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Table of Contents

CHAPTER I.......................................................................................................................2

INTRODUCTION.............................................................................................................2

1.1 Abstract..............................................................................................................2

1.2 Description Of Genus Edwardsiella...................................................................2

1.3 Species Of Genus Edwardsiella...........................................................................3

1.3.1 Edwardsiella hoshinae................................................................................3

1.3.2 Edwardsiella ictaluri...................................................................................3

1.3.3 Edwardsiella tarda......................................................................................4

CHAPTER II......................................................................................................................6

SPECIFICATION OF GENUS EDWARDSIELLA..........................................................6

2.1 EDWARDSIELLA HOSHINAE................................................................................6

2.1.1 Description.................................................................................................6

2.2 EDWARDSIELLA ICTALURI...................................................................................6

2.2.1 Description.................................................................................................6

2.2.2 Signalment..................................................................................................7

2.2.3 Clinical Signs.............................................................................................8

2.2.4 Epidemiology.............................................................................................8

2.2.5 Distribution................................................................................................9

2.2.6 Pathology....................................................................................................9

2.2.7 Diagnosis....................................................................................................9

2.2.8 Treatment.................................................................................................10

2.2.9 Control......................................................................................................10

ESC may be controlled through reducing the amount of stress in fish stocks and cessation of feeding when outbreaks occur. A killed bacterin vaccine is available and administered in water by bath immersion.....................................................................10

2.3 EDWARDSIELLA TARDA.....................................................................................10

2.3.1 Description...............................................................................................10

2.3.2 Signalment................................................................................................12

2.3.3 Clinical Signs...........................................................................................12

2.3.4 Epidemiology...........................................................................................13

2.3.5 Distribution...............................................................................................13

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2.3.6 Pathology..................................................................................................14

2.3.7 Diagnosis..................................................................................................14

2.3.8 Treatment.................................................................................................15

2.3.9 Control......................................................................................................15

CHAPTER III..................................................................................................................16

EDWARDSIELLA INFECTIONS OF FISHES.................................................................16

3.1 Introduction.....................................................................................................16

3.3 Pathology Edwardsiella tarda...........................................................................17

3.4 Pathology Edwardsiella ictaluri........................................................................18

3.5 Host and Geographic Range.............................................................................19

3.6 Source and Reservoir of Infection....................................................................21

3.7 Incubation Period.............................................................................................21

3.8 Control.............................................................................................................21

3.9 Treatment........................................................................................................22

CHAPTER IV..................................................................................................................23

EDWARDSIELLA TARDA SEPTICEMIA WITH UNDERLYING MULTIPLE LIVER ABSCESSES...................................................................................................................23

4.1 Abstract............................................................................................................23

4.2 Introduction.....................................................................................................23

4.3 Case Report......................................................................................................24

4.4 Discussion.........................................................................................................26

CHAPTER V...................................................................................................................29

NATURAL ANTIBIOTIC SUSCEPTIBILITIES OF EDWARDSIELLA TARDA, E. ICTALURI, AND E. HOSHINAE.....................................................................................29

5.1 Abstract............................................................................................................29

5.2 MATERIALS AND METHODS..............................................................................31

5.2.1 Bacterial strains........................................................................................31

5.2.2 Identification............................................................................................31

5.2.3 Antibiotics and antibiotic susceptibility testing........................................32

5.2.4 Evaluation of natural antibiotic susceptibility...........................................33

5.2.5 β-Lactamase testing..................................................................................34

5.3 RESULTS............................................................................................................34

5.3.1 Identification............................................................................................34

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5.3.2 Natural antibiotic sensitivity and resistance..............................................35

5.3.3 Quality assurance.....................................................................................35

5.3.4 β-Lactamase testing..................................................................................36

CHAPTER VI..................................................................................................................37

CONCLUSION................................................................................................................37

LIST OF PICTURE.........................................................................................................39

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CHAPTER I

INTRODUCTION

1.1 Abstract

The genus Edwardsiella was first described in 1965, with E. tarda as the type

species (Ewing et al. 1965). A second species was isolated from reptiles and birds,

and was characterized by Grimont et al. (1980) as E. hoshinae. E. ictaluri, the

causal agent of ESC was first isolated in 1976 (Hawke 1979); however, the

bacterium was not characterized and classified until 1979 (Hawke et al. 1981).

1.2 Description Of Genus Edwardsiella

Edwardsiella is a genus of small, straight gram-negative rods which are

facultatively anaerobic bacteria (family Enterobacteriaceae) containing motile,

chemoorganotrophic, peritrichous, nonencapsulated rods. Members of this genus

are usually found in the intestines of cold-blooded animals and in fresh water.

They are pathogenic for eels, CATFISHES, and other animals and are rare

opportunistic pathogens for humans. The type species is Edwardsiella tarda,

which is occasionally isolated from the stools of both healthy humans and those

with diarrhea, from the blood of humans and other animals, and from human

urine. Edwardsiella tarda is an etiologic agent of gastroenteritis in humans. The

two other species in this genus are Edwardsiella hoshinae and Edwardsiella

ictaluri.

Kingdom : Bacteria

Phylum : Proteobacteria

Class : Gamma Proteobacteria

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Order : Enterobacteriales

Family : Enterobacteriaceae

Genus : Edwardsiella

Species : E. hoshinae

E. ictaluri

E. tarda

(R. Sakazaki et al., 1962)

1.3 Species Of Genus Edwardsiella

1.3.1 Edwardsiella hoshinae

Grimont. 1981, sp. Nov. Edwardsiella hoshinae a motile species that, isolated

from animals and humans, does not produce indole. This is a straight rod that is

motile by peritrichous flagella. Growth is best at 35-37 C. It is a Gram negative

organism. Gas is often produced and hydrogen sulfide is common. It is rarely

found in the feces of healthy people, and is characterized as an infrequent

opportunistic pathogen. It has also been isolated from birds, reptiles, and the

environment.

Type strain : strain 2 – 78 = ATCC 33379 = CIP 78.56 = DSM 13771 = JCM

1679 = NCTC 12121.

Etimology : N.L. gen. n. hoshinae, of Hoshina; named after Toshikazu Hoshina,

the Japanese bacteriologist who was one of the first to describe an organism that

was probably an Edwardsiella.

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1.3.2 Edwardsiella ictaluri

Hawke et al. 1981, sp. nov. Edwardsiella ictaluri a nonmotile species that does not

produce indole, occurring as a pathogen of catfish. Edwardsiella ictaluri (also

known as Enteric Septicaemia of Catfish, Hole in the Head Disease, and ESC) is a

member of the Enterobacteriaceae family.

The bacterium is a short, gram negative, pleomorphic rod with flagella. It causes

the disease enteric septicaemia of catfish (ESC), which infects a variety of fish

species (including many catfish species, knifefish and barbs). The bacteria can

cause either acute septicaemia or chronic encephalitis in infected fish. Outbreaks

normally occur in spring and autumn. E. ictaluri can be found in Asia and the

United States, being of particular economic importance in the U.S. It is not a

zoonosis.

Type strain (see also StrainInfo.net) : strain SECFDL (Southeastern Cooperative

Fish Disease Laboratory) GA 77-52 = ATCC 33202 = CDC 1976-78 = CCUG

18764 = CIP 81.96 = DSM 13697 = JCM 1680 = JCM 16934 = NCTC 12122.

