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Detection of Edwardsiella infections in Opsanus tau by Polymerase Chain Reaction.

Opsanus tau, the oyster toadfish, is an important laboratory animal used in hearing and balance research at the Marine Biological Laboratory (MBL) (1). Wild-caught and cultured fish are maintained year-round in both recirculation and flow-through tanks in the Marine Resources Center (MRC) at the MBL. A major cause of disease in toadfish held at the MRC is Edwardsiella tarda (1, 2). A member of the Enterobacteriaceae, E. tarda is a natural inhabitant of fresh and marine water and causes gastrointestinal and extraintestinal disease in humans (3). Published reports of fish disease caused by E. tarda involve cultured warm-water fish, but the disease can also affect cold-water salmon (4). Poor water quality, high water temperatures, feces, and decaying organic matter likely contribute to the onset and severity of the disease and probably allow for occurrence and proliferation of the bacteria in the fish's environment. These factors, combined with capture and holding-induced stresses, may account for the high levels of disease caused by E. tarda in the toadfish in our facility.

Currently, the only diagnostic test available for identification of E. tarda in diseased fish is bacterial culture and identification using traditional biochemical tests. At the MRC, these tests are sent to an outside laboratory, and the process takes about three weeks. Treatment is often attempted before verification that the lesions identified at necropsy were caused by E. tarda. Using the PCR methods described here, we can specifically and rapidly identify E. tarda, resulting in timely and appropriate management procedures and treatments for infected fish.

The polymerase chain reaction (PCR) is a DNA-based method that can be used for the quick and sensitive detection of microorganisms in both antemortem and necropsied tissues. PCR primers specific for E. tarda isolates from Japanese eels have been derived from an anonymous species-specific sequence (5) or from the hemolysin gene (6). Here we describe PCR primers based on Edwardsiella small subunit (ssu) RNA genes for direct identification of E. tarda. Primer development will be described in detail elsewhere.

The type strains E. tarda ATCC 15947 and E. ictaluri ATCC 33202 were obtained from the American Type Culture Collection. E. ictaluri is an important pathogen of cultured catfish and produces very similar lesions to those caused by E. tarda in toadfish. In addition, E. tarda has been shown to cause disease in catfish (7). Cultures were grown in Difco marine brain heart infusion broth. One milliliter of the liquid broth culture was transferred aseptically to 1.5-ml microfuge tubes, which were then centrifuged at 2790 x g for 4 min to pellet cells. Pellets were resuspended in 0.6 ml of lysis buffer (10 mM Tris-Cl pH 8.0, 5 mM EDTA, and 1% SDS). Proteinase K (50 [micro]g) was added, and tubes were incubated overnight at 50[degrees]C. Genomic DNA was precipitated by the addition of 0.55 ml of isopropanol; the liquid portion was removed completely. The DNA pellet was rinsed twice with 0.8 ml of 70% ethanol. After the ethanol was evaporated, the DNA was dissolved in 0.8 ml of TE (Tris-EDTA; 8).

Forward primer EtaI-363f (5'-GTG TRC GTG TTA ATA GCA-3') was designed to amplify E. tarda from human sources (biotype 1), represented by the type strain ATCC 15947. Forward primer Eta2-351 (5'-TAG GGA GGA AGG TGT GAA-3') was designed to amplify E. tarda strains isolated from fish (biotype 2). An Edwardsiella genus-specific reverse primer Edwsp-780r (5'-CTC TAG CTT GCC AGT CTT-3') was used with the forward primers.

PCR amplification was performed with an Applied Biosystems GencAmp 9700. The reaction mixture (10 [micro]l) contained l x Taq polymerase buffer (Promega), 1.5 mM Mg[Cl.sub.2], 1 mM dNTP mix, 100 nM each of forward and reverse primer, 0.5 units Taq polymerase (Promega) and template DNA. The thermal cycle profile commenced with an initial denaturation for 1 min at 94[degrees]C; 30 cycles of denaturation (1 min at 94[degrees]C), annealing (1 rain at 58[degrees]C), and extension (1 min at 72[degrees]C); and a final extension at 72[degrees]C for 7 min. Amplification products were electrophoresed in a 1.5% agarose gel (Fisher Scientific) in 0.5 x Tris-Borate-EDTA (TBE) buffer (8). Gels were stained with 1 [micro]g/ml ethidium bromide and digitally photographed. Images were manipulated in Adobe Photoshop. Some amplifications were carried out directly from bacterial colonies, in which case the initial denaturation time in the PCR profile was increased to 5 min at 94 [degrees]C.

The results, as determined by electrophoresis, showed that DNA of E. tarda and E. ictaluri could be distinctly amplified by the appropriate primers (Fig. 1). The Etal-363f-Edwsp-780r primers amplified E. tarda ATCC 15947, yielding a product of 216 bp, and did not amplify E. ictaluri ATCC 33202 (Fig. 1A, lanes 1-4) or MRC fish isolates biochemically identified as E. tarda (not shown). As would be expected, Eta2-351f-Edsp-780r failed to amplify both type strains (Fig. 1A, lanes 5-8). However, Eta2-351f-Edwsp-780r primers amplified the E. tarda fish isolates (Fig. 1B, lanes 1-6), demonstrating that the strains isolated at the MRC were clearly of fish origin, and biotype 2. The biotype 2 amplifications were performed directly from bacterial colonies, showing that direct identifications were indeed possible.


Future work will involve development of a direct assay method for E. tarda from both postmortem toadfish tissues and, more importantly, antemortem tissues. The ability to amplify E. tarda strains from toadfish is the first step to being able to understand, isolate, and control future outbreaks of E. tarda.

R. Smolowitz acknowledges support from NIH/NIDCD grant 5P01 DC001837-09. H. Chikarmane acknowledges support from the Wilkens Foundation and the Cape Cod Community College Educational Foundation.

Literature Cited

(1.) Smolowitz, R., and R. Bullis. 1997. Biol. Bull. 193: 270-271.

(2.) Wenuganen, S., R. Bullis, R. Smolowitz, and E. Barbieri. 1997. Biol. Bull. 193: 269-270.

(3.) Janda, J. M., and S. L. Abbott. 1993. Clin. Infect. Dis. 17: 742-748.

(4.) Plumb, J. A. 1999. Pp. 479-490 in Fish Diseases and Disorders Vol. 3, Viral Bacterial, and Fungal Infections, P. T. K. Woo and D. W. Bruno, eds. CABI Publishing, New York.

(5.) Aoki, T., and I. Hirono. 1995. Asian Fisheries Society Special Publication. 10: 135-146.

(6.) Chen, J. D., and S. Y. Lai. 1998. Zool. Stud. 37: 169-176.

(7.) Darwish, A., J. A. Plumb, and J. C. Newton. 2000. J. Aquat. Anim. Health 12: 255-266.

(8.) Sambrook, J., and D. Russell. 2001. Molecular Cloning: A Laboratory Manual, 3rd ed. Cold Spring Harbor Laboratory Press, New York.

Krystal D. Baird (1,2), Hemant M. Chikarmane (1,3), Roxanna Smolowitz (1) *, and Kevin R. Uhlinger (1)

(1) Marine Biological Laboratory, Woods Hole, MA

(2) Barnstable County AmeriCorps Cape Cod, Barnstable, MA

(3) Cape Cod Community College, W. Barnstable, MA

* Corresponding author:
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Author:Baird, Kyrstal D.; Chikarmane, Hemant M.; Smolowitz, Roxanna; Uhlinger, Kevin R.
Publication:The Biological Bulletin
Date:Oct 1, 2003
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