CONCURRENT INFECTIONS (PARASITISM AND BACTERIAL DIESEASE) in
TILAPIA
Craig A. Shoemaker, De-Hai Xu, Phillip H. Klesius and Joyce J. Evans
Aquatic Animal Health Research Laboratory
USDA-Agricultural Research Service
990 Wire Rd
Auburn, Alabama, USA 36832
Abstract
Most laboratory disease studies in tilapia to date have focused on a single parasite or a single bacterial pathogen. In
intensive tilapia aquaculture, the reality of a single disease agent resulting in death-loss may be small. More likely,
multiple disease agents are present (i.e., parasites, bacteria and/or a combination) and responsible for disease
losses. This paper will focus on concurrent infections or the potential for concurrent infections in tilapia aquaculture.
We will highlight a recent study completed at our laboratory on parasitism with a monogenetic trematode and
subsequent bacterial infection with Streptococcus iniae in Nile tilapia (Oreochromis niloticus). Concurrent
experimental infection with Gyrodactylus niloticus and S. iniae resulted in significantly higher mortality in tilapia
(about 42%) as compared to immersion infection with S. iniae alone (7%) and parasitism with G. niloticus only (0%).
Gyrodactylus niloticus presumably provided a portal of entry for invasive bacteria due to damage of the fish
epithelium. Interestingly, G. niloticus was also found to harbor viable S. iniae at 24 and 72 h post infection
suggesting that G. niloticus may vector S. iniae from fish to fish.
INTRODUCTION
Tilapia aquaculture has expanded rapidly in the last ten years. The trend of increased production
is expected to continue due to the increased demand for tilapia in the international market. To meet the
increased demand, intensification of production will undoubtedly occur. Intensive fish production often
results in increased disease due to poor water quality and high stock densities used. Tilapia are
susceptible to a number of infectious agents including bacteria and parasites (Shoemaker et al. 2006).
Research is typically aimed at a single disease agent and not at concurrent infections. In reality, most

Aquatic Animal Health Research Laboratory, United States Department of Agriculture-Agricultural
Research Service, 990 Wire Rd, Auburn, Alabama, USA 36832
Tel.: (334) 887-3741; Fax (334) 887-2983; E-mail: craig.shoemaker@ars.usda.gov
1
intensive tilapia production systems probably have multiple disease agents resulting in death-loss. Two
important pathogens of tilapia are Streptococcus iniae (Figure 1), a gram positive bacteria responsible for
significant losses in intensive culture (Perera et al. 1994; Stoffregen et al. 1996; Shoemaker and Klesius
1997; Bowser et al. 1998; Klesius et al. 1999; Shoemaker et al. 2000; 2001), and Gyrodactylus niloticus,
a monogenetic trematode that can cause problems in young fish stocked at high numbers in eutrophic
(nutrient rich) waters (Klesius and Rogers 1995; Shoemaker et al. 2006).
Cusack and Cone (1986) reviewed the limited information on the ability of parasites to vector
viral and bacterial diseases of fish. They concluded that parasite vectors increase the transmission
efficiency of pathogens by creating portals of entry and/or by having the ability to transfer pathogens
directly from fish to fish. Recent studies have demonstrated that Cusack and Cone’s hypothesis was
correct (Kanno et al. 1990; Pylkkö et al. 2006; Bandilla et al. 2006; Evans et al. 2007). Other studies
suggest different mechanisms that increase host susceptibility; for example, parasitism has been shown
to result in increased stress responses believed to be linked to decreased disease resistance (Bowers et
al. 2000; Tully and Nolan 2002). This manuscript will highlight a recent study where we evaluated
concurrent G. niloticus and S. iniae infections in Nile tilapia (Oreochromis niloticus). We will further
discuss trends in examining concurrent parasitism and bacterial infection in fish.
Figure 1. A) Streptococcus iniae infected tilapia showing spinal curvature and erratic swimming and B)
positive starch reaction of Streptococcus iniae. (arrow shows the zone of hydrolysis).
2
MATERIALS AND METHODS
Fish and parasite
Nile tilapia, Oreochromis niloticus, reared in tanks using filtered recirculated water at the Aquatic
Animal Health Research Laboratory, Auburn, Alabama were used as experimental animals. Upon
examination of the gills and fin under a light microscope, this stock had a mean intensity of Gyrodactylus
of less than 10 per fish. The parasites were identified as G. niloticus (Cone et al. 1995) (Figure 2).
