Cumitech 37: Laboratory Diagnosis of Bacterial and Fungal Infections Common to Humans, Livestock, and Wildlife

Laboratory Diagnosis of Bacterial and Fungal Infections Common to Humans, Livestock, and Wildlife TTONE, GARY D. CAGE, M...

Author: Thomas J. Inzana

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Laboratory Diagnosis of Bacterial and Fungal Infections Common to Humans, Livestock, and Wildlife TTONE, GARY D. CAGE, MAY C. CHU, DIANE FABIAN, BRUCE HANNA, FRANK HOLLIS, TAO HONG, THOMAS J. INZANA, HASSAN NAMDARI, AND ROBBIN S. WEYANT COORDINATING

EDITOR

THOMAS J. INZANA

Cumitech

CUMULATIVE TECHNIQUES AND PROCEDURES IN CLINICAL MICROBIOLOGY

Cumitech 1B Cumitech 2B

Blood Cultures 111 Laboratory Diagnosis

Cumitech Cumitech Cumitech Cumitech

3A 5A 6A 7A

Quality

Cumitech Cumitech Cumitech Cumitech

12A 13A 16A 18A

Cumitech 19A Cumitech 21 Cumitech 23 Cumitech 2-l Cumitech 25 Cumitech 26

Control

oi Urinary

and Quality

Tract Infections

Assurance Practices in Clm~cal 1licrobiolog!-

Practical Anaerobic Bacteriolog! New Developments in hntimicrobial Agent Susceptibility Testing: a IVactiial Laboratory Diagnosis of Lower Respiratory Tract Infections Laborator\- Diagnosis of’ Bacterial Diarrhea Laborator)- Diagnosis of’ Ocular Infections Laboratory Laboratory

Diagnosis Diagnosis

Guide

of’ the .1Iycobacterioses of Hepatitis Viruses

Laborator! Diagnosis of C/?l;imy~L7 t~~7c/mm~7tis Infections Laborator!- Diagnosis of \-irA Respiratory Disease Infections of the Skin and Subcutaneous Tissues Rapid Detection of \‘iruses by Imlnilnofluorescer7ce Current Concepts and Approaches to ;\ntimicrobial A1gent Susceptihilit!. Tf,ting Laboratory Diagnosis of \‘iral Infections Producing Enteritis Laboratory Diagnosis of Zoonotic Infections: Bacterial Infections Obtained from Companion oratory Animals

,~nd I ‘xl>-

Cumitech

27

Cumitech

28

Laboratory Diagnosis ot’ Zoonotic Infections: Chlamydial, mined from Companion and Laboratory Xnimals

Cumitech Cumitech

29 30

Cumitech

31

Cumitech

32

Cumitech

33

Laboratory Safety in Clinical Alicrohiolog! Selection and Use ot’ Laboratory Procedures for Diagnosis of Parasltlc Infections of the (~,~strointestin,l Tract Verification and \.‘alidation of Procedures in the Clinical .\licrobiolog! Laborator!Laboratory Diagnosis of Zoonotic Infections: \kal. Rickettsial, and Parasitic Int~
Cumitech 34 Cumitech 35 Cumitech 36 Cumitech 37

Fungal.

\.iral.

and I’,lrasitic

Postmortem ,\Iiirohiolog! Biosafet!- Considerations for Large-Scale Production of .\licroorganism~ Laborator!- Diagnosis of Bacterial and Fungal Infections Common to Humans,

Infections Ob

I.~\estock. and Wildlife

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Reserved

Laboratory Diagnosis of Bacterial and Fungal Infections Common to Humans, Livestock, and Wildlife Edward The Mount

Sinai Hospital,

Mount

Sinai School

of Medicine,

Bottone

One Gustave L. Levy Pl., New York, NY 10029

Gary D. Cage Arizona

Department

Infectious

Disease

of Health Microbiology

Services,

Bureau

Section,

2520

of State Laboratory W. Adams,

Phoenix,

Services, AZ 85007

May C. Chu Division

of Vector-Borne Diseases, Centers

Infectious for Disease

Diseases, National Center for Infectious Control and Prevention, P.O. Box 2087, Ft. Collins, CO 80522-2087

Diane Fabian Department

of Pathology,

Hackensack

University

Medical

Center, 30 Prospect Ave., Hackensack, NJ 07601

Bruce Hanna Department

of Pathology,

Belleuue

Hospital,

NYU

School of Mediczne, New York, NY 10016

Frank Hollis Department

of Pathology,

Hackensack

University

Medical

Center, 30 Prospect Ave., Hackensack, NJ 07601

Department

of Pathology,

Hackensack

University

Medical

Center, 30 Prospect Ave., Hackensack, NJ 07601

Tao Hong

Thomas J. Inzana Center for Molecular Medicine Regional College of Veterinary

and Infectious Diseases, Virginia-Maryland Medicine, Virginia Polytechnic Institute and State Unzuersity, Blacksburg, VA 24061

Hassan Namdari Clinical

Laboratories,

Inc., 901 Keystone

Industrial

Park,

Throop,

PA 18512

Robbin S. Weyant Special Branch,

Bacteriology Reference Laboratory, Meningitis and Special Pathogens National Centers for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

COORDINATING Center for Molecular Medicine Regional College of Veterinary

EDITOR: Thomas J. Inzana

and Infectious Diseases, Virginia-Maryland Medicine, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061

Introduction ... . ..................................................................................... Bacterial Infections ............................................................................... Eryslpelo

thrix

rhuslopa

thlae ................................. 1

.......................................................