Etimology : N.L. n. Ictalurus, the genus name for catfish; N.L. gen. n. ictaluri, of

Icatlurus, of catfish.

1.3.3 Edwardsiella tarda

Ewing and McWhorter 1965, species. (Type species of the genus). This bacterium

is a facultatively anaerobic, small, motile, gram negative, straight rod with

flagella. Edwardsiella tarda is a member of the Enterobacteriaceae family. The

bacterium is a facultatively anaerobic, small, motile, gram negative, straight rod

with flagella.

Infection causes Edwardsiella septicemia (also known as ES, edwardsiellosis,

emphysematous putrefactive disease of catfish, fish gangrene and red disease) in

channel fish, eels and flounder.

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It is a zoonosis and can infect a variety of animals including fish, amphibians,

reptiles and mammals. E.tarda has a worldwide distribution - it is found in mud

and the intestine of fish and other marine animals. It is spread by carrier animal

faeces.

Type strain (see also StrainInfo.net) : strain ATCC 15947 = CCUG 1638 = CIP

78.61 = DSM 30052 = JCM 1656 = LMG 2793 = NCCB 73021 = NCTC 10396.

Etimology : L. fem. adj. tarda, slow (intended meaning was “inactive,” referring

to the fermentation on only a few carbohydrates compared to many other

Enterobacteriaceae).

Edwardsiella tarda Ewing and McWhorter 1965 (Approved Lists 1980) and

Edwardsiella anguillimortifera (Hoshina 1962) Sakazaki and Tamura 1975

(Approved Lists 1980) have the same type strain and Edwardsiella

anguillimortifera (Hoshina 1962) Sakazaki and Tamura 1975 (Approved Lists

1980) is the earlier synonym. However, Farmer et al. 1976 have submitted a

Request for an Opinion to conserve the specific epithet tarda over the specific

epithet anguillimortifera. No action was taken on the request, because the request

has been withdrawn. Consequently, the two names remain homotypic synonyms.

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CHAPTER II

SPECIFICATION OF GENUS EDWARDSIELLA

2.1 EDWARDSIELLA HOSHINAE

2.1.1 Description

Edwardsiella hoshinae a motile species that, isolated from animals and humans,

does not produce indole. This is a straight rod that is motile by peritrichous

flagella. Growth is best at 35-37 C. It is a Gram negative organism. Gas is often

produced and hydrogen sulfide is common. It is rarely found in the feces of

healthy people, and is characterized as an infrequent opportunistic pathogen. It has

also been isolated from birds, reptiles, and the environment.

2.2 EDWARDSIELLA ICTALURI

2.2.1 Description

Edwardsiella ictaluri belongs to the Enterobacteriaceae family and is a Gram

negative, short, pleomorphic rod, measuring 0.75 × 1.5-2.5 µm, which is weakly

motile at 25-30°C, but not at higher temperatures. It has peritrichous flagella and

occasionally pili that can be seen with a scanning electron micrographs and can

have between one to three plasmids depending on their molecular mass. It is

generally considered an obligate pathogen, although it can survive in sterilised

pond bottom mud for over 90 days but does not compete well with other

microbes.

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The organism is lactose negative, catalase-positive, cytochrome oxidase-negative,

glucose fermentative and reduces nitrate to nitrite.[1]

E. ictaluri affects fish species only and causes enteric septicaemia of catfish

(ESC) and various other species of fish. ESC is considered one of the most

important infectious disease problems in the commercial catfish industry in the

USA. Other species of Edwardsiella include E. tarda, which causes septicemia in

fish and can affect other animals, whereas E. hoshinae infects birds and reptiles.

Within channel catfish species the bacteria cause two forms of ESC; an acute

septicaemia and chronic encephalitis. In the latter form the infection spreads from

the olfactory sacs, and migrates along the olfactory nerves to the brain, generating

granulomatous inflammation. In the acute form of ESC, the disease is thought to

develop from the intestinal mucosa causing a bacteremia.

2.2.2 Signalment

Channel Catfish

Wild hosts include white, bullhead, blue, and wels catfish species and Japanese

eel, Glass knifefishes, Tadpole Madtom, Rosy barb (minnow family), and species

of carp called Devario devario. Domestic hosts include white, walking, channel

and sutchi catfish species and under experimental setting rainbow trout and

chinook salmon.

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2.2.3 Clinical Signs

With the chronic form of ESC clinical signs include, altered mentation,

listlessness and chaotic swimming with ‘head-up, tail-down’ posture, circling and

mortality. In later stages, the dorsum of the head swells and ulcerates revealing

areas of the brain (hence the name ‘hole in the head disease’).

With acute forms of ESC you can see petechial haemorrhages around the buccal

area, throat, abdomen and the fin base, that progress to depigmented ulcers. Fish

generally suffer from moderate pale inflamed gills, exophthalmia, anaemia,

haemorrhagic enteritis, systemic oedema, dropsy, ascites and splenomegaly.

General behavioural changes include loss of balance, swimming near the surface,

lethargy and cessation of feeding.

2.2.4 Epidemiology

The bacteria can survive in pond sediment and once a population of fish have

recovered from an infection of ECS, they can become carriers. They can be found

in the kidneys of fishes and are thought to be shed in the faeces of fish.

Outbreaks are mainly seasonal and occur within a set temperature range of 18-

28°C, primarily in spring and autumn. This temperature limitation precludes the

bacterium from being a pathogen for humans or other warm-blooded animals [2]

and is not therefore zoonotic. Other environmental factors have been linked to

outbreaks and include poor water quality, high stocking density and other

stressors. E.ictaluri can invade the, gill mucosa, olfactory organ and nasal

epithelium and nerve, brain meninges, skull and capillaries in the dermis of the

skin.

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2.2.5 Distribution

E.ictaluri is mainly found in the USA, Asia and Thailand. The continual

worldwide dissemination of channel catfish for aquaculture purposes may increase

its future distribution.

2.2.6 Pathology

Histological examination reveals a systemic infection of all organs and skeletal

muscles, with the most severe changes being diffuse interstitial necrosis of the

anterior and posterior kidney and systemic haemorrhages. Focal necrosis in the

liver and spleen are also generally seen as pale grey/white lesions.

Skeletal muscle and areas of necrosis within internal organ tissue can be

infiltrated with macrophages, that phagocytose the bacteria but do not destroy

them.

2.2.7 Diagnosis

Clinical signs are quite pathognomonic for ESC but PCR is used to confirm the

presence of E. ictaluri in blood and tissues but other methods have been used such

as indirect FAT (detecting antibodies) and ELISA test.

The organism is slow growing and forms small, translucent, greenish colonies on

Edwardsiella isolation media (EIM), while inhibiting Gram-positive and most

Gram-negative contaminating organisms. E. ictaluri can be separated from

E.tarda because it is indole-negative and does not produce H2S on triple sugar

iron (TSI) agar.

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2.2.8 Treatment

Potentiated sulphonamide, sulfadimethoxine, methoprim or oxytetracycline have

been used to treat ESC, but resistance has been recorded.

2.2.9 Control

ESC may be controlled through reducing the amount of stress in fish stocks and

cessation of feeding when outbreaks occur. A killed bacterin vaccine is available

and administered in water by bath immersion.