Intense infections were developed by holding 50 or more fish in 57-L aquaria for 1-2 weeks. Dead fish
killed by heavy parasite burden were removed and naïve individuals added back to maintain the parasite
population (Busch et al. 2003). During the trial, the mean ± standard deviation of dissolved oxygen (DO)
was 6.5 ± 0.7 mg L-1, temperature was 26.4 ± 0.6 °C, pH was 7.4 ± 0.2, ammonia was 0.2 ± 0.1 mg L -1,
and hardness was 91.9 ± 12.3 mg L-1. Nitrite concentrations were below the threshold for detection.
Figure 2. Gyrodactylus niloticus shown associated and attached to gill filaments from a parasitized
tilapia.
3
Examination of Gyrodactylus infection of fish
Four wet mount samples were prepared from each fish to assess intensity, two from the caudal
fin and two from the gill. Mucus was scraped with a glass cover slip from entire caudal fin, each side of
the fin representing one sample. Gill filaments (5 × 5 mm) were clipped from the left and right branchial
arches, placed in a wet mount and compressed by applying pressure using a cover slip. These samples
were examined using a compound microscope (Olympus, Orangeburg, New York) at low magnification.
The entire wet mount was scanned from left to right and from top to bottom to enumerate the parasites.
Bacterial isolation
An isolate of S. iniae (ARS-98-60), originally isolated from a hybrid striped bass ( Morone chrysops
X M. saxatilis), was obtained from diseased tilapia in the laboratory and identified biochemically
(Shoemaker and Klesius 1997). The isolate was grown on a sheep blood agar plate and then cultured in
tryptic soy broth (Difco, Becton Dickinson, Sparks, MD) for 24 h at 28°C and used to challenge the tilapia.
Dead or moribund tilapia were removed twice daily during the trial and bacterial samples aseptically
obtained from brain of tilapia were streaked onto 5% sheep blood agar plates. Bacterial colonies with
beta-hemolysis, testing negative for catalase production, positive by Gram-stain and having a coccoid
morphology were considered S. iniae positive. Forty fish (20 Gyrodatylus infected and 20 non-infected
fish) were cultured to verify the S. iniae free status prior to each trial.
Streptococcus iniae infection trial
Tilapia infected with G. niloticus were divided into 2 groups, one group treated with formalin and
potassium permanganate and the other received no treatment. The treated group of fish was immersed
in formalin at 100 mg L-1 for one hour on Day 1. Potassium permanganate was added to tanks at 5 mg
L-1 to treat fish for 1 h on Day 2 and Day 3, respectively. After treatment, flowing water was provided to
each tank at 0.5 L min-1. The treated fish were allowed one week to recover from parasite infection and
chemical treatment. A total of 200 tilapia infected with G. niloticus and 200 fish parasite free were used
[fish ranged from 9.0 ± 0.6 (mean ± SD) cm in length and 11.7 ± 2.6 g in body weight (N=20)]. Ninety
fish were used in each of four groups: 1) G. niloticus infection and challenged with S. iniae (G-S), 2) no
parasite infection and challenged with S. iniae (N-S), 3) G. niloticus infected and no S. iniae (G-N) and 4)
4
no G. niloticus and no S. iniae (N-N), with 30 fish per tank and 3 tanks per group. The remaining fish (20
infected with G. niloticus and 20 treated fish without parasite) were examined for parasite load and S.
iniae infection prior to the trial. Twelve buckets, one for each aquarium, were filled with 2-L tank water
and 30 fish per bucket with aeration. For fish challenged with S. iniae, the bacterial suspension was
added to the bucket at the rate of 107 colony-forming units (CFU) mL-1. After immersion for 1 h, the fish
from each bucket were moved to a 57-L aquarium with flowing water at 0.4-0.5 L min-1 with aeration.
Mortality of fish was recorded, and dead or moribund fish were examined for parasite load and S. iniae
infection twice daily for 2 weeks. Five surviving fish from each tank were sampled for S. iniae at trial
termination.
