Journal of Invertebrate Pathology 106 (2011) 79–91
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Journal of Invertebrate Pathology
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Diseases of spiny lobsters: A review
J.D. Shields ⇑
Department of Environmental and Aquatic Animal Health, Virginia Institute of Marine Science, The College of William & Mary, Gloucester Point, VA 23062, USA
a r t i c l e
i n f o
Keywords:
Panulirus argus virus 1
Palinurid
Jasus
Panulirus
Symbiont
Pathogen
Fishing
a b s t r a c t
Spiny lobsters have few reported pathogens, parasites and symbionts. However, they do have a diverse
fauna comprised of a pathogenic virus, several bacteria, protozoans, helminths and even symbiotic crustaceans. A few idiopathic syndromes have also been reported, but these appear correlated with lobsters
held in poor conditions. Fungal and bacterial pathogens present significant threats for rearing spiny lobsters in aquaculture settings, but only one pathogen, Panulirus argus virus 1, is thought to have damaged a
fishery for a spiny lobster. No doubt others will emerge as lobsters are brought into aquaculture setting
and as fishing pressure intensifies with stocks become more susceptible to anthropogenic stressors.
Ó 2010 Elsevier Inc. All rights reserved.
Contents
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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Panulirus argus virus 1 (PaV1) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
White spot syndrome virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Gaffkemia – Aerococcus viridans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Shell disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Vibriosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Fouling bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Water molds and Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Oomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Fusarium solani . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Peritrich ciliates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Microsporidia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Helminths . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Digenetic trematode infections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Cestoda . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Nemertea – Carcinomertes spp. and Pseudocarcinonemertes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Miscellaneous metazoan symbionts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Nicothoidae – parasitic copepods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Amphipods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Fouling organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Disease syndromes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
21.1.
Turgid lobster syndrome. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
21.2.
Pink lobster syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
21.3.
Mass mortalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
22. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
⇑ Fax: +1 804 684 7186.
E-mail address: [email protected]
0022-2011/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved.
doi:10.1016/j.jip.2010.09.015
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J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
1. Introduction
Spiny lobsters in the genera Jasus, Panulirus, and Palinurus support worldwide fisheries with annual landings of over 70,000 m
(Booth, 2006; Phillips and Melville-Smith, 2006). There are surprisingly few diseases reported from these lobsters given the size and
value of their fisheries. However, recent attempts to culture lobsters have failed because of mortalities associated with potential
agents, most notably bacterial infections (Kittaka, 1997; Bourne
et al., 2004, 2006). That, plus the emergence of a pathogenic virus
in the Caribbean spiny lobster (Shields and Behringer, 2004), highlight the potential damage that can be caused by unrecognized
pathogens. I review the microbial diseases, parasites, and symbionts of the spiny lobsters, Panulirus spp., Palinurus spp., and Jasus
spp. For earlier reviews of the diseases of spiny lobsters see Evans
and Brock (1994) and Evans et al. (2000). For a broader perspective
on the diseases of clawed and spiny lobsters, see Shields et al.,
(2006). Several overviews and syntheses of crustacean diseases
are also available (Couch, 1983; Johnson, 1983; Overstreet, 1983;
Brock and Lightner, 1990; Sindermann, 1990; Meyers, 1990;
Shields and Overstreet, 2007), as well as those presented in this issue of the Journal of Invertebrate Pathology.
2. Panulirus argus virus 1 (PaV1)
PaV1 is the first naturally occurring pathogenic virus found
infecting a lobster. As its name implies, it infects P. argus. The virus
has a tropism for mesodermal cells, namely certain hemocytes
(hyalinocytes and semi-granulocytes), soft connective tissue cells,
some hematopoietic tissues, and fixed phagocytes (Shields and
Behringer, 2004; Li et al., 2008). PaV1 is currently an unclassified
virus, but it shares several morphological features with the Herpesviridae and the Iridoviridae (Shields and Behringer, 2004). For
example, it is an unenveloped, icosahedral, DNA virus with a nucleocapsid approximately 182 nm in size that develops within the nuclei of host cells. The Herpesviridae are icosahedral, but they are
enveloped and have a conserved size of approximately 125 lm.
The Iridoviridae are icosahedral and unenveloped, but they are
usually formed within the cytoplasm, whereas PaV1 is formed
within the nucleus.
Lobsters heavily infected with PaV1 are often lethargic or slow.
Their hemolymph does not clot and is milky in color, from cellular
debris and exudates (Shields and Behringer, 2004). PaV1 initially
infects the fixed phagocytes in the hepatopancreas (Li et al.,
2008). These cells line the hepatic arterioles and, as implied by
their name, they phagocytize non-self particles, including virions
of PaV1. The infected fixed phagocytes lyse, and the virus spreads
to spongy connective tissues as well as hemocytes. In late stages
of the infection, virtually all of the spongy connective tissue cells
become infected, with some apparently proliferating around the
hepatopancreas (Fig. 1). The tubules of the hepatopancreas atrophy
in late stages of infection (Li et al., 2008). Reserve inclusion cells in
the hepatopancreas and other tissues lose their glycogen reserves
and the host probably dies from metabolic wasting. Few lobsters
have concurrent bacterial infections (Shields and Behringer, 2004,
Shields, unpubl. data), which is surprising considering that the primary defensive cells of the host, the hyalinocytes and semigranulocytes, are targets for the virus.
Until recently, histology has been the primary tool for diagnosing PaV1 infections in P. argus. Pathological alterations to the fixed
phagocytes and spongy connective tissues, with Cowdry-like inclusions and enlarged nuclei, are the primary pathologies used in
diagnosis. Histopathological changes can be observed in the tissues
within 10–15 days after infection. Using a 110-bp piece of the viral
genome, a DNA probe and dot-blot assay were developed for use in
diagnostics (Li et al., 2006). A fluorescent-in situ-hybridization
(FISH) technique was used in conjunction with the probe to visualize infected tissues. The probe was specific to the virus and sensitive to 10 pg. Infected cells were visualized in many tissues. A
primer set for the polymerase chain reaction (PCR) has also been
developed for diagnosis (Montgomery-Fullerton et al., 2007). The
primer set amplified a 499-bp piece of viral DNA. It was specific
for the virus and sensitive to 1.2 fg. The primer set showed a high
acuity with histological findings. A primary cell-culture method
has been used to quantify the virus in infected hemolymph as a
TCID50 (Li and Shields, 2007). For cultures, naïve hemocytes, particularly hyalinocytes and semi-granulocytes, were separated using
density centrifugation, then cultured in modified Leibovitz medium, prior to exposure to the virus. Virally infected cells were demonstrated in cultures using an in situ hybridization technique (Li
and Shields, 2007). Uninfected cell cultures retained a high viability through 15 or more days.
