalence of infection. In moderate to heavy infections, normal gametogenesis was either impaired or totally absent in 6% to 11% of the infected mussels.

The prevalence of P. maculatus in this study showed seasonality with the trematode absent until August leading to epizootic prevalences in fall and early winter (see Fig. 6). A similar trend was described by Lang and Dennis (1976) in New Jersey and Tripp and Turner (1978) in Delaware. Uzmann (1953) noted that although mussels were extensively studied in the higher latitudes for years, the presence of P. maculatus had not been reported. Based on the results of this study, those of Lang and Dennis (1976), and Tripp and Turner (1978), we conclude that P. maculatus appears at high seasonal prevalences in the mussels close to the southern border of their distribution and could likely affect the potential of mussel culture in this region.

Mussels also acquired infestation with pea crabs, Pinnotheres maculatus (see Table 1). Pea crabs were absent or occurred at low prevalences (3.3%) from February until November when their prevalence reached 20% and peaked to 60% in January. Pea crabs impacted a large portion of the mantle cavity causing deformed gills and a weak and watery visceral mass in infested specimens. Seasonal prevalences of the pea crabs followed what was previously reported about the life cycle of pea crabs in the area (Kelly 1988). The larval stages infest the mussels in mid-September following initial planktonic stages. After several molts, the adult pea crabs leave the host to mate in a copulatory swarming in open water by mid-October, and inseminated females reinfest the host for the winter. Few males are found in mussels following the mating. Males live 1 year or less, and are significantly smaller, while females live 3 to 4 years (Kelly 1988, Pearce 1964).

Proctoeces maculatus infection and Pinnotheres maculatus infestation clearly pose a threat to the health of mussels and subsequently to the potential of commercial culture in LIS. Both parasites lowered the product quality and would likely affect consumer perception. In addition, human consumption of trematode-infested mollusks may be hazardous due to the accumulation of toxic metabolites (butyric and other short-chain fatty acids) resulting from degeneration of the hosts' neutral fats by parasite-secreted enzymes (Bower & Figueras 1989). A third parasite of regulatory significance was Steinhausii mytilovum (Mussel Egg Disease). This can be classified as a parasite that can damage the mussel, but is unlikely to be lethal to the host (Villalba et al. 1997). Microscopical details of the parasite were presented by Sprague (1965) and Sindermann (1990) assigned the protozoan to the phylum Microspora. The condition, naturally, affects only females. The parasite was first observed in maturing ova in the beginning of gametogenesis (Fig. 5). Prevalence decreased in June after the first spawning, but increased again with the second spawning peak (Fig. 6). The infection is believed to affect the fecundity of mussels (Villalba et al. 1997), but this view was disputed by Robledo et al. (1994b) who stated that the low number of infected ova would be an insignificant factor because of the large number of ova produced by the animals.

There were several other parasites and pathologic conditions present that are considered of no commercial importance (Table 2). Prokaryotic inclusions were observed inside digestive epithelial cells of one specimen (0.3%, Table 2). These can be caused by Rickettsia, Chlamydia, or Mycoplasma, which are obligate intracellular organisms indistinguishable from each other under light microscopy. While they cause serious diseases in humans and domestic animals, micro-organisms from these three groups are considered ubiquitous and of low pathogenicity to marine bivalves (Harshbarger et al. 1977, Lauckner 1983). Finally, there was a low prevalence (3.1%, Table 2) of the ciliate Ancistrum mytili on the gills of the mussels. However, this species is considered ubiquitous and not of regulatory significance (Moret et al. 1999).

Three of the mussels contained pearls (Table 2), a condition generally attributed to the trematode metacercaria in the family Gymnophallidae. The number of pearls in the mussels in Maine increased with age from zero in 2-year class animals up to an average of 43 pearls/mussel in 8-year-old mussels (Lutz & Hidu 1978). Pearl formation may seriously affect marketability of these animals, but because of the fast growth rate of the mussels, pearl formation would not have time to reach prevalences that would affect a LIS suspended culture.

Abnormal byssal thread formation was observed in one specimen in the present experiment. The condition was previously reported by Sunila (1987). In the affected mussel, byssus threads formed inside the byssus glands as massive concentric structures may impair the attachment of the mussel. This condition is insignificant because of the low prevalence.