Collection of G. niloticus from S. iniae infected tilapia
Tilapia infected by G. niloticus at 40 (SD 8, N=10) parasites per wet mount sample from caudal
fin and 20 (SD 9) from gill were distributed into 3 buckets, 35 fish per bucket. Fish in each bucket
received one of following treatments: 1) S. iniae IP injected, 2) S. iniae immersion exposed, and 3) no
treatment (control). Each fish in the injected group was IP injected with 0.1 ml of S. iniae, equal to 108
CFU fish-1. Thirty-five tilapia were immersed in 2-L water with 7 × 107 CFU mL-1 S. iniae for one hour.
Fish in each bucket were then moved to a 57-L aquarium and ten fish in each group were sampled 24
and 72 h post exposure to S. iniae. Gyrodactylus was collected only from live or moribund fish since the
parasites leave the fish shortly after death. At sampling, one tilapia at a time was placed in 50 ml cold
MS-222 solution (300 mg mL-1) in a 500 mL beaker. Fish body surface was washed with MS-222 solution
using a Pasteur pipette. The solution containing the parasite was passed through a screen with an
aperture of 425 µm, transferred to a 50-mL centrifuge tube and centrifuge at 90 g for 3 min. All
parasites collected from an individual fish were pooled and counted as one sample. The parasites were
treated with 300 IU of penicillin mL-1 and 300 µg streptomycin mL-1 (Sigma) for 15 min, washed 3 times
with sterile water in 15 mL centrifuge tubes centrifuged at low speed (90 g). The washed G. niloticus
was transferred to a 1.5 mL centrifuge tube containing 0.5 mL sterile water, vortexed at high speed for
one min, inoculated onto Columbia CNA 5% sheep blood agar and incubated at 28 °C for 24-48 h to
identify S. iniae colonies (Shoemaker & Klesius 1997).
5
Statistical analysis
Percentage of fish positive for S. iniae and mortality of fish from different treatments were
analyzed with Duncan multiple range tests (SAS Institute 1989). Median days to death were calculated
and compared by Lifetest procedure (SAS Institute 1989) (Kaplan-Meier method). Probabilities less than
0.05 were considered significant.
RESULTS
Streptococcus iniae infection trial
Mortality was 42.2% in the G-S group, which was significantly higher (P < 0.05) than fish
exposed to S. iniae but not G. niloticus (N-S group=6.7%, Table 1). No G-N or N-N control fish died
during the trial. S. iniae was isolated from all dead fish with SBA except 3 out of 38 fish from the G-S
group. The percentages of fish culture positive for S. iniae were 92.1% and 100% for fish in the G-S and
the N-S groups, respectively. Bacteriologic examination revealed no S. iniae from fish prior to the trial or
from surviving fish.
Gyrodactylus niloticus was found on the gills and fins of all fish in groups G-S and G-N
throughout the trial (Table 2). Intensity of infection was higher at the start of the experiments but
lower on surviving fish. The parasite was rarely found on fish that had been treated with formalin and
K2MnO4.
Table 1 Cumulative mortality of Gyrodactylus niloticus infected Nile tilapia 14 days post-S. iniae
immersion challenge. Within a given column, means (± SD) followed by different superscript letters
are statistically different (P < 0.05) (Xu et al. 2007).
Parasite Infection
Challenge
Mortality (%)
Mean days to death
7.8 ± 0.3
a
7.8 ± 0.1
a
(N=90)
Gyrodactylus
S. iniae (G-S)
42.2 ± 10.7
No infection
S. iniae (N-S)
6.7 ± 0
Gyrodactylus
None (G-N)
0±0
b
NA *
No infection
None (N-N)
0±0
b
NA
* Not available.
6
b
a
Isolation of S. iniae from infected tilapia and G. niloticus
In this trial, 36.7% and 40.0% of fish infected by G. niloticus died 72 h post-S. iniae challenge by
IP injection and immersion, respectively. All dead fish from IP injected and 40% fish from immersion
exposure were positive for S. iniae 24 h post challenge. Fish were 44.4% and 37.5% positive for S. iniae
when challenged by IP injection and immersion, respectively, 72 h post challenge. In tilapia co-infected
with S. iniae and G. niloticus, S. iniae was isolated from 60% and 11% of G. niloticus collected 24 and 72
h, respectively, from fish IP injected with S. iniae (Table 3). Forty and 25% of parasites cultured from
immersion exposed fish were positive for S. iniae when collected at 24 and 72 h.