PaV1 has been shown to induce a remarkable behavior in
healthy, uninfected lobsters. Such lobsters have the ability to detect and avoid diseased lobsters (Behringer et al., 2006). This is
the first example of avoidance of diseased individuals by healthy
animals other than in humans. Moreover, the avoidance behavior
occurs around the time infected lobsters become infectious! This
has important consequences for the denning behavior of P. argus,
because the animals are normally gregarious and the ability to
detect and avoid disease may serve to limit the spread of the
disease in the host population (Behringer et al., 2006). The nature of the avoidance is likely chemical detection as spiny lobsters have an exquisite sense of smell. PaV1 has been
transmitted to uninfected lobsters via inoculation, ingestion, contact and through seawater (Butler et al., 2008). Contact and
water-borne transmission are apparently the primary modes of
transmission in nature. Lobsters inoculated with infected hemolymph can develop acute infections and die within 30–90 days,
but some take much longer to show signs of infection. Lobsters
infected more naturally, via contact or water-borne transmission,
die within 200 days, with mortality highly correlated with smaller sized animals. Infectivity declines significantly with lobster
size. In lobsters exposed via contact, 63% of animals <25 mm carapace length (CL) became infected, as opposed to 33% of those
30–40 mm CL, and 11% of the large juveniles (40–50 mm CL),
respectively (Butler et al., 2008). Transmission coefficients used
in mass-balance models were calculated for the different modes
of infection. Contact transmission among the smallest juveniles
had the highest transmission coefficient; that for water-borne
transmission was about five times lower. Thus, the virus appears
to be transmitted by direct contact among small juveniles and
over short distances through the water column.
In short-term mark-recapture studies, overtly diseased lobsters had a lower survival rate than animals without disease
(Behringer et al., 2008). In the laboratory, lightly infected lobsters showed no significant differences in movement compared
to uninfected animals, but as the disease progressed, infected
lobsters were much less active than healthy animals. The physiological state of intermolt lobsters from the field was assessed
using hemolymph serum proteins as a general indicator of
health. Overtly diseased lobsters had significantly lower hemolymph protein concentrations than presumably uninfected lobsters, by as much as 4–5 mg/dl protein levels (Behringer et al.,
2008). Interestingly, starvation did not increase the susceptibility
of animals to viral infection. Given that lightly infected animals
were still highly active, they were thought to facilitate dispersal
of the virus to new habitats.
The discovery of PaV1 coincided with a decline in the landings
of P. argus from the Florida Keys fishery (Shields and Behringer,
2004). Whether the virus caused this decline or not remains to
J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
81
Fig. 1. PaV1 infections in lobsters. (A) Heart of a juvenile lobster infected with PaV1. Note the hypertrophied, infected connective tissue cells lining the arterioles (arrows).
Bar = 100 lm. (B) Hepatopancreas from a heavily infected juvenile lobster. The hemal sinus (star) shows heavy infiltration by infected hemocytes and spongy connective
tissue cells. Bar = 50 lm. (C) Fixed phagocytes (arrow) around an arteriole (lumen marked with star) in the hepatopancreas of an uninfected lobster. Bar = 50 lm. (D) Fixed
phagocytes around an arteriole (lumen marked with star) in the hepatopancreas of an infected lobster. Note the hypertrophied cells with emarginated chromatin in infected
cells (arrows). Bar = 50 lm.
be determined, but there is no doubt that the agent is widespread
and pathogenic to lobsters in their early benthic juvenile stage.
Lobsters infected with PaV1 have been found in several places
around the Caribbean Sea, including the Florida Keys, US Virgin Islands, Mexico, and Belize (Butler et al., 2008; Huchin-Mian et al.,
2008, 2009). PaV1 is widespread in the Florida Keys and Florida
Bay, particularly in the shallow juvenile nurseries. In field studies,
the mean prevalence was 7%, with local ‘‘hot spots” of up to 30%
(Shields and Behringer, 2004). The size predilection for the virus
stands out in the field studies because prevalence was 16% among
the smallest juveniles (<20 mm CL), whereas it was 5% among the
largest juveniles (>40 mm CL); in field surveys of adults, <1% of
adults were visibly infected. Off of Puerto Morelos and the Chinchorro Bank, Mexico, prevalence was as high as 10.9% in juvenile
lobsters, but there was no apparent change in host density with increased prevalence of disease. (Lozano-Álvarez et al., 2008). The
differences in the nursery habitat between the Florida Keys and
the area around Puerto Morelos are large enough to warrant further attention to density issues and how habitat restrictions potentially alter the host-pathogen association.
PaV1 appears to be specific to P. argus (Butler et al., 2008). In
infection trials, three potential alternate hosts (the spotted lobster,
P. guttatus, the stone crab, Menippe mercenaria, and the channel
crab, Mithrax spinosissimus) were inoculated with hemolymph
from an infected Caribbean spiny lobster. After 80 days, none of
the three alternate hosts showed signs of disease using histology.
This is particularly interesting because the spotted lobster, P. guttatus, is a close congener with P. argus (Ptacek et al., 2001), and its
lack of observable disease suggests that PaV1 is highly specific to P.
argus.
The recent emergence of PaV1 and its potential association with
a decline in the spiny lobster fishery in the Florida Keys is cause for
some concern. Several viruses have severely damaged the shrimp
aquaculture and fishery industries (Flegel, 1997; Lightner and
Redman, 1998); therefore, programs should be developed to monitor for PaV1 in populations of the spiny lobster from around the
Caribbean Sea to establish baselines and to document its potential
effect on artisanal and regional fisheries. Infections have been
identified in two culture facilities in Florida (Shields, unpubl. data),
and we know that it occurs in several countries around the Caribbean Sea. Given that lobsters and lobster tails are shipped internationally (see Huchin-Mian et al., 2009), plus the fact that nascent
aquaculture ventures have been attempting to culture spiny lobsters from enzootic areas, lobsters should be screened for the virus
as well as other microbial infections to control for infections as
well as to prevent their potential spread to other areas (see
Table 1).
3. White spot syndrome virus
Spiny lobsters are not naturally infected with the white spot
syndrome virus (WSSV), but with epidemics of WSSV occurring in
native shrimp stocks, it seems inevitable that it will spread into
new host species. Indeed, at least 42 crustaceans, including three
species of Panulirus, can serve as experimental reservoir hosts for
WSSV (see Supamattaya et al., 1998; Rajendran et al., 1999; Musthaq et al., 2006). In experimental infections, a DNA probe was used
to detect WSSV in the tissues of P. versicolour and P. penicillatus
(Chang et al., 1998). Lobsters survived for at least 70 days, with
the virus present in their tissues, but they were not examined histologically for signs of disease. That is, the lobsters were infected
but were not obviously diseased. PCR has also been used with a
virus-specific primer set to detect WSSV in several species of lobster
(P. versicolour, P. penicillatus, P. ornatus and P. longipes) fed infected
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J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
Table 1
Parasites and symbionts naturally occurring in spiny lobsters, including the tissues in which they are found.
Parasite
Host
Tissue
Key reference
Virus
PaV1
P. argus
Connective tissues,
hemocytes
Shields and Behringer (2004)
H. americanus, H. gammarus, P.
argus
P. argus, P. ornatus
J. edwardsi
Panulirus homarus
P. argus, P. laevicauda
Several genera
Systemic
Snieszko and Taylor (1947), Rabin (1965),
Stewart et al. (1966), Stewart et al. (1980), Bobes et al. (1988)
Porter et al. (2001), Bourne et al. (2006), Payne et al. (2007)
Diggles et al. (2000)
Hameed (1994), Abraham et al. (1996)
Silva dos Fernandes Vieira et al. (1987)
Nilson et al. (1975), Steenbergen and Schapiro (1976), Harper and Talbot
(1984)
P.
P.
P.
P.
