A One-Year Surveillance Program for Erysipelothrix rhusiopathiae: Methodology, Findings, And Recommendations
IAAAM Archive
Martin G. Greenwell; Jeffrey R. Boehm; Brigita M. Harris
John G. Shedd Aquarium, Chicago, IL, USA


A twelve month study (January 2001 to December 2001) was undertaken in which feed fish (capelin, herring and squid), potential reservoir hosts (pinnipeds and cetaceans) and environmental samples (habitat water, pool surfaces) were sampled and cultured for the presence or absence of Erysipelothrix rhusiopathiae. The working hypothesis of this pilot study was that the organism would be found with the greatest frequency on or in feed fish, with moderate to low frequency in animals and that the organism would rarely if ever be isolated from environmental samples.


Erysipelothrix rhusiopathiae can cause a peracute to acute septicemia as well as a more subacute to chronic dermatological disease in cetaceans. The cutaneous form is usually amenable to treatment by prompt intervention with antibiotics. The septicemic form is often fatal due to the rapid temporal sequence wherein vague, nonspecific clinical signs, i.e., anorexia, lethargy, lead to septic shock and then death in the span of hours. Erysipelothrix rhusiopathiae has been repeatedly isolated from food items (fish and invertebrates) that are routinely fed to captive cetaceans.1 Presumably, the organism is carried on food fish and is then introduced through a breech in the oral cavity, esophagus, or gastrointestinal tract of a susceptible animal.2,3,4 This study represents a prospective investigation that was undertaken to both standardize microbiological isolation techniques and to increase understanding of the prevalence of E. rhusiopathiae in feed fish, potential reservoir animals, and on pool surfaces in the captive environment.


The animals sampled in this study included five beluga whales (Delphinapterus leucas), four Pacific white-sided dolphins (Lagenorhynchus obliquidens), and four harbor seals (Phoca vitulina). All of these animals are housed in an indoor, artificial saltwater system of 2.7 million gallons. The system is composed of five pools that can be connected in a variety of arrays or separated by netted gates. The three species above are kept separated at all times. The water temperature is maintained between 55 and 65 degrees. Water quality is maintained by mechanical and biological filtration through rapid sand filters and then disinfection with ozone. All of the animals were in good health throughout the study. Restaurant-quality fish and squid are fed to the animals. Feed fish are obtained frozen from established vendors. Accurate records were kept as to the lots of the fish that were fed. Herring, capelin, and squid were the only food items being offered during this time period and all three were randomly sampled. Proximate nutritional analyses of these feed fish were consistent with previous lots.

Materials used for isolation of E. rhusiopathiae included the following:

1.  Erysipelas enrichment broth5,6

a.  Disodium phosphate 12.02 g

b.  Monopotassium phosphate 2.09 g

c.  Beef extract 3.00 g

d.  Tryptose 15.00 g

e.  Sodium Chloride 5.00 g

f.  Distilled water 1.00 liter

g.  Filter through cotton or filter paper and autoclave. Cool and add aseptically:

i.  Bovine, horse or other serum 50.00 ml

ii.  Kanamycin 400.00 mg

iii.  Neomycin 50.00 mg

iv.  Vancomycin 25.00 mg

h.  Dispense aseptically. Store at 4-5 degrees C for no longer than 2 weeks.

2.  Sterile, cotton-tipped swabs and CulturettesTM

3.  35 degrees C incubator

4.  Blood agar and Columbia C.N.A. agar plates

5.  TSI agar slants, OF glucose tubes, gelatin infusion medium, oxidase and catalase reagents

6.  API Coryne StripTM for confirmation

Sampling techniques were as follows: Recently thawed fish (typically covered with a thin layer of ice, but not solidly frozen) were sampled early on the morning of testing. Sterile cotton swabs were inserted among fish in several places and the swab was immediately placed into the enrichment broth. This was repeated with a second swab from a separate area with each of the study species of fish (herring, capelin, and squid). In addition, samples were obtained from the bottoms of feed buckets immediately following a feeding session. Duplicate samples were obtained of the fluid remaining from the assortment of herring, capelin and/or squid that had been fed. Swabs were wiped around the edges of the bucket or over any leftover fish and placed into the enrichment broth. Samples of feces (L. obliquidens, D. leucas) and saliva from gingival surfaces (P. vitulina) were obtained via CulturetteTM and inoculated into the enrichment broth. System water from the animals' habitat was collected via a standard technique from a pre-determined site in sterile polypropylene bottles for quantitative cultures. One milliliter was removed from each bottle with a sterile pipette and placed into nine milliliters of enrichment broth. Concurrently, samples were obtained at the interface of the tank wall and the water surface. Samples were obtained from two standard locations in a habitat and swabs were then inoculated into enrichment media.

Isolation and identification of E. rhusiopathiae proceeded as follows. The tubes containing erysipelas enrichment broth were warmed to room temperature and then inoculated with cotton-tipped sample swabs followed by incubation for 48 hours at 35 degrees C. Tubes were examined for turbidity at the end of the incubation period and an inoculum was then plated on to C.N.A. agar plates. Plates were incubated for 24 to 48 hours. After 24 hours, C.N.A. plates were examined for growth and colony morphology was noted; at 48 hours, the presence or absence if alphaehemolyses was noted. Positive samples were inoculated onto a TSI slant and into an O-F glucose tube and incubated for an additional 48-72 hours. A gelatin tube was also inoculated and incubated at room temperature for 48 to 72 hours. Following incubation, the stab line of the TSI agar slant was evaluated for the presence or absence of hydrogen sulfide, the gelatin tube was evaluated for the presence or absence of "feather" growth, and the O-F glucose tube was evaluated for the presence or absence of a weak acid reaction. Suspect colonies were confirmed with an API Coryne StripTM. Positive isolates were then transferred to another C.N.A. agar plate. Following culture on C.N.A. agar, the entire growth was then inoculated in to 2 mls of skim milk in a cryovial and stored at minus 70 degrees Celsius.

