Concurrent Infections (parasitism and bacterial … A... · Web viewTilapias are susceptible to a number of infectious agents including bacteria and parasites (Shoemaker et al. 2006)

8th International Symposium on Tilapia in Aquaculture 2008

CONCURRENT INFECTIONS (PARASITISM

Aquatic Animal Health Research Laboratory, USDA-Agricultural Research Service, 990 Wire Rd, Auburn, Alabama, USA 36832

AbstractMost 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.

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: [email protected]

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. Tilapias 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 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

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mailto:[email protected]

CONCURRENT INFECTIONS (PARASITISM AND BACTERIAL DIESEASE) in TILAPIA

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).

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CRAIG A. SHOEMAKER et al.

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.

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

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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 was 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) 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

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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 tilapiaTilapia 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).

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

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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

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

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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

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

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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 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

(grayling,

Thymallus thymallus)

Diplostomum spathaeceum

Aeromonas salmonicida

Presence of D. spathaeceum

invasion in fish increased the

proportion of fish carrying A.

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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.

REFERENCES

1. Bandilla M., E. T. Valtonen, L-R Suomalainen, P. J. Aphalo and T. Hakalahti. 2006. A link between ectoparasite infection and susceptibility to bacterial disease in rainbow trout. Int. J. Parasitol. 36, 987-991.

2. Bowers J. M., A. Mustafa, D. J. Speare, G. A. Conboy, M. Brimacombe, D. E. Sims and J. F. Burka. 2000. The physiological response of Atalntic

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salmon, Salmo salar L., to a single experimental challenge with sea lice, Lepeophtheirus salmonis. J. Fish Dis. 23, 165-172.

3. Bowser P. R., G. A. Wooster, R. G. Getchell and M. B. Timmons. 1998. Streptococcus iniae infection of tilapia Oreochromis niloticus in a recirculation production facility. J. World Aquac. Soc. 29, 335–339.

4. Buchmann K. and J. Brescian. 1997. Parasitic infection in pond-reared rainbow trout Oncorhynchus mykiss in Denmark. Dis. Aquat. Org. 28, 125-138.

5. Buchmann K. and T. Lindenstrøm. 2002. Interactions between monogenean parasites and their fish hosts. Int. J. Parasitol. 32, 309-319.

6. Busch S., I. Dalsgaard and K. Buchmann. 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., J. R. Arthur and M. G. Bondad-Reantaso. 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 P. H. Odense 1984. Pathology of five species of Gyrodactylus Nordmann, 1832 (Monogenea). Can. J. Zool. 62, 1084-1088.

9. Cusack R. and D. K .Cone. 1985. A report of bacterial microcolonies on the surface of Gyrodactylus (Monogenea). J. Fish Dis. 8, 125-127.

10. Cusack R. and D. K. Cone. 1986. A review of parasites as vectors of viral and bacterial diseases of fish. J. Fish Dis. 9, 169-171.

11. Evans J. J., P. H. Klesius., D. J. Pasnik and C. A. Shoemaker. 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., T. Nakai and K. Muroga. 1990. Scanning electron microscopy on the skin surface of ayu Plecoglossus altivelis infected with Vibrio anguillarum. Dis. Aquat. Org. 8, 73-75.

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13. Kabata Z. 1985. Parasites and diseases of fish cultured in the tropics. Taylor and Francis, Philadelphia. 318p.

14. Klesius P. and W. Rogers. 1995. Parasitisms of catfish and other farm raised food fish. J. Amer. Vet. Med. Assoc. 207, 1473-1478.

15. Klesius P. H., C. A. Shoemaker and J. J. Evans. 1999. Efficacy of a killed Streptococcus iniae vaccine in tilapia (Oreochromis niloticus). Bull. Eur. Assoc. Fish Pathol. 19(1), 39-41.

16. Perera R. P., S. K. Johnson, M. D. Collins and D. H. Lewis. 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., L-R Suomalainen, M. Tiirola and E. Tellervo Valtonen. 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 C. Sommerville. 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., J. J. Evans and P. H. Klesius. 2000. Density and dose: factors affecting mortality of Streptococcus iniae infected tilapia (Oreochromis niloticus). Aquaculture 188: 229-235.

23. Shoemaker C. A., P. H. Klesius and J. J. Evans. 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, J. J. Evans and P. H. Klesius. 2006. Parasites and Diseases. In C. Lim and C.D. Webster (eds) Tilapia:

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Biology, Culture and Nutrition (pp. 561-582). The Haworth Press, Inc., Binghamton, NY.

25. Stoffregen D. A., S. C. Backman, R. E. Perham, P. R. Bowser and J. G. Babish. 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 D. T. Nolan. 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, C. A. Shoemaker and P. H. Klesius. 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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Documents

Concurrent Infections (parasitism and bacterial … A... · Web viewTilapias are susceptible to a number of infectious agents including bacteria and parasites (Shoemaker et al. 2006)