Kidney concretions were also observed in some mussels (Table 2). The concretions are formed principally of amorphous calcium phosphate (Doyle et al. 1978) and are larger and more numerous in quahogs (Mercenaria mercenaria) from polluted areas than in kidneys of quahogs obtained from more pristine areas (Rheinberger et al. 1979). Presence of kidney stones in LIS mussels is not surprising considering the anthropogenic stress in the area. Kidney stones in mussels were previously reported by Sunila (1987) who observed higher prevalences in sampling locations closer to an iron and steel factory compared with animals sampled from more pristine bottoms in the Baltic Sea.

While evaluating biologic potential for mussel longlines in LIS, two opposite forces clearly came together in the area. The first, a positive factor, was the exceptional growth rate that occurred compared with rates previously published for commercial mussel operations. The second, a negative factor, was the epizootic prevalence of Proctoeces maculatus, which is a species usually characteristic of tropical and temperate regions. Because of its high prevalence in sampled animals P. maculatus could be considered an economically important parasite of mussel culture in the area. Figure 8 demonstrates the size of mussels and pathology prevalences for each sampling month. The presence of fouling organisms was also included in the pathology prevalence data. There was a 5-month "window of opportunity" for mussel culture between February and June, during which mussels were already of marketable size, free of fouling organisms and parasites and had good quality meats. Based on histologic sections, mussels were also healthy in July and August, but at that time they were extensively fouled with Crepidula fornicata. Providing a marketable product during those months would require the use of cleaners and debyssers. No investments were made to import such commercially available equipment to the area during the present, experimental stage of mussel culture. However, the fast growth, good quality of meat and the unfouled shells between February to June suggest the potential for a seasonal product.

The reproduction data of this study indicates that seed lines in the Milford area should be deployed in June. Although spawning specimens dominated in June (Fig. 4), the gonad index had already decreased, indicating that the spawning peak was in May (Fig. 3). When predicting the right timing for future seed line deployments it must be kept in mind that mussel spawning and setting times differ significantly in different areas in LIS as well as interannually (Fell & Balsamo 1985, Brousseau 1983, Newell et al. 1982). The eastern end of LIS is more oceanic and differs significantly from the western, estuarine end of LIS. Accordingly, in the eastern end of LIS, mussel spawning followed the pattern of more oceanic mussel populations (Fell & Balsamo 1985). This is in accordance with Newell et al. (1982) who observed much later spawning in Shinnecock, NY (more oceanic south shore of Long Island, Atlantic Ocean) than in Stony Brook, NY (more estuarine north shore of Long Island, LIS). Therefore, providing seed for commercial operations would require the deployment of multiple seed lines, timed differently in different parts of the sound.

Apart from the biologic factors, expansion of mussel longline culture in LIS could meet serious regulatory obstacles. LIS, "The Urban Sea," is a heavily populated and highly prized body of water, and any attempt to use the water column would conflict with the needs of different user groups, such as recreational boating and fishing in inshore areas and commercial ship traffic in deeper water. However, small-scale culture of mussels could provide supplemental income to shellfishermen seeking to diversify their business.

Sampling times, average size of mussels (n = 30) collected, seawater
temperature, fouling with Crepidula fornicata and the prevalence of
Pinnotheres maculatus (% of specimens affected) in a mussel
(Mytilus edulis) longline culture over the course of 1 year in Long
Island Sound, Connecticut.

Sampling Size Temperature Crepidula Pinnotheres
 Time (mm) [degrees]C fornicata % maculatus %

 2.20.02 56 4.9 0 3.3
 3.21.02 52 6.0 0 3.3
 4.23.02 61 11.4 0 0
 5.21.02 60 12.2 0 0
 6.27.02 68 22.3 0 0
 7.30.02 66 23.7 86.6 0
 8.30.02 72 23.2 70 0
 9.30.02 62 20.8 93 3.3
10.31.02 70 12.4 100 0
11.19.02 70 10.3 100 20
12.16.02 69 5.9 100 23.3
01.16.03 66 2.5 100 60.0


A list of infectious organisms, pathologic responses and their
average prevalences based on histopathological observations over
the course of 1 year in mussels (Mytilus edulis, n = 360) from a
longline culture in Long Island Sound, Connecticut.

