A description of the pathology of epizootic shell disease in the American lobster, Homarus americanus, H. Milne Edwards 1837.

ABSTRACT Epizootic shell disease is responsible for great economic losses in the commercial lobster fishing industry in the northeastern United States. Histologic examination of the carapace from affected American lobsters showed the disease is primarily caused by bacteria invading the carapace from the surface of the shell. The carapace itself appears normal in areas that are not eroded and there seems to be an appropriate inflammatory response mounted by the lobsters' hemocytes and tissues to the erosive lesions.

KEY WORDS: lobster, epizootic shell disease, carapace, bacteria, Homarus americanus, erosion, inflammatory cuticle, melanization, pathology

INTRODUCTION

The American lobster (Homarus americanus H. Milne Edwards, 1837) is an important part of the New England commercial fisheries. In 1999, Maine harvested 20,000 metric tons yielding $133 million in profits (Floreto et al. 2000). In Rhode Island the value of the catch decreased from $24.8 million in 1999 to $19.7 million in 2000, and in 2001, $15.6 million (National Marine Fisheries Service, 2001, unpublished report). The Massachusetts fishery decreased from $59.5 million in 1999 to $54.9 million in 2000, $45.5 million in 2001 (Dean et al. 2002) and $49.4 million in 2002 (Dean et al. 2004). Decrease in catch value is partially due to the increased fishing pressure on the resource and an associated decreased average size of the catch. However, off the shores of New England epizootic shell disease has significantly affected the commercial value, if not the abundance, of lobsters.

Three main types of shell disease are recognized and two are well described. The first well-described form of shell disease, impoundment shell disease, occurs in American lobsters and was described by Smolowitz et al. (1992). Early lesions are bilaterally symmetrical and are centered around setal cores on the dorsum of the animal (Bullis 1989). In this form, lesions consist of round blackened focal erosions in the carapace that coalesce as the disease worsens, thus increasing the surface area of infected carapace. In its most severe form the lesions cover the entire surface of the carapace (Smolowitz et al. 1992). A common histologic characteristic of impoundment shell disease is the scooped out appearance of the eroded cuticle. The onset of the disease is related to overcrowding, poor water quality, and inadequate diet associated with winter impoundments (Fisher 1988, Smolowitz et al. 1992, Prince et al. 1995). Bacteria were the most common organisms identified on the surface of the lesions (Smolowitz et al. 1992).

The second form of well-described shell disease, burn spot or rust spot shell disease appears as individual circular blackened lesions at various locations on the body, and is attributed to invasions by several different fungi (Stewart 1980, Burns et al. 1979) and/or bacteria (Rosen 1970, Sindermann 1979). Burn spot has been reported in American lobsters in offshore canyons at a prevalence up to 8% in some areas (Ziskowski et al. 1996) and may be similar to early forms of impoundment shell disease. It is believed that many of the lesions seen in this disease begin at the pores and pits in the carapace (Malloy 1978). Researchers have suggested that burn spot shell disease could be used as an environmental health indicator because it is commonly associated with pollution (Ziskowski et al. 1996).

Shell disease of inshore populations of H. americanus have existed at enzootic levels for decades (B. Estrella, Massachusetts Dept. of Marine Fisheries, pers. comm., Estrella 1991). It was first noted at epizootic levels in 1999. This third form of shell disease is poorly described and presents as severe erosive shell disease affecting the dorsal carapace of Homarus americanus. The range of epizootic shell disease extends from eastern Long Island Sound to the near shore waters of southeastern Massachusetts. Ongoing surveys (Castro & Angell 2000, CT Dept. of Environmental Protection 1999, Estrella 1991) found that the prevalence of shell disease in 1996 and 1997 was relatively low (0% to 5.6%) in southern New England. Yet by the year 2000, the percentage of diseased lobsters approached or exceeded 20% in near shore coastal areas from southern Massachusetts to eastern Long Island Sound. Percentage of affected animals in Rhode Island rose to 42.9% by the year 2001. Areas with a high prevalence of diseased populations were localized to Rhode Island, eastern Long Island Sound (22.7% in 2001) and Buzzards Bay (11.6% in 2001). This is in contrast to western and central Long Island Sound and the offshore canyons where shell disease prevalence in 2001 was less than 3%. Recently, still low, but increasing, numbers of animals affected by the disease have been seen in Cape Cod Bay, more northern waters along the coast of Massachusetts (Bruce Estrella, MA Dept of Marine Fisheries, pers. comm.) and one area on the coast of Maine (Carl Wilson, Maine Dept. of Marine Resources, pers. comm.).