Addition of the vaccine to feed may serve as a booster after vaccination as higher

survival rates of fish given immersion plus oral applications than fish given

double immersions. Age-related factors and the induction of a cell mediated

response are important in eliciting protection.

2.3 EDWARDSIELLA TARDA

2.3.1 Description

Edwardsiella tarda was the first species identified of the genus Edwardsiella, and

was named after a renowned microbiologist P. R. Edwards (Janda, 1991). E. tarda

was originally named Edwardsiella anguilimortifera, but it was ultimately

changed to E. tarda because this name was used more often in scientific reports.

E. tarda is a Gram-negative bacilli that belongs to the Enterobacteriaceae family

and was first characterized in 1965 (Health, 2001).

E. tarda has many traits that are characteristic of many enterobacteria such as E.

coli. These characteristics include it being a facultative anaerobe, rod-shaped, and

motile (Health 2001). Its motility is due to peritrichous flagella.

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Although Edwardsiella tarda was initially characterized more than thirty years

ago, there is still very little known about this bacterium.

E. tarda is known for causing diseases in both humans and fish, both of which can

potentially be fatal if untreated. Though this may be the case, the likelihood of a

serious infection is very slim.

As a fish pathogen, it is of particular importance to aquaculture and the fishing

industry, especially commercial fish farms. It may become more of a significant

health issue to fish and humans alike, especially in light of emerging and

increasing antibiotic resistance in fish pathogens, due in large part to overuse of

antibiotics in fish farming (Greenlees et al., 1998; Lehane and Rawlin, 2000).

Some studies have focused on using proteomics and molecular techniques to

elucidate the mechanism of pathogenesis in Edwardsiella tarda (Rao et al., 2004).

Studies such as these have allowed the characterization of novel toxin secretion

pathways, such as the discovery of a type VI secretion system essential for E.

tarda pathogenesis (Zheng and Leung, 2007). These types of analyses help us

better understand bacterial pathogenesis in general, as well as provide new

insights for fighting disease.

Edwardsiella tarda belongs to the Enterobacteriaceae family and is a motile Gram

negative, small, straight rod with peritrichous flagella and measures 1 × 2-3 mm.

It is cytochrome oxidase negative, and ferments glucose and is classified as

facultatively anaerobic.

Edwardsiella tarda infects freshwater and marine fishes, reptiles and amphibians

and mammals throughout the world. It causes Edwardsiella septicemia (ES)

which is also known as fish gangrene, emphysematous putrefactive disease of

catfish or red disease in eels. It causes serious systemic infection in cultured

channel fish in the USA and in eels and flounders in Japan. Pale skin, petechiation

and necrotic abscesses within the muscle of fish (that have a putrid odour when

incised) are characteristic of ES.

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Mortality rates can depend on the amount of stress that the fish are kept under and

high temperature, poor water quality and high organic fertility probably contribute

to the onset and severity of the disease.

Unlike E. ictaluri, E. tarda is zoonotic and can infect humans. E. ictaluri causes

enteric septicaemia of catfish (ESC) and only infects fish species, whereas

Edwardsiella hoshinae infects birds and reptiles.

2.3.2 Signalment

Wild hosts include European and Japanese eels, largemouth bass, striped sea bass,

Atlantic salmon, Marble goby, snakes and birds. Domestic hosts include Japanese

Eels, channel catfish, Siamese fighting fish, carp species including catla and rosy

barb, crimson and European seabass, black tetra, Asian seabass (barramundi),

rainbow trout, chinook salmon, Nile tilapia, red seabream, turbot, and Angel fish.

Other fish hosts that have been documented are perch-like species including

Cichlidae, Chrysophrys unicolor, flathead mullet , bastard halibut, flounders, and

mozambique tilapia.

E. tarda can also be found in zoo animals, zebu, cattle, pigs, reptiles, marine

mammals, members of the Alligatoridae family (alligators and caimans) and

humans.

2.3.3 Clinical Signs

Clinical signs vary between fish species; consequently they are generally of little

use except to indicate a bacterial infection. All life stages of fish are affected by E.

tarda and haemorrhaging of the body cavity, muscle, and organs including liver

and kidneys are commonly seen. Within the kidneys and spleen, necrotic

white/grey lesions can be seen on the surface of the organs.

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In adult fish, a variety of clinical sign can be seen including organomegaly, pale

inflamed gills, exophthalmia and cataracts, haemorrhagic red lesions (ecchymosis)

on the skin and fins, erosion of the skin, systemic oedema and ascites.

The anal region of certain species can become swollen and hyperaemic and rectal

prolapses can occur. General behavioural changes include loss of balance, bursts

of abnormal activity, and increased food consumption.

In humans it causes diarrhoea, gastroenteritis, while extraintestinal infections may

produce typhoid-like illness, peritonitis with sepsis, cellulitis and meningitis.

2.3.4 Epidemiology

E.tarda commonly resides in the intestine of fish and other aquatic animals and in

the bottom mud of many bodies of water. Within the USA, E.tarda has been

isolated from the mud, water samples, frogs, turtles and crayfish from catfish

ponds. The bacteria are transmitted through infected water and mud from carrier

animal faeces, and most probably infect susceptible fish through trauma of the

epithelium or via the intestines. The infection can be enhanced by water

temperatures of 20-30°C. Humans have been known to be infected with E. tarda

by eating infected fish meat.

2.3.5 Distribution

E.tarda is a ubiquitous organism and is predominantly found in fish cultured in

the USA, Venezuela, Japan, Taiwan, Korea, India, Thailand, Egypt, Israel and

many developing countries including Africa and South and Central America. It

has also been found in wild fish from Canada, USA and Australia.

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2.3.6 Pathology

Histopathology shows suppurative interstitial nephritis in adult eels, with masses

of degenerate neutrophils containing bacteria. Within early stages of infection

small abscesses are present.

These enlarge and liquefy, spreading bacteria to surrounding tissues and vessels,

causing ulceration of the dermis and emboli and infecting the spleen, liver,

epicardium, stomach, gill and musculature.

In the hepatitis form, micro-abscesses can also develop in the liver and in different

species, such as Japanese flounders, red sea bream, Japanese eels and tilapia,

show predominantly granulomatous inflammation.

At least some E. tarda isolates produce toxic extracellular products (ECP) which

may play a role in their virulence. Its haemolytic activity, which is partially

regulated by iron concentration, could contribute to the pathogenicity of this

bacteria to humans.

2.3.7 Diagnosis

E. tarda can be isolated on brain–heart infusion (BHI) agar or trypton soya agar

(TSA) with inocula from infected internal organs or muscle. It forms small, round,

convex transparent colonies (0.5 mm in diameter) after 24-48 hours. On

Edwardsiella isolation media (EIM), it forms small green colonies with black

centres.

Indirect FAT (detecting antibodies) and enzyme-linked immunosorbent assay

(ELISA) test is used to confirm the presence of E. tarda. There is no serological

cross-reactivity between E. tarda and E. ictaluri. More recently, a loop-mediated

isothermal amplification (LAMP) for rapid and sensitive detection of E. tarda has

been developed.

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2.3.8 Treatment

Oxytetracycline, sulfadimethoxine or methoprim have been used to treat ES. The

latter two can cause cessation of feeding in some fish species. Antibiotic resistant

strains have been isolated e.g. in Taiwan. Some of these resistant strains can be

treated with the addition of oxalinic acid or miloxacin in their feed.

