a. Uninucleate plasmodial Hematodinium (arrows) in antennal gland. Note diffuse (D) and condensed (C) nuclei. Parasites
proliferate and replace host tissue. Bar = 20 pm.
*. b. Uninucleate plasmodial Hematodinium in hemolymph smear. Note diffuse (D) and condensed (C) chromosomes. Bar = 20

N.

ym.
¢. Uninucleate plasmodial Hematodinium in hemolymph smear. Note lack of nuclear membranes (arrow). Bar = 10 pm.
d. Long-form multinucleate plasmodial Hematodinium in hemolymph smear (arrow). Bar = 20 pm.



Page 8
|
[eet

¢

Figure 1. Viral infections causing mortalities in blue crabs. - .

a, Reo-like - rhabdo-like virus A (RLV-RhVA) combination in hemopoietic tissue of intermolt crab. Note increased
cytoplasmic volume (v), inclusions (arrows), and normal cells (c). Bar = 10 pm.

b. RLV-RhVA infection in young hemocytes released from hemopoietic tissue. Note increased cytoplasmic volume (v), viral
inclusions (arrows), nuclei of various sizes (n), and normal young hemocytes (y). Bar = 10 pm. .

c. Bi-facies virus (BFV) infecting hemocytes. Note hypertrophy of nuclei (h), refractive cytoplasmic inclusions (arrows), and
normal hemocytes (c). Bar = 20 pm. :

4, BFV infecting hemocytes in hemal space of antennal gland. Note hypertrophy of nuclei (h), cytoplasmic inclusions (arrow),
and cells about to lyse (L). Bar = 20 pm.

e. Chesapeake Bay virus (CBV) infecting thoracic ganglion of central nervous system. Note extensive destruction of
neurosecretory cells (v), and normal neurosecretory cells (arrows). Bar = 50 um.

f. CBV infecting epithelial cells of bladder. Note cytoplasmic inclusions in infected cells (arrows). Bar = 50 ym.



Reproduced from
best available copy.

|

Figure 12. Leech (Myzobdella lugubris) on blue crab. Top, several specimens of the leech on the crab’s anterior. Bottom, several
specimens on crab’s posterior, most individuals deposit cocoons near the rear margin. (Photographs courtesy of Dr. R.M.
Overstreet.)



CONTENTS

Abstract ...
Introduction
Viral Diseases ..
Bacterial Diseases
Fungal Diseases...

Protozoan Diseases ...... 9
Diseases Caused by Helminths .. 13
Ectocommensal and Parasitic Barnacles 15
Summary

Acknowledgments
References Cited ..

LIST OF TABLES

* Table 1. Principal pathogens of blue crabs
Table 2. Viral diseases of blue crabs

LIST OF FIGURES

Figure 1. Viral infections causing mortalities in blue crabs
Figure 2, Shell disease in the blue crab....
Figure 3. Pamlico River blue crab with carapace lesions .
Figure 4. Bacterial infections in blue crab tissue
Figure 5. Microsporidan infection in blue crab muscle .
Figure 6. Paramoeba perniciosa in blue crab tissues
Figure’7.° Hematodinium perezi in’ blue'crab hemolymph ..
Figure 8. Haplosporidan-like organisms in blue crab
Figure 9. Ciliates infesting blue crab gills .......
Figure 10, Microphallus basodactylophallus hyperparasitized by
Figure 11. Ribbon worms between gill lamellae ...
Figure 12, Myzobdella lugubris attached to blue crab .
Figure 13. Loxothylacus texanus infecting blue crab

Urosporidium crescens.



metacercariae, although they may debilitate them. The
hyperparasites are only released upon death of the crab due
to rupture of the metacercarial cyst (Couch 1974b).

NEMERTEANS

The ribbon worm, Carcinonemertes carcinophila, is
found encysted between gill lamellae of female blue crabs in
high-salinity waters (Figure 11). Humes (1942) described
the morphology and life cycle of this parasite. Asa juvenile,
the worm migrates from the gills to the host’s egg mass to
become sexually mature, lays eggs of its own, and returns to
the gills as a mature adult (Hopkins 1947). Crab eggs
provide nourishment for the worm, anda color change from
cream to orange can be observed due to the ingestion of crab
egg-yolk material. The parasite builds a mucus tube as a
habitat in the egg mass. After mating, the female worm
leaves her eggs and mucus tube and returns to the gills as an
adult (Overstreet 1978). The larvae escape from the egg
membranes through the action of enzymes and may remain
in the tube, swim among the eggs, or leave the host and
change to a free-living form, enabling them to infest other
crabs (Davis 1965).

Since worms change color as a result of feeding on crab
eggs, and since adults return to the gills, C. carcinophila can
be used as a biological indicator. Nonovigerous female
crabs that have already spawned once can be distinguished
from nonspawning females by examining the gills of these
crabs with the unaided eye. Only large bright-red adult
worms occur in crabs that have spawned and only immature
worms, light in color, are found in gills of females that have
not spawned. This method of diagnosing spawned blue
crabs is about 97-percent accurate (Hopkins 1947; Pyle and
Cronin 1950; Overstreet 1978, 1982).

ANNELIDS (leeches)

The leech, Myzobdella lugubris, is often found attached
to blue crabs from low-salinity waters, but is not considered
fatal. Leeches have been implicated inconclusively in
mortalities of blue crabs in Florida where 50 percent of crabs
sampled had leeches attached near perforations on the
carapace and articulations of the pereiopods (Figure 12)
(Hutton and Sogandares-Bernal 1959). The life cycle of M.
lugubris involves two hosts, the white catfish, Ameiurus
catus, which provides most of the leech’s nutrition, and the
blue crab, on which it deposits cocoons near the rear margin
of the crab's carapace. In addition to providing a surface for
the cocoon toattach, the crab also acts as a means of dispersal
for the young (Overstreet 1978; Millikin and Williams
1984). The parasite is common along the Atlantic and Gulf

* coasts of the United States and is usually not considered fatal
(Moore 1946; Sawyer et al. 1975).

Figure 8. Haplosporidan-like organisms in hemolymph of blue
crab.

a, Uninucleate (u) and multinucleate (m) plasmodial
forms of haplosporidan-like parasite. Note individual
spores (arrows). Bar = 20 ym.

b. Vascular space of gill filled with haplosporidan-like
plasmodia (arrows); (c) is anormal hemocyte. Bar =
20 pm.

ECTOCOMMENSAL AND PARASITIC
BARNACLES a

Ectocommensal barnacles are observed frequently on
blue crabs. The gooseneck or pedunculate barnacle,
Octolasmis mulleri, is found on gills of blue crabs in high-
salinity waters. These may interfere with crab respiration in
heavy infestations due to inhibited scaphognathite move-
ment, but usually they cause no harm to the host (Walker
1974; Overstreet 1978; Millikin and Williams 1984). An-
other ectocommensal, the acorn barnacle Chelonibia patula,
is found attached to the carapace of blue crabs, especially
older crabs, after their terminal molt. The only hardship
these barnacles may present to the host is excess weight on
the carapace which in heavy infestations may inhibit crab
mobility (Overstreet 1978).


Page 21



Figure 10. Microphallus basodactylophallus hyperparasitized by

Urosporidium crescens.

a. Hyperparasitized metacercariae (m) and
unparasitized metacercariae (arrow). Bar = 100
um.

b. Pockets of haplosporidans U. crescens (arrows)
parasitizing M. basodactylophallus metacercariae.
Bar = 50 pm.

Systemic bacterial infections are usually manifested in
crabs that have been stressed by one or more factors. By
reducing stress, both in the shedding system and during
capture, these infections could be lowered. Crabbers should
be advised to treat peeler crabs with better care by providing
shade, reducing handling stress, and trying to reduce crowded
conditions, if possible. Crab shedders could reduce stress to
captive crabs by providing them with good water flow and
well oxygenated water, and by keeping their systems clear
of sediments and high levels of metabolic byproducts.

Although diseases which occur frequently in captive
crabs have the potential for being treated, diseases which

Figure 11. Ribbon worms between blue crab gill lamellae.
a. Long orange ribbon worm (Carcinonemertes
carcinophila) pulled out from between gill lamellae.
Other light-colored regions on gills are additional
specimens. Also note bamacle on gills. Both
bamacle and ribbon worm indicate that the crab
had inhabited high-salinity water. (Photograph
courtesy of Dr. R.M. Overstreet.)
b. Histologic section showing nemerteans encysted
between gill lamellae. Bar = 100 pm.

occur most often in wild populations cannot usually be
treated or controlled. Unfortunately, these are diseases that
may cause the most harm to crab populations. One of these,
the fungus Lagenidium callinectes, attacks egg masses, thus
causing reduced fecundity. Protozoan diseases can induce
mass mortalities in wild populations of blue crabs. The most
detrimental protozoans found infecting blue crabs are
microsporidans, including Nosema and Pleistophora. Itis
known that these pathogens are transmitted directly when
healthy crabs ingest dead sick crabs, so crab fishermen have
been told not to return sick crabs to the waters as they had
done previously. Other protozoans of significance are two


Page 12

Paramoebais found inthe hemolymphand nonepithelial
tissues of blue crabs. The amoebae range in size from 3 to
35 ym, are round to elongate, contain a well defined nucleus
with a large central endosome, plus a morphologically
different secondary nucleus-like body inthe cytoplasm. The
major diagnostic characteristic of P. perniciosa is the pres-
ence of this secondary body in the cytoplasm (Figure 6). In
heavily infected individuals, Paramoeba disease causes
drastic changes in muscle tissues. In very heavy cases, most
blood cells are replaced by the amoebae, but in light infec-
tions few amoebae may be apparent. Not all crabs infected
with the parasite exhibit the grey sterna, but most of the crabs
which have the grey ventral surface are lethargic and heavily
infected with amoebae (Johnson 1977b; Couch 1983).

Most crabs infected by Paramoeba demonstrate some
type of defense reaction, usually noted by the presence of
degraded lysed amoebae being phagocytized by hyaline

hemocytes; occasionally, there is also encapsulation of ©

amoebae by hemocytes (Figure 6c). Hemocytes are usually
seen concentrated in tissues containing amoebae, especially
in the midgut and hemal spaces of the antennal gland
(Johnson 1977b). The cause of death in Paramoeba infec-
tions is unknown. It is suggested that loss of vital tissue,
such as blood cells and muscle, may lead to a state of
inanition and loss of function. Blood sinuses in infected gills
may become so congested with amoebae that respiratory
function may fail. Glucose levels in the blood of infected
ctabs are reduced, perhaps due to competition between P.
perniciosa and the infected tissue for the limited amount of
available glucose. Serum protein in infected crabs is also
reduced which probably causes the weakened, lethargic
response (Pauley et al. 1975; Campbell 1984).