Larva
Pueruli, small juveniles
Cuticle
Gills
Bacteria
Aerococcus viridans
Several genera
Vibrio harveyi
V. alginolyticus
Vibrio spp.
Leucothrix mucor
Fungia
Atkinsiella panulirata
Haliphthoros sp.
Fusarium solani
Didymaria palinuri
Ramularia branchiales
Protozoans
Microsporidia
Panulirus spp.
P. argus
Muscle? Muscle
Peritrich ciliates
Jasus edwardsii
Helminths
Turbellaria
Thulakiotrema genitale
Cymatocarpus solearis
Tetraphyllid cestode
Carcinonemertes
wickhami
Carcinonemertes
australiensis
Carcinonemertes sp.
Crustaceans
Choniomyzon panuliri
Paramphiascopsis sp.
Parapleustes commensalis
Gitanopsis iseebi
Octolasmis californiana
Octolasmis spp.
Trilasmis fissum
Balanomorphs
Paralepas spp.
a
japonicus
argus,J. verreauxi
cygnus
elephas
External
Systemic, phyllosoma
Systemic
Systemic
Carapace, eggs
Kitancharoen and Hatai (1995)
Fisher et al. (1975), Diggles (2001)
McAleer and Baxter (1983)
Sordi (1958)
Bach and Beardsley (1976), Dennis and Munday (1994), Kiryu et al.
(2009)
Kittaka (1997), Handlinger et al. (1999), Bourne et al. (2004)
Panulirus spp.
P. cygnus
P. argus
Panulirus spp.
P. interruptus
On mouthparts
Gonad
Abdominal muscle
Foregut
Eggs
Shields (unpubl. data)
Deblock et al. (1991)
Gómez del Prado-Rosas et al. (2003)
Shields (unpubl. data)
Shields and Kuris (1990)
P. cygnus
Eggs
Campbell et al., (1989)
J. edwardsii
Eggs
Frusher and Shields (unpubl. data)
Panulirus spp.
J. edwardsii, J. verreauxi
P. interruptus
P. japonicus
P. interruptus
P. polyphagus
P. japonicus, P. penicillatus
P. argus
Spiny lobsters
Eggs
Gills
Eggs
Branchial chamber
Gills, carapace
Gills, carapace
Mouthparts
Carapace
Carapace
Pillai (1962)
Booth (pers. comm.)
Shoemaker (1952)
Yamato (1993)
Newman (1960)
Jeffries et al. (1982)
Bowers (1968)
Eldred (1962)
Jones (2003)
The Phylum Fungi no longer includes the fungus-like Phycometes and other classical forms that are now placed in a separate Kingdom, the Chromista.
shrimp (Wang et al., 1998). The virus was detectable in all of the
lobsters, but none developed disease. Lobsters (Panulirus homarus)
inoculated with WSSV-infected shrimp tissues died, presumably
from the infection (Rajendran et al., 1999; Musthaq et al., 2006),
but the question is unresolved because animals fed infected tissues
did not develop infections (Musthaq et al., 2006), and controls consisting of uninfected tissue homogenates were not used. However,
in the latter study, all of the lobsters injected with the virus died
over 160 days.
It is surprising that no other viruses are known from lobsters.
Shrimp have at least 20 and the blue crab, Callinectes sapidus, has
at least eight pathogenic viruses (Lightner and Redman, 1998;
Shields and Overstreet, 2007). Several shrimp viruses have spread
as pandemics through the Americas and Asia with catastrophic results for shrimp aquaculture (Flegel, 1997; Lightner and Redman,
1998). Perhaps more viruses are known from shrimp because they
are intensively cultured; hence, there are more opportunities for
outbreaks under crowded conditions. Nonetheless, clawed and
spiny lobsters support large fishing industries in which diseased
animals would be observed or recognized. Perhaps the paucity of
viruses in lobsters is simply a reflection of the fact they are only
now being investigated for intensive aquaculture, and that most
of the focus on lobsters has been on their fisheries and not on
the more susceptible larval and juvenile stages.
4. Gaffkemia – Aerococcus viridans
Gaffkemia, or red-tail disease, is a serious disease of clawed lobsters, primarily Homarus americanus, but also H. gammarus. Outbreaks usually occur in holding facilities; it is not common in
natural populations of lobsters (Stewart et al., 1966; Keith et al.,
1992; Lavallée et al., 2001). Aerococcus viridans, a Gram-positive
bacterium, is the causative agent of gaffkemia, and it is a common
soil microbe with many variants. The variant that is pathogenic to
lobsters is known as A. viridans var. homari. The pathogen can be
spread via shipment and holding of diseased clawed lobsters;
therefore, further risk assessment should be initiated in areas
where there could be likely introductions of contaminated water
or discarded lobsters.
Aerococcus viridans reportedly occurs naturally in P. argus from
Cuba. Bobes et al. (1988) isolated and cultured A. viridans from six
J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
animals that presented signs of disease. The animals had reddish
discolored exoskeletons, lethargy, pink hemolymph and coagulopathies. The bacterium was susceptible to 10 of 13 antibiotics in culture (Bobes et al., 1988). While Koch’s postulates were not
attempted, this finding has further implications for the transportation of spiny lobsters. Aerococcus viridans var. homari has been isolated from other crustaceans from Massachusetts (Rabin and
Hughes, 1968) and the Gulf of Mexico (Liuzzo et al., 1965). While
other crustaceans can be carry the bacterium, clawed lobsters are
apparently the only hosts in which it is highly pathogenic (Cornick
and Stewart, 1975).
The route of transmission of Aerococcus viridans is through damage to the host exoskeleton, not through intact cuticle or ingestion
(Stewart et al., 1969). The bacterium is not part of the natural
microflora of lobsters (Stewart, 1975). It has been isolated from
lobster holding facilities; and the isolation of the agent usually
indicates the presence of diseased lobsters (Goggins and Hurst,
1960; Kellog et al., 1974; Stewart, 1980). Unfortunately, it can remain viable in the benthos for long periods where it can act as a
reservoir of infection (Stewart, 1984). Aerococcus viridans has also
been documented in muds adjacent to a holding facility for clawed
lobsters in Anaheim Bay, California, and was likely transported
there through diseased clawed lobsters (Kellog et al., 1974).
Gaffkemia can be experimentally induced in P. interruptus, but
the lethal dose is higher than that for H. americanus (Schapiro
et al., 1974; Steenbergen and Schapiro, 1974). That is, spiny lobsters show a high degree of resistance to experimental infections
(Schapiro et al., 1974; Schapiro and Steenbergen, 1974). Because
most studies of gaffkemia have been carried out in the clawed lobsters, there will be no further treatment of it here. For reviews of
gaffkemia, see Stewart (1980) and Brock and Lightner (1990).
5. Shell disease
Shell disease, or shell disease syndrome, is a progressive chitinolysis and necrosis (bioerosion) of the exoskeleton of crustaceans
(Rosen, 1970). Virtually all crustaceans are susceptible to shell disease, but it is most notable in large decapods. The syndrome first
appears as small pits or erosions in the cuticle and progresses to
large eroded lesions (Rosen, 1970; Getchell, 1989; Noga et al.,
1994). The bacterial form, which is referred to as classical, or enzootic, shell disease, was first described from the American lobster
(Hess, 1937). Recently, an epizootic form of the syndrome has
emerged in American lobsters from Rhode Island (see Castro
et al. (2006), for review). This new form of the disease has not been
reported from palinurid lobsters. For comprehensive reviews of
shell disease in crabs and lobsters see Shields et al. (2006) and
Shields and Overstreet (2007).