Table 2 Number of Gyrodactylus niloticus in fin and gill samples of Nile tilapia (Xu et al. 2007).
Fish group
Day 0
Day 1-13
Day 14
No.
Fin
Gill
No.
Fin
Gill
No.
Fin
Gill
G – S*
40
19 ± 4
29 ± 7
32
45 ± 10
20 ± 5
30
3±1
4±1
N–S
40
0±0
0±0
4
0±0
0±0
30
0±0
0±0
G–N
40
19 ± 4
29 ± 7
NA
NA
NA
30
4±2
1±0
N–N
40
0±0
0±0
NA
NA
NA
30
0±0
0±0
* G-S: Gyrodactylus niloticus infected and challenged with S. iniae, N-S: no parasite infection and
challenged with S. iniae, G-N: Gyrodactylus niloticus infection and no S. iniae, and N-N: no G. niloticus
and no S. iniae infection.
Table 3 Number (n) of Gyrodactylus niloticus samples positive for Streptococcus iniae at 24 or 72 h post
challenge with S. iniae. All Gyrodactylus niloticus collected from an individual fish were pooled and
counted as one sample (Xu et al. 2007).
Gyrodactylus niloticus
Streptococcal Challenge
IP injection
Immersion
Not exposed
n/ samples
%
n/ samples
%
N/samples
%
24 h S. iniae positive
6/10
60
4/10
40
0/10
0
72 h S. iniae positive
1/9
11
2/8
25
0/10
0
7
DISCUSSION
Anecdotal data have suggested the possible role of parasites in enhancing infections of fish with
bacterial pathogens. For example, Plumb (1997) reported that in a recirculation tilapia production facility,
presence of Trichodina spp. presumably caused epidermal injuries that lead to streptococcal and
edwarsiellosis infections that could not be controlled by antibiotics. Control of the parasite with formalin
resulted in a decrease in overall death loss. Cusack and Cone (1985) demonstrated the presence of
bacteria in close association with monogenetic trematodes using scanning electron microscopy and
suggested a vectoring role for ectoparasites. Others (Roberts and Summerville 1982; Kabata 1985;
Buchmann and Bresciani 1997) also suggest the enhancement of secondary bacterial infections due to
presence of ectoparasites. However, until recently, few studies documented these interactions. The
present study demonstrated that captive tilapia with a single infection of G. niloticus or of S. iniae had
less than 7% total mortality. However, during co-infection, mortality increased significantly. The present
experimental challenge study helps to confirm earlier impressions that under farm conditions these two
diseases can occur together with devastating results.
Gyrodactylus spp. may serve as a vector for S. iniae as well as damage fish epithelium and create
portals of entry for bacterial invasion. G. niloticus attaches to fish gills, fins and skin by a posterior
attachment haptor with one large anchor and 16 marginal hooklets (Hoffman 1985). The invasion and
movement of parasites from one location to another on fish cause mechanical injuries to the epithelium
(Cone and Odense 1984). These injuries may serve as portals of entry for bacterial invasion making fish
with G. niloticus infection more susceptible to S. iniae. In the present study, S. iniae was isolated from
Gyrodactylus collected not only from fish exposed to S. iniae by immersion but also from fish infected by
IP injection. The material ingested while Gyrodactylus feeds on fish epithelia (blood or tissues) passes to
the parasite gut (Buchmann and Lindenstrøm 2002). The results suggest that S. iniae was ingested by G.
niloticus and survived in the parasite for approximately 72 hours. Busch et al. (2003) studied
concomitant infection of G. derjavini and Flavobacterium psychrophilum in trout. Results of their study
suggested that invasion by F. psychrophilum was only slightly enhanced by presence of G. derjavini and
mortality was correlated to gyrodactylid infection (Table 4). Interestingly, the highest mortality occurred
8
in the group with highest number of parasites concurrently infected with F. psychrophilum. These
authors (Busch et al. 2003) did not try and isolate F. psychrophilum from G. derjavini.
Table 4. Recent literature documenting concurrent infections between parasites and bacterial diseases
in fish.
Study
Parasite/Bacteria
Result
Evans et al. 2007
Trichodina spp.