Peak prevalences of the parasite occur during May and
June (Campbell 1984) and in dredged hibernating crabs
from October to February. The disease in epizootic form
could be a major factor in annual winter and late ‘spring
mortalities of blue crabs (Newman and Ward 1973; Couch
1983), and the cause of chronic low-level mortalities in late
spring of nonepizootic years. Marked annual and geo-
graphic variations in the prevalences of Paramoeba have
been noted and mortalities are most commonly reported
from high-salinity coastal bays from Maryland to Georgia.
The parasite appears to be opportunistic by invading the
hemolymph via injuries to the exoskeleton, especially in
newly molted crabs. Paramoeba has also been reported to
infect Atlantic rock crabs, Cancer irroratus, American
lobsters, Homarus americanus (Sawyer and MacLean 1978),
and green crabs, Carcinus maenas (Campbell 1984).

Another member of the protozoan phylum
Sarcomastigophora recognized as causing disease in blue
crabs is the dinoflagellate Hematodinium perezi. This

. Parasite was first reported in portunid crabs from France
(Chatton and Poisson 1931), but was not reported as a
parasite in other Crustacea until found in blue crabs sampled
for P. perniciosa (Newman and Johnson 1975). Hematodinium
causes a fatal disease, but infected crabs exhibit no external
signs except lethargy. Drawn blood from heavily infected

Figure 6. Paramoeba perniciosa infection in blue crab tissues.

a. Hemal space of antennal gland filled with parasites
(arrows). Note halo surrounding individual amoebae
and secondary bodies in cytoplasm of parasites. Bar
= 50 um.

b. Note nucleus with large endosome (e), secondary
bodies in cytoplasm (arrow), and normal hemocytes
(©). Bar = 20 um.

¢c. Encapsulation (E) of degraded lysed amoebae (arrows)
being phagocytized by hyaline hemocytes. Bar = 20
pm.



Page 4

Table 1. Continued

Pathogen Histopathology and Effect Gross Signs Geographic Location Reference
Tissues Infected on Host of Disease and Prevalence
Lagenophrys Ectocommensal Secretes a No gross Chincoteague Couch 1966,
callinectes living in protective signs; and 1967, 1983
(protozoan, lorica found lorica on diagnosis Chesapeake
ciliate) onflat gill lamellae, through Bays; Gulf of
surfaces of may inter- microscope Mexico; peak
gill lamellae fere with for presence prevalence
respiratory of lorica during summer
and excretory months
function of
gills
Microphallus Hyperpara- Pigmented Crab tissues Chesapeake DeTurk 1940;
basodacty- sitized _ spores contain large Bay to Texas Couch 1974;
lophallus Microphallus debilitate brownish-black Overstreet
(digenean, cysts present and enlarge metacercasia 1978, 1982
fluke, hyper- in thoracic worm cyst} called
parasitized by muscles, actual effect “pepper spot”
Urosporidium hepatopancreas, on crabs or “buckshot”
crescens, and ventral slight; causes
haplo- ganglion reduced market
sporidan) value of crabs
Carcino- Externally Feeds on Destruction Gulf coast; Humes 1942;
nemertes infests gills host's egg of egg mass; infects crabs Hopkins 1947;
carcinophila and egg mass mass; causes worms seen. from high Pyle & Cronin
(nemertean) reduction in grossly on salinity 1950; Over-
reproductive gills between habitats street 1978,
potential; lamellae 1982
cements gill .
lamellae
together
Myzobdella Symbiont; Unknown, Presence Atlantic and Moore 1946;
lugubris attaches to associated of cocoons, Gulf coasts; Hutton &
(annelid, external in some usually near low-salinity Sogandares-
leech) carapace reports with posterior habitats; Bernal 1959;
mortalities, margin of male crab Sawyer et
but no carapace usual host al, 1975;
lesions Overstreet
observed 1978
Loxothylacus Internal Inhibits Parasite’s Gulf of Reinhard
texanus Parasite crab growth, externa Mexico; more 1950, 1956;
Girriped, sends root- terminates protrudes prevalent Adkins 1972;
parasitic like system reproduction, under crab's in higher Ragan &
bamacle, throughout removes apron; male salinity Matheme
thizocephalan) host's muscle; individuals erabs acquire waters 1974; Over-
develops an. from fishery; secondary street 1978
external sac may reduce adult female
which serves up to 50% of sexual
as brood sac commercial qualities
for nauplii stocks in
larvae some areas



Page 24

Sindermann, C.J., and A. Rosenfield. 1967. Principal
diseases of commercially important marine bivalve
Mollusca and Crustacea. Fish. Bull., U.S. 66: 335-385.

Sizemore, R.K., and J.W. Davis. 1985. Source of Vibrio
spp. found in the hemolymph of the blue crab, Callinectes
sapidus. J. Invertebr. Pathol. 46: 109-110.

Sizemore, R.K., R.R. Colwell, H.S. Tubiash, and T.E.
Lovelace. 1975. Bacterial flora of the hemolymph of the
blue crab, Callinectes sapidus: numerical taxonomy. Appl.
Microbiol. 29(3): 393-399.

Sparks, A.K. 1985. Synopsis of invertebrate pathology
exclusive of insects. Elsevier Scientific Publishers,
Amsterdam, The Netherlands. 423 pp.

Sprague, V. 1950. Notes on three microsporidian parasites
of decapod Crustacea of Louisiana coastal waters. Occas.
Pap. Mar. Lab., La. State Univ. No. 5: 1-8.

. Sprague, V. 1963. Minchinia louisianan. sp. (Haplosporidia,
Haplosporididae), a parasite of Panopeus herbstii. J.
Protozool. 10: 267-274.

Sprague, V. 1965. Nosemasp. (Microsporida, Nosematidae)
in the musculature of the crab Callinectes sapidus. J.
Protozool. 12: 66-70. :

Sprague, V. 1966. Two new species of Pleistophora
(Microsporida, Nosematidae) in decapods, with particu-
lar reference to one inthe blue crab. J. Protozool. 13: 196-
199... ,

Sprague, V. 1970. Some protozoan parasites and hyper-
parasites in marine decapod Crustacea. Pages 416-430 in
S.F. Snieszko, ed. A Symposium on Diseases of Fishes
and Shellfishes. Amer. Fish. Soc. Spec. Publ. No. 5.

Sprague, V. 1977. Systematics of Microsporidia. Pages 1-
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Sprague, V. 1978. Comments on trends in research on
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Sprague, V.,andR.L. Beckett. 1966. A disease of blue crabs
(Callinectes sapidus) in Maryland and Virginia. J.
Invertebr. Pathol. 8: 287-289.

Sprague, V., and R.L. Beckett. 1968. The nature of the
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Sprague, V., and J. Couch. 1971. An annotated list of
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Sprague, V.,R.L. Beckett, and T.K. Sawyer. 1969. Anew
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Tharp, T.P., and C.E. Bland. 1977. Biology and host range
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Yudin, A.L, and W.H. Clark, Jr. 1979, A description of
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BY U.S, GOVERNMENT PRINTING OFFICE: 1992--500-031--40007



Page 1

\ ABSTRACT
Sj

~ Bluecrabs, Callinectes sapidus, are an important ecdnomic and recreational resource onthe Atlantic and Gulf coasts.
Accordingly, a number of pathological agents which may decrease the availability or demand for blue crabs have been
studied. Investigations have emphasized diseases and parasites that may either cause mortalities or reduce market value
of blue crabs. Diseases and parasites are known to cause mortalities in both wild and captive crabs. Some of the most
harmful pathogens infecting wild popilations of blue crabs are several microsporidans, including Ameson michaelis; an
amoeba, Paramoeba perniciosa; and a dinoflagellate, Hematodinium perezi. Crabs held in captivity are subject to
systemic bacterial infections, including Vibrio parahaemolyticus, and to various viral infections, including a reo-like
virus, a picorna-like virus, and a bi-facies virus. Other pathogens may decrease fecundity, including the fungus
Lagenidium callinectes, the nemertean Carcinonemertes carcinophila, and the parasitic barnacle Loxothylacus texanuse
Finally, still other diseases can reduce marketability of infected crabs. Chitinoclastic bacteria including species of Vibrio im
and Pseudomonas produce unattractive necrotic lesions on the exoskeleton of crabs; and a trematode, Microphallus
basodactylophallus, hyperparasitized by the haplosporidan Urosporidium crescens, produces undesirable black pig- \
mented spots in the musculature of cooked crabs. ‘

INTRODUCTION rized in Table 1, and is discussed briefly in the sections that
follow.

The blue crab, Callinectes sapidus, is the most valuable
commercial and recreational species of crab along the Atlan-
tic and Gulf coasts. The significance of this fishery is
demonstrated by the many papers and books written onthe © @ Reo-like virus (RLV) + rhabdo-like virus A (RhVA)
subject. Asearly as 1905, thelifehistory ofthe bluecrabwas  @ Chesapeake Bay virus (CBV), a picorna-like virus
described (Hay 1905). Since then, many experimental  @ _ Bi-faciesvirus (BFV), formerly herpes-like virus (HLV)
findings and generalized data have been reported; a few of
the comprehensive reports available deal with general Bacteria
anatomy (Pyle and Cronin 1950), form and growth
(Newcombe et al. 1949), and histology of the blue crab
(Johnson 1980). An extensive synopsis of biological data

Viruses

@ = Chitinoclastic bacteria, including Vibrio and Pseudo-

monas
has been published (Millikin and Williams 1984) which = @ ~— Syctemicbacteria, including V. haemolyticus, Pseudo-
includes most of the pertinent references to blue crab biol- Systemicbacterasinuding ¥ parahaemabyicus Peed?
ogy. . - heterogenous group of coliforms, including Escherichia -
Although the blue crab has consistently been a major coli
fishery resource in both the hard-shell and soft-shell stages,
the fishery has experienced major fluctuations in landings.” Fungus

Outbreaks of disease combined with environmental factors
have been blamed for some of the variations in crab produc-
tion. Diseases have been established as the cause of
mortalities, both in wild populations and in crabs contained
in holding facilities (Sprague 1965; Overstreet 1977; John- Protozoans
son 1983).

