Concurrent Infections (parasitism and bacterial … · Web viewCraig A. Shoemaker(, De-Hai Xu,...
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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: [email protected]
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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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