Histopathological Observations Prevalence %

Infectious organisms
 Prokaryotic inclusions in digestive cells 0.3
 Ancistrum mytili on the gills 3.1
 Steinhausii mytilovum in the ova 5.8
 Proctoeces maculatus 20.8
Pathologic responses
Acute inflammation:
 Multifocal aggregation of hemocytes in the byssus 0.3
 Focal aggregation of hemocytes in the digestive 0.3
 Abscesses in the digestive diverticula or the kidney 1.1
Chronic inflammation:
 Pearl formation 0.8
Degenerative responses:
 Degeneration of digestive tubules with collagen 0.3
 Abnormal byssal thread formation 0.3
 Kidney concretions 3.3
 Hemorrhage in the stomach 0.8


The authors thank Mr. Walter Canzonier from Port Norris, New Jersey and Mr. James Winstead from EPA, Florida for their constructive and extensive reviews of the draft manuscript. Mr. Winstead also helped in the diagnosis of Proctoeces maculatus trematode. Dr. Lillemor Svardh from Tjarno Marine Biological Laboratory, Sweden is appreciated for her input in the early stages of the manuscript. Shannon Kelly is acknowledged for detailed editing of the manuscript. This work was supported in part by Connecticut Sea Grant, Development Project M/PD-1 (2001).


Bayne, B. L. 1964. Primary and secondary settlement in Mytilus edulis L. (Mollusca). J. Anim. Ecol. 33:513-524.

Bower, S. M. & A. J. Figueras. 1989. Infectious diseases of mussels, especially pertaining to mussel transplantation. World Aquaculture 20: 89-93.

Brousseau, D. J. 1983. Aspects of reproduction of the blue mussel Mytilus edulis (Pelycypoda; Mytilidae) in Long Island Sound. Fish. Bull. 81: 733-739.

Canzonier, W. J. 1972. Cercaria tenuans, larval trematode parasite of Mytilus and its significance in mussel culture. Aquaculture 1:267-278.

Chew, K. K. & T. L. King. 2000. Molluscan culture. In: R. R. Stickney, editor. Encyclopedia of Aquaculture. New York. Chichester. Weinheim. Brisbane. Singapore. Toronto: John Wiley & Sons, Inc. pp. 540-548.

Chipperfield, P. N. J. 1953. Observations on the breeding and settlement of Mytilus edulis (L.) in British waters. J. Mar. Biol. Assoc. U.K. 32:449-476.

Corayer, T. 2003. Deep water, longline shellfish farming in Narragansett Bay. J. Shellfish Res. 22:291-292.

De Zwaan, A. 1977. Anaerobic energy metabolism in bivalve mollusks. Oceanogr. Mar. Biol. Ann. Rev. 15:103-187.

Doyle, L. J., N. J. Blake, C. C. Woo & P. Yevich. 1978. Recent biogenic phosphorite: concretions in mollusk kidneys. Science 199:1431-1433.

Elston, R. A., J. D. Moore & K. Brooks. 1992. Disseminated neoplasia of bivalve mollusks. Rev. Aqua. Sci. 6:405-406.

Emmen, B., K. Thompson & J. D. Popham. 1987. The reproductive and energy storage cycles of two populations of Mytilus edalis (Linne) from British Columbia. J. Shellfish Res. 6:29-36.

Engle, J. B. & V. L. Loosanoff. 1944. On season of attachment of larvae of Mytilus edulis Linn. Ecology 25:433-440.

Fell, P. E. & A. M. Balsamo. 1985. Recruitment of Mytilus edulis L. in the Thames estuary, with evidence for differences in the time of maximal settling along the Connecticut shore. Estuaries 8:68-75.

Feng, S. Y. 1988. Host response to Proctoeces maculatus infection in the blue mussel, Mytilus edulis L. J. Shellfish Res. 7:118.

Figueras, A. J., C. F. Jardon & J. R. Caldas. 1991. Diseases and parasites of mussels (Mytilus edulis, Linneaus, 1758) from two sites on the east coast of the United States. J. Shellfish Res. 10:89-94.

Gosner, K. L. 1978. A field guide to the Atlantic seashore from the Bay of Fundy to Cape Hatteras. Boston. New York: Houghton Mifflin Company. 327 pp.