Berried female lobsters are more severely affected by epizootic shell disease. This finding was first reported when this form of shell disease was considered enzootic (Estrella 1991). The increased prevalence and severity of erosive shell lesions seen in berried females are believed to develop because of the long periods between molting that results from egg incubation (vs. nonberried females or males). However, some berried females with severe erosive lesions molt with eggs still attached (Laufer et al. 2005). To better understand epizootic shell disease, we evaluated lobster populations from the northeast using gross and microscopic methods of examination. Bacterial sampling was also conducted (Chistoserdov et al. 2005).

MATERIALS AND METHODS

Sampling

Thirty-seven American lobsters (Homarus americanus) with severe erosive lesions characteristic of epizootic shell disease were collected from areas reported to contain a high prevalence of shell diseased lobsters. Lobsters were collected by the Connecticut Department of Environmental Protection, Massachusetts Division of Marine Fisheries and New York Department of Environmental Conservation. In total, 25 animals from Eastern Long Island Sound, NY; 5 animals from Buzzards Bay, MA and 1 animal from Vineyard Sound, MA were examined in 2001. Six animals without shell disease were also collected from Long Island Sound by the CT DEP. Lobsters were held in flowing seawater for 1 to 2 wk before necropsy. Necropsies occurred on April 5, 2001 and June 29, 2001.

As a part of A. Hsu's Masters Thesis work, an additional 27 lobsters were sampled from various locations along the New England coast from May 2002 to August 2002. These animals were collected by the Maine, Atlantic Offshore, Massachusetts, Rhode Island and Connecticut Commercial Lobstermans' Associations. They were kept cool and necropsied immediately upon arrival to prevent change in species or loss of potential infectious agents on the surface of the carapace.

Necropsy

At necropsy, each animal was given a unique identification number and was evaluated for carapace lesion prevalence and severity following the methods developed by Bruce Estrella, MA Dept. of Marine Fisheries (Estrella 1991, Estrella, Lobster Shell Disease Workshop, Millstone Environmental Laboratory, Waterford, CT, June 15, 2000) (Table 1). The dorsal carapace of each necropsied lobster was photographed. A midsagittal cut was made from the posterior of the cephalothorax to just behind the rostrum allowing the animal to exanguinate. One half of the carapace and the underlying connective tissues was carefully removed and fixed in 10% formalin in seawater. Additional lesions from other areas of the carapace, such as the dorsal abdomen and rostrum, were removed with the underlying epithelium and connective tissues, and fixed immediately. Internal organs were examined grossly and any abnormalities were noted. Internal tissues selected for examination in all animals included sections of gonad, hepatopancreas, kidney, neural cord, gill, pyloric stomach, abdominal muscle and sections of any gross abnormalities identified.

Tissue Processing

After fixation of greater than 1 wk, the carapace samples were decalcified in a 5% formic acid solution. All tissues were trimmed so that sections contained all layers of the carapace and the attached underlying epithelium and connective tissues. Tissues were processed in paraffin using standard histologic methods (Luna 1992). Tissue sections of 6 x 11 thickness were cut and stained with hematoxylin and eosin stains (Bullis & McCafferty 1995). Sections from selected blocks were also stained with Gram or Gomori methemine-silver nitrate (GMS) stains (Luna 1992, Presnell & Schreibman 1997).

Microscopic Evaluation

Lesion appearance/progression was divided into 3 stages based on the depth of the erosions into the carapace. Characteristics of each stage are described. Other tissues collected from each animal were histologically evaluated for occurrence of any lesions or other abnormalities. Normal appearing portions of the carapace, from the same lobsters, and sections of hard carapace from unaffected lobsters were used as microscopic controls.