This synopsis provides a summary of diseases which
may affect the blue crab industry by either causing mortali-
ties, decreasing fecundity, or decreasing demand. Included
are photographs, when available, for better visualization and
identification of the various diseases and parasites infecting
blue crabs.

Many microbial diseases have been reported from blue
crabs, including those of viral, bacterial, fungal, and proto- —_ He{minths -
zoan etiologies. A few helminths and a parasitic crustacean
have also been implicated as causing either disease or
reduced fecundity. The following organisms have been
reported to cause mortalities or significant pathology in blue
crabs. Information about each of these pathogens is summa-

@ Lagenidium callinectes (phycomycetous)

Ameson michaelis (Microspota)

Paramoeba perniciosa (Sarcomastigophora)
Hematodinium perezi (Sarcomastigophora)
Haplosporidan (Ascetospora)

Lagenophrys callinectes (Ciliophora)

@ = Microphallus spp. (trematode metacercariae)
hyperparasitized by Urosporidium crescens
(haplosporidan)




animals is milky, due to the presence of many uninucleate
dinoflagellates; in some instances, tissues may have a pink
opalescent color. The disease has been noted only in blue
crabs from salinities above 11 ppt, although samples from
lower salinities have been examined. Diagnosis is depen-
dent on the presence of uninucleate or multinucleate
plasmodial cells ranging in size from 5.8 to 64 um. Nuclei
are large in proportion to cytoplasmic volume and chromo-
somes can be either condensed or diffused. This character-
istic dinokaryon nucleus is a major diagnostic indicator of
the dinoflagellate (Figure 7). The parasite is found primarily
in the hemolymph, but is also seen in and between muscle
fibrils, gonad, and in the hepatopancreas. Reported geo-
gtaphic distribution in blue crabs includes the Atlantic and
Gulf coasts of the United States. The parasite has been
detected in crabs in all seasons except late winter and early
spring (Newman and Johnson 1975; Newman 1977; Couch
1983). Hematodinium perezi hasalso been detected in other
crabs, including Carcinus maenas and Portunus depurator
from Europe (Chatton and Poisson 1931), the Atlantic rock
crab, Cancer irroratus, the Jonah crab, C. borealis, and the
lady crab, Ovalipes ocellatus, from the Middle Atlantic
Bight of the U. S. east coast (MacLean and Ruddell 1978).
The protozoan phylum Ascetospora includes the very
pathogenic haplosporidan Haplosporidium nelsoni which,
since the 1960s, has been known to cause considerable
mortalities in oysters. This phylum is poorly represented as
a crustacean pathogen; only Atlantic mud crabs, Panopeus
herbstii, from Louisiana infected with H. louisiana (Sprague
1963, 1978), flatback mud crabs, Eurypanopeus depressus,
from Chincoteague Bay, Virginia, and Harris mud crabs,
Rhithropanopeus harrisii, from Chesapeake Bay, Mary-
land, have been found infected with ascetosporans
(Rosenfield et al. 1969). Two blue crabs from Virginia and
North Carolina were reported infected witha haplosporidan-
like organism (Newman et al. 1976). The prevalence of the
teported disease was rare, but apparently lethal. Drawn
blood from the infected crabs was opaque white, of low
viscosity, and failed to clot properly. Masses of uninucleate
or multinucleate plasmodial bodies filled vascular spaces
and interstitial tissues (Figure 8). Uninucleate cells ranged
from 3.4 to 7.3 ym, and their nuclei ranged from 1.1 to 2.2
um in diameter. Electron microscopy revealed that these
organisms were not the same as those found infecting either
oysters or mud crabs, but a new species parasitizing blue
crabs (Newman et al. 1976). Although reportedly a rare
disease of blue crabs, it has the ability to proliferate in
vascular spaces, thus causing fatalities in infected crabs.
Lagenophrys callinectes is an ectocommensal ciliate
found attached to flat surfaces of blue crab gills (Couch
1966, 1967). The colorless, transparent, semihemispherical
lorica which cements itself to the crab gill measures 48-59
um long by 48-57 ym wide. Mortalities caused by
Lagenophrys are most common among crabs about to molt
and those being held in holding facilities, although crabs in
the wild have also been reported to be heavily infested. The
ciliate is most prevalent from July through November,

Page 13

although a decreased prevalence observed during May and
June is probably due to a higher incidence of molting
activity, thereby freeing infested crabs of the attached
ectocommensals (Couch 1983), Although these ciliates are
technically not parasites (since they do not derive nourish-
ment directly from the host), they can debilitate crabs by
preventing proper gas exchange between the water and gill
tissue (Couch 1983).

Another ciliate, Epistylis sp., observed infesting blue
crabs attaches to the margins and stems of the gill lamellae
rather than the flat portions. The effect of this stalked ciliate
on the health of infested crabs is unknown (Overstreet 1978)
(Figure 9).

DISEASES CAUSED BY HELMINTHS

Marine decapod crustaceans are parasitized by an array
of helminths. Few helminths are truly pathogenic, except in
isolated instances of heavy infections in individual hosts.
These usually immature worms form a spectrum of para-
sites, with a usually modest effect on the host. For a review
of metazoan parasites in blue crabs and other crustaceans,
see Overstreet (1982, 1983). Of those helminths listed,
larval trematodes, nemerteans, and leeches have been re-
ported to either reduce the market value of the captured .rabs
or reduce the fecundity of heavily infected populatioi...

TREMATODES

Metacercariae of digeneans Microphallus
basodactylophatlus and Megalophallus sp. often parasitize
the musculature of blue crabs. These larval flukes are not
easily seen unless hyperparasitized by the haplosporidan
Urosporidium crescens. The metacercariae are easily rec-
ognized when hyperparasitized because they become en-
larged (410-654 ym), darkly pigmented, and rupture easily;
whereas, uninfected or lightly infected metacercariae are
smaller (189-269 um), white to cream color, and can with-
stand some mechanical pressure (DeTurk 1940; Couch
1974b; Overstreet 1978; Millikin and Williams 1984).
Masses of dark pigmented spores in the metacercaria pro-
duce a conspicuous black spot seen easily by the unaidedeye
(Figure 10). Fishermen call the cysts “buckshot” or “pepper
spots” and the infected crabs “pepper crabs.” The presence
of these “pepper spots” in cooked crab meat is objectionable
and can decrease the desirability and market value of in-
fected crabs. Very heavy infections may result in lysis of
host muscle tissue surrounding the parasite. Hyperparasitized
metacercariae of Microphallus basodactylophallus infect
thoracic muscles, hepatopancreas, and ventral ganglia of
blue crabs from Chesapeake Bay to Texas (Overstreet
1982). If U. crescens spores were released within the live
crab, hemocytes or other defense mechanisms would prob-
ably kill the spores; so, as a protective measure, the spores


Table 1. Continued

Page 3

Pathogen Histopathology and Effect Gross Signs Geographic Location Reference
Tissues Infected on Host of Disease and Prevalence
Lagenidium Phycomycete Eggs fail Diseased Chesapeake Couch 1942;
callinectes zoospores to hatch or portions of Bay; east and Sandoz et
(fungus) settle abnormal egg mass Gulf coasts; al, 1944;
on eggs, larvae are appear either 5-30% Newcombe &
germinate, produced; brownish prevalence Rogers 1947;
and extend may also kill or grey, Rogers-Talbert
germ tubes newly hatched depending on 1948; Bland &
which develop larvae within maturity of Amerson 1973,
into branched 48 hr infected egg 1974
septate mass
mycelia
Ameson Micro- Muscle lysis; Lethargy; Delaware and Sprague 1950,
michaelis sporidans parasite muscle has Chesapeake 1965, 1970,
(protozoan, within blood destroys and chalky opaque Bays 1977; Over-
microsporida cells and replaces appearance; southward street &
+ muscle tissue musculature abdomen may to Gulf of Whatley 1975;
due to have white Mexico Overstreet
biochemical or grey color 1977, 1978;
imbalance Couch 1983
Paramoeba Organisms Amoeba fills Grey-colored From Sandy Sprague &
perniciosa with well tissues, abdomen and Hook Bay to Beckett.1966,
(protozoan, defined teplaces appendages; Chincoteague 1968; Sawyer
amoeba, nucleus plus hemocytes, called “grey Bay; usually 1969; Sprague
Sarcomas- a secondary and alters crab” disease seen in etal. 1969;
tigophora) body in hemolymph; higher Newman &
tissues; probably salinity Ward 1973;
large halos causes winter waters Pauly et al.
may surround mortalities 1975; Johnson.
individual 1977b; Couch
amoeba 1983
Hematodinium _ Uninucleate Debilitating Lethargy and Maryland Chatton &
perezi and and lethal weakness, to Gulf of Poisson 1931;
(protozoan, binucleate due to withdrawn, Mexico; Newman &
parasitic plasmodial ability to blood milky found in Johnson 1975;
dino- parasites in proliferate or opaque waters with Newman 1977;
flagellate, hemolymph; and replace in heavy salinities Couch 1983
Sarcomas- chromosomes host tissues; infections, greater than
tigophora) condensed or laboratory- slow to clot, 11 ppt
diffused with infected and contains
no nuclear crabs die few hemocytes
membrane
Haplosporidan _— Uninucleate Causes Stuggishness; Reported in Newman et
(protozoan, and hemocytic hemolymph. two crabs al. 1976
ascetosporan) __plasmodial dysfunction opaque white from Chinco-
stages of and of low teague Bay,
haplosporidan viscosity Virginia, and
parasites coastal North
in tissues Carolina


members of the phylum Sarcomastigophora, Paramoeba
perniciosa and Hematodinium;, a member of the phylum
Ciliophora, Lagenophrys callinectes, and a member of the
phylum Ascetospora, a haplosporidan.

Some diseases do not cause mortalities, but reduce the
demand for crabs because of the unappetizing features they
produce in the crabs. Besides shell disease, mentioned
earlier, a digenean trematode, Microphallus
basodactylophallus, hyperparasitized by the haplosporidan
Urosporidium crescens can cause reduced demand for in-
fected crabs. These cause enlarged, darkly pigmented
“pepper spots” in muscle tissue of infected crabs. Cooked
crab meat with these foreign bodies appears unappetizing.
Another parasite which removes crabs from the fishery and
causes an interesting morphological change in young crabs
is caused by a parasitic barnacle, Loxothylacus texanus.
This parasite attacks young crabs and causes a marked
change in secondary sexual characteristics, making infected
individuals appear as miniature adult females. The parasite
castrates infected males, thus reducing potential reproduc-
tive abilities of the population.