Classical shell disease is caused by a number of chitinoclasitic,
Gram-negative bacteria. Vibrios (Vibrio vulnificus, V. parahaemolyticus, V. alginolyticus) are the most common bacteria associated
with shell disease, but other species have also been isolated from
the lesions, including Shewanella spp. and Aeromonas hydrophila
(Geddes et al., 2003; Porter et al., 2001; Reuter et al., 1999). Both
V. alginolyticus and a V. harveyi-like bacterium were isolated from
the hemolymph and lesions occurring on P. homarus reared in
the laboratory (Abraham et al., 1996). Using the 16s and 23s rDNA
intergene regions, Porter et al., (2001) examined the bacterial fauna of lobsters with and without classical shell disease. DNA fingerprinting found no specific etiological agent associated with the
lesions, but bacteria in several genera were identified, including
Vibrio, Pseudoaltermomonas, Pseudomonas and Shewanella. The native bacterial flora was thought to be responsible for shell disease
lesions in spiny lobsters (Porter et al., 2001), and many of these are
indeed chitinoclastic.
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Classical shell disease requires a portal of entry through the epicuticle, and mechanical abrasion is the usual mode of transmission.
Shell disease starts as small pits or ‘‘burn” marks in the cuticle. The
lesions occur on the sternum or legs of American lobsters (Rosen,
1967; Johnson, 1983), but they are more commonly found on the
uropods of spiny lobsters (Porter et al., 2001; Geddes et al.,
2003). As the pits coalesce, they develop into large, brown or black,
irregularly shaped lesions (Rosen, 1967). The lesions can penetrate
through the cuticle into underlying tissues of the lobster, but they
rarely do so; instead they develop laterally with further erosion of
the cuticle (Overstreet, 1978; Johnson, 1983). The exposed cuticle
is weakened and brittle, and discolored from the deposition of melanin (Johnson, 1983; Smolowitz et al., 1992). Melanization and cellular infiltration occur as part of the host response to shell disease
in American lobsters (Smolowitz et al., 1992, Smolowitz et al.,
2005). Pseudomembrane formation can occur in particularly severe cases of shell disease, particularly in epizootic shell disease
(Smolowitz et al., 2005). It is no doubt the same process in spiny
lobsters. Individuals usually overcome the disease by molting
(Rosen, 1967; Castro and Angell, 2000), but animals that molt less
frequently can presumably succumb to infection.
In spiny lobsters, P. argus, Panulirus cygnus and Jasus edwardsii,
another form of shell disease occurs. It is known as ‘‘tail fan necrosis”, an aptly named syndrome because the uropod and telson appear melanized and partly or fully eroded away (Porter et al., 2001;
Geddes et al., 2003). In clawed lobsters, shell disease lesions are
more frequently seen on the ventral side of the claws, tail and carapace (Estrella, 1991). Shell disease is normally found at low (<1%)
levels in healthy wild populations of crustaceans. Iversen and
Beardsley (1976) examined 9000 P. guttatus, and found only two
infected individuals. They cite several unpublished reports indicating a very low prevalence of shell disease in P. argus.
Classical shell disease is often associated with pollution
(Gopalan and Young, 1975; Young and Pearce, 1975; Couch,
1983; Morado et al., 1988; Gemperline et al., 1992; Weinstein
et al., 1992; Ziskowski et al., 1996), and spiny lobsters require relatively pristine water quality parameters; therefore, shell disease
is not usually found at a high prevalence in palinurid lobsters.
However, clawed lobsters held in crowded pounds with poor water
quality can develop fatal shell disease (Taylor, 1948, in Sindermann, 1989). Therefore, in culture systems, efforts should be directed at maintaining good water quality and reducing crowding
effects. Lobsters with moderate to severe forms of shell disease
are not esthetically pleasing and are not used in the ‘‘live food”
trade. Indeed, epizootic shell disease has had an economic impact
on the clawed lobster fishery off Rhode Island, because the cuticle
of affected animals is severely compromised making the animals
unsuitable for live trade (Castro et al., 2006).
Antibiotics have been used to treat shrimp and lobsters with
shell disease, but none are currently registered for use in crustacean aquaculture in the USA. Some such as furanace may be used
in other countries. Juvenile lobsters have been successfully treated
with Malachite green (Fisher et al., 1976a,b, 1978). Shrimp have
been treated with Malachite green, formalin, or antibiotic baths
(penicillin–streptomycin, furanace, erythromycin, oxolinic acid)
(Tareen, 1982; Brock, 1983; El-Gamal et al., 1986). Contaminated
aquaria should be disinfected with bleach solutions, and it is a
good practice to do this prior to any laboratory culture. Animals
with severe lesions should be destroyed to prevent further spread
in impoundments.
6. Vibriosis
Vibriosis is an infection by any species of Vibrio. Vibrios are
ubiquitous in the marine environment, and several species cause
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serious diseases to invertebrates, fishes and even humans. Lobsters, in turn, can be exposed to several pathogenic species of
Vibrio. Vibrio alginolyticus, V. harveyi, V. parahaemolyticus and
V. anguillarum are facultative, opportunistic pathogens that have
been implicated in disease in clawed and spiny lobsters (Bowser
et al., 1981; Brinkley et al., 1976; Jawahar et al., 1996). Vibrios
can also contaminate seafood products including lobsters (Wong
et al., 1999); and there are many cases of human food poisoning
from improper handling and preparation of marine crustaceans.
Lobsters should be properly cooked prior to eating or freezing,
and all surfaces used in the preparation of lobsters should be
cleaned and disinfected afterwards.
Vibrio infections can cause serious disease in spiny lobsters, but
few outbreaks have been reported. This is probably because spiny
lobsters are usually kept in short-term culture, but not long-term
culture where there would be more opportunity for disease under
crowded conditions. However, outbreaks are known to occur in larval cultures. Indeed, vibriosis may be the single most important
impediment to successful larval culture (Bourne et al., 2004). Cultures of phyllosoma larvae of Sagmariasus(=Jasus) verreauxi experienced significant mortalities due to a luminescent strain of V.
harveyi in New Zealand (Diggles et al., 2000). Infected larvae were
opaque and had small red spots throughout the body. The hepatopancreas was atrophied and the hepatopancreatic tubules had
numerous bacterial plaques. Mortalities occurred over 4 weeks.
The bacterium is an opportunistic pathogen and attacked injured
larvae more than uninjured ones. Two drugs, sulfadimidine and trimethoprim, were found to enhance survival the lobster larvae
(Diggles et al., 2000).