Normal carapace formation is described in stages. The stage designation depends on the number of carapace layers that have been laid down by the cuticular epithelium (Aiken 1980). The last stage, before ecdysis begins, is termed C4/D0. At this stage, the cuticular epithelium has finished laying down all five layers of the current carapace (Fig. 1). The outermost layer, the epicuticle, is divided into the cement layer (deposited post ecdysis by the tegmental glands of the subcutis) and a wax layer, which is deposited just before ecdysis by secretions from the epidermal cells. The innermost layers of the epicuticle consist of an outer and an inner porous trilaminarate, membranous tissue. The wax secretions, produced by underlying epithelial cells, arrive at the surface by way of the wax canals. Wax canals are small tributaries that originate from larger pore canals. Pore canals transverse the inner layers of the cuticle. Wax and pore canals are formed by the epidermal cells in a vertical fashion as the cuticle is produced (Neville 1975).

[FIGURE 1 OMITTED]

The exocuticle is the only other layer produced before ecdysis begins (Stage D1) (and is calcified after ecdysis has ended). The layers internal to the exocuticle consists of the calcified endocuticle, uncalcified endocuticle and finally the membranous layer. These layers are laid down in a sequential manner after ecdysis (Aiken 1980). All of these layers (except the epicuticle) can be simplistically described as being composed of a woven, spiraling lattice of chitin crystals within which a protein matrix is deposited. The configuration of the chitin crystals and the types of protein forming the matrix between the chitin crystalline lattice vary within the carapace layers (Neville 1975). Importantly, the entire carapace is laid down between molts by the simple columnar epithelium that underlays the cuticle.

The uncalcified endocuticle is laid down in C2/C3 and the membranous layer is secreted in stage C4. The membranous layer is produced last and is absorbed before ecdysis allowing separation between the old carapace and the newest portions of the new carapace (epicuticle and exocuticle). It is often termed the ecdysial membrane (Aiken 1980). In stage C2, the epicuticle, exocuticule and calcified endoculticle is formed. In C3, the uncalcified endocuticle is completed Histologically, it is often hard to identify the end of stage C2 and the beginning of stage C3.

The inner epicuticle contains extracellular dihydrophenols that were deposited as the epicuticle was produced. Phenol oxidases (also deposited during formation in the epicuticle as an extracellular enzyme) can oxidize the phenols resulting in "tanning" of the normal (uninfected) cuticle or can form melanized brown/black hardened areas in the cuticle when the cuticle is damaged. The melanizing reaction can be identified histologically by a yellow/ brown discoloration of the affected tissues (Bang 1983, Ghidalia 1985, Lightner & Redman 1977). Melanizing proteins are also present to varying extent in the exocuticle and calcified endocuticle of the carapace (Neville 1975). Melanization of inner layers of cuticle can occur by deliverance of substrate and the phenol oxidase activator through the pore canals (Unestam & Ajaxon 1976).

RESULTS

Gross Morphologic Findings

The gross ratings for the affected lobsters examined ranged from 2 to 3 with 12/25 of the animals examined in 2001 and 17/22 of animals examined in 2003 rated as 3. Six animals examined in 2003 were rated 0. Grossly, epizootic shell disease was characterized by dorsal midline erosions into the carapace of the cephalothorax (Fig. 2). In extreme cases, lesions extend laterally from the midline to cover the opercula and/or extended along, and laterally from, the dorsal midline of the abdominal segments. Lesions affected the hard portions of the carapace and rarely affected the arthrodial membranous joints that separate them. Only in the most severe cases were lesions noted on the claws and the ventral carapace. Erosions were characterized grossly by brown/tan/black, irregular, granular surfaces. Granular tissues at the base of the erosive lesions varied from firm/hard, to rubbery, to soft and thin in texture. Underlying internal soft connective tissues were not seen from the surface of the lesions in almost all cases.

[FIGURE 2 OMITTED]

Microscopic Findings

Evaluation of lesions from all animals show a progressive pattern of erosion from the shell surface into deeper layers of the carapace. All animals examined appeared to be in C3 or C4/DO stages of the molt cycle (Aiken 1980).

Erosions were grouped into three general categories based on microscopic evaluation of the depth of the erosions into the preexistent hard carapace. Associated characteristic inflammatory responses to the erosions were determined for each category. True ulceration of the carapace (total destruction of the carapace and its epithelium with exposure of the underlying connective tissues), which occurred rarely was also described. Arthrodial membranes (the joints) were not significantly affected in epizootic shell disease.