In conclusion, blue crabs are subject to a number of
diseases, some with a high potential to cause mass mortali-
ties and others which seem to be more a nuisance than a
threat to the crab’s health. Methods have been devised to
reduce the potential risk of some of these diseases and
further research should provide additional information, lead-
ing to better control--especially in captive populations.

ACKNOWLEDGMENTS

We wish to thank Fred Kem, Dr. Eric May, and Dr.
Aaron Rosenfield for manuscript review and helpful com-
ments, and Martin Newman and Dr. Phyllis Johnson for use
of archived specimens. We are also indebted to Jane Keller
for diligent typing, editing, and counsel, to Susie Hines for
assistance in obtaining library materials, and to Jay. Lewis
for photography advice. '

REFERENCES CITED

Adkins, G. 1972. Notes on the occurrence and distribution
of the rhizocephalan parasite (Loxothylacus texanus
Boschma) of blue crabs (Callinectes sapidus) in Louisi-
ana estuaries. [La. Wildl. Fish. Comm.,] Tech. Bull. No.
12. 13 pp.

Anonymous. 1972. Vibrioparahaemolyticus gastroenteritis-
-United Kingdom. [U.S. Public Health Serv.,] Morbid.
Mortal. Week Rep., Mar. 25: 99, 104.

Bazin, F., P. Monsarrat, J.R. Bonami, G. Croizier, G.
Meynadier, J.M. Quiot, and C. Vago. 1974. Particules
virales de type Baculovirus observees chez le Crabe
Carcinus maenas. Rev. Trav. Inst. Peches. Marit. 38:
205-208.

Page 19

Benton, A.G. 1935. Chitinovorous bacteria. A preliminary
survey. J. Bacteriol. 29: 449-463.

Blake, P.A., D.T. Allegra, J.D. Snyder, T.J. Barrett, L.
McFarland, C.T. Caraway, J.C. Feeley, J.P. Craig, J.V.
Lee, N.D. Puhr, and R.A. Feldman. 1980a. A possible
endemic focus in the United States. New. Engl. J. Med.
302(6): 305-309.

Blake, P.A., R.E. Weaver, and D.G. Hollis. 1980b. Dis-
eases of humans (other than cholera) caused by vibrios.
Annu. Rev. Microbiol. 34: 341-367.

Bland, C.E., and H.V. Amerson. 1973. Observations on
Lagenidium callinectes Couch: isolation and sporangial
development. Mycologia 65: 310-320.

Bland, C.E., and H.V. Amerson. 1974. Occurrence and
distribution in North Carolina waters of Lagenidium
callinectes Couch, a fungal parasite of blue crab ova.
Chesapeake Sci. 15: 232-235.

Bland, C.E., D.G. Ruch, B.R. Salser, and D.V. Lightner.
1976. Chemical control of Lagenidium, a fungal patho-
gen of marine Crustacea. [Univ. N.C. Sea Grant Prog.,]
Sea Grant Publ. UNC-SG-76-02. North Carolina State
University, Raleigh, N.C. 38 pp.

Campbell, D.G. 1984. The abundance and distribution of
Paramoeba perniciosa. Ph.D. Thesis. Johns Hopkins
University, Baltimore, Md. 188 pp.

Chatton, E., and R. Poisson. 1931. Sur l’existence, dans le
sang des Crabes, de Peridiniens parasites: Hematodinium
perezin.g.,n. sp. (Syndinidae). C.R. Soc. Biol. 105: 553-
557.

Colwell, R.R., T.C. Wicks, and H.S. Tubiash. 1975. A
comparative study of the bacterial flora of the hemolymph
of Callinectes sapidus. Mar. Fish. Rev. 37(S-6): 29-33.

Cook, D.W.,and'S.R. Lofton. 1973. Chitinoclastic bacteria
associated with shell disease in Penaeus shrimp and the
blue crab. J::Wildl. Dis. 9: 154-159.

Couch, J.A. 1966. Two peritrichous ciliates from the gills
of the blue crab. Chesapeake Sci. 7: 171-176.

Couch, J.A. 1967. A new species of Lagenophrys (Ciliata:
Peritrichida: Lagenophryidae) from a marine crab,
Callinectes sapidus. Trans. Amer. Microsc. Soc. 86: 204-
211. .

Couch, J.A. 1974a. Anenzootic nuclear polyhedrosis virus
of pink shrimp: ultrastructure, prevalence, and enhance-
ment. J. Invertebr. Pathol. 24: 311-331.

Couch, J.A. 1974b. Pathological effects of Urosporidium
(Haplosporida) infection in microphallid metacercariae.
J. Invertebr. Pathol. 23: 389-396.

Couch, J.A. 1981. Viral diseases of invertebrates other than
insects. Pages 127-160 in E.W. Davidson, ed. Pathogen-
esis of invertebrate microbial diseases. Allanheld Osmun
& Co. Publishers, Totowa, N.J.

Couch, J.A. 1983. Diseases caused by Protozoa. Pages 79-
111 in A.J. Provenzano, ed. The biology of Crustacea.
Vol. 6. Pathobiology. Academic Press, New York.

Couch, J.A.,and S. Martin. 1982. Protozoan symbionts and
telated diseases of the blue crab, Callinectes sapidus
Rathbun from the Atlantic and Gulf coasts of the United


Synopsis of Principal Diseases

of the

Blue Crab, Callinectes sapidus

U. S. DEPARTMENT OF COMMERCE
National Oceanic and Atmospheric Administration
National Marine Fisheries Service
Northeast Region
Northeast Fisheries Science Center
Woods Hole, Massachusetts

January 1992

REPRODUCED BY
U.S DEPARTMENT OF COMMERCE,

NATIONAL TECHNICAL INFORMATION ceayi
SPRINGFIELD, VA 22161 Seavice



Page 6

Table 2. Viral diseases of blue crabs

Virus Name Histopathology and Effect Gross Signs Geographic Location Reference
and Type Tissues Infected on Host of Disease and Prevalence
Bi-facies Nuclei of Nuclear Just before Assawoman Johnson
(BFV); form- hemocytes hypertrophy death crabs and 1976b,
erly herpes- hypertrophied; followed by become Chincoteague 1978b,
like (HLV); refractile cell lysis; inactive; Bays only; 1983, 1984,
fatal; DNA cytoplasmic death due to withdrawn, captive and 1988
virus; re- inclusions hemocytic blood milky wild crabs;
lated to dysfunction and clots teported
Trido-, Pox-, improperly infections
and Baculo- upto 13% -
viridae
Baculo-A; Nuclei of Benign, since Crabs Widespread Johnson.
benign; epithelial hepatopan- appear along 1976a,
DNA virus cells of creatic cells healthy Atlantic 1983, 1984;
(non- hepatopancreas constantly coast; Johnson &
occluded) hypertrophied, replaced infections Lightner
focal from 4 to 20% 1988
infections in adults
and juveniles
Baculo-B; Nuclear Lysing and Occurs Tred Avon Bazin et
DNA virus; hypertrophy dysfunction in either River, al, 1974;
similar to of hemopoietic of hemocytes, normal- Chesapeake Johnson
baculovirus tissue and otherwise appearing Bay 1983, 1984,
of Carcinus hemocytes effect crabs or in 1986
maenas unknown those with
other viral
infections
Reo-like Cytoplasmic Rhabdo-like Slowness, Actual Vago 1966;
LY); fatal; inclusions; A (RbVA) al- paralysis; prevalence Johnson
RNA virus; increased ways associ- withdrawn, unknown, 1977a, c,
similar to cytoplasmic ated with blood does infects 1983, 1984,
paralysis volume in RLV; syner- not clot crabs from . 1986
virus of hemocytes, gistically completely various
Macropipus hemopoietic causes par- salinities
depurator tissue, alysis and from Chesapeake
and glial death due to and Chinco-
nerves nerve and teague Bays
hemocyte
dysfunction
Rhabdo- Always seen Stress No reported Atlantic Jahromi
like A with other telated; gfoss signs and Gulf 1977; Yudin
(RbVA); viruses; may have coasts; & Clark
synergistic; infects synergistic may be 1978, 1979;
RNA virus cytoplasm effect with ubiquitous Johnson.
of nerve other virus 1978a, b,
ganglia, diseases 1983, 1984,
1986

hemopoietic
tissue



@ Carcinonemertes carcinophila (nemertean)
@ = Myzobdella lugubris (leech)

Parasitic Crustacean
‘

@ Loxothylacus texanus (thizocephalan)

VIRAL DISEASES

Before 1966, when a virus was first reported from a
European shore crab (Vago 1966), viruses from marine
invertebrates were unknown. Since the 1960s, with in-
creased availability of transmission electron microscopy,
other mortality-associated viruses have been discovered in
crustaceans (Johnson and Bodammer 1975; Couch 1981;
Johnson 1984). Today there are seven, possibly eight,
viruses reported infecting blue crabs, although not all are
pathogenic or cause mortalities. Viral effects on the host
may be lethal, the result of synergism with other viruses,
relatively benign, or even unknown (Johnson 1978b). Of the
viruses known to infect blue crabs (Table 2), three are
relatively benign: baculoviruses A and B, and rhabdo-like
virus B (RhVB). Two are found to be associated with other
viruses and perhaps cause a synergistic effect: paramyxo-
like enveloped helical virus (EHV) and RhVA. Three others
are considered lethal: RLV, BFV, and CBV (Figure 1). The
RhVA apparently acts synergistically with the RLV and is
responsible for hemocyte and nerve tissue damage which
may result in death, especially in premolt crabs about to go
through ecdysis (molt), since this process is controlled by
the nervous system (Johnson 1986). Multiple viral infec-
tions are frequently discovered within the same crab. Viral
infections may become patent only when crabs have been
exposed to stress factors such as captivity, crowding, or a
degraded environment (Couch 1974a; Johnson 1977a, 1978b,
1984, 1986; Yudin and Clark 1979; Messick and Kennedy
1990).