Harshbarger, J. C., S. C. Chang & S. V. Otto. 1977. Chlamydia (with phages), mycoplasms, and rickettsia in Chesapeake Bay bivalves. Science 196:666-668.

Hrs-Brenko, M. 1971. The reproductive cycle of the Mytilus galloprovincialis Lamk. in the Northern Adriatic Sea and Mytilus edulis L. at Long Island Sound. Thalassia Jugoslavica 7:533-542.

Incze, L. S., R. A. Lutz & L. Watling. 1980. Relationships between effects of environmental temperature and seston on growth and mortality of Mytilus edulis in a temperate northern estuary. Mar. Biol. 57:147-156.

Jones, J. B., P. D. Scotti, S. C. Dearing & B. Wesney. 1996. Virus-like particles associated with marine mussel mortalities in New Zealand. Dis. Aqua. Org. 25:143-149.

Kautsky, N. 1982. Quantitative studies on gonad cycle, fecundity, reproductive output and recruitment in a Baltic Mytilus edulis population. Mar. Biol. 68: 143-160.

Kelly, M. S. 1988. Aspects of the life history of Pinnotheres maculatus and its effects on Mytilus edulis. Master's Thesis. The University of Connecticut. 76 pp.

King, G. & J. Cortes-Monroy. 2002. Mussel farming in the United Sates. Bull. Aquacul. Assoc. Canada 102-3:49-57.

Koehn, R. K., J. G. Hall, D. J. Innes & A. Z. Zera. 1984. Genetic differentiation of Mytilus edulis in eastern North America. Mar. Biol. 70: 117-126.

Lane, D. J., A. R. Beaumont & J. R. Hunter. 1985. Byssus drifting and the drifting threads of the young postlarval mussel Mytilus edulis. Mar. Biol. 84:301-308.

Lang, W. H. & E. A. Dennis. 1976. Morphology and seasonal incidence of infection of Proctoeces maculatus (Looss, 1901) Odhner, 1911 (Trematoda) in Mytilus edulis L. Ophelia 15:65-75.

Langan, R. 2000. Submerged longline culture of blue mussels (Mytilus edulis) at an open site in the Gulf of Maine. J. Shellfish. Res. 19:575.

Langan, R. & C. Horton. 2002. Submerged longline culture of blue mussels in exposed oceanic environments. Bull, Aquacul. Assoc. Canada 1023:96.

Lauckner, G. 1983. Diseases of mollusca: Bivalvia. Ch.13. In: O. Kinne, editor. Diseases of marine animals, vol. 2. Introduction: Bivalvia to Scaphopoda. New York. Chichester. Brisbane. Toronto: John Wiley & Sons, Inc. pp. 477-879.

Lubet, P. 1957. Cycle sexuel de Mytilus edulis et M. galloprovincialis (Lmk) dans le basin d'Arcachon (Gironde). Annie Biol. 33:19-29.

Lutz, R. A. & H. Hidu. 1978. Some observations on the occurrence of pearls in the blue mussel, Mytilus edulis L. Proc. Natl. Shellfish. Assoc. 68:17-37.

Mallet, A. L. & C. E. Carver. 1995. Comparative growth and survival patterns of Mytilus edulis in Atlantic Canada. Ca. J. Fish. Aquac. Sci. 52:1873-1880.

McDonald, C., R. Gallant & C. Couturier. 2002. Canadian mussel aquaculture; an industry with room to grow. Bull. Aquacul. Assoc. Canada 102-3:87-92.

McDonald, J. H., R. Seed & R. K. Koehn. 1991. Allozymes and morphological characters of three species of Mytilus in the northern and southern hemispheres. Mar. Biol. 111:323-333.

McLeod, D. 2002. The life and times of the "Myti" mussel. Bull. Aquacul. Assoc. Canada 102-3:8-16.

Moret, K., C. Couturier, G. J. Parsons & K. Williams. 1999. Monitoring shellfish health in Newfoundland. J. Shellfish. Res. 18:297-298.

Munford, J. G., L. Da Ros & R. Strada. 1981. A study on the mass mortality of mussels in Laguna Veneta. J. Worm Maricul. Soc. 12:186199.