Erosions

Category 1: Shallow Epicuticular and Exocuticular Lesions

Shallow erosions in the epicuticle and exocuticle showed either multifocal pitting or more extensive areas of tissue loss in superficial carapace layers. Erosions extended from the surface into the underlying, normal appearing exocuticule. The leading edge of many erosions appeared to originate from the surface and progress into the exocuticle through the cuticular pores/wax canals and more rarely through setal canals or tegmental gland ducts (Fig. 3A).

[FIGURE 3 OMITTED]

The ventral and lateral edges of the erosions were characterized by a brown/gold color indicating activation of the melanistic (phenolic) inflammatory response. Rarely was significant inflammation or other abnormality noted in cuticular layers internal to the exocuticle or in the epithelium and underlying connective tissues of the erosive lesions. Bacterial colonies, appearing as stacks of short rods, were commonly noted within erosions in the crystalline chitin lattice and were present at the leading edge of the lesions (Fig. 3B).

Category 2: Moderately Deep Erosions Into the Calcified Endocuticle

Moderate lesions consisted of erosions into the calcified endocuticle. Bacteria, often present in large numbers, were the primary organisms found at the leading edge of the lesions. The bacteria did not distort the crystalline chitin lattice as it invaded but instead removed and replaced the protein matrix between the lattice crystals. This resulted in "pillars" of remaining lattice that projected from the floors (leading edge) of the lesions (Fig. 4A). In other areas, vertical shafts surrounded by melanized endocuticle, penetrated into the underlying intact cuticular layers (Fig. 4B and 4C). Hyperplasia and hypertrophy of the underlying cuticular epithelium was evident and moderate numbers of inflammatory cells consisting of granular and agranular hemocytes were present in the underlying connective tissues.

[FIGURE 4 OMITTED]

In many affected animals, identified as being in stage C4/D0 in unaffected portions of the cuticle, the occurrence of a de novo inflammatory cuticle was identified between the cuticular epithelium and the older uncalcified endocuticle/membranous layer. The inflammatory cuticle was present only in deeply eroded areas of the cuticle, varied in thickness (up to 160 [micro]m) and was composed of pale eosinophilic, lamellar tissue similar to the uncalcified cuticle/membranous layer (Fig. 4A). Animals that were not in stage C4/D0 did not produce the inflammatory cuticle in eroded areas of their carapace.

Often in the layered area between the older cuticle and the inflammatory cuticle or between the eroded cuticle and the cuticular epithelium were accumulations of necrotic hemocytes and edematous, fibrin-like material (Fig. 4A). In these areas sometimes also present were foci of melanized debris that occurred often in a half moon fashion, around an origin point underlying the degenerating cuticle. In some instances such areas could be associated with pores, seta and tegmental gland canals that traversed the overlying carapace. Occasionally, small round granulomas with melanized centers were noted in the subepithelial connective tissues.

The most commonly identified secondary organisms in the category two erosions were small protistans (Fig. 4C and 4D) that appeared to invade the eroded carapace after or (in low numbers) with the bacteria. In rare cases the organisms appeared to invade ahead of the bacteria once the bacteria had eroded the epicuticle and portions of the exocuticle. Protistians caused a disruption of the crystalline lattice producing a swiss cheese effect in portions of the remaining exocuticular and calcified endocuticular layers (Fig. 4D).

Category 3: Severe Deep Erosions Into the Uncalcified Endocuticle/Membranous Layer

Erosions in this category were characterized by extension of the lesions into the deeper uncalcified layers of the endocuticle with loss of most of the overlying carapace (Fig. 5). Pillars of chitin lattice with and without remaining matrix were sometimes present in portions of these erosions, but in general were not present on these severely eroded surfaces. The exposed surfaces of these deep layers were melanized. Inflammatory cuticle, as described in category 2, was often present in these deep lesions and, in some cases, was the only tissue remaining between the cuticular epithelium and the external environment. The cuticular epithelium was hyperplastic and hypertrophic. Intense inflammation consisting of large numbers of granulocytic and agranulocytic hemocytes were present in the subepithelial connective tissues. Small, centrally melanized granulomas were common in the underlying connective tissues. No pathogens were identified in the inflamed connective tissues.

[FIGURE 5 OMITTED]

Ulceration

Ulcerations of the carapace were characterized by total loss of all cuticular material and the cuticular epithelium, thus exposing the connective tissues of the body to the environment (Fig. 6). In such areas focally extensive hemocytic infiltration expanded the connective tissues. Degranulated hemocytes at the surface of the wound formed a pseudomembrane over the exposed connective tissues. The external surface of the pseudomembrane was necrotic and melanized. Small granulomas in deeper connective tissues and other adjacent organs were noted in these lesions and in one animal contained fungal hyphae.