2 2 2

2

Bottone

et al.

CUMITECH

37

Group C Streptococci ................................................................................................. 1 is teria monocytogenes .............................................................................................. Rhodococcus equi ...................................................................................................... Nontuberculous Mycobacterium spp. ........................................................................... Brticella spp. ............................................................................................................ Burkholderia pseudomallei and Burkholderia mallei ...................................................... Francisella tularensis ................................................................................................ Leptospira spp. ........................................................................................................ Yersinia pestis ......................................................................................................... Rare, Miscellaneous Bacterial Agents .........................................................................

5 7 8 9 11 13 16 20 23 28

Fungal

29

Infections

Dermatophytes

References

................................................................................

........................................................................................................

.....................................

INTRODUCTION nfectious diseasescommon to humans and animals have been prevalent since humans domesticated I wild animals and began using them for food, clothing, and companionship. Over 100 years ago Koch, Loeffler, Pasteur, and others investigated the causesof infectious diseasesof animals, some of which also occurred in humans or were transmissible to humans. While some bacterial infections associated with animals are common, such as those due to Salmonella enterica, Campylobacter jejuni, and someserotypes of Escherichia coli, many infections are rare, may be associated with trauma or bite wounds, or may be obtained from the same sources as animal infections. However, the numbers of such affected individuals have increased significantly due to infection with human immunodeficiency virus (HIV) or human T-cell lymphotrophic viruses, cancer chemotherapy, bone marrow transplantation, and other medical treatments resulting in severe suppression of the immune response. As a result, infections with zoonotic and environmental pathogens are becoming more common, and the accurate identification of these agents becomesmore critical to the management and care of patients with these infections. Cumitech 32 described the diagnosis of zoonotic infections due to viral, rickettsial, and parasitic agents associatedwith food animals and wildlife (83). The intention of this Cumitech is to review the habitat, description, diseases,diagnosis, and treatment of some of the bacteria and fungi commonly found associated with livestock and wildlife that may also cause infections in humans. In addition, some infectious agents that are not directly transmissible from animals to humans but that are closely associatedwith animals, for which animals act asa reservoir, or that causeinfections in both animals and humans will also be described. Although susceptibility to antimicrobial agents is described, it must be emphasized that standardized antimicrobial susceptibility tests have not been established for most of these agents. The antimicrobial susceptibilities that have been reported should be considered as a guide only.

. .................................

29

. . .................

31

For applicable agents, a brief description of laboratory safety and precautions is also provided. Agents that are commonly isolated in the clinical microbiology laboratory, though they may be zoonotic in nature, will not be described here (e.g., Salmonella). This Cumitech is not intended as a comprehensive text on zoonotic or animal-related infections. Rather, this Cumitech focuses on the description and diagnosis of those agents that may causeinfections in both humans and animals and be isolated only rarely or cause infections under specific circumstances. Therefore, these agents may be misidentified or not identified at all if laboratory personnel are not familiar with their characteristics. A brief discussion of the disease (if any) in animals is also included, as animals may serve as sentinels for some diseasesand offer important clues in diagnosis when an adequate clinical history is not known for the patient. A summary of each group of organisms, their natural host, diseasein animals, mode of transmission, diseasein humans, diagnosis, antibiotic susceptibility, and laboratory safety level is given in Table 1.

BACTERIAL Erysipelothrix

INFECTIONS rhusiopathiae

Description, Natural Habitat, and Diseasein Animals First described by Koch in 1878, E. rhusiopathiae was one of the earliest recognized agents of bacterial zoonosis (139). In 1886 LoeMer identified this organism as the agent responsible for swine erysipelas, a disease of significant economic importance (148). Its pathogenicity in humans was first establishedby Rosenbach in 1909 (205). Excellent reviews of the epidemiology, bacteriology, and clinical significance of E. rhusiopathiae have been published by Gorby and Peacock in 1988 (100) and by Reboli and Farrar in 1989 (199). E. rhusiopathiae is a facultatively anaerobic, grampositive, thin, pleomorphic, non-spore-forming rod or filament. Individual cell sizes range from 0.2 to 0.4 Frn in diameter and 0.8 to 2.5 pm in length. Filament

Table

1.