2.3.9 Control

ES may be controlled by the immersion of fish in formalin-killed whole cells

(FKC), lipopolysaccharide (LPS) culture filtrates or whole cell bacterins vaccines.

The two former vaccination may be administered via intramuscular injection and

can cause death to some fish species.

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CHAPTER III

EDWARDSIELLA INFECTIONS OF FISHES

3.1 Introduction

The genus Edwardsiella was suggested by Ewing et al. (1965) to encompass a

group of enteric bacteria generally described under vernacular names such as

paracolon. The type species is E. tarda, which is an opportunistic pathogen of

many animals. Meyer and Bullock (1973) reported E. tarda as a pathogen of

channel catfish (Ieta/urus punetatus) and named the disease emphysematous

putrefactive disease of catfish. However, the organism described by Hoshina

(1962) as the fish pathogen Parae%baetrum anguillimortiferum is now

recognized as beingE. tarda (Wakabayashi and Egusa 1973). Hawke (1979)

isolated several strains of a bacterium closely resembling E. tarda from diseased

cultured channel catfish, but later research showed it to be a distinct new species

named E. ieta/uri (Hawke et al. 1981). Accordingly, the name applied to E.

ieta/urus infections in catfish is enteric septicemia.

3.2 Etiology and Diagnosis

Edwardsiella tarda and E. icta/uri are both gramnegative motile rods that are

cytochrome oxidase negative and ferment glucose with production of acid and

gas. The two species can be differentiated biochemically in that E. tarda produces

both indol and hydrogen sulfide whereas E. ieta/uri produces neither.

Additionally, the two species do not cross-react serologically. Presumptive

diagnosis of E. tarda or E. ieta/uri is based on clincal signs and on isolation and

serological identification of the causative agents.

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A positive slide agglutination test with antiserum specific for E. Tarda or E.

ieta/uri provides a confirmatory diagnosis. Rogers (1981) developed a fluorescent

antibody test and enzyme immunoassay that identify E. tarda and E. ieta/uri, both

in culture and in infected tissues. Horiuchi et al. (1980) also demonstrated that an

indirect fluorescent antibody test in which tissue impressions are used was

effective in detecting and diagnosing E. tarda in Japanese eels (Anguilla

japoniea).

3.3 Pathology Edwardsiella tarda

Fish infected with E. tarda sometimes become lethargic, "hang" at the surface,

and swim in a spiraling or erratic pattern. Gross external lesions vary with species.

Channel catfish often develop small, cutaneous ulcerations; in advanced cases,

however, larger depigmented areas mark the sites of deep muscle abscesses

(Meyer and Bullock 1973). The flounder Para/iehthys olivaeeus and the cichlid

Ti/apia nUotiea develop swollen abdomens due to ascites (Nakatsugawa 1983;

Kubota et al. 1981), and the bream Evynnis japonicus develops ulcers on the head

(Kusuda et al. 1977). Diseased common carp (Cyprinus carpio), Japanese eel, and

striped bass (Morone saxatilis) show hemorrhages on the body and fins (Miyazaki

and Egusa 1976b; Sae-Oui et al. 1984).

In eels, lesions on internal organs may perforate the body wall, and in striped

bass, epithelial hyperplasia sometimes gives the fish a tattered appearance.

Internally, the most common gross lesion consists of light-colored nodules on the

kidneys, spleen, or liver.

Histologically such lesions are focal necrotic areas, often with abundant bacteria,

both free and within macrophages. These lesions may be walled off by fibrocytes

and epitheloid cells, or be invasive and spread into adjacent skeletal muscle. Two

forms of the disease have been described from Japanese eels (Miyazaki and Egusa

1976a, b): in the more common form the initial lesions occur in the kidneys

(suppurative interstitial nephritis) and in the second form the liver is the primary

organ affected (suppurative hepatitis).

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Histopathology of internal organs is generally similar in Japanese eels, tilapia, and

striped bass. Tilapia sometimes also shows intestinal abscesses and gill

inflammation. Striped bass have epidermal hyperplasia and necroses (particularly

in the cephalic canals of the lateral line system) in which masses of E. Tarda may

occur. Large abscesses that develop in muscles of channel catfish and striped

mullet (MugU eepha/us), and in internal organs of Japanese eels emit a

malodorous gas when punctured.

3.4 Pathology Edwardsiella ictaluri

Channel catfish infected with E. Ictaluri refuse feed, tend to hang at the surface,

and swim with a spiral movement that includes erratic bursts. Gross external

lesions include hemorrhages around the mouth, on the lateral and ventral portions

of the body, and on the fins. Other signs include pale gills, exophthalmia, and

small ulcerations on the body. Ulceration in the fontanelle of the frontal bones

gives the disease one of its common names, "hole-in-the-head disease." Inter

nally, petechiae occur or develop throughout the visceral mass and in the

peritoneum and body musculature. Some fish develop ascites, and the liver,

kidneys, and spleen are commonly enlarged (Plumb and Schwedler 1982; Rogers

1983).

Danios (Dania devaria) infected with E. ictaluri swim erratically in a spinning

pattern, but gross lesions have not been observed in this species. Histopathology

has been described for both natural and experimental infections of channel catfish

(Areechon and Plumb 1983; Jarboe et al. 1984; Blazer et al. 1985). Chronic

natural infections are characterized by infiltrates of mononuclear cells that include

bacteria-laden macrophages, and diffuse necrosis and inflammation occur in

visceral organs.

Inflammation of the intestinal submucosa and mucosa is common. Blazer et al.

(1985) reported diffuse inflammation of the olfactory bulb and telencephalon, and

considered the nares a possible route of infection. Jarboe et al. (1984) detected no

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lesions in the brain but did not examine the olfactory tract. Areechon and Plumb

(1983) found necrotic lesions in the liver, spleen, kidneys, and pancreas of

channel catfish that had been injected with E. ictaluri; due to the acute course of

the experimental infection, the intestine did not become involved

.

3.5 Host and Geographic Range

Edwardsiella tarda has been isolated from many warm water fishes and some

coldwater fishes, whereas E. ictaluri has been isolated only from a few species of

warm water fishes (Table I). Additionally, E. Tarda causes disease in such other

animals as marine mammals, pigs, turtles, alligators, ostriches, skunks, and

snakes. It has also occasionally infected humans (Clarridge et al. 1980; Nagel et

al. 1982). In contrast, E. ictaluri is limited to fish, and survivors of epizootics

probably become carriers. The geographic range of E. tarda is worldwide,

whereas that of E. ictaluri is still confined to the catfish growing areas of the

United States (Rogers 1983).

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Table 1. Fish hosts of Edwardsiella tarda and Edwardsiella ictaluri.

Edwardsiella tarda

Atlantic salmon

Black skirted tetra

Brown bullhead

Channel catfish

Chinook salmon

Japanese eel

Emerald shiner

Hirame flounder

Goldfish

Grass carp

Largemouth bass

Striped mullet

Striped bass

Nile tilapia

Yellowtail

Edwardsiella ictaluri

Brown bullhead

Channel catfish

Dania

Green knifefish

Blue tilapia

White catfish

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Salmo salar

Gymnocorymbus sp.

lctalurus nebulosis

letalurus punctatus

Oncorhynchus tshawytscha

Anguilla japonica

Notropis atherinoides

Paralichthys olivaceus

Carassius auratus

Ctenopharyngodon idella

Micropterus sa/moides

Mugil cephalus

Morone saxatilis

Tilapia nilotica

Seriola lalandei

letalurus nebulosis

lctalurus punctatus

Danio devario

Eigenmannia virescens

Tilapia aurea

Letalurus catus

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3.6 Source and Reservoir of Infection

Because E. tarda is ubiquitous, many animals can serve as reservoirs of infection.