Concurrent infection made catfish
(channel catfish, Ictalurus
Streptococcus iniae
susceptible to streptococcal
punctatus)
S. agalactiae
disease
Bandilla et al. 2006
Argulus coregoni
Parasitic infection increased the
(rainbow trout,
Flavobacterium
susceptibility of trout to
Oncorhynchus mykiss)
columnare
columnaris disease
Pylkkö et al. 2006
Diplostomum
Presence of D. spathaeceum
(grayling,
spathaeceum
invasion in fish increased the
Thymallus thymallus)
Aeromonas salmonicida
proportion of fish carrying A.
(Fish species)
salmonicida
Busch et al. 2003
Gyrodactylus derjavini
Correlated host mortality to
(rainbow trout)
F. psychrophilum
gyrodactylid infection level
Studies in other fish species have recently linked parasitic disease and bacterial infection (Table
4). Evans et al. (2007) showed that parasitism of channel catfish ( Ictalurus punctatus) fry with Tricodina
sp. resulted in increased susceptibility of catfish to streptococcal disease caused by either S. iniae or S.
agalactiae. These authors further suggested that parasite-induced mechanical injury increased fish
susceptibility. Mechanical injury due to invasion and/or feeding of parasites has also been shown in
coldwater fish species. Bandilla et al. (2006) demonstrated an increase in disease due to Flavobacterium
columnare, if fish were co-infected with Argulus coregoni. They suggested that the parasite feeding on
9
the skin was responsible for damage and possibly aided bacterial attachment. Another study suggested
that infection with a digenetic trematode (Diplostomum spathaceum) resulted in more Aeromonas
salmonicida cells in heart tissue than in fish without trematode infection (Pylkkö et al. 2006). However,
increased mortality due to the presence of both infectious agents was not observed. While many of the
studies have demonstrated an association between concurrent infectious agents and disease in fish, some
have produced unequivocal results. This area of research is relatively little studied in tilapia. Due to the
intensification of culture conditions, studies examining multiple pathogens will be invaluable to solve
tilapia producers’ current and future problems.
Acknowledgements
We thank Drs. Tom Welker and David Pasnik (USDA-ARS) for critical review of the manuscript.
10
REFERENCES:
1.
Bandilla M., Valtonen E.T., Suomalainen L-R., Aphalo P.J. and Hakalahti T. 2006. A link between
ectoparasite infection and susceptibility to bacterial disease in rainbow trout. Int. J. Parasitol. 36, 987991.
2.
Bowers J.M., Mustafa A., Speare D.J., Conboy G.A., Brimacombe M., Sims D.E. and Burka J.F.
2000. The physiological response of Atalntic salmon, Salmo salar L., to a single experimental challenge
with sea lice, Lepeophtheirus salmonis. J. Fish Dis. 23, 165-172.
3.
Bowser P.R., Wooster G.A., Getchell R.G. and Timmons M.B. 1998. Streptococcus iniae infection
of tilapia Oreochromis niloticus in a recirculation production facility. J. World Aquac. Soc. 29, 335–339.
4.
Buchmann K. and Bresciani J. 1997. Parasitic infection in pond-reared rainbow trout
Oncorhynchus mykiss in Denmark. Dis. Aquat. Org. 28, 125-138.
5.
Buchmann K. and Lindenstrøm T. 2002. Interactions between monogenean parasites and their
fish hosts. Int. J. Parasitol. 32, 309-319.
6.
Busch S., Dalsgaard I. and Buchmann K. 2003. Concomitant exposure of rainbow trout fry to
Gyrodactylus derjavini and Flavobacterium psychrophilum: effects on infection and mortality of host. Vet.
Parasitol. 117, 117-122.
7.
Cone D.K., Arthur J.R. and Bondad-Reantaso M.G. 1995. Description of two new species of
Gyrodactylus von Nordmann, 1832 (Monogenea) from cultured Nile tilapia, Tilapia nilotica (Cichlidae), in
the Philippines. Proc. Helminthol. Soc. Wash. 62, 6-9.
8.
Cone D.K. and Odense P.H. 1984. Pathology of five species of Gyrodactylus Nordmann, 1832
(Monogenea). Can. J. Zool. 62, 1084-1088.
9.
Cusack R. and Cone D.K. 1985. A report of bacterial microcolonies on the surface of
Gyrodactylus (Monogenea). J. Fish Dis. 8, 125-127.
10.
Cusack R. and Cone D.K. 1986. A review of parasites as vectors of viral and bacterial diseases
of fish. J. Fish Dis. 9, 169-171.