[FIGURE 6 OMITTED]

Internal Lesions

Granulomas and diffuse hemocytic inflammation were associated with connective tissues underlying the most severe erosions and in ulcerated areas. However, only in 3 animals were any lesions (granulomas) noted in the connective tissues distant from the carapace erosions or in other organs examined histologically.

The Histopathologic Identification of the Cause of Shell Disease

The predominate organisms seen at the leading edge/interface between degraded carapace and underlying intact carapace were bacteria. This finding was most prominent in tissues from animals examined in the early spring. Abundant colonies of bacteria (mostly appearing as stacks of short rods) were noted on the surface, and deep within shaft-like erosive extensions into the crystalline lattice of the carapace. Gram stains showed that the bacteria were Gram negative, but identification of the bacterial genus/ species was not possible from examination of histologic sections.

Other organisms observed in the lesions included tree living nematodes, filamentous algae, large and small protozoa, protistans and barnacles. The second most commonly observed organisms in the lesions were small, similarly sized protistans. Protistians were associated with portions of the lesions in many of the animals and occurred in eroded carapaces of animals from all areas sampled. They were commonly associated with animals collected in late spring and summer. Unlike bacteria that were found at all levels of erosion, these secondary organisms were seldom found in the lesions that extended into the uncalcified endocuticle or membranous layers. The secondary organisms (especially the small protistans) were not the primary agents responsible for the lesions but did contribute, sometimes markedly, to the ongoing primary bacterial degeneration of the epi- and exocuticles of the infected carapaces.

Molting of Eroded Carapaces

Histologic examination of one animal in molt showed new epicuticle and exocuticle was formed internal to the eroded carapace and associated inflammatory cuticle. Histologic sections showed a lytic space formed between the inflammatory cuticle and the new epicuticle thus allowing for molt to occur.

DISCUSSION

Shell disease has reached epizootic proportions in eastern Long Island Sound and along the coast lines of Rhode Island, southern Massachusetts and off the coast of Kittery, Maine. Epizootic shell disease most likely represents an extreme presentation of enzootic shell disease, which had been identified in previous years during the yearly inshore lobster monitoring work conducted by many northeastern states. Epizootic shell disease is characterized by severe deep dorsal midline erosions that most often affect the cephalothorax and abdomen and that extend laterally and irregularly over the dorsal carapace of the animals. In many cases the rostrum was severely necrotic or missing. Other portions of the animals carapace were affected, but with markedly less number and size of lesions. Arthrodial membranes (the joints) were not significantly affected in epizootic shell disease.

Bacteria are identified at the leading edges of the invasive lesions in the affected carapace confirming that cause of the disease is bacterial in origin, but histology alone could not identify bacteria responsible for the erosions. Importantly, at the histologic level the carapaces appeared to be normal (lesions did not result from poorly formed carapace) and the disease resulted from invasion of the carapace from the carapace surface (not from an internal disease or abnormality).

One of the common histologic characteristics resulting from bacterial erosion in this disease was the occurrence of pillars composed of chitin crystalline lattice that either lacked the protein and lipid matrix normally filling the interstices within the lattice or contained bacteria in place of matrix. This lesion is very rarely identified in impoundment shell disease and never identified in burn spot shell disease. The occurrence of pillars suggested that the invading bacteria were not primarily chitin feeders, but instead were attracted to the proteins and lipids making up the matrix. Secondary invading organisms were noted in the lesions, but when present, they invaded areas previously invaded by bacteria. Potentially significant is that secondary infections did contribute to the shell erosion in the epicuticular, exocuticular and calcified endocuticular layers.

The lobsters' inflammatory response to bacterial invasion of the carapace appeared appropriate and effective. In fact, the effectiveness of the inflammatory response is demonstrated by the ability of infected lobsters to molt out of the disease. Histologically, the hemocytic portion of the inflammatory response that occurred in erosions classified as categories 2 and 3 was appropriate, and demonstrated that the innate immune system was functioning well. Small granulomas, present in the tissues of severe shell disease of all forms, were identified within the connective tissues underlying the epithelium in areas of eroded carapace. The presence of granulomas suggested migration of an irritating substance or infective organism (possibly bacteria) occurred through the pores, tegumental glands and setal canals that transverse the cuticle. Histologic sections stained with Gram and GMS did not identify the cause of these granulomas indicating that, in most instances, the cause was no longer present in the tissues (the inflammatory response was able to effectively eliminate potential invading organisms and/or substances).