Furthermore, the environment can be a source of infectivity because this

bacterium survives as long as 76 days in pond water and mud (Ishihara and

Kusuda 1982; Minagawa et al. 1983). Fish that survive epizootics serve as carriers

and, because E. tarda is prevalent in the intestines of turkey vultures (Cathartes

aura), birds may also be an important reservoir of infection (Winsor et al. 1981).

Catfish that survive epizootics of E. ictaluri probably serve as reservoirs of

infection, since fish are the only known host and the bacterium survives less than

8 days in pond water (Rogers 1983).

3.7 Incubation Period

Incubation time is temperature related; channel catfish that were infected with E.

tarda and held at 27 DC died within 10 days (Meyer and Bullock 1973). In studies

at the National Fish Health Research Laboratory, striped bass held at 22 DC began

dying within 72 h after a 90-s bath exposure. Hawke (1979) reported that channel

catfish injected with E. ieta/uri died within 96 h, and that fish exposed to this

bacterium in aquarium water died within 2 weeks.

3.8 Control

Prevention

Because both E. tarda and E. ictaluri are principally pathogens of warmwater

fishes held in ponds, it is difficult to prevent disease outbreaks by following

specific management procedures. At present, E. ictaluri is more damaging than E.

tarda as a cause of mortality of cultured catfishes (J. A. Plumb, personal

communication).

Outbreaks of E. ictaluri infections occur at water temperatures of 24-28 °C, and

are thus restricted essentially to May-June and SeptemberOctober.

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Management procedures that reduce stress during these months may lessen the

severity of outbreaks. An experimental E. ieta/uri vaccine produced high titers in

channel catfish (Rogers 1983). Commercial production of vaccines for both

Edwardsiella pathogens is feasible.

3.9 Treatment

Outbreaks of E. tarda or E. ieta/uri can be controlled by feeding Terramycin at

the rate of 2.5 - 3.0 g/lOO lb of fish per day for 10 days. However, a strain of

Terramycin-resistant E. tarda from channel catfish was reported by Hilton and

Wilson (1980). Additionally, the potentiated sulfonamide Romet has proved

effective in controlling E. ieta/uri outbreaks, and the drug is in the process of

registration with the U.S. Food and Drug Administration for use on E. icta/ uri

infections in catfishes.

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CHAPTER IV

EDWARDSIELLA TARDA SEPTICEMIA WITH UNDERLYING MULTIPLE LIVER ABSCESSES

4.1 Abstract

Edwardsiella tarda has recently been described as a member of the family

Enterobacteriaceae. The genus Edwardsiella contains three species; E. hoshinae,

E. ictaluri and E. tarda. Edwardsiella tarda is the only species which has been

recognised as pathogenic to humans, especially in those with an underlying

disease. The most common presentation is watery diarrhoea. Extra intestinal

infections have been reported infrequently. Humans seem to be infected or

colonised with Edwardsiella through ingestion or inoculation of a wound. This

report is of a patient with multiple liver abscesses due to E. tarda who later

developed bacterial peritonitis and septicaemic shock.

Key words: Edwardsiella tarda, liver abscesses

4.2 Introduction

The genus Edwardsiella was first described by Ewing in 1965 and consisted of a

single species, Edwardsiella tarda, until 1980-1981 when two other species,

Edwardsiella hoshinae and Edwardsiella ictaluri, were added to the genus. E.

tarda is the most common of the three, and is the only species which has recently

been implicated in human disease. The organism is widely distributed in nature. It

is common in tropical and subtropical environments and appears to be spread by

contact with infected marine life, including ornamental fish and turtles, or by

eating raw fish.

E. tarda is an oxidase-negative, catalase-positive, facultative, anaerobic, motile,

Gram-negative bacillus. Certain biochemical properties are useful in

distinguishing E. tarda from other Enterobacteriaceae such as Salmonella and

Proteus species.

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The non-lactose fermenting colonies of E. tarda produce hydrogen sulphide and

indole but do not produce D-manitol, urease, oxidase and D-sorbital. E. tarda

causes illness in both humans and animals.

The asymptomatic carrier state is rare but documented. Humans are regarded as an

occasional host, and are prone to suffer from serious disease. E. tarda most

frequently causes gastroenteritis with acute watery diarrhoea3, but dysentery-like

presentations also occur.4 We recently encountered a case of multiple liver

abscesses complicated by peritonitis due to E. tarda infection without any

predisposing illness.

4.3 Case Report

A 27-year-old Indonesian male presented at the Accident and Emergency

Department of the Hospital Tengku Ampuan Afzan, Kuantan, Pahang with a two-

week history of fever, chills and rigors associated with right upper abdominal

pain. The fever was intermittent and was associated with generalised body

weakness. He had about four to five loose stools/day alternating with constipation

but gave no history of vomiting or yellow discoloration of the eyes. He was

admitted to the hospital for further investigation. He did not have any history of

chronic illnesses like liver disease, diabetes mellitus or renal problems. No other

co-worker living in the immediate environment suffered from a similar illness.

On examination, he looked ill and was drowsy and febrile, with a temperature of

37.80C, pulse rate of 112/minute, blood pressure of 110/70 mm Hg and

respiratory rate of 26/minute. He was dehydrated with mild pallor and jaundice,

but no lymphadenopathy was evident. His abdomen was distended with diffuse

tenderness but no rigidity. Bowel sounds were sluggish. Chest examination

revealed decreased air entry over the base of the right lung with bi-basal

crepitations. Both heart sounds were heard with no added sounds. There was no

neck stiffness and the musculo-skeletal examination was normal. A provisional

diagnosis of liver abscesses was made and he was treated empirically with

intravenous ceftazidime, metronidazole and intravenous fluids.

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Initial investigations revealed a haemoglobin level of 9.2 g/dl, a white blood cell

count of 12.3x109 /L with neutrophils 86%. His platelet count was normal. Blood

urea was raised (11.1 mmol/L), but serum creatinine was normal. Blood film for

malarial parasites was negative. Liver function tests were remarkable with a total

bilirubin of 24.3 μmol/L (direct 12.7 μmol/L, indirect 11.6 μmol/L), albumin 15.5

g/L, globulin 45.3g/L, alkaline phosphatase 516 U/L, alanine amino transferase

(ALT) 492 IU/L, aspartate aminotransferase (AST) 284 IU/L, prothrombin time

17.5 with INR 1.43, activated partial thromboplastin time 34.7 sec, glucose 6.3

mmol/L. Renal function was normal after hydration. Serology for hepatitis B and

C was negative. Chest X-ray showed raised right hemi-diaphragm with basal

consolidation. The blood culture grew E. tarda which was sensitive to Ampicillin,

gentamicin, Cefroxime, Cefperozone, Ceftriaxone and Ciprofloxacin. As a

consequence of this finding, the antibiotic was changed to Ampicillin 2 gm 6

hourly.