Myrand, B. & J. Gaudreault. 1995. Summer mortality of blue mussels (Mytilus edulis Linneaus, 1758) in the Magdalen Islands (southern Gulf of St. Lawrence, Canada). J. Shellfish. Res. 14:395-404.

Nelson, T. C. 1928. Pelagic dissoconchs of the common mussel, Mytilus edulis, with observations on the behavior of the larvae of allied general. Biol. Bull. 55:180--192.

Newell, R. I. E., T. J. Hilbish, R. K. Koehn & C. J. Newell. 1982. Temporal variation in the reproductive cycle of Mytilus edulis L. (Bivalvia, Mytilidae) from localities on the east coast of the United States. Biol. Bull. 162:299-310.

Pearce, J. B. 1964. On reproduction in Pinnotheres maculatus. Biol. Bull. 127:384.

Penney, R. W. & M. J. Hart. 1999. Distribution, genetic structure, and morphometry of Mytilus edulis and M. trossulus within a mixed species zone. J. Shellfish Res. 18:367-374.

Rasmussen, L. P. D. 1986. Virus-associated granulocytomas in the marine mussel, Mytilus edulis, from three sites in Denmark. J. Invert. Pathol. 48:117-123.

Rawson, P. D., S. Hayhurst & B. Vanscoyoc. 2001. Species composition of blue mussel populations in the northeastern Gulf of Maine. J. Shellfish Res. 20:31-38.

Rheinberger, R., G. Hoffman & P. Yevich. 1979. The kidney of the quahog (Mercenaria mercenaria) as a pollution indicator. In: Animals as monitors of environmental pollutants. Washington, DC: National Academy of Sciences. pp. 119-133.

Robledo, J. A. F., V. Boulo, E. Mialhe, B. Despres & A. Figueras. 1994a. Monoclonal antibodies against sporangia and spores of Marteilia sp. (Protozoa: Ascetospora). Dis. Aquac. Org. 18:211-216.

Robledo, J. A. F., M. M. Santardm & A. Figueras. 1994b. Parasite loads of rafted blue mussels (Mytilus galloprovincialis) in Spain with special reference to the copepod, Mytilicola intestinal&. Aquaculture 127:287-302.

Seed, R. 1969. The ecology of Mytilus edulis L. on exposed rocky shores. I. Breeding and settlement. Oecologia 3:277-316.

Seed, R. 1975. Reproduction in Mytilus (Mollusca: Bivalvia) in European waters. Publ. Staz. Zool. Napoli. 39:317-335.

Sindermann, C. J. 1990. Principal diseases of marine fish and shellfish, vol. 2. Diseases of marine shellfish. San Diego: Academic Press. 516 pp.

Sprague, V. 1965. Observations on Chytridiopsis mytilovum (Field), formerly Haplosporidium mytilovum Field, (Microsporida?). J. Protozool 12:385-389.

Stunkard, H. W. & J. R. Uzmann. 1959. The life-cycle of the digetic trematode, Proctoeces maculatus (Looss 1901) Odhner, 1911 Syn. P. subtenuis (Linton, 1907; Hanson, 1960) and description of Cercaria adranocerca N.S.P. Biol. Bull. 116:184-193.

Sunila, I. 1981. Reproduction of Mytilus edulis L. (Bivalvia) in a brackish water area, the Gulf of Finland. Ann. Zool. Fennici. 18:121-128.

Sunila, I. 1987. Histopathology of mussels (Mytilus edulis L.) from the Tvarminne area, the Gulf of Finland (Baltic Sea). Ann. Zool. Fennici. 24:55-69.

Sunila, I. 2001. Mussel beaching in Long Island Sound. Long Island study fall update. NY Sea Grant Extension Program. Stony Brook, New York. 4 pp.

The Island Institute 1999. The Maine guide to mussel raft culture. Island Institute. Rockland, Maine. 32 pp.

Thompson, R. J. 1984. The reproductive cycle and physiological ecology of the mussel Mytilus edulis in a subarctic, nonestuarine environment. Mar. Biol. 79:277-288.

Thompson, R. J. & R. Newell. 1985. Physiological responses to temperature in two latitudinally separated populations of the mussels, Mytilus edulis. In: Proceedings of the 19th European Marine Biology Symposium. Cambridge. United Kingdom: Cambridge University Press. pp. 481-495.