Ulceration (loss of the entire carapace and its epithelium) and exposure of soft tissues to the external environment was only rarely noted in histologic sections. Ulcers most likely resulted in an inability to molt because adhesion of the internal tissues to the old cuticle is a common sequella to ulcerative events. Additionally, an ulcer would allow for invasion of the internal soft tissues by bacteria or fungus leading to systemic infection and death.

The inflammatory mechanisms used by the lobsters to combat the erosive lesions are similar to those described in other forms of shell disease. Melanization (more correctly termed phenolic oxidation) causes cross-linking of proteins in the layers of the carapace thus forming a wall that inhibits invasion by pathogenic organisms (Lightner & Redman 1977, Soderhall & Unestam 1979, Unestam & Ajaxon 1976). Melanization also occurs as an inflammatory response mounted by hemocytes when forming granulomas (to cross-link tissue proteins and wall off the invading microbes) (Soderhall & Unestam 1979). Phenolic oxidase and its associated phenolic substrates are present in the upper layers of the epicuticle and exocuticle and may be supplied through pore canals for endocuticular layers (Unestam & Ajaxon 1976). Melanization of the irregular, exposed surfaces of the eroded cuticle accounted for the grossly observable brown/black granular appearance of the lesions (tanning of the cuticle to this extent does not normally occur in these areas of the carapace). The degree of melanization seen in epizootic shell disease did not differ from that seen in other forms of shell disease.

Previous studies of other forms of lobster shell disease, and shell disease in other crustaceans, have noted that bacterial-caused erosions rarely penetrate through the uncalcified endocuticle. But only in a few studies has the production of inflammatory cuticle secondary to injury or erosions been described (Neville 1975, Dillaman & Roer 1980, Smolowitz et al. 1992). This author (Smolowitz) has not before seen production of inflammatory cuticle to the extent present in the erosions of epizootic shell disease. Inflammatory cuticle production occurred in animals in C4/D0 stage of cuticle formation. The method by which this inflammatory cuticle is formed (how the epithelial cells are stimulated to produce new carapace only where erosions occur) is not known. However, this method is extremely effective in preventing ulceration into the underlying tissues. Also, its production provides an extra layer of protection during development of the new epicuticle and exocuticle that must be produced internally to the inflamed cuticle. As noted histologically, during molt the inner layers of the inflammatory cuticle dissolve forming a space between the inflammatory cuticle and the new epicuticle allowing lobsters to "molt out" of the disease. The production of this inflammatory cuticle in combination with the stubble of the remaining eroded original cuticle resulted in the soft rubbery feel of the pitting erosions.

Whereas not seen in the sections, it is also possible that in some areas, inflammatory cuticle was the only new cuticle formed by the epithelium before molt, and the inner layers of the inflammatory cuticle could dissolve, allowing molting of the old shell. This condition could be the cause of the "glass panes" (thin, soft, white to clear areas) seen grossly in the new carapace of some affected animals that had recently molted out of the disease.

The reasons for the occurrence of epizootic shell disease are still not understood, but environmental factors may play a roll. Certainly bacteria have been identified as the cause of shell disease in previous work (Smolowitz et al. 1992, Fisher et al. 1978, Getchell 1989, Malloy 1978). Epizootic shell disease is unusual in the severity and extent of the erosive carapace lesions and the primary location of lesions on the dorsum of the cephalothorax. It is possible that increased environmental temperatures may promote the growth of bacteria in the marine environment and/or on the shell resulting in disease. It is possible that the ability of the lobster to effectively remove such bacteria, especially along the dorsal midline, may be compromised by increased temperatures that may slow down the lobster's normal responses (an effect of thermal stress). The bacteria causing the lesions may be more aggressive than the flora that normally resides on the carapace surface. Such changes in pathogenicity might occur through plasmid or phage transfer between bacteria. Changes in environment might also produce increased invasions by secondary organisms resulting in increased degeneration of the superficial layers of the carapace. Whereas the carapace and its inflammatory response appear normal in the histologic sections, it is possible that at the molecular level there is an abnormality in formation of the cuticle (especially in formation of the protein/lipid matrix or in inclusion of phenolic substances or phenol oxidases). Any one or a combination of these possible causes could be at work in this disease. These possibilities warrant further study, not only to answer the important questions about shell disease in lobsters but to better understand how changes in the marine environment can interact with an animal, such as a lobster, to cause disease.