By the third day following admission, the patient had improved clinically and was

alert. However, his abdomen remained distended and was tender with sluggish

bowel sounds. Ultrasonography (Fig.1) revealed multiple well-defined

hypoechoic lesions in the liver. In view of the ultrasonography finding, a CT scan

of the abdomen was performed. This revealed multiple, well-circumscribed,

hypodense, cystic-like lesions disseminated in both hepatic lobes (see Figs. 2a and

b). Ultrasound guided drainage of liver abscesses was performed and 25 ml of

thick pus was drained, and sent for culture and sensitivity. A drain was inserted

and connected to a bag.

Stool examination revealed hook worm infection; no E. tarda was reported. Pus

culture from the liver abscesses did not grow any bacteria. The patient’s abdomen

distended further with very sluggish bowel sounds. A repeat CT scan of the

abdomen was done (Figs. 3a and b). This showed an increase in the free fluid in

the abdomen. An exploratory laparotomy was performed which revealed

substantial amounts of slough and pus in the peritoneal cavity with sections of

bowel adherent to one another. Multiple loculated abscesses were found in the

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liver. There was a collection of pus in the pelvic cavity. Drainage and peritoneal

toilet was performed. Post-operatively, the patient went into shock and was

treated in the intensive care unit.

He was ventilated and the antibiotics were changed to intravenous amoxicillin-

clavulanic acid and intravenous metronidazole. In spite of this, the patient

deteriorated subsequently and died of septic shock two weeks later.

4.4 Discussion

The most common manifestation of E. tarda infection is a gastrointestinal disease

causing watery diarrhoea, but cases of invasive enterocolitis4 have been reported

suggesting that this pathogen can invade cells to spread systemically and cause

tissue damage in vivo. Risk factors for E. tarda infections include exposure to

aquatic environments or exposure to exotic animals (e.g., reptiles or amphibians),

pre-existing liver disease, and dietary habits (e.g. ingestion of raw fish).

A number of serious, extra-intestinal infections have been reported such as

septicaemia with a mortality rate close to 50%,5 meningitis, peritonitis, septic

arthritis,6 myo-necrosis,7 tubo-ovarian abscess,8 liver abscesses and wound

infections.9 Humans seem to be infected or colonised with Edwardsiella through

either ingestion or inoculation of a wound. Although the gut is probably the portal

of entry in most cases of extra-intestinal infections, E. tarda has been isolated

only rarely from the stools of such patients.

Our patient presented with an episode of gastroenteritis which was the most likely

cause of his sepsis and multiple liver abscesses. He later developed bacterial

peritonitis most likely due to liver abscesses that might have spontaneously

ruptured into the peritoneal cavity before surgery. Stool culture failed to isolate E.

tarda as he had received antibiotics for over a week.

Open surgical drainage along with antimicrobial chemotherapy has long been

regarded as standard treatment for pyogenic liver abscesses.

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However, in many centres it is being replaced by percutaneous drainage under the

guidance of computed tomography or ultrasound. Percutaneous drainage has been

successful in patients with a single abscess, but the outcome has been less

favourable in those with multiple abscesses.

Open surgical drainage is reserved for patients in whom treatment fails or who

have complications. We initially performed percutaneous drainage in our patient.

Pus was sterile on culture, possibly because he had been on antibiotics for more

than one week. He had to undergo exploratory laparotomy due to development of

peritonitis and further deterioration. Several researchers have obtained satisfactory

results in selected patients with pyogenic liver abscess who received only medical

therapy. On review of the literature, we could trace only reported cases of liver

abscess due to E. tarda. Wilson and colleagues reviewed cases of serious

infections due to Edwardsiella tarda, two of which had liver abscess.

A 71-year-old Panamanian woman with liver abscesses had E. tarda isolated from

a specimen of blood as well as from pus obtained by needle aspiration of the liver

abscess. She died of septicaemia despite receiving prolonged antibiotic therapy. A

14-year-old female Nicaraguan immigrant to the United States presented with

septicaemia due to E. tarda. Exploratory laparotomy revealed a liver abscess,

which was drained. The pus revealed E. tarda on culture. The patient recovered

after antibiotic therapy.

From India, Koshi and Lalitha reported a case of liver abscess due to E. tarda in a

patient who had hepatoma.1 This patient died despite receiving antibiotics and

undergoing surgical drainage. Zighelboim and associates from Baylor College of

Medicine, Houston, Texas reported a case of multiple liver abscesses due to E.

tarda, which was successfully managed with antibiotic therapy alone.14 Of the

total five cases of liver abscesses due to E. tarda including our case, the overall

mortality was 60 % and most of the patients were treated by both drainage and

antibiotics.

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E. tarda is susceptible in vitro to a wide range of antibacterial agents.15 Most

strains of E. tarda are sensitive to Ampicillin, most lactam antibiotics, quinolones,

chloramphenicol, tetracycline, and aminoglycosides. Our isolates showed the

same pattern. Our patient received a variety of antibiotics and death was related to

consequences of sepsis.

Although extra-intestinal human infection with E. tarda has been reported

infrequently, the recent identification of the first such case at our hospital suggests

the need to consider such unusual pathogens in patients who present with febrile

diarrhoea and consequent bacteraemia. Early empiric therapy for infections may

prevent the isolation and recognition of E. tarda because of its susceptibility to

numerous antibiotics. Therefore blood cultures should be taken before giving

antibiotics.

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CHAPTER V

NATURAL ANTIBIOTIC SUSCEPTIBILITIES OF EDWARDSIELLA TARDA, E. ICTALURI, AND E. HOSHINAE

5.1 Abstract

The natural antibiotic susceptibilities to 71 antibiotics of 102 Edwardsiella strains

belonging to E. tarda (n = 42), E. ictaluri (n = 41), and E. hoshinae (n = 19) were

investigated. MICs were determined using a microdilution procedure according to

NCCLS criteria and German standards. All edwardsiellae were naturally sensitive

to tetracyclines, aminoglycosides, most β-lactams, quinolones, antifolates,

chloramphenicol, nitrofurantoin, and fosfomycin. Edwardsiella species were

naturally resistant to macrolides, lincosamides, streptogramins, glycopeptides,

rifampin, fusidic acid, and oxacillin.

Although slight species-dependent differences in natural susceptibilities to some

antibiotics (e.g., macrolides and cefaclor) were seen, differences in natural

susceptibility affecting clinical assessment criteria were only seen with

benzylpenicillin. Whereas E. tarda was naturally resistant to benzylpenicillin, E.

hoshinae was naturally sensitive.

Natural sensitivity and resistance to this penicillin were found among the strains

of E. ictaluri. The observed oxacillin sensitivity of E. ictaluri was attributed to the

failure of the species to grow at higher salt concentrations found in oxacillin-

containing microtiter plates.

The present study describes a database concerning the natural susceptibility of

Edwardsiella species to a wide range of antibiotics, which can be applied to

validate forthcoming antibiotic susceptibility tests of these microorganisms.

The genus Edwardsiella comprises a genetically distinct taxon weakly related to

other members of the Enterobacteriaceae. It consists of bacteria differing strongly

in their biochemical and physiological features, natural habitats, and pathogenic

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properties. The most common species of the genus is E. tarda, which was already

described in 1965 (8). Although it has been recovered from a variety of

environmental and animal sources (for a review, see reference 13), E. tarda is

predominantly found in freshwater and fish.

