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 LaboratoryUSDA-Agricultural Research Service990 Wire RdAuburn, 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 36832Tel.: (334) 887-3741; Fax (334) 887-2983; E-mail: craig.shoemaker@ars.usda.gov

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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.

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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).

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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.

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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

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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) 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

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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).

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

(N=90)

Mortality (%) Mean days to death

Gyrodactylus S. iniae (G-S) 42.2 ± 10.7 a 7.8 ± 0.3 a

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No infection S. iniae (N-S) 6.7 ± 0 b 7.8 ± 0.1 a

Gyrodactylus None (G-N) 0 ± 0 b NA *

No infection None (N-N) 0 ± 0 b NA

* Not available.

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.

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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

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

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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 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

(Fish species)

Parasite/Bacteria Result

Evans et al. 2007

(channel catfish,

Ictalurus punctatus)

Trichodina spp.

Streptococcus iniae

S. agalactiae

Concurrent infection made

catfish susceptible to

streptococcal disease

Bandilla et al. 2006

(rainbow trout,

Oncorhynchus mykiss)

Argulus coregoni

Flavobacterium

columnare

Parasitic infection increased

the susceptibility of trout to

columnaris disease

Pylkkö et al. 2006 Diplostomum Presence of D. spathaeceum

invasion in fish increased the

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(grayling,

Thymallus thymallus)

spathaeceum

Aeromonas

salmonicida

proportion of fish carrying A.

salmonicida

Busch et al. 2003

(rainbow trout)

Gyrodactylus derjavini

F. psychrophilum

Correlated host mortality to

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 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.

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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, 987-991.

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.

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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. Lowe-McConnel (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: 174-177.

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.

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