ACKNOWLEDGMENTS

The authors thank Erin Estrada and Kevin Uhlinger for technical support. They also thank the lobstermen's associations and the state fisheries biologists for their willingness to provide needed samples. This publication was supported in part by the National Sea Grant College Program of the US Department of Commerce's National Oceanic and Atmospheric Administration under award #NA16RG1354 to the Research Foundation of State University of New York for New York Sea Grant. The views herein do not necessarily reflect the views of any of those organizations. This work was also partially funded by the Woods Hole Oceanographic Institution Sea Grant Program, under a grant from the National Oceanic and Atmospheric Administration, US Department of Commerce, Grant No. NAI6RG2273, project no. R/B-167.

LITERATURE CITED

Aiken, D. E. 1980. Molting and growth. In: J. S. Cobb & B. F. Phillips, editors. The biology and management of lobsters, physiology and behavior, vol. 1. New York: Academic Press. pp. 91-163.

Bang, F. B. 1983. Crustacean disease responses. In: D. E. Bliss & A. J. Provenzano, Jr., editors. The biology of crustacea, vol. 6. New York: Academic Press. pp. 114 147.

Bullis, R. A. 1989. Shell disease in impounded American lobsters, Homarus americanus. Bio. Bull. 177:327.

Bullis, R. A. & M. McCafferty. 1995. A method for the histological processing of crustacean exoskeleton. In: J. S. Stolen, T. C. Fletcher, S. A. Smith, J. T. Zelikoff, S. L. Kaattari, R. S. Anderson, K. Soderhall & B. A. Weeks-Perkins, editors. Techniques in fish immunology-4, immunology and pathology of aquatic invertebrates. Fair Haven, N J: SOS Publications.

Burns, C. D., M. E. Berrigan & G. E. Henderson. 1979. Fusarium sp. infection in the freshwater prawn Macrobrachium rosenbergii (De Man). Aquaculture 16:193-198.

Castro, K. M. & T. E. Angell. 2000. Prevalence and progression of shell disease in American lobsters from Rhode Island waters and offshore canyons. J. Shellfish. Res. 19:691-700.

Chistoserdov, A. Y., R. Smolowitz, F. Mirasol & A. Hsu. 2005. Culture-dependent characterization of the microbial community associated with epizootic shell disease lesions in American lobster, Homarus americanus. J. Shellfish Res. 24(3):741-747.

Connecticut Department of Environmental Protection, Bureau of Natural Resources, Marine Fisheries Office. 1999. Information regarding the impact of 1999 lobster mortalities in Long Island Sound. Hartford, CT. 48 pp. Available at: http://dep.state.ct.us/burnatr/fishing/marineinfo/ lobrep.htm#report

Dean, M. J., K. A. Lundy & T. B. Hoopes. 2002. 2001 Massachusetts lobster fishery statistics. Massachusetts Division of Marine Fisheries, Technical Report #13. Available at: http://www.state.ma.usldfwele/ dmf/index.html

Dean, M. J., K. A. Lundy & T. Hoopes. 2004. 2002 Massachusetts lobster fishery statistics. Massachusetts Division of Marine Fisheries Technical Report TR-20. Available at: http://www.mass.gov/dfwele/dmf/ Publications/lobster_report_2002 tr20.pdf

Dillaman, R. M. & R. D. Roer. 1980. Carapace repair in the green crab, Carcinus maenas (L.). J. of Morphology 163:135-155.

Estrella, B. 1991. Shell disease in American lobster (Homarus americanus, H. Milne Edwards, 1937) from Massachusetts coastal waters with considerations for standardizing sampling. J. Shellfish Res. 10:483-488.

Fisher, W. S. Shell disease of lobsters. 1988. In: C. J. Sindermann & D. V. Lightner editors. Disease diagnosis and control in North American marine aquaculture. Amsterdam: Elsevier.