Humans are regarded to be occasional hosts but are prone to serious diseases due

to this organism. Most frequently, E. tarda causes gastroenteritis presenting as

acute watery diarrhea resembling that produced by other toxigenic

enteropathogens, but dysentery - like courses also occur. Immunocompromised

patients, older adults, and children are predominantly affected. Extraintestinal

infections such as septicemia—with a mortality rate near 50%—and wound

infections have also been reported . Exceptionally, E. tarda has also been found to

cause meningitis, peritonitis, osteomyelitis, and liver abscesses.

In 1980, a second Edwardsiella species was proposed by Grimont et al. and was

named E. hoshinae . In contrast to E. tarda, E. hoshinae is found in relatively few

ecological niches (i.e., birds, reptiles, and water). Although E. hoshinae has been

isolated from human feces, its role as a human or animal pathogen has not been

established. The third Edwardsiella species was created in 1981 and was called E.

ictaluri. E. ictaluri shows unusual properties: Apart from having a low optimal

growth temperature, this organism has been predominantly isolated from channel

catfish, in which it causes fatal systemic infections known as enteric septicemia.

Human infections due to E. ictaluri are not known; however, virulence-associated

properties such as serum resistance, indicating the potential to cause human

disease, have been documented for all Edwardsiella species.

The aim of the present study was to create a database concerning the natural

susceptibilities to a wide range of antibiotics of all known Edwardsiella species

originating from different areas and sources. Particularly, we investigated whether

there are species-related differences in natural antimicrobial susceptibility that

affect the clinical assessment criteria for the MICs.

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5.2 MATERIALS AND METHODS

5.2.1 Bacterial strains

A total of 103 strains labeled as E. tarda, E. ictaluri, or E. hoshinae originating

from European countries, Japan, and different areas in the United States were

examined. E. tarda strains were predominantly isolated from clinical specimens or

were taken from several fish species. All but one E. ictaluri strain derived from

channel catfish and E. hoshinae strains were mainly isolated from reptiles and

water. An overview of the origin of the Edwardsiella strains examined is shown in

Table Table1. 1 . Escherichia coli ATCC 25922 (derived from the Deutsche

Stammsammlung für Mikroorganismen und Zellkulturen, Braunschweig,

Germany) and Yersinia pseudotuberculosis ATCC 29833 (kindly provided by H.

Neubauer, Munich, Germany) served as controls for antibiotic susceptibility

testing.

5.2.2 Identification

All strains were identified to the species level with a commercial identification

system for Enterobacteriaceae (Micronaut-[MCN]-E; Merlin-Diagnostika,

Bornheim, Germany) and additional conventional tests. The inoculum for the

commercial test reactions was a suspension from an overnight culture on solid

medium in physiological saline solution at a concentration of 106 (E. tarda and E.

hoshinae) or 108 (E. ictaluri) CFU/ml. Regarding E. tarda and E. hoshinae,

incubation times for MCN-E tests were 24 h at 36 ±1°C. MCN-E tests for E.

ictaluri were read after 24 h at 25 and 36°C, 48 h at 25 and 36°C, and 72 h at

25°C.

Fermentation of trehalose and d-mannitol was tested on bromcresol purple agar

(Difco Laboratories, Detroit, Mich.) supplemented with trehalose (3 g/liter) and

mannitol (4 g/liter). H2S production was tested on triple sugar iron (TSI) agar

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(Merck, Darmstadt, Germany) and with the MCN-E test; citrate assimilation was

examined on Simmons citrate agar (Oxoid, Basingstoke, United Kingdom) and

with the MCN-E test. Agar plate tests were incubated at 36°C (E. tarda and E.

hoshinae) and at 25 and 36°C (E. ictaluri) and were read after 24, 48, and 72 h.

5.2.3 Antibiotics and antibiotic susceptibility testing

The natural susceptibilities to 71 antibiotics were investigated. All antibiotics

were kindly provided to Merlin-Diagnostika's disposal by their manufacturers.

The following concentrations were included: 0.01 to 32 mg/liter (for

benzylpenicillin, ciprofloxacin, sparfloxacin, ofloxacin, enoxacin, fleroxacin,

pefloxacin, lincomycin, clindamycin, rifampin, and fusidic acid), 0.03 to 64

mg/liter (for tetracycline, doxycycline, minocycline, oxacillin, cefuroxime,

cefotiam, cefoxitin, cefixime, cefpodoxime, cefdinir, cefoperazone, cefotaxime,

ceftibuten, ceftriaxone, ceftazidime, cefepime, imipenem, meropenem, aztreonam,

norfloxacin, erythromycin, roxithromycin, clarithromycin, azithromycin,

dalfopristin, quinupristin, dalfopristin-quinupristin, trimethoprim, and

vancomycin), 0.06 to 128 mg/liter (for gentamicin, netilmicin, tobramycin,

apramycin, ribostamycin, lividomycin, amoxicillin, amoxicillin-clavulanic acid,

ampicillin-sulbactam, pipemidic acid, teicoplanin, and chloramphenicol), 0.125 to

256 mg/liter (for amikacin, streptomycin, kanamycin, neomycin, spectinomycin,

piperacillin, piperacillin-tazobactam, ticarcillin, mezlocillin, cefaclor, loracarbef,

cefazolin, co-trimoxazole, nitrofurantoin, and fosfomycin, and 0.25 to 512

mg/liter (for azlocillin and sulfamethoxazole).

Antibiotic susceptibilities were tested by a microdilution procedure in Iso-

Sensitest broth (Oxoid) (used for E. tarda and E. hoshinae strains) and in cation-

adjusted Mueller-Hinton broth (CAMHB) (Difco) (used for E. ictaluri strains).

Six strains of each of E. tarda and E. hoshinae were also tested using CAMHB.

After inoculation of antibiotic-containing microtiter plates (Merlin-Diagnostika)

with 100 μl of the appropriate bacterial suspension (3 × 105 to 5 × 105 CFU/ml)

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and incubation for 20 h at 36°C (E. tarda and E. hoshinae) and for 48 h at 25°C

(E. ictaluri), MICs were determined with a photometer for microtiter plates

(Labsystems Multiscan Multisoft, Helsinki, Finland). MIC data were evaluated

with Excel (Microsoft).

5.2.4 Evaluation of natural antibiotic susceptibility.

Plotting the MIC of a particular antibiotic for one species against the number of

strains found with the respective MIC usually results in a bimodal distribution.

One peak with relatively low MICs represents the natural population, and one

peak with higher MICs represents the strains with acquired (secondary) resistance.

Analysis of the MIC distribution of all strains of one species for each antibiotic

permitted the determination of the biological thresholds, i.e., the thresholds which

limit the natural population at high MICs but not those strains with secondary

resistance.

We investigated whether the MICs for the natural population were above or below

the breakpoints of the standards used to assess clinical susceptibility. When the

natural population was sensitive or intermediate according to the cited standard, it

was described as naturally sensitive or naturally intermediate, respectively. When

the natural population was clinically resistant, it was described as naturally

(intrinsically) resistant. The method has been described in detail previously. In the

present study, breakpoints according to the American standard (NCCLS) valid for

Enterobacteriaceae, Pseudomonas aeruginosa and other non-Enterobacteriaceae,

Neisseria gonorrhoeae, and Staphylococcus species were applied. For antibiotics

for which NCCLS clinical assessment criteria do not exist, breakpoints according

to German, French, or Swedish standards were employed. Breakpoints for

ribostamycin, apramycin, and lividomycin were used as published recently.