The dynamics of bacterial flora have been studied in cultures of
larval spiny lobsters. Larval cultures of Panulirus ornatus had a high
microbial diversity, with representatives from many bacterial taxa
(Bourne et al., 2006; Payne et al., 2007). Denaturing gradient gel
electrophoresis (DGGE) was used to study diversity in larval cultures (Bourne et al., 2004; Payne et al., 2007) as well as biofilms
in culture aquaria (Bourne et al., 2006). There were few changes
in the bacterial flora in relation to mortality. In fact, bacterial densities decreased significantly over time and had no relationship
with mortality (Bourne et al., 2004). However, larval mortality
was high, reaching 100% in 13–24 days. Vibrios were visualized
and enumerated using a fluorescence in situ hybridization (FISH)
technique (Webster et al., 2006). Vibrios increased both on and
within phyllosoma larvae starting about seven to ten days after
hatching, and reached high levels 18 days after hatching. Two isolates of V. harveyi were used in a challenge study and one strain
caused more mortality than the controls (Bourne et al., 2006). Several species of Vibrio and other bacteria have been identified from
cultures of the phyllosoma of J. edwardsii (Handlinger et al., 1999).
An isolate of V. harveyi was obtained from the diseased gut of an
infected larva. More research is needed in this area because it is
not clear how larval nutrition, host defenses, temperature, culture
stress and disease agents interact in culture systems.
Vibrio alginolyticus, V. parahemolyticus and a V. anguillarumlike species were isolated from P. argus and P. laevicauda from
commercial landings in Brazil. Prevalence varied from 2.8% in V.
anguillarum to as high as 45.7% in P. alginolyticus (Silva dos Fernandes Vieira et al., 1987). Unfortunately, the disease status of the lobsters was not reported. Vibrio alginolyticus and a V. harveyi-like
bacterium were isolated from shell disease lesions and hemolymph
of juvenile and adult P. homarus from India (Hameed, 1994). In
inoculation trials, the Vibrios became systemic and induced shell
disease-like lesions in larvae of Penaeus indicus and P. homarus
(Hameed, 1994). Several tissues showed histopathological alterations. In a separate study by Abraham et al., (1996), mortality
reached 100% in P. homarus injected with 108 bacterial cells, but
there was no mortality in lobsters inoculated with 107 bacterial
cells. I have obtained similar results in inoculation trials with Vibrios in J. edwardsi (Shields, unpubl. data).
Vibrios are frequently isolated from the hemolymph of crustaceans; and the fact is that crustacean hemolymph is not necessarily
sterile. In a debate in the 1970s, Bang (1970), Colwell et al., (1975)
and Tubiash et al., (1975) found a high prevalence of bacteria, particularly Vibrios in the hemolymph of apparently healthy blue
crabs. These were thought to have been acquired through abrasions or other wounds in the cuticle that occurred because of the
rough handling of crabs en route to markets (Johnson, 1976). Indeed, crabs are handled roughly during capture and transport to
processing plants (Shields, pers. obs.). However, in a definitive
study, species of Vibrio were shown to occur in the hemolymph
and various other tissues of freshly caught, unstressed crabs (Davis
and Sizemore, 1982). Nonetheless, we rarely isolate bacteria from
the hemolymph of spiny lobsters. Thus, while it is possible that
spiny lobsters can carry background levels of septicemia, there
are few reports of such in adults, nor have I observed many bacterial isolates in routine health examinations of P. argus and J. edwardsii (Shields, unpubl. data).
7. Fouling bacteria
Filamentous bacteria are ubiquitous on the eggs of marine crustaceans, and spiny lobsters are no exception (Johnson et al., 1971;
Bland and Brock, 1973). Leucothrix mucor is a common filamentous
species that has been observed on or isolated from cultured larvae
of the American lobster (Nilson et al., 1975; Dale and Blom, 1988).
It and other fouling agents are common fouling organisms in larval
culture of spiny lobsters (e.g., Kittaka, 1997; Bourne et al., 2004,
2006; Payne et al., 2007). Leucothrix mucor is very common in seawater systems. It is a saprophyte of dead algae. I have observed it
on crustacean eggs and limbs, including amphipods, crabs and lobsters. Filamentous bacteria can be difficult to control because they
are so common and because they grow readily in the organically
enriched environments encountered in culture systems. Heavy
infections are thought to contribute to larval mortality. Control requires careful site selection and system design, good water quality
parameters and strict attention to nutrition. Outbreaks of L. mucor
have been controlled with antibiotics (Steenbergen and Schapiro,
1976; Fisher et al., 1976a,b; Sadusky and Bullis, 1994).
8. Water molds and Fungi
The ‘‘lower fungi” have been reclassified as protists in the Kingdom Chromista (Cavalier-Smith, 1993). The Oomycota include the
Oomycetes and Phycomycetes after Margulis et al., (1990). They
are now referred to as water molds. The true Fungi include Deuteromycetes, Ascomycetes and Basidomycetes and a few other classes
of ‘‘higher” fungi.
9. Oomycetes
A few oomycetes are known from spiny lobsters. These typically
infect the eggs or larvae, but a few infect adult lobsters (e.g.,
Alderman, 1973). Atkinsiella panulirata is an oomycete described
from phyllosoma of P. japonicus (Kitancharoen and Hatai, 1995).
Haliphthoros mildfordensis has been implicated in mortalities in
aquaculture facilities for postlarval spiny and clawed lobsters (Fisher et al., 1975, 1978; Nilson et al., 1975 Diggles, 2001). An outbreak of
Haliphthoros sp. caused mortalities in a grow-out facility in New Zealand (Diggles, 2001). The disease infected the pueruli and juveniles
of J. edwardsii, which exhibited morbidity, including lethargy and
loss of appetite. Only lobsters less than 30 mm CL became infected.
Perhaps this was due to larger animals having a thicker or more resil-
J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
ient cuticle. Antibiotic treatment using Malachite green and formalin was successful in preventing the spread of disease. The outbreak
was associated with poor water quality (Diggles, 2001).
Melanization of infected tissues is a common defense against
Haliphthoros and other mold and fungal infections in crustaceans.
Therefore, black or brown spots are commonly seen in necropsies
involving mold infections. The quinones and melanin, which are
activated by the prophenoloxidase pathway, cause these black spots.
Prophenoloxidase occurs naturally in the cuticle and its activated
enzyme, phenoloxidase, is activated through a well known cellular
process involving hemocytes. Melanin is highly toxic to many microbial pathogens (for review, see Jiravanichpaisal et al. (2006)).
Maintaining animals in conditions of excellent water quality is
the preferred method to control water mold outbreaks in cultures
of lobsters. Ozonated or dechlorinated seawater should be used in
cultures. Water changes to maintain low organic loads and low
ammonium levels is also critical. Furanace and other antimold
drugs, such as Malachite green, can be used to treat infected animals (Abrahams and Brown, 1977; Fisher et al., 1978), but these
drugs are not approved for use in aquaculture by the Food and
Drug Administration in the USA and in several other countries.
10. Fusarium solani
A deuteromycete fungus, F. solani, causes disease in cultured
crustaceans. This species and other unidentified species have
caused disease in shrimp (Lightner and Fontaine, 1975), spiny lobsters (McAleer and Baxter, 1983) and clawed lobsters (Alderman,
1981). Black lesions resembling shell disease occur on the cuticle
of infected hosts. An outbreak of F. solani was reported from P. cygnus off Western Australia (McAleer and Baxter, 1983). The lesions
occurred on the abdomen, uropods, telson and pereopods. The fungus was isolated from lobsters and from seawater samples in areas
with diseased lobsters. Environmental factors were thought to
have facilitated the outbreak, but they were not identified. Given
the location of the lesions around the uropods and telson, water
quality was likely an issue.