Fisher, W. S., E. H. Nilson, J. F. Steenbergen & D. V. Lightner. 1978. Microbial diseases of cultured lobsters: A review. Aquaculture 14:115-140.

Floreto, E. A. T., D. L. Prince, P. B. Brown & R. C. Bayer. 2000. The biochemical profiles of shell-diseased American lobsters, Homarus americanus Milne Edwards. Aquaculture 188:247-262.

Getchell, R. G. 1989. Bacterial shell disease in crustaceans: a review. J. Shellf. Path. 8:1-6.

Ghidalia, W. 1985. Structural and biological aspects of pigments. In: D. E. Bliss & L. H. Mantel, editors. The biology of crustacea, vol. 9. pp 347-363.

Laufer, H., N. Demir & W. Biggers. 2005. Response of the American lobster to the stress of shell disease. J. Shellfish Res. 24(3):757-760.

Lightner, D. V. & R. Redman. 1977. Histochemical demonstration of melanin in cellular inflammatory processes of Penaeid shrimp. J. Invert. Path. 30:298 302.

Luna, L. G. 1992. Histopathologic methods and color atlas of special stains and tissue artifacts. Downers Grove, IL: Johnson Printers.

Malloy, S. C. 1978. Bacteria induced shell disease of lobsters (Homarus americanus). J. Wildl. Dis. 14:2-10.

Neville, A. C. 1975. Biology of the arthropod cuticle. In: D. S. Fames, editor. Zoophysiol. ecol., Vol 4/5. New York: Springer-Verlag.

Presnell, J. K. & M. P. Schreibman. 1997. Humanson's animal tissue techniques. 5th ed. W. H. Freedman and Co. pp. 314-317.

Prince, D. L., R. C. Bayer, M. L. Gallagher & M. Subrainanyam. 1995. Reduction of shell disease with an experimental diet in a Nova Scotian lobster pound. J. Shellf Res. 14:205-207.

Rosen, B. 1970. Shell disease of aquatic crustaceans. In: S. F. Snieszko, ed. Symposium on diseases of fishes and shellfishes, special publication, No. 5. Washington, DC: American Fisheries Society. pp. 409-415.

Sindermann, C. J. 1979. Pollution associated disease and abnormalities of fish and shellfish-review. Fishery Bulletin, US. 76:171-749.

Smolowitz, R., R. A. Bullis & D. A. Abt. 1992. Pathologic cuticular changes of winter impoundment shell disease preceding and during intermolt in the American Lobster, Homarus americanus. Biol. Bull. 183:99-112.

Soderhall, K. & T. Unestam. 1979. Activation of serum prophenoloxidase in arthropod immunity. The specificity of cell wall glucan activation and activation by purified fungal glycoproteins of crayfish phenoloxidase. Can. J. Microbiol. 25:406-414.

Stewart, J. E. 1980. Diseases. In: J. S. Cobb & B. F. Phillips, editors. The biology and management of lobsters, Vol. 1. Physiology and behavior. New York: Academic Press.

Unestam, T. & R. Ajaxon. 1976. Phenol oxidation in soft cuticle and blood of crayfish compared with that in other arthropods and activation of the phenol oxidases by fungal and other cell walls. J. Invert. Path. 27:287-295.

Ziskowski, J., R. Spallone, D. Kapareiko, R. Robohm, A. Calabrease & J. Pereira. 1996. Shell disease in American lobster in the offshore Northwest-Atlantic region around the 106-mile sewage-sludge disposal site. J. Mar. Environ. Eng. 3:247-271.

ROXANNA SMOLOWITZ, (1) * ANDREI Y. CHISTOSERDOV (2) AND ANDREA HSU (1)

(1) Marine Biological Laboratory, 7 MBL Street, Woods Hole, Massachusetts 02536, (2) Department of Biology, University of Louisiana, Lafayette, Louisiana 70504

* Corresponding author. E-mail: [e-mail]

TABLE 1.
Shell disease index used to assess the severity and proportion of
lobsters affected. *

0 = no shell disease symptoms
1 = shell disease symptoms on 1-10% of the shell surface
2 = shell disease symptoms on 11-50% of the shell surface
3 = shell disease symptoms on >50% of the shell surface.

* Presented by B. Estrella to the Lobster Shell Disease Workshop,
Connecticut Sea Grant College Program, Waterford, CT, June 15, 2000.