Table 1. Characteristics of E. rhusiopathiae and differentiation from other commonly encountered gram-positive, catalase-negative bacteria7


E. rhusiopathiae


A. pyogenes

OF Glucose

Weak acid

Strong acid

Weak acid


No liquefication

No liquefication



H2S positive

H2S negative

H2S negative


Feed fish samples yielded one positive isolate of E. rhusiopathiae from herring (n=24), three isolates from capelin (n=24), and one isolate from squid (n=14). From the samples obtained from fish feed buckets post-feeding, a total of ten isolates were recovered (n=72). Fecal samples (n=24) from Pacific white-sided dolphins (L. obliquidens) resulted in two positive isolates from the same animal. Fecal samples (n=24) taken from five different beluga whales (D. leucas) yielded no growth. Gingival swabs (n=25) from four different harbor seals (P. vitulina) resulted in five positive E. rhusiopathiae isolates. Four of the five positive seal gingival isolates were from the same two individual seals, i.e., two positive isolates per seal, and the fifth gingival isolate was from a third harbor seal. Environmental samples yielded no positive isolates (n=96).


This study confirms that E. rhusiopathiae can be consistently albeit sporadically isolated from food items. In contrast to an earlier pilot study at Shedd Aquarium8, the organism was not predominantly isolated from one particular species of fish. It has been shown in previous studies that the pathogen can be isolated from both freshwater and marine fish as well as marine shellfish.9 In the present study, E. rhusiopathiae was more frequently isolated from the accumulated debris from thawing food fish that remains in the feed bucket (external body mucus, scales, melted ice, etc.) after the fish themselves have been fed out. This debris originates from several whole, thawed fish; thus, it is feasible that the organisms are concentrated here. It would certainly appear to be a better source for erysipelas isolation than samples from single, isolated food fish. Having confirmed the work of previous investigators documenting the presence of E. rhusiopathiae in food fish, sporadic recovery of the organism from the oral cavity and feces of marine mammals should be expected. Indeed, the agent has been recovered in clinically normal captive and free-ranging cetaceans and pinnipeds alike in previous studies.8,10 The Pacific white-sided dolphin and the beluga whale that both succumbed to erysipelas septicemia at the John G. Shedd Aquarium had small, single, yet quite distinct, rents in their esophageal mucosae. These findings support the consensus opinion, which maintains that the alimentary tract is the probable portal-of-entry for the pathogen.2,3,4 The inability to isolate E. rhusiopathiae from environmental samples in this study is consistent with having a very small sample size in comparison to the total water volume and surface area of the animal habitats, i.e., a tremendous dilution factor, as well as the effectiveness of ozone disinfection of the water. Excellent reviews of the current state of knowledge regarding the prevention and diagnosis of erysipelas as well as current and future research priorities are available.1,4 The current clinical approach at the Shedd Aquarium is to promptly and aggressively initiate antibiotic therapy with any cetacean that suddenly and dramatically exhibits behavioral changes, i.e., sudden, complete anorexia, acute weakness/lethargy, especially if there are no evident social reasons for such changes. Concurrently, blood samples are collected for an in-house CBC, chemistry profile (i-STATTM), and blood culture (Septi-ChekTM BHI).


1.  Dunn JL, JD Buck, TR Robeck. 2001. Bacterial diseases of cetaceans pinnipeds. In: CRC Handbook of Marine Mammal Medicine. 2nd Edition. Dierauf, L.A. and F.M.D. Gulland, eds., CRC Press, Boca Raton, pp.309-335.

2.  Geraci JG, RM Sauer, W Medway. 1966. Erysipelas in dolphins. Am. J. Vet. Res., 27:597.

3.  Kinsel MJ, JR Boehm, BM Harris, RD Murnane. 1997. Fatal Erysipelothrix rhusiopathiae septicemia in a captive Pacific white-sided Dolphin (Lagenorhynchus obliquidens). J. Zoo. Wildl. Med. 28: 494-497.

4.  Boehm JR, G Lacave, RA Patterson. 2001. Proceedings of the First International Workshop on Erysipelas in Cetaceans.

5.  Blair JE, EH Lennette, JP Truant. 1970. Manual of Clinical Microbiology American Society for Microbiology. P. 650.

6.  National Veterinary Services Laboratory, Ames, Iowa, USA. 2000. Personal Communication.

7.  Wood RL. 1970. In: Manual of Clinical Microbiology. Blair, J.E., E.H. Lennette and J.P. Truant, eds. American Society for Microbiology. Pp. 101, 899.

8.  Harris BM, JR Boehm. 2001. A pilot surveillance program for E. rhusiopathiae in a public aquarium. In: Proceedings of the 32nd Annual Conference of the IAAAM. F.M.D.Gulland, ed. Pp. 104-107.

9.  Lauckner G. 1985. Diseases of mammalia: pinnipedia. In: Diseases of Aquatic Animals. O. Kinne, ed. Biologische Anstalt Helgoland, Hamburg, 654 Pp.

10. Suer LD, NV Vedros. 1988. E. rhusiopathiae. I. Isolation and characterization from pinnipeds and bite/abrasion wounds in humans. Dis. Aquat. Organisms. 5:1-5.

Speaker Information
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Martin G. Greenwell
John G. Shedd Aquarium
Chicago, IL, USA

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