BACTERIAL DISEASES

SHELL DISEASE

The disease syndrome known variously as shell dis-
ease, rust disease, black spot, or burnt spot was first discov-
ered in crustaceans in 1900 in Germany by Happich (Rosen
1970), and was first reported in the Chesapeake Bay in 1967
(Rosen 1967). The syndrome, which is manifested as
various kinds of exoskeletal erosions and lesions, is caused
by chitinoclastic bacteria (and frequently other chitin-de-
stroying microorganisms). Since there are numerous spe-
cies of chitinoclastic bacteria found in marine environments
(Benton 1935; Zobell and Rittenberg 1938; Hock 1940,
1941; Lear 1963; Hood and Meyers 1974), shell disease is

Page 5

one of the most common and widespread bacterial diseases
of crustaceans. Chitinoclastic bacteria play an important
role inthe breakdown and recycling of chitin in exoskeletons
of dead animals and chitinous casts from developmental
stages of crustaceans in the sea (Hock 1940). Although
beneficial in recycling chitin, these bacteria can be detri-
mental to live crustaceans (Sindermann 1989). The bacteria
can cause deep, necrotic, blackened lesions on various
portions of the crab’s exoskeleton, thus decreasing its mar-
ket value, making it unmarketable, or even allowing second-
ary infections to invade tissues and kill crabs.

Shell disease in blue crabs occurs naturally in all
populations and has been reported from such varied geo-
graphic locations as the Louisiana and Mississippi coasts,
Biscayne Bay (Florida), the South Carolina coast, Pamlico
River, and the Chesapeake Bay. A large portion of blue
crabs reported with shell disease has been subjected to
abnormal environmental conditions (Sindermann 1989),
either in the wild by being exposed to deteriorated water
quality, as was thought to be the case in coastal North
Carolina (Engel and Noga 1989, McKenna et al. 1990), or
by being held in crowded conditions such as shedding or
holding tanks for extended periods (Figures 2 and 3).

Most chitinoclastic bacteria isolated from lesions in
blue crabs belong to several genera including Vibrio and
Pseudomonas (Cook and Lofton 1973; Johnson 1983). For
a detailed report on shell disease, see Sindermann (1989).

SYSTEMIC BACTERIAL INFECTIONS

Many blue crab mortalities are attributed to systemic
bacterial infections, especially when the animals are sub-
jected to crowded, confined conditions. Mortalities of 50
percent or higher have been reported in crabs held in
commercial shedding facilities (Krantz et al. 1969). Gross
signs of infection include lethargy, weakness, and possibly
an enlarged chalky white area on a fifth pereiopod or the
gills. Upon dissection, a premortem plasma clot can often
be seen in anterodorsal and frontal blood sinuses. Hemo-
lymph from infected crabs may have reduced clotting abili-
ties and diminution of hemocyte numbers. The formation of
hemocyte aggregations can be seen histologically in arteries
and hemal spaces of various tissues, including the heart; as
the infection progresses, nodules can be seen in gills, heart,
antennal gland, and other organs (Johnson 1976c) (Figure
4). Apparently healthy crabs may exhibit a hemocytic
response seen histologically which may be the result of
recovery from a previous light bacterial infection (Messick
and Kennedy 1990). Various species of bacteria have been
isolated from the hemolymph of moribund and apparently
healthy crabs, including Vibrio parahaemolyticus, a patho-
gen to both crabs and humans (Krantz er al. 1969; Davis and
Sizemore 1982; Sizemore and Davis 1985), also Pseudo-
monas, Acinetobacter, Bacillus, Flavobacterium, and a




ae

(continued from inside front cover)

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Division, Northeast Fisheries Center. December 1989. iv + 110p.,50 figs., 93 tables. NTIS Access. No. PB90-170622/
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and Tamara J. Holzwarth. December 1989. iii + 32 p., 31 figs. NTIS Access. No. PB90-252172/AS.

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December 1989. iii+14 p., 11 tables, 1 app. NTIS Access. No. PB91-128389/AS.

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Page 2

Table 1. Principal pathogens of blue crabs

Pathogen _ Histopathology and Effect Gross Signs Geographic Location Reference
Tissues Infected on Host of Disease and Prevalence
Reo-like . Rhabdolike RLV and Sluggishness, Chincoteague Johnson
virus (RLV); virus A (RhVA) RhVA act paralysis; and 1977a, ¢,
infects RNA always found synergis- withdrawn Chesapeake 1983, 1984,
with RLV; tically, blood clots Bays; infects 1986
causes cyto- causing incompletely crabs from.
plasmic inclu- necrosis of high and low
sions, in- hemopoietic salinities;
creased cyto-. tissue, actual
plasmic volume hemocytes, prevalence
in hemocytes, and CNS;* unknown
hemopoietic death due
tissue, and to nerve and
glial nerves hemocytic
dysfunction
Chesapeake Causes focal Extensive Abnormal Chesapeake Johnson
Bay virus infections; destruction behavior, Bay; infects 1978a, b
(CBV), cytoplasmic of gill and erratic captive 1983, 1984,
infects inclusions in bladder swimming, juveniles and 1986
RNA; a epithelium of epithelium blindness probably wild
picoma- ill, gut, and neuro- populations
like virus bladder, CNS secretory
cells,and = - cells,
epidermis blindness,
and death
Bi-facies Hypertrophy Nuclear Crabs become Chincoteague Johnson
virus (BFV); of nuclei hypertrophy inactive; and Assawoman 1976b, 1978b,
infects DNA; of hemocytes; followed by withdrawn, Bays; infects 1983, 1984,
formerly infected lysis of cell; blood is captive 1988 ©
herpes-like cells have death due to milky and and wild
virus (HLV) refractive hemocytic clots populations
cytoplasmic dysfunction improperly
inclusions
Chitino- Erosion of Decreased Exoskeleton. Ubiquitous, Rosen 1967,
clastic exoskeleton, salability; blackened particularly 1970; Cook
bacteria, especially secondary and pitted; apparent when & Lofton 1973;
principally chelipeds infections condition crabs have Johnson 1983;
Vibrio and and carapace may cause called “shell been injured Engel &
Pseudomonas, _ overlying internal disease” or or held in Noga 1989;
plus others gills necrosis of “box bumt prolonged Sindermann
tissue and disease” captivity 1989
death
Vibrio Hemolymph Formation Lethargy and Injured or Krantz et
parahaemo- contains of hemocytic weakness due stressed al, 1969;
lyticus and bacteria; aggregations to systemic crabs prone Sizemore et
related causes and nodules; infections; to disease; al, 1975;
facultative hemocytic causes withdrawn mortalities Tubiash et
bacteria aggregations internal blood clots reported 50% al. 1975,
in gills, clotting of incompletely or higher Johnson
heart, and hemolymph in shedding 1976¢; Blake
other tissues tanks etal, 1980a,
b; Messick &

Kennedy 1990



Page 16

A truly parasitic sacculinid barnacle infecting blue
crabs is the rhizocephalan Loxothylacus texanus. Postmolt
juvenile crabs are infected by swimming L. texanus cypris
larvae. The interna or “root system” of the barnacle pen-
etrates the soft abdominal joint of the crab and is extended
into internal tissues when the crab molts. The externa of the
barnacle, consisting of a brood pouch for gonads and larvae,
protrudes from under the crab’s abdomen, appearing as a
bulge which can be seen grossly. The barnacle causes a
restriction in growth; infected individuals are only between
3 and 8 cm wide, thereby making them unavailable to the
blue crab fishery. Infected crabs are often termed “button
crabs” and appear as miniature adult females, with second-
ary sexual characteristics transformed to those of a female
(Figure 13). The abdominal segments of infected males
become wider; six distinct segments develop instead of the
normal four; and abdominal musculature appears similar to
that of a mature female. The aprons of infected immature
females are transformed to broader, rounded aprons similar
to mature females (Reinhard 1950, 1956; Adkins 1972).

Castration may result from a malfunction of the andro-
genic gland in males; in experiments in which the organ was
removed from a young male, female characteristics devel-
oped, including production of ovarian tissue in the testes. It
is conceivable that L. texanus causes malfunction to this
gland, or possibly hormones produced by the barnacle’s
ovaries cause over-feminization of infected crabs (Overstreet
1978). Infections have been reported from the Gulf coast
only and are usually seen in high-salinity waters (Ragan and
Mathere 1974). Epizootics were reported in Mississippi in
1965 and 1977 (Overstreet 1978). Since diseased crabs are
below market size, actual prevalence of the parasite is hard
toestimate, and because the parasite removes crabs from the
fishery, it causes significant economic losses during epizoot-
Ics.

SUMMARY

From the abundant literature available on diseases of
blue crabs, it becomes obvious that there are several severe
pathogens that can cause mortalities in both wild and captive
blue crabs. Because of the economic importance of the blue
crab fishery, it is apparent that diseases should be controlled
whenever possible. Through increased research, better
methods of control can be devised to help reduce the damage
diseases can cause, as was the case with “sick crab” disease
caused by microsporidans. After finding that this disease is
transmitted by crabs ingesting infected tissues from dead
diseased crabs, watermen were told to stop crushing dis-
eased crabs and throwing them back into the water. Instead,
crabbers were told to dispose of the crabs properly on shore,
thus reducing some of the potential spread of the parasite.

Diseases commonly found in crabs held in captivity
such as bacterial and viral infections can be aggravated by
stress. The viruses considered pathogenic in blue crabs are:

Figure 9. Ciliates infesting blue crab gills.
a. Stalked ciliate, Epistylis, attached to stem (S) of blue
crab gill. Bar = 20 pm.
b. Lagenophrys ciliates cemented to flat surfaces of gill.
Note semitransparent lorica. Bar = 40 pm.
c. Infestation of the gills of a blue crab by the ciliate

Lagenophrys.

Couch.)
(1) the reo-like virus (RLV) acting synergistically with
thabdo-like virus A (RhVA), (2) the Chesapeake Bay virus
(CBV), and (3) the bi-facies virus (BFV). The RLV-RhVA
combination is known to cause mortalities in shedding
systems. Perhaps by devising better methods of reducing
stress in these animals, mortalities could be lowered, thus
allowing for increased productivity and profits for the soft-
shell crab producer.

Shell disease, caused by chitinoclastic bacteria, occurs
frequently in crabs held in captivity for prolonged periods.
This disease could be limited by reducing captivity time;
when this is impossible, some kind of prophylactic treat-
ment to exclude chitinoclastic bacteria would help reduce
the incidence of this disfiguring disease which may make,
crabs unmarketable.

(Photograph courtesy of Dr. J.A.



Page 22

States. Pages 71-81 in H.M. Perry and W.A. Van Engel,
eds. Proceedings of the Blue Crab Colloquium, Biloxi,
Miss. Gulf States Marine Fisheries Commission, Ocean
Springs, Miss.