The pathology of Fusarium infections has been studied in
shrimp and clawed lobsters. The fungus penetrates through the
cuticle, which causes the formation of lesions (Lightner and Fontaine, 1975; Fisher et al., 1978). The lesions often become melanized through the phenoloxidase pathway. Lesions containing
cellular aggregates, necrotic tissues, and fungal hyphae can be observed in histology. The hyphae of Fusarium produce diagnostic micro- and macroconidia (Lightner and Fontaine, 1975).
Two other deuteromycetes, Didymaria palinuri and F. (Ramularia) brachialis were isolated from moribund P. elephas and H. gammarus from the Mediterranean Sea (Sordi, 1958), but no attempts
were made to infect healthy lobsters. It is not clear if the fungi
were primary pathogens or opportunists. Little data were reported
from these infected animals.
11. Peritrich ciliates
Peritrich and suctorian ciliates are often found as commensals
on the external surfaces and embryos of crustaceans. Heavy infestations often indicate poor water quality or the presence of a disease. They are not usually pathogenic but heavy infections can
impose a respiratory debt to the host (Schuwerack et al., 2001).
They can be quite abundant in larval cultures of spiny lobsters
(e.g., Handlinger et al., 1999; Bourne et al., 2004). Their presence
on crustacean embryos generally indicates that a female host is
not feeding well or is not preening or otherwise taking care of herself. Several studies have found peritrichs in the genera Vorticella
and Zoothamnium, suctorian ciliates Acineta and Ephelota, several
85
cyanobacteria and diatoms in the clutches of clawed lobsters
(Dannevig, 1928, 1937; Nilson et al., 1975; Harper and Talbot,
1984; Dale and Blom, 1988).
Vorticella sp., Navicula sp., L. mucor and other organisms fouled the
larvae of J. edwardsii (Kittaka, 1997). Fouled phyllosoma larvae died,
but their cause of death was not documented. Interestingly, other fouling agents, including Saprolegnia sp., hindered fouling by the ciliates.
Chilodonella-like protists and Epistylis spp. have also been reported as
fouling agents (Handlinger et al., 1999; Bourne et al., 2004). Wescodyne
and Malachite green have been used to control Vorticella infestations on
clawed lobsters (Boghen, 1982). Baths of formalin, oxytetracycline,
erythromycin and streptomycin were used to control fouling organisms on P. ornatus (Bourne et al., 2004). Weak formalin solutions have
been used to effect pathogen control in many culture situations, including ciliate infections in fishes.
12. Microsporidia
The Microsporidia is a phylum of small, intracellular and intranuclear parasites. The phylum is named after the small, highly
refractory spore produced as the transmissive stage. All members
of the phylum have an extrusible polar tube that is used to inject
the sporoplasm into a new host cell. The Microsporidia were once
considered a phylum within the Protozoa, but they are now considered to be closely related to the true Fungi (for review, see Lee et al.
(2008)). Some species, such as Nosema apis, can survive in their
cadavers for very long periods.
Microsporidia are very rare in lobsters. A single infection was
reported from P. argus from Florida (Bach and Beardsley, 1976),
but no details were given. More recently, Kiryu et al., (2009) found
two lobsters with Ameson-like microsporidiosis from Florida. The
microsporidians are aptly named as those in the lobster were
1.4 1.0 lm in size (Kiryu et al., 2009). In Australia, at least one
species of Ameson sp. is known to infect spiny lobsters, P. cygnus
and P. ornatus (Dennis and Munday, 1994; Owens and Glazebrook,
1988). In both systems, infections occur in the muscles. They cause
a distinctive liquefactive and coagulative necrosis of the muscles,
with heavy infections causing the muscle fibers to turn creamy
white in color. A similar condition in crabs and shrimp is known
as ‘‘cotton” disease. The flesh of these animals when cooked has
a consistency like that of cotton; and it is very unappetizing
(Shields, pers. obs.). Very little is known about infections in lobsters. In blue crabs, Ameson michaelis infections are transmitted
via cannibalism (Overstreet, 1978), but their rarity in lobsters suggests that other pathways are involved or that lobsters may be
accidental hosts. Microsporidia are known to damage prawn aquaculture (Flegel et al., 1992; Hudson et al., 2001) so there is a precedent for their potential infection of lobster populations. However,
in field surveys from the Florida Keys that involved several hundred animals, there was no obvious histological infection by any
microsporidian (Shields, unpubl. data). Given the live market for
lobsters and their shipment to international markets, the potential
transport of microspores in infected hosts should be investigated
as a risk factor, albeit many microsporidians show high host specificity, which could lower the risk to other hosts.
13. Helminths
Few platyhelminthes reportedly use lobsters as hosts. There are
no published reports of turbellarians on spiny lobsters, but I have
observed them on the mouthparts and gills of several species of
Panulirus, from the Great Barrier Reef (Shields, unpub. data). Turbellaria have been reported from the egg clutches of crabs (Kuris
et al., 1991), and it would not be surprising to find them in the
clutches of other crustaceans, including lobsters. There are a few
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reports of digenetic trematodes from palinurid lobsters, but there
are no published reports of cestodes from them other than my
anecdotal observations below.
14. Digenetic trematode infections
Lobsters serve as the second intermediate host to digenetic
trematodes. They bear the metacercaria, or the encysted stage.
The metacercarial cysts are usually quite small (<1 mm) and can
be detected using microscopy. As the name implies, the microphallid trematodes are known for their small phallus. Several species
use crustaceans as intermediate hosts. While the metacercariae
of most trematodes are difficult to identify to the species level,
the microphallids develop sexual characteristics as metacercariae
making it possible to identify and describe them. Thulakiotrema
genitale is a microphallid found encysted as metacercaraie in the
gonads of Panulirus cygnus (Deblock et al., 1991). The prevalence
in Western Australia was quite high, ranging from 47% to 87%.
The vertebrate definitive host is unknown, but it is likely to be a
fish. Cymatocarpus solearis(=C. undulatus) is a brachycoeliid trematode. Metacercariae infect the abdominal muscles of P. argus
(Gómez del Prado-Rosas et al., 2003). The cysts of C. solearis are
large enough to see by eye (1.5 mm in size). The parasite was relatively abundant in lobsters, with a prevalence of 35.8% and a
mean intensity of 26 cysts per host. The loggerhead turtle Caretta
caretta serves as the definitive host, where it lives in the intestines
(Linton, 1910; Caballero, 1959). Light infections of metacercariae
probably do not result in significant pathology, albeit in other systems, particularly those involving the nervous systems of their
hosts, they can cause alterations to host behaviors.
15. Cestoda
Lobsters may serve as an intermediate host for cestodes, bearing the metacestode, which is usually encysted in the connective
tissues. Metacestodes vary in size, some are microscopic whereas
others can be quite large. I have observed metacestodes of a tetraphyllidean cestode in the foreguts of Panulirus spp. and Scyllarides
sp. from the Great Barrier Reef (Shields, unpub. data). The tetraphyllidean cestodes use elasmobranchs as definite hosts. Tetraphyllidean metacestodes cannot be identified to species unless
fed to and recovered from an elasmobranch host. This can be difficult to do in the laboratory, but it is possible (see Sakanari and
Moser, 1989). There are no published records of metacestodes from
lobsters. This is intriguing because they no doubt serve as intermediate hosts for other parasites that use elasmobranchs, other fishes,
and turtles as definitive hosts.