11.
Evans J.J., Klesius P.H., Pasnik D.J. and Shoemaker C.A. 2007. Influence of natural Trichodina
sp. parasitism on experimental Streptococcus iniae or Streptococcus agalactiae infection and survival of
young channel catfish Ictalurus punctatus (Rafinesque). Aquaculture Res. 38, 664-667.
12.
Kanno T., Nakai T. and Muroga K. 1990. Scanning electron microscopy on the skin surface of
ayu Plecoglossus altivelis infected with Vibrio anguillarum. Dis. Aquat. Org. 8, 73-75.
13.
Kabata Z. 1985. Parasites and diseases of fish cultured in the tropics. Taylor and Francis,
Philadelphia. 318p.
14.
Klesius P. and Rogers W. 1995. Parasitisms of catfish and other farm raised food fish. J. Amer.
Vet. Med. Assoc. 207, 1473-1478.
15.
Klesius P.H., Shoemaker C.A. and Evans J.J. 1999. Efficacy of a killed Streptococcus iniae
vaccine in tilapia (Oreochromis niloticus). Bull. Eur. Assoc. Fish Pathol. 19(1), 39-41.
11
16.
Perera R.P., Johnson S.K., Collins M.D. and Lewis D.H. 1994. Streptococcus iniae associated
mortality of Tilapia nilotica × T. aurea hybrids. J. Aquat. Anim. Health 6, 335–340.
17.
Plumb J.A. 1997. Infectious diseases of tilapia. In: Tilapia aquaculture in the Americas (ed by
B.A. Costa-Pierce & J.E. Rakocy), pp 212-228, World Aquaculture Society, Bataon Rouge, Louisiana.
18.
Pylkkö P., Suomalainen L-R, Tiirola M.and Tellervo Valtonen E. 2006. Evidence of enhanced
bacterial invasion during Diplostomum spathaceum infection in European grayling, Thymallus thymallus
(L.). J. Fish Dis. 29: 79-68.
19.
Roberts R.J. and Sommerville C. 1982. Diseases of tilapias. In R.S.V. Pullin and R. H. LoweMcConnel (Eds.), The biology and culture of tilapias (pp 247-263). Manila, Philippines: ICLARM
Conference Proceedings 7.
20.
SAS Institute. 1989. SAS/Stat user΄s guide, Ver. 6, 4th edn. SAS Institute, Cary, NC, USA.
21.
Shoemaker C.A. and Klesius P.H. 1997. Streptococcal disease problems and control: a review.
In: Tilapia Aquaculture (ed. by K. Fitzsimmons), pp 671-680. Northeast Regional Aquacultural
Engineering Service, Ithaca, New York.
22.
Shoemaker C.A., Evans J.J. and Klesius P.H. 2000. Density and dose: factors affecting mortality
of Streptococcus iniae infected tilapia (Oreochromis niloticus). Aquaculture 188: 229-235.
23.
Shoemaker C.A., Klesius P.H. and Evans J.J. 2001. Prevalence of Streptococcus iniae in tilapia,
hybrid striped bass and channel catfish from fish farms in the United States. Amer. J. Vet. Res. 62: 174177.
24.
Shoemaker C.A., Xu De-Hai, Evans J.J. and Klesius P.H. 2006. Parasites and Diseases. In C.
Lim and C.D. Webster (eds) Tilapia: Biology, Culture and Nutrition (pp. 561-582). The Haworth Press,
Inc., Binghamton, NY.
25.
Stoffregen D.A., Backman S.C., Perham R.E., Bowser P.R. and Babish J.G. 1996. Initial disease
report of Streptococcus iniae infection in hybrid striped (sunshine) bass and successful therapeutic
intervention with fluoroquinolone antibacterial enerofloxacin. J. World Aquac. Soc. 27: 420–436.
26.
Tully O. and Nolan D.T. 2002. A review of the population biology and host-parasite interactions
of the sea louse Lepeophtheirus salmonis (Copepoda: Caligidae). Parasitology 124: 165-182.
27.
Xu De-Hai, Shoemaker C.A. and Klesius P.H. 2007. Evaluation of the link between gyrodactylosis
and streptococcosis of Nile tilapia, Oreochromis niloticus (L.). J. Fish Dis. 30: 233-238.
12