Couch, J.N. 1942. A new fungus on crab eggs. J. Elisha
Mitchell Sci. Soc. 58: 158-162.

Davis, C.C. 1965. Astudy of the hatching process in aquatic
invertebrates: XX. The blue crab, Callinectes sapidus,
Rathbun, XXI. The nemertean, Carcinonemertes
carcinophila (Kolliker). Chesapeake Sci. 6(4) 201-208.

Davis, J.W., and R.K. Sizemore. 1982. Incidence of Vibrio
species associated with blue crabs (Callinectes sapidus)
collected from Galveston Bay, Texas. Appl. Environ.
Microbiol. 43(5): 1092-1097.

DeTurk, W.E. 1940. The occurrence and development of
ahyperparasite Urosporidium crescens sp.nov. (Sporozoa,
Haplosporidia), which infests the metacercariae of
Spelotrema nicolli, parasitic in Callinectes sapidus. J.
Elisha Mitchell Sci. Soc. 56: 231-232.

Engel, D.W., and E.J. Noga. 1989. Shell disease in the blue
crabs of the Pamlico River. Environs, Feb. 1989: 3-5.
Fisher, W.S., E.H. Nilson, and R.A. Shleser. 1975. Effect
of the fungus Haliphthoros milfordensis on the juvenile
stages of the American lobster, Homarus americanus. J.

Invertebr. Pathol. 26: 41-45.

Fisher, W.S., E.H. Nilson, L.F. Follett, and R.A. Shleser.
1976. Hatching and rearing lobster larvae (Homarus
americanus) in a disease situation. Aquaculture 7: 75-80.

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

Hay, W.P. 1905. The life history of the bluecrab (Callinectes
sapidus). Rep. U.S. Bur. Fish. 1905: 395-413.

Hock, C.W. 1940. Decomposition of chitin by marine
bacteria. Biol. Bull. (Woods Hole) 79: 199-206.

Hock, C.W. 1941. Marine chitin-decomposing bacteria. J.
Mar. Res. 4: 99-106.

Hood, M.A., and S.P. Meyers. 1974. Distribution of
chitinoclastic bacteria in natural estuarine waters and
aquarial systems. Pages 115-121 in R.S. Amborski, M.A.
Hood, and R:R. Miller, eds. Proceedings of Gulf Coast
Regional Symposium on Diseases of Aquatic Animals.
La. State Univ. Publ. No, LSU-SG-74-05.

Hopkins, S.H. 1947. The nemertean Carcinonemertesasan
indicator of the spawning history of the host, Callinectes
sapidus. J. Parasitol. 33: 146-150.

Humes, A.G. 1942. The morphology, taxonomy and
bionomics of the nemertean genus Carcinonemertes. Ill.
Biol. Monogr. 18(4): 1-105.

Hutton, R.F.,and F. Sogandares-Bemal. 1959. Notes onthe
distribution of the leech, Myzobdella lugubris Leidy, and
its association with mortality of the blue crab, Callinectes
sapidus Rathbun. J. Parasitol. 45: 384, 404, 430.

Jahromi, $.S. 1977. Occurrence of rhabdovirus-like par-
ticles in the blue crab, Callinectes sapidus. J. Gen. Virol.
36: 485-493.

Johnson, P.T. 1976a. A baculovirus from the blue crab,
Callinectes sapidus. Page 24 in Proceedings of the First
International Colloquium on Invertebrate Pathology and
IXth Annual Meeting of the Society for Invertebrate
Pathology, Kingston, Ont. Queen’s University, Kingston,
Ont. .

Johnson, P.T. 1976b. A herpeslike virus from the blue crab,
Callinectes sapidus. J. Invertebr. Pathol. 27: 419-420.

Johnson, P.T. 1976c. Bacterial infection in the blue crab,
Callinectes sapidus: course of infection and
histopathology. J. Invertebr. Pathol. 28: 25-36.

Johnson, P.T. 1977a. A viral disease of the blue crab,
Callinectes sapidus: histopathology and differential diag-
nosis. J. Invertebr. Pathol. 29: 201-209.

Johnson, P.T. 1977b. Paramoebiasis in the blue crab,
Callinectes sapidus. J. Invertebr. Pathol. 29: 308-320.

Johnson, P.T. 1977c. Reovirus-like virus of blue crabs.
Pages 103-105 in C.J. Sindermann, ed. Disease diagnosis
and control in North American marine aquaculture.
Elsevier, New York.

Johnson, P.T. 1978a. New information on viral diseases of
the blue crab, Callinectes sapidus. Page 55 in S.E.
Milligan, ed. International Colloquium on Invertebrate
Pathology and XIth Annual Meeting of the Society for
Invertebrate Pathology, Prague, Czechoslovakia.

Johnson, P.T. 1978b. Viral diseases of the blue crab,
Callinectes sapidus. Mar. Fish. Rev. 40(10): 13-15.

Johnson, P.T. 1980. Histology of the blue crab, Callinectes
sapidus: a model for the Decapoda. Praeger, New York.
440 pp:

Johnson, P.T. 1983. Diseases caused by viruses, rickettsiae,
bacteria, and fungi. Pages 1-78 in A.J. Provenzano, ed.
The biology of Crustacea. Vol. 6. Pathobiology. Aca-
demic Press, New York.

Johnson, P.T. 1984. Viral diseases of marine invertebrates.
Pages 65-98 in O. Kinne and H.P. Bulnheim, eds. Dis-
eases of marine organisms. Helgol. Meeresunters. 37.

Johnson, P.T. 1986. Bluecrab (Callinectes sapidus Rathbun)
viruses and the diseases they cause. Pages 13-19 in H.M.
Perry and R.F. Malone, eds. Proceedings of the National
Symposium on the Soft-Shelled Blue Crab Fishery. Gulf
Coast Research Laboratory, Biloxi, Miss.

Johnson, P.T. 1988. Development and morphology of an
unusual nuclear virus of the blue crab Callinectes sapidus.
Dis. Aquat. Org. 4: 67-75.

Johnson, P.T., and J.E. Bodammer. 1975. A disease of the
blue crab, Callinectes sapidus, of possible viral etiology.
J. Invertebr. Pathol. 26: 141-143.

Johnson, P.T., and C.A. Farley. 1980. A new enveloped
helical virus from the blue crab, Callinectes sapidus. J.
Invertebr. Pathol. 35: 90-92.

Johnson, P.T., and D.V. Lightner. 1988. Rod-shaped
nuclear viruses of crustaceans: gut-infecting species. Dis.
Aquat. Org. 5: 123-141.

Krantz, G.E., R.R. Colwell, and E..Lovelace. 1969. Vibrio
parahaemolyticus from the blue crab Callinectes sapidus



NOTE ON SPECIES NAMES

The Northeast Fisheries Science Center's policy on the use of species names in technital publications and reports is to follow
the American Fisheries Society’s (AFS) lists of common and scientific names for fishes (Robins et al. 1991)*, mollusks

(Turgeon et al. 1988), and decapod crustaceans (Williams et al. 1989)*. This policy applies to all issues of the NOAA
Technical Memorandum NMFS-F/NEC series.

* Robins, C.R. (chair), R.M. Bailey, C.E. Bond, J.R. Brooker, E.A. Lachner, R.N. Lea, and W.B. Scott. 1991. Common and scientific names 'of fishes
from the United States and Canada, 5th ed. Amer. Fish. Soc. Spec. Publ. 20. 183 pp.

* Turgeon, D.D. (chair), A.E. Bogan, E.V. Coan, W.K, Emerson, W.G. Lyons, W.L. Pratt, C.F.E. Roper, A. Scheltema, F.G. Thompson, and J.D.
Williams. 1988, Common and scientific names of aquatic invertebrates from the United States and Canada: mollusks. Amer. Fish. Soc. Spec. Publ.
16. 277 pp.

* Williams, A.B. (chair), L.G, Abele, D.L. Felder, H.H. Hobbs, Jr., R.B. Manning, P.A. McLaughlin, and I. Pérez Farfante. 1989. Common and
scientific names of aquatic invertebrates from the United States and Canada: decapod crustaceans. Amer. Fish, Soc. Spec. Publ. 17. 77 pp.



contact with crab eggs (Couch 1942). Infected eggs are
usually below their normal size and either fail to hatch or
Ptezoea are produced which are abnormal and rarely survive
(Sandoz et al. 1944). Infected eggs have an atypical brown
or grey color, depending on the maturity of the eggs infected.
Heaviest infections occur on sponge crabs that are exposed
to the fungus for the longest time, regardless of the develop-
mental stage of the eggs (Newcombe and Rogers 1947).
Infection spreads rapidly over the periphery of the egg mass,
but penetration into the sponge is slow and rarely over 3-mm
deep. Peripheral infections donot retard the development of
eggs in the interior of the sponge; therefore, less than 25
percent of the eggs of heavily diseased sponges are infected.
Itis not unusual to find 80-90 percent of the crabs ina sample
population to have some degree of infection. The fungus has
a broad tolerance for salinity change and, experimentally,
can develop in salinities from 5 to 30 ppt. Lagenidium can
also infect larvae of blue crabs, causing extensive mortali-
ties. Under laboratory conditions, transmission of the
fungus is extremely rapid, with infections apparent within
two days (Rogers-Talbert 1948).

Lagenidium callinectes has been transmitted experi-
mentally to eggs of the oyster pea crab, Pinnotheres ostreum,
and the mud crab, Dyspanopeus (Neopanope) texana, which
inhabit the same areas as the blue crab (Rogers-Talbert
1948). Other strains of Lagenidium have been isolated in
larvae from cultured shrimp in Florida, Texas, Mexico,
Tahiti, and Honduras; in Dungeness crab, Cancer magister,
larvae from Oregon; and American lobster, Homarus
americanus, larvae from California. The fungus is consid-
ered so potentially pathogenic that attempts have been made
to devise chemical control methods (Bland et al. 1976).

Another phycomycetous fungus, Haliphthoros
milfordensis, was isolated initially from ova of the Atlantic
oyster drill, Urosalpinx cinerea, but also infects penaeid
shrimp, lobsters (Fisher et al. 1975, 1976, 1978), and,
experimentally, blue crab ova. Infected ova of the blue crab
can be identified by a color change; normally brown, mature
ova appear dark orange-yellow and are filled with multi-
branched hyphae. During sporulation, long unbranched,
extramatrical discharge tubes grow through the surface of
the ova to the external medium. Zoospores from one
infected egg are capable of infecting 20-30 other eggs in the

_ Same sponge within 24 hours. Haliphthoros milfordensis is
morphologically similar to L. callinectes (Tharp and Bland
1977).