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5.2.5 β-Lactamase testing

Two methods were applied to detect β-lactamase. All the strains were tested using

a conventional nitrocefin colony testing procedure (Carr-Scarborough

Microbiologicals, Inc., Decatur, Ga.). The tests were performed according to the

manufacturer's instructions. Four strains each of E. hoshinae and E. ictaluri were

also tested as described previously (29), with CAMHB as the medium. The latter

tests were performed in the absence of an inducer at temperatures of 36°C (E.

hoshinae and E. ictaluri) and 25°C (E. ictaluri); E. tarda ATCC 15947 served as a

positive control.

5.3 RESULTS

5.3.1 Identification

The identification of all but one of the received strains was confirmed. Although

the MCN-E system was able to identify Edwardsiella strains to the species level,

additional tests were helpful for discrimination. Apart from hydrogen sulfide

production, the examined strains showed the expected phenotypic properties. E.

hoshinae was metabolically the most active species, being able to ferment

sucrose, mannitol, and trehalose, and E. ictaluri showed some temperature-

dependent features, being metabolically more active with several substrates at low

temperatures (i.e., β-glucuronidase test, malonate and citrate assimilation,

ornithine decarboxylase test, and hydrogen production on TSI agar). Numerous

strains of each species were able to produce hydrogen sulfide, dependent on the

applied test and on the incubation time (and temperature for E. ictaluri). Classical

biovar 1 strains of E. tarda (hydrogen sulfide-negative and sucrose- and d-

mannitol-fermenting edwardsiellae) were not found. An overall view of the

phenotypic properties of the examined Edwardsiella strains is shown in Table

Table22.

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5.3.2 Natural antibiotic sensitivity and resistance

To most antibiotics there were only minor differences in natural susceptibility

among the species which were not affected by clinical assessment criteria. All

edwardsiellae were naturally sensitive to tetracyclines, aminoglycosides, most β-

lactam antibiotics, quinolones, antifolates, chloramphenicol, nitrofurantoin and

fosfomycin. Edwardsiella species were naturally resistant to macrolides,

lincosamides, streptogramins, glycopeptides, rifampin and fusidic acid. Species-

dependent differences in natural susceptibility affecting clinical assessment

criteria were seen with benzylpenicillin. Additionally, oxacillin susceptibility was

likely to be species-associated.

E. tarda was naturally resistant to benzylpenicillin and oxacillin, whereas E.

hoshinae was naturally sensitive to the former. E. ictaluri seemed to be highly

susceptible to oxacillin and was naturally sensitive and naturally resistant to

benzylpenicillin. An overall view of the antibiotic susceptibilities of E. tarda, E.

ictaluri, and E. hoshinae is shown in Fig. Fig.1. 1 . MICs are presented separately

for each species for which distinctive patterns were demonstrated. Natural

antibiotic sensitivities and intrinsic resistances are summarized in Fig. Fig.2. 2 .

5.3.3 Quality assurance

Apart from the MICs of tetracyclines, which were one or two dilution steps higher

in Iso-Sensitest broth than in CAMHB, there were no significant differences in

antibiotic susceptibility dependent on the medium (data not shown). Susceptibility

testing of E. ictaluri was only performed in CAMHB, because the species grows

poorly in Iso-Sensitest broth. The prolonged incubation time and the lower

incubation temperature used for the determination of MICs for E. ictaluri did not

significantly affect the MICs (data not shown). The MICs for E. coli ATCC 25922

in CAMHB and Iso-Sensitest broth were within the control limits for

susceptibility testing according to NCCLS criteria (22) (data not shown).

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Penicillin MICs for Y. pseudotuberculosis ATCC 29833 (the MIC range of

benzylpenicillin was 0.5 to 1 mg/liter) were in agreement with the data of a

previous study (31).

5.3.4 β-Lactamase testing

All strains of E. tarda gave weakly positive or positive results for β-lactamase

production using nitrocefin β-lactamase disks. No strain of E. hoshinae or E.

ictaluri exhibited any detectable β-lactamase activity. The latter results were also

obtained with the second procedure applied. β-Lactamase activity of E. tarda

ATCC 15947 was slightly enhanced at 36°C (data not shown).

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CHAPTER VI

CONCLUSION

The role of the genus Edwardsiella in human illness is reviewed. Of the three

recognized species, only Edwardsiella tarda has been demonstrated to be

pathogenic for humans. Chief infections associated with this species include

bacterial gastroenteritis, wound infections such as cellulitis or gas gangrene

associated with trauma to mucosal surfaces, and systemic disease such as

septicemia, meningitis, cholecystitis, and osteomyelitis. Risk factors that are

associated with E. tarda infections include exposure to aquatic environments or

exotic animals (e.g., reptiles or amphibia), preexisting liver disease, conditions

leading to iron overload, and dietary habits (e.g., raw fish ingestion).

Although studies indicate that this bacterium is susceptible to most commonly

prescribed antibiotics, fatal gastrointestinal and extraintestinal infections have

been described.

Edwardsiella species were naturally resistant to macrolides, lincosamides,

streptogramins, glycopeptides, rifampin and fusidic acid. Species-dependent

differences in natural susceptibility affecting clinical assessment criteria were seen

with benzylpenicillin. Additionally, oxacillin susceptibility was likely to be

species-associated.

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[ ] November 8, 2012

BIBLIOGRAPHY

http://www.whonamedit.com/synd.cfm/3123.html

http://www.rightdiagnosis.com/medical/edwardsiella.htm

http://digitalcommons.unl.edu/cgi/viewcontent.cgi?

articleuXsWTAwfzJJQg#search=%22edwardsiella%22

http://www.mjpath.org.my/past_issue/MJP2006.1/07Liver%20absecess.pdf

http://etd.auburn.edu/etd/bitstream/handle/10415/894/

ZHANG_YINFENG_23.pdf?...

http://www.vumicro.com/vumie/help/VUMICRO/Edwardsiella_hoshinae.htm

http://en.wikipedia.org/wiki/Edwardsiella_ictaluri

http://en.wikivet.net/Edwardsiella_ictaluri

http://www.bacterio.cict.fr/e/edwardsiella.html

http://europepmc.org/articles/PMC3349661/

reload=0;jsessionid=2fCesajF1wvAUJMraUmV.4

LIST OF PICTURE

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[ ] November 8, 2012

Mixture of three biotypes of Edwardsiella tarda on John L's MacConkey-based

"ET Agar."

Transmission electron micrograph if Edwardsiella ictaluri strain 93-146.

Scanning electron micrograph if Edwardsiella ictaluri strain 93-146.

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[ ] November 8, 2012

Adult channel catfish displaying ascites, one of the clinical signs associated with

the acute form of ESC. Other external lesions include white punctate spots on the

skin, petechial hemorrhages, and exophthalmia

Petechial hemmorrhages on the ventral abdomen of an adult channel catfish with

ESC.

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[ ] November 8, 2012

Gross internal lesions associated with acute ESC. Hemorrhagic ascites,

dark macropapular lesions on the liver, and splenomegaly are visible on this fish.

Other internal lesions include petechial hemorrhages on the liver, intestine, and

abdominal serosa, and renomegaly.

OF CENDRAWASIH UNIVERSITY 43