16. Nemertea – Carcinomertes spp. and Pseudocarcinonemertes
Two species of Carcinonemertes are known from spiny lobsters.
They are egg predators and, at least on crabs, they can occur in epidemics where they damage reproduction on a population level
(Wickham, 1979, 1986; Hobbs and Botsford, 1989; Shields and
Kuris, 1988; Kuris et al., 1991). Carcinonemertes can be difficult to
detect on lobsters because they only infest the egg masses and only
during reproduction; that is, one must assess their presence when
lobsters carry eggs.
Panulirus interruptus from Southern California are infested by
Carcinonemertes wickhami (Shields and Kuris, 1990). It is a relatively large carcinonemertid worm, 30–50 mm in length. Panulirus
cygnus from Western Australia are infested by C. australiensis
(Campbell et al., 1989). It is a relatively small worm for the genus
at 7 mm in length. Jasus edwardsii from Tasmania is also infested by
a carcinonemertid worm, but it has yet to be identified (Frusher
and Shields, unpubl. data). It is a larger, more robust species than
C. australiensis.
Both C. wickhami and C. australiensis are monostyliferous hoplonemerteans; that is, they each bear a single stylet on the proboscis
armature. They are dioecious, and use internal fertilization for
reproduction. Other members of the Carcinonemertidae can be
hermaphroditic (Roe, 1986; Shields, 2001). The eggs are extruded
into a mucilaginous egg strand, which adheres to a seta on the pleopod within the clutch. The egg strands can also be used to indicate
the presence of the worms. The biology and ecology of these nemerteans is unknown. Comparative studies on the life history of
these species would be very interesting.
Clawed lobsters bear a nemertean that is unlike the carcinonemertids. It is known as Pseudocarcinonemertes homari and it probably belongs in the family Tetrastemmidae. Charmantier et al.,
(1991) used freshwater baths to rid clawed lobsters of infestations.
Freshwater killed worms after only 4 min. Freshwater baths and
Malachite green have been used to control infestations of C. errans
on Cancer magister (Wickham, 1988). The fact that carcinonemertids can occur in epidemics on crab species indicates that these
worms have the potential to occur in outbreaks on lobsters. Care
should be taken in transporting ovigerous lobsters to new locations
to avoid the introduction of their worms and other parasites.
Broodstock should also be examined prior to establishing colonies
of lobsters.
17. Miscellaneous metazoan symbionts
As yet, spiny lobsters are not known to be infested with any
member of the Cycliophora. Symbion pandora and S. americanus
are the only described members of the phylum Cycliophora (Obst
et al., 2006). The phylum is highly specialized with both S. pandora
and S. americanus living on the mouthparts of the Norway lobster
and American lobster, respectively (Funch and Kristensen, 1995;
Obst et al., 2006). However, I have observed rotifers and rotifer-like
animals on the mouthparts and gills of spiny lobsters from the
Great Barrier Reef, and given what is now known about the Cycliophora, these organisms may be of further scientific interest.
18. Nicothoidae – parasitic copepods
Nicothoid copepods are a family of highly specialized symbionts
that live on other crustaceans. Nicothoidae were once in the family
Choniostomatidae (see Boxshall and Lincoln, 1983). The copepods
live in the egg clutches or marsupia of their hosts. They are egg
predators and in some cases they are very similar to host eggs;
in fact, they are considered egg mimics (Hansen, 1897; Bowman
and Kornicker, 1967). Choniomyzon panuliri is a nicothoid copepod
that infests the clutches of Panulirus spp. (Pillai, 1962). Several species of Panulirus from the Great Barrier Reef, Australia, have a similar species in their egg clutches (Shields, unpubl. data). The larval
stages of species of Choniosphaera occur in the gill chambers of
non-ovigerous crab hosts, but this has not been examined for
Choniomyzon. The feeding habits of at least one species of Nicothoidae can cause significant damage to the fecundity of individual
hosts (Shields and Wood, 1993).
Two other copepods have been reported from spiny lobsters.
The harpacticoid copepod Paramphiascopsis sp. is apparently symbiotic in the gills of male and female J. edwardsi and S. verreauxi
(Booth, personal communication). Members of this genus apparently occur in symbiotic relationships with other invertebrates
(Soyer, 1973; Hicks, 1986). Another harpacticoid copepod occurs
in the gills of Panulirus spp. from the Great Barrier Reef, Australia.
It is similar to Sunaristes (Shields, personal observation).
J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
19. Amphipods
Spiny lobsters often have amphipods living in close association
with them, either on their egg clutches or within their branchial
chambers. The relationship is likely commensal, but those found
on the egg clutches are probably predatory on the eggs of their
hosts. Parapleustes commensalis occurs on the pleopods and egg
clutches of P. interruptus and Cancer anthonyi from Southern
California (Shoemaker, 1952; Shields pers. observation) and Paralithodes californiensis (Wicksten, 1982). It ate crab eggs in experimental systems (Shields, unpubl. data). Ischyroceros sp. infests
the egg clutches of Paralithodes camtschaticus (Kuris et al., 1991).
It was correlated with significant egg mortality on that host; hence,
the potential of amphipods as egg predators on lobsters. Gitanopsis
iseebi has been reported from the branchial chamber of Panulirus
japonicus (Yamato, 1993). Other members in the family of amphiolochid amphids are commensals; therefore, it is likely to be a
commensal. Isaea elmhirsti lives as a commensal on the mouthparts
of H. gammarus (McGrath, 1981, in Costello et al. (1990)). Amphipods are highly motile and they can swim away from a host when
it is captured; therefore, some of these relationships may be hard
to study. Symbiotic amphipods warrant attention, particularly in
studies of fecundity or reproductive behaviors or when establishing broodstock for nascent culture studies.
20. Fouling organisms
A plethora of organisms can foul the exoskeleton of lobsters and
other crustaceans. Some are opportunists that simply require a
hard surface. Others are obligate symbionts. Some are also useful
as indicators of the health of their host. The gooseneck barnacles,
Octolasmis spp., use a number of decapod crustaceans as hosts,
including panulirid and scyllarid lobstes (Jeffries et al., 1982).
These barnacles live on the exoskeleton and within the gill chambers of their hosts. They are highly adapted to the life history patterns of their hosts. One species, O. bullata, had a prevalence of 48%
on Panulirus polyphagus off Singapore (Jeffries et al., 1982). No
other species were found on the lobster, but other species were recorded from a number of different decapods from the area. Octolasmis angulata has been recorded from P. versicolor from Western
Australia (Jones, 2004). Octolasmis californiana was first reported
from the California spiny lobster, P. interruptus (Newman, 1960),
but it is known to occur on several decapods (Newman and Abbott,
1980). Another lepadomorph barnacle, Trilasmis fissum hawaiiensis
is specific to P. japonicus and P. penicillatus where it lives on the setae of the mouthparts (Bowers, 1968). This relationship is quite
intriguing and indicative of the specialization that can occur in
barnacles.