PROTOZOAN DISEASES

There are many protozoans known to cause diseases in
blue crabs; some occur in epizootic proportions which result
in mass mortalities, but often these diseases are reported as
being enzootic, killing only an occasional individual. Dis-
eases caused by protozoans often become patent only when
crabs are held in artificial environments, such as peeler
shedding tanks or holding facilities for market. There are

Figure 2. Shell disease in the blue crab -- early lesions (above) and
advanced lesions (below). (Photographs courtesy of Dr.
RM. Overstreet, Gulf Coast ResearchLaboratory, Ocean.

Springs, Mississippi.)

several reviews of protozoan pathogens, including those
infecting blue crabs (Sindermann and Rosenfield 1967;
Sindermann 1990; Sprague 1970; Sprague and Couch 1971;
Overstreet 1978; Couch and Martin 1982, Couch :1983;
Millikin and Williams 1984, Sparks 1985). Some of the
most pathogenic protozoans found infecting blue crabs are
microsporidans, including Ameson, Nosema, and
Pleistophora, two members of the Sarcomastigophora,.
Paramoeba and Hematodinium, a haplosporidan-like
ascetosporan; and a ciliophoran, Lagenophrys.

* At least five different species of microsporida infect
blue crabs; one that is widespread and causes considerable
harm to the host is A. michaelis (Overstreet 1978). This


| [Reproduced trom

best available copy. ©)

Figure 4. Bacterial infections in blue crab tissue.

: a. Hemocyte aggregation (h) and nodule formation (n)
in heart tissue. Note pyknotic nuclei in center of
nodule and layer of flattened hyaline hemocytes
encapsulating nodule (arrow). Bar = 20 um.

b. Nodule formation in heart tissue (arrow). Bar = 20
ym.

populations (Sprague 1966, 1970; Couch and Martin 1982).

Besides the Microspora, there are other pathogenic
protozoan phyla recognized as causing diseases in blue
crabs. From the phylum Sarcomastigophora, there are an
amoebic parasite, Paramoeba perniciosa, and a parasitic
dinoflagellate, Hematodinium perezi. Both these parasites
can cause lethal infections in blue crabs along the Atlantic
coast of the United States.

A well documented disease of blue crabs is the “grey
crab disease” caused by P. perniciosa. First reported in blue
crabs by Sprague and Beckett (1966, 1968), the parasite was
identified as the causative agent in crab shedding mortalities
in Chincoteague Bay, Virginia. The condition had long been
detected and reported by fishermen from Maryland and
Virginia who noticed that some crabs which appeared
sluggish would often havea grey ventral surface during May
and June--hence, the term “grey crab disease.” These crabs
had reduced clotting abilities and would often die after being
stressed by handling (Sawyer 1969; Sprague et al. 1969;
Johnson 1977b; Couch 1983).

Figure 5. Microsporidan infection in blue crab skeletal muscle
tissue.

a. Muscle lysis (arrow), and tissue destruction and
replacement by spores (S). Bar = 50 ym.

b. Muscle lysis (L), and bundles of spores (arrows). Bar
= 50pm.

c. Note individual spores in each bundle (arrow).
Bar = 10 pm.


NOAA Technical Memorandum NMFS-F/NEC-88

This TM series is used for documentation and timely communication of preliminary results,
interim reports, or special purpose information, and has not undergone external scientific
review.

Synopsis of Principal Diseases
of the
Blue Crab, Callinectes sapidus

Gretchen A. Messick and Carl J. Sindermann
Oxford Lab., National Marine Fisheries Serv., Oxford, MD 21654

U. S. DEPARTMENT OF COMMERCE
‘ Rockwell A. Schnabel, Acting Secretary

National Oceanic and Atmospheric Administration
John A. Knauss, Administrator

National Marine Fisheries Service
William W. Fox, Jr., Assistant Administrator for Fisheries

. Northeast Region
Northeast Fisheries Science Center
Woods Hole, Massachusetts

January 1992

jee


Page 20

Figure 13. Sacculinid bamacle, Loxothylacus texanus, infecting blue crab. (Photographs courtesy of Dr. R.M. Overstreet.)

a. A blue crab infected with the rhizocephalan L. texanus (bottom), compared to a normal egg-bearing female (top).

b. The protruding pouch (externa) under the crab’s abdomen contains larvae and gonads (ovary and testes), Long, narrow
extensions invade most other tissues.

¢. Topcrab has dark extema, indicating an older infection than that of the yellowish externa of the middle crab. The larger
lower crab reveals the small size of infected crabs. It is an immature female, whereas the aprons on the infected crabs
reveal the apron of a mature crab. Infections also modify the shape of the male's abdomen into that of a female,

d. Crab with three developing externae. Usually only one extema occurs per crab, but occasionally over five may be present.



Recent issues in this series

53. Contaminants in Hudson-Raritan Estuary Water and Influence of Cold Storage upon Its Chemical
Composition. By Anthony Calabrese, Lawrence J. Buckley, and J. Christopher Powell. May 1988. vii + 37 p., 10 figs.,
11 tables. NTIS Access. No. PB88-225628/AS.

54. Epizootic Ulcerative Syndromes in Coastal/Estuarine Fish. By Carl J. Sindermann. June 1988. v + 37 p., 8 figs.,
1 table. NTIS Access. No. PB89-110803/AS.

55. A Plan for Study: Response of the Habitat and Biota of the Inner New York Bight to Abatement of Sewage
Sludge Dumping. By Environmental Processes Division, Northeast Fisheries Center. June 1988. iii + 34 p., 5 figs., 3
tables, 4 app. NTIS Access. No. PB89-100903/AS.

56. Characterization of the Middle Atlantic Water Management Unit of the Northeast Regional Action Plan. By
Anthony L. Pacheco, ed. July 1988. v + 322 p., 136 figs., 21 tables. NTIS Access. No. PB89-145262/AS.

57. An Analysis and Evaluation of Ichthyoplankton Survey Data from the Northeast Continental Shelf
Ecosystem. By Wallace G. Smith, ed. August 1988. xiii + 132 p., 53 figs., 12 tables, 1 app. NTIS Access. No. PB89-
122501/AS. .

58. An Indexed Bibliography of Northeast Fisheries Center Publications and Reports for 1987. By Jon A. Gibson.
August 1988. iii + 20 p. NTIS Access. No. PB89-113013/AS.

59. Surveys of Breeding Penguins and Other Seabirds in the South Shetland Islands, Antarctica, January-
February 1987. By W. David Shuford and Larry B. Spear. September 1988. vii + 27 p., 14 figs., 1 table. NTIS Access.
No. PB89-141311/AS.

60. Survey of Antarctic Fur Seals in the South Shetland Islands, Antarctica, during the 1986-1987 Austral
Summer. By John L. Bengtson, Lisa M. Ferm, Tero J. Harkonen, Everett G. Schaner, and Brent S. Stewart. September
1988. vii + 8 p., 1 fig., 3 tables. NTIS Access. No. PB89-141303/AS.

61. Fish as Sentinels of Environmental Health. By Robert A. Murchelano. September 1988. iii + 16 p., 4 figs. NTIS
Access. No. PB89-139737/AS.

62. The Effects of Density Dependent Population Mechanisms on Assessment Advice for the Northwest Atlantic
Mackerel Stock. By W. J. Overholtz, S.A. Murawski, W.L. Michaels, and L.M. Dery. October 1988. v + 49 p.,7 figs.,
20 tables. NTIS Access. No. PB89-151948/AS.

63. Status of the Fishery Resources Off the Northeastern United States for 1988. By Conservation and Utilization
Division. October 1988. iii + 135 p., 51 figs., 52 tables. NTIS Access. No. PB89-130819/AS.

64. The Shell Disease Syndrome in Marine Crustaceans. By Carl J. Sindermann. February 1989. v + 43 p., 5 figs.,
2 tables. NTIS Access. No. PB89-162523/AS.

65. Stock Assessment Information for Pollock, Pollachius virens (L.), in the Scotian Shelf, Georges Bank, and Gulf
of Maine Regions. By Ralph K. Mayo, Stephen H. Clark, and M. Christina Annand. April 1989. vi + 14 p., 6 figs.,
14 tables. NTIS Access. No. PB90-120676/AS. ;

66. Guidelines for Estimating Lengths at Age for 18 Northwest Atlantic Finfish and Shellfish Species. By Judith
A. Pentilla, Gary A. Nelson, and John M. Burnett, III. May 1989. iii + 39 p., 18 figs., 19 tables. NTIS Access. No., PB90-
120675/AS.

67. Response of the Habitat and Biota of the Inner New York Bight to Abatement of Sewage Sludge Dumping.

Second Annual Progress Report - 1988. By Environmental Processes Division, Northeast Fisheries Center. July
1989. vii + 47 p., 39 figs., 11 tables, 3 app. NTIS Access. No. PB90-125444/AS.

(continued on inside back cover)

es



in Chesapeake Bay. Science 164: 1286-1287.

Lear, D.W., Jr. 1963. Occurrence and significance of
chitinoclastic bacteria in pelagic waters and zooplankton.
Pages 594-610 in C. Oppenheimer, ed. Symposium on
Marine Microbiology. Thomas, Springfield, Ill.

MacLean, S.A., and C.L. Ruddell. 1978. Three new
crustacean hosts for the parasitic dinoflagellate
Hematodinium perezi (Dinoflagellata: Syndinidae). J.
Parasitol. 64: 158-160.

McKenna, S., M. Jansen, and M.G. Pulley. 1990. Shell
disease of blue crabs, Callinectes sapidus in the Pamlico
River, North Carolina. [N.C. Div. Mar. Fish.,] Spec. Sci.
Rep. No. 51. 30 pp.

Messick, G.A., and V.S. Kennedy. 1990. Putative bacterial
and viral infections in blue crabs, Callinectes sapidus
Rathbun, 1896 held in a flow-through or a recirculation
system. J. Shellfish Res. 9: 33-40.

Millikin, M.R., and A.B. Williams. 1984. Synopsis of
biological data on the blue crab, Callinectes sapidus
Rathbun. [U.N.,] FAO [Food Agr. Org.] Fish. Synop. No.
138, NMFS-S-138. 39 pp.