Thorarinsdottir, G. G. 1996. Gonad development, larval settlement and growth of Mytilus edulis L. in a suspended population in Hvalfjordur, south-west Iceland. Aquacult. Res. 27:57-65.

Tripp, M. R. & R. M. Turner. 1978. Effects of the trematode Proctoeces maculatus on the mussel Mytilus edulis. In: L. A. Bulla & T. C. Cheng, editors. Comparative Pathobiology, vol. 4. New York: Plenum Press. pp. 74-84.

Uzmann, J. R. 1953. Cercaria milfordensis Nov. sp., a microcercous trematode larva from the marine bivalve, Mytilus edulis L. with special reference to its effects on the host. J. Parasit. 39:445-45 h

Villalba, A., S. G. Mourelle, M. J. Carballal & C. L6pez. 1997. Symbionts and diseases of farmed mussels Mytilus galloprovincialis throughout the culture process in the Rias of Galicia (NW Spain). Dis. Aquac. Org 31:127-139.

Villalba, A., S. G. Mourelle, M. C. L6pez, M. J. Carballal & C. Azevedo. 1993. Marteiliasis affecting cultured mussels Mytilus galloprovincialis of Galicia (NW Spain). I. Etiology, phases of the infection, and temporal and spatial variability in prevalence. Dis Aquac. Org. 16:61-72. Wardle, W. J. 1980. On the life cycle stages of Proctoeces maculatus (Digenea; Fellodistomidae) in mussels and fishes from Galveston Bay, Texas. Bull. Mar. Sci. 30:737-743.

Weiss, H. M. 1995. Marine animals of Southern New England and New York. State geological and natural history survey of Connecticut. Department of Environmental Protection. Bulletin 115.


* Corresponding author. E-mail:

(1) State of Connecticut, Department of Agriculture, Bureau of Aquaculture, P.O. Box 97, Milford, Connecticut 06460; (2) Jessie D., Inc., 68 Anchorage Drive, Milford, Connecticut 06460; (3) Cornell University, 126C Valentine Place, Ithaca, New York 14850; (4) Connecticut Sea Grant, University of Connecticut, 1084 Shennecossett Road, Groton, Connecticut 06340
COPYRIGHT 2004 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2004, Gale Group. All rights reserved. Gale Group is a Thomson Corporation Company.

 Reader Opinion




Article Details
Printer friendly Cite/link Email Feedback
Author:Getchis, Tessa
Publication:Journal of Shellfish Research
Geographic Code:1USA
Date:Dec 1, 2004
Previous Article:Reproduction of the lion's paw scallop Nodipecten subnodosus sowerby, 1835 (bivalvia: pectinidae) from Laguna Ojo de Liebre, B.C.S., Mexico.
Next Article:First occurrence of the nonindigenous green mussel, Perna viridis (Linnaeus, 1758) in coastal Georgia, United States.

Related Articles
Reproductive cycle of coexisting mussels, Mytilus californianus and Mytilus galloprovincialis, in Baja California, New Mexico.
Localized synchronous spawning of Mytilus californianus conrad in Barkley Sound, British Columbia, Canada.
Mini-review: distribution of the Mediterranean mussel Mytilus galloprovincialis (Bivalvia: Mytilidae) and hybrids in the Northeast Pacific.
First occurrence of the nonindigenous green mussel, Perna viridis (Linnaeus, 1758) in coastal Georgia, United States.
Evaluation of the neutral red retention assay as a stress response indicator in cultivated mussels (Mytilus spp.) in relation to seasonal and...
Selection response for growth rate (shell height and live weight) in the Chilean blue mussel (Mytilus chilensis Hupe 1854).
Particular aspects of gonadal cycle and seasonal distribution of gametogenic stages of Mytilus galloprovincialis cultured in the Estuary of Vigo.
Distribution of Mytilus edulis and M. trossulus on the Gaspe Coast in relation to spatial scale.
Allozyme identification of mussels (Bivalvia: Mytilus) on the pacific coast of South America.
Valve-gape response times in mussels (Mytilus edulis)--effects of laboratory preceding-feeding conditions and in situ tidally induced variation in...

Terms of use | Copyright © 2014 Farlex, Inc. | Feedback | For webmasters