There are few reports of balanomorph, or acorn barnacles, on
spiny lobsters. However, Chelonibia patula and a few other species
sometimes infest them. Chelonibia patula is a host generalist, found
on turtles, drift wood, decapods and other hard surfaces. It will settle on virtually any hard surface. Species of Paralepas occur on
spiny lobsters from Australia (Jones, 2003). Several species of balanomorph barnacles have been found on spiny lobsters P. argus
(Eldred, 1962). Heavy infestations of acorn barnacles may indicate
that the host lobster is not in good health and may be suffering an
underlying infection. Such hosts should receive priority for health
examinations for rare infections.
Bryozoans, hydroids, serpulorbid molluscs, bivalve molluscs
and many other animals can be found as fouling organisms on
the shells of spiny lobsters, but there are few records of their
occurrence. In horseshoe crabs and true crabs, the fouling community has been used to address ecological questions on succession (Key et al., 1996, 1997, 1999). I have seen fewer fouling
87
organisms on lobsters than on crabs, perhaps because many crab
species have terminal molts whereas lobsters often do not.
21. Disease syndromes
21.1. Turgid lobster syndrome
Turgid lobster syndrome is an unusual condition that occurs
in J. edwardsii from New Zealand (Diggles, 1999) and Panulirus
ornatus from Australia (Evans, pers. communication). The etiology of the syndrome is unknown, but the condition is easy to
identify because the thin arthrodial membranes between the
tergites swell from fluid pressure, causing the animals to become
stiff and lifeless. Affected animals have difficulty moving so they
stop feeding and become lethargic. It occurs in short-term aquaculture facilities and is probably the result of poor water quality,
but that remains to be determined. Up to 50% of the affected
animals can die (Diggles, 1999). Animals can be bled with a syringe to relieve fluid pressure, and those treated this way often
survive.
A similar condition was described in lobsters from Japan
(Wada et al., 1994). Lobsters were lethargic and had visible
swelling of the arthrodial membranes. Histology revealed an
inflammation of the myocardium as well as infiltration of hemocytes into affected cardiac muscles. Interestingly, the changes to
the heart were visible as white nodules in the heart. The etiology
of the syndrome was not determined, but it was not a microbial
agent. Both wild and laboratory-held animals showed signs of
the syndrome.
21.2. Pink lobster syndrome
An unusual syndrome occurs in P. cygnus from Western Australia. The condition is known as pink lobster syndrome and the flesh
of affected animals becomes pink or orange (see Shields et al.,
2006). Affected lobsters usually die from the syndrome. They show
signs of lethargy and have a bitter flavor, making them unpalatable. Very little information is available on this syndrome, but it
appears to occur in holding facilities as well as in nature. This condition should be further documented to determine if it is caused by
an infectious or non-infectious agent.
21.3. Mass mortalities
Several mass mortalities have been documented in spiny lobster populations. An interesting mortality event was recorded from
a tagging study that was undertaken to estimate the size of the
population of P. ornatus at Yule Island, Papua New Guinea (Dennis
et al., 1992). The mortality of adult lobsters was high, 95%, during
the fishing period; however, only a third of the mortality was
attributed to fishing pressure, the rest was thought to be due to
physiological stress from reproduction and migration, but disease
agents were not examined. These events are different from those
that alter habitat, thereby forcing migrations such as observed
with hypoxia events. Lobster mortalities are relatively common
along the coast of South Africa (see Cockcroft, 2001). Hypoxia
caused by harmful algal blooms develops in deep waters and forces
lobsters, Jasus lallandii, to move into extremely shallow littoral
areas, which are subject to rapid tidal changes. Lobsters cannot escape exposure to air, and when caught in the intertidal zone, they
die in mass strandings. Ecological disturbances to the Florida Keys
have altered the habitats of the juvenile lobsters causing them to
move into less impacted areas (Butler et al., 1995). The potential
for increased exposure to pathogens has not been studied in these
88
J.D. Shields / Journal of Invertebrate Pathology 106 (2011) 79–91
systems, but the emergence of PaV1 shortly after these events is
very intriguing.
22. Conclusions
Spiny lobsters have few reported parasites, diseases and symbionts. Several species of spiny lobster have supported fisheries for
many years, so one would expect their diseases to be well documented in their fished populations. However, this is not the case.
This is primarily because fisheries focus on animals of legal size,
which are typically adults. In my experience, diseases are more
common and more serious in postlarval and early juvenile stages,
a segment of the population that is not sampled by the fishery. Further, it is likely that new emerging diseases will become significant
issues in nascent aquaculture industries for spiny lobsters. In fact,
several microbial pathogens are known to be problematical to cultures of larval and juvenile clawed lobsters (see Fisher et al.,
1976a,b, Nilson et al., 1975, 1976; Brock and Lightner, 1990). These
include bacterial (Vibrio spp., L. mucor), fungal (Lagenidium spp.)
and protozoal (peritrich ciliates) agents, all of which can be difficult to control in closed systems. No doubt more symbionts and
disease agents will emerge as fisheries become more intensively
managed and aquaculture production force animals into high density, stressful conditions.
The presence of emerging pathogens and disease outbreaks are
often signs that a population is under significant stress. Identifying
the stressors that ultimately lead to or contribute to outbreaks can
be very difficult. Such factors as degrading water quality, seasonal
hypoxia or increased temperatures have been linked to disease or
death in other systems; and they are likely to operate in populations of spiny lobsters. While such stressors have yet to be implicated in disease in spiny lobster populations, several have been
implicated in mortalities of clawed lobsters. For example, a major
mortality occurred in Long Island Sound in 1999. Several factors
were thought to have contributed to the mortality, including temperature stress, hypoxia, increased manganase concentrations,
pesticides, other pollutants, Neoparamoeba sp. or combinations of
these factors (Pearce and Balcom, 2005; Howell et al., 2005). It
can be very difficult to undertake multifactorial studies on multiple
stressors, but in the case of the 1999 mortality, experiments indicated that lobsters cannot tolerate combinations of severe hypoxic
conditions, poor water quality and temperature stress (Dove et al.,
2004, 2005; Draxler et al., 2005a,b; Robohm et al., 2005). Thus, it is
important to understand both the proximate and ultimate causes
that give rise to emerging pathogens to determine their potential
threat to lobster fisheries.
At present, the virus PaV1 in P. argus represents the most significant pathogen to any spiny lobster. It infects the smallest
juveniles and, therefore, would normally go unnoticed in populations. However, fishermen in the Caribbean spiny lobster fishery
use juveniles as social attractants (bait) in fished traps and
lethargic diseased lobsters would be discarded. Thus, fishermen
may spread diseased animals, or at least the virus, into new fishing grounds (Shields and Behringer, 2004). While contact transmission is the most efficient mode for the spread of PaV1,
water-borne transmission can also occur, albeit at a much lower
efficiency (Butler et al., 2008). However, healthy lobsters can detect and avoid diseased animals, which makes contact transmission less likely to occur in nature (Behringer et al., 2006). The
ramifications of transmission to the spread of PaV1 is unclear,
but it is clear that the virus can threaten the valuable Caribbean
fisheries (Shields and Behringer, 2004). Recent emerging diseases, such as PaV1 in P. argus and Hematodinium in the Norway
lobster (e.g. Field et al., 1992), highlight the fact that we know
very little about the pathogens in our lobster stocks.
Acknowledgments
This work was partially supported by NSF Grants OCE-0136894
and OCE BE-UF-0723662. This is contribution #3101 from the Virginia Institute of Marine Science.
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