Moore, J.P. 1946. The anatomy and systematic position of
Myzobdella lugubris Leidy (Hirudinea). Not. Nat. (Phila.)
184: 1-12.

Newcombe, C.L., and M.R. Rogers. 1947. Studies of a
fungus parasite that infects blue crab eggs. Turtox News
25(9): 1-7.

Newcombe, C.L., F. Campbell, and A.M. Eckstine. 1949.
Astudy of the form and growth of the blue crab Callinectes
sapidus Rathbun. Growth 13: 71-96.

Newman, M.W. 1977. Hematodinium disease of blue
crabs. Pages 211-213 in C.J. Sindermann and D.V.
Lightner, eds. Disease diagnosis and control in North
American marine aquaculture. Elsevier, New York.

Newman, M.W., and C.A. Johnson, III. 1975. A disease of
blue crabs (Callinectes sapidus) caused by parasitic di-
noflagellates, Hematodinium sp. J. Parasitol. 61: 554-
555. ut

Newman, M.W., and G.E. Ward, Jr. 1973. An epizootic of
blue crabs, Callinectes sapidus, caused by Paramoeba
perniciosa. J. Invertebr. Pathol. 22: 329-334.

Newman, M.W.,C.A. Johnson, III, and G.B. Pauley. 1976.
A Minchinia-like haplosporidan parasitizing blue crabs,
Callinectes sapidus. J. Invertebr. Pathol. 27: 311-315.

Overstreet, R.M. 1977. Microsporosis of blue crabs. Pages
200-203 in C.J. Sindermann and D.V. Lightner, eds.
Disease diagnosis and control in North American marine
aquaculture. Elsevier, New York.

Overstreet, R.M. 1978. Marine maladies? Worms, germs,
and other symbionts from the northern Gulf of Mexico.
Blossman Printing, Ocean Springs, Miss. 140 pp.

Overstreet, R.M. 1982. Metazoan symbionts of the blue
crab. Pages 81-87 in H.M. Perry and W.A. Van Engel,
eds. Proceedings of the Blue Crab Colloquium, Biloxi,
Miss. Gulf States Marine Fisheries Commission, Ocean
Springs, Miss.

Page 23

Overstreet, R.M. 1983. Metazoan symbionts of crusta-
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Overstreet, R.M., and E.C. Whatley. 1975. Prevention of
microsporidosis in the blue crab, with notes on natural
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Pauley, G.B., M.W. Newman, and E. Gould. 1975. Serum
changes in the blue crab, Callinectes sapidus, associated
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Pyle, R., and E. Cronin. 1950. The general anatomy of the
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Reinhard, E.G. 1950. An analysis of the effects of a
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Reinhard, E.G. 1956. Parasitic castration of Crustacea.
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Rogers-Talbert, R. 1948. The fungus Lagenidium callinectes
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Biol. Bull. (Woods Hole) 95: 214-228.

Rosen, B. 1967. Shell disease of the blue crab, Callinectes
sapidus. J. Invertebr. Pathol. 9: 348-353.

Rosen, B. 1970. Shell disease of aquatic crustaceans. Pages
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of Fishes and Shellfishes. Amer. Fish, Soc. Spec. Publ. 5.

Rosenfield, A., L. Buchanan, and G.B. Chapman. 1969.
Comparison of the fine structure of spores of three species
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Parasitol. 55: 921-941.

Sandoz, M.D., M.R. Rogers, and C.L. Newcombe. 1944.
Fungus infection of eggs of the blue crab, Callinectes
sapidus Rathbun. Science 99: 124-125.

Sawyer, R.T., A.R. Lawler, and R.M. Overstreet. 1975.
Marine leeches of the eastern United States and the Gulf
of Mexico with a key to the species. J. Nat. Hist. 9: 633-
667.

Sawyer, T.K. 1969. Preliminary study on the epizootiology
and host-parasite relationship of Paramoeba sp. in the
blue crab, Callinectes sapidus. Proc. Nat. Shellfish.
Assoc. 59: 60-64.

Sawyer, T.K., and S.A. MacLean. 1978. Some protozoan
diseases of decapod crustaceans. Mar, Fish. Rev. 40(10):
32-35. °

Sindermann, C.J. 1989. The shell disease syndrome in
marine crustaceans. NOAA [Nat. Ocean. Atmos. Admin.]
Tech. Mem. NMFS [Nat. Mar. Fish. Serv.]-F/NEC-64.

Sindermann, C.J. 1990. Principal diseases of marine fish
and shellfish. Academic Press, New York. 1037 pp.


Table 2. Continued

- Page 7

Virus Name Histopathology and Effect Gross Signs Geographic Location Reference
and Type Tissues Infected on Host of Disease and Prevalence
Enveloped Infects Effect No reported Chesapeake Johnson
helical cytoplasm of unreported; gfoss signs and Chinco- & Farley
(EHV); RNA. hemocytes and always teague Bays; 1980;
virus; hemopoietic associated east coast Johnson
paramyxo- tissue with other of Florida; 1983, 1984,
like virus viruses which prevalence 1986
synergisti- low
cally cause
pathology
Rhabdo- Associated “Infected” No reported Found only Yudin &
like B extra- glands normal gfoss signs in 1 of 60 Clark 1979;
(RhVB); cellularly and crabs (3%) confined Johnson
RNA virus; with basal showed no erabs'‘from 1983
formerly lamina of sign of Galveston,
labeled mandibular abnormal Texas
EGV-I gland behavior,
very similar
to EHV, prob-
ably same
Chesapeake Cytoplasm of Causes Abnormal Juvenile Johnson
Bay (CBV); epithelium destruction. behavior crabs from 1978a,b
fatal; RNA of bladder, of epithelium. and blindness Tangier Sound, 1983, 1984,
virus; epidermis, of bladder, probably 1986
picorna- gill, gut, and gill, and other areas
like virus nerve cells neuro- also
filled with secretory
viral material; cells of
infections CNS; may
are usually produce
focal blindness
FUNGAL DISEASES

heterogenous group of coliforms including Escherichia coli
(Colwell et al. 1975; Sizemore et al. 1975; Welsh and
Sizemore 1985)'.

Bacterial infections appear to be acquired during cap-
ture and transport, suggesting that potentially pathogenic
bacteria in the water or on the exoskeleton may be intro-
duced into tissues by wounding or other means under
stressful conditions. Physical factors contributing to stress
include the process of capture and handling, crowding in
traps and/or holding tanks, transport out of water, exposure
to elevated temperatures, and wounding by fellow captives
(Johnson 1976c).

Crabs may often contain low bacterial levels in their
blood due to exposure to bacteria during ecdysis, as well as
their day-to-day contact with ubiquitous bacteria in the
marine environment; but, their defense response would
probably remove these bacteria, sometimes leaving histo-
logic signs of the previous infections in the tissues (Messick
and Kennedy 1990).

In the early 1940s, there were marked annual fluctua-
tions in commercial catches of blue crabs noted in the
Chesapeake Bay. In the same period, a parasitic fungus was
discovered on egg masses of many blue crabs taken from
natural spawning grounds in the lower Chesapeake Bay
(Sandoz et al. 1944; Rogers-Talbert 1948). Samples from
infected animals were examined and a new phycomycetous
fungus species, Lagenidium callinectes Couch, was de-
scribed (Couch 1942). It was believed this fungus parasite
could limit the production of crab larvae and result in yearly
fluctuations in crab populations. This same species of
fungus was later found infecting crabs from coastal North
Carolina (Bland and Amerson 1973, 1974).

Infection takes place when fungal zoospores germinate
oncrab egg surfaces, sending in a germ tube which develops
into branched, septate mycelium. Lagenidium callinectes
zoospores are apparently incapable of germinating without

* Vibrio cholerae and other vibrios, including V. parahaemolyticus and V. vulnificus, can cause human disease when humans
ingest improperly cooked (Blake et al. 1980a,b) or handled crabs (Anonymous 1972).


Page 10

Figure 3. Blue crab from the Pamlico River, North Carolina, with extensive carapace lesions (arrows). (Photograph courtesy of Dr. D.

Engel.)

microsporidan was first reported and classified as a species
of Nosema (Sprague 1950, 1965), but was later placed in the
genus Ameson (Sprague 1977). The disease-inducing spores
are ovoid when live and measure 2.2 pm by 1.7 ym (Sprague
1965). Heavily infected crabs are called “sick crabs,” since
they are sluggish and have an abnormal appearance; they are
often seen in shallow water and, in heavy infections, can be
identified by an opaque white-to-grey ventral sternum
(Sprague 1970; Overstreet 1977). Infected animals die
rapidly if stressed (Overstreet and Whatley 1975). This
parasite is considered highly host specific since only
Callinectes sapidus is infected (Overstreet 1978).

Under the right conditions, spores infect host tissue by
everting a long internal tube or polar filament. The single-
celled spore squeezes through this extended pliable filament
and invades the host's blood cells; the spore subsequently
undergoes vegetative growth and produces cells which in
turn produce strings of eight individual spores along muscle
tissue. In heavy infections, enormous numbers of these
resistant spores lyse and destroy adjacent muscle tissue and
teplace the normal musculature (Overstreet 1978) (Figure
5). Ameson michaelis may be more pathogenic than other

microsporidans because it lacks pansporoblast membranes,
allowing it direct contact with crab tissue. These mem-
branes normally prevent harmful substances such as en-
zymes from being released to solubilize host tissue which is
consequently used for nutrition by the parasite (Couch
1983). This parasite is potentially very pathogenic, espe-
cially when considering the loss of economic potential of
reared crabs for thesoft-shell industry. Some measures have
been taken to find chemical deterrents to help reduce the
spread of the parasite under controlled conditions (Overstreet
and Whatley 1975).

Two other microsporida, A. sapidus and Pleistophora
cargoi, can also be found parasitizing blue crabs. Ameson
sapidus can induce extensive destruction of muscle tissue,
leading to white and opaque muscles. Spores are oval and
larger than A. michaelis at 3.5 um by 2.13 ym (Sprague
1970; Couch and Martin 1982; Sparks 1985). Another
species of microsporidan, P. cargoi, is characterized by its
development into 16 spores; when fresh, these are ellipsoi-
dal and measure 5.1 sm by 3.3 pm. These spores have been
found in skeletal (Figure 5) and cardiac muscle of blue crabs;
little is known of the effect of P. cargoi on blue crab