Summary

lnfectlous agent(s)

of selected

zoonotic

Reservorrs mammals, fish

thlae

Swine, birds,

E rhwopa

occaslonally anrmals

agents

that or In host

are associated

Diseases outcome(s) Septlcemra, skin lesions, endocardrtrs, arthritis Resprratory InfectIons, strangles, abortion, septicemia Septrcemra, abortron, menrngoencephalltrs, necrosis of internal organs

with

food

Mode(s) transmission

animals, of

tract

Contact with Infected urine, feces, saliva, or tissues Aerosol, direct contact, trauma Gastrorntestrnal

herbr-

contact wrth or soil

Horses, other

C streptococcr

other

Aerosol, animals

organ

Foal pneumonia, lymphadenrtrs, abscesses

Group

Horses, vores

direct

bite,

Contact spores

Flea

with

lnfectrous

aerosol

Tick bites, contact wrth or ingestion of infected tissue, laboratory aerosol Exposure to Infected urine, milk, or tissues

Consumption of contaminated dart-y products, contact with Infected tissues Aerosol, direct contact with Infected tissues

contact,

animals,

cattle, sheep, animals

Myco-

Birds, marine mammals

Aerosol, wounds

Swine, other

spp

Systemic and cutaneous infections, granulomas, lymphadenopathy Abortion, stenllty

L monocytogenes

R equl

spp

Nontuberculous bacterium

Brucella

Systemic and pulmonary lesions, ulceration of lymph nodes, death

Cattle, goats, swine, dogs, camels, wild deer, horses

B. pseudomallel B. malIe/

Soil (B pseudoma//e/), horses, other equines

and

F. tularensls

Systemic, lesions;

plague-like death

Rabbits, beavers, muskrats, voles, sheep, many others

Abortion, kidney tion, systemic tion, death

spp

Leptosplra

Wide variety, lncludlng birds and reptiles

Buboes, lessons In liver and spleen, pneumonia, death Alopecra, erythema

rnfecinfec-

Y. pest6

Rodents (rats, ground squirrels, chipmunks, voles) Various mammals, birds, sod

Dermatophytes

horses,

and wildlife

Diseases or outcome In humans

Confirmation infection

of

producstain

Culture, tion,

H,S Gram

Skin lesions, endocardltrs, meningitis

Culture, detection of group C antigen, blochemical tests Cold enrichment culture, tumbling motrlrty at 20°C, blochemlcal tests

Containment level and other recommendations

BSL-2, no special cautions

Treatment

Penlcrllln

BSL-2

G, others

Penicillin, vancomycin, cephalosporrns, SXT

BSL-2

amrno-

Penicillin plus glycoslde

pre-

BSL-2 for cllnlcal speclmens, BSL-3 for cultures BSL-2

BSL-2 for cllnlcal specrmens, BSL-3 for work with pure cultures; avoid aerosol BSL-2, use caution and gloves with Infected fluids

BSL-2 for clrnrcal speclmens, pure cultures should be handled under BSL-3 BSL-2 for cllnrcal specrmens, BSL-3 for handling cultures

BSL-2

BSL-2

+ nfampln + gentamrcln

Erythromycrn + nfampin, amrnoglycosldes, vancomycin, ciprofloxacin Variable susceptrbrlrty to tuberculosis drugs

Doxycyclrne or SXT

Streptomycin, gentamitin, fluoroquinolones, tetracycline

SXT + ceftazrdrme, tetracycline, broad-spectrum cephalosponns, amlnoglycosrdes

Gram stain of abscesses, culture (not recommended If facrlltres are rnadequate), serology Culture, fluorescence assay of tissue, serology

toprcal terblnaflne

Streptomycin, gentamitin, tetracyclines

doxycyclrne

Tolnaftate, azoles,

Penrclllrns,

Culture and dark-field exam of blood, CSF, and urine, silver start-r or DFA of tissues; serology DFA, PCR, culture, bacteriophage lys~s, anlmal inoculation Culture, Txhophyton agar

Culture, brochemrcal tests, hrgh-performance lrqurd chromatography Blood culture, brochemical tests, serology

Culture, pinkish colony morphology, may be partrally acid-fast

Sepsis, cellulitis, pneumonia, abscesses, pharyngltls Neonatal InfectIons, central nervous system infections, sepsis, focal infections of various organs Vanable. pneumonia, cutaneous infections, septrcemra, lymphadenitis Systemic, pulmonary, soft tissue, and cutaneous infections

sep-

Systemic infections, splenomegaly, hepatomegaly, encephalrtls

Pulmonary lesions, ticemia, drssemlnated abscesses

Ulceratron and lymphadenrtls, sepsis, pneumonia, flu-like Illness, death Flu-like Illness, liver and kidney InfectIon, death

Buboes, bacteremla, pneumonia, skin lesions, death Erythema, hyperkeratosis, alopecia

Bottone

et al.

length may exceed 60 pm (13 1). E. rhusiopathiae cells decolorize readily and may appear gram negative. When grown on plated medium, two colony types may be observed: a smooth type, which contains predominately rods, and a rough type, which contains predominately filaments (248). Molecular taxonomic studies (228, 229) indicate that the genus Erysipelothrix contains two named species, E. rhusiopathiae and E. tonsillarum, along with at least two unnamed genomic groups. E. tonsillarum includes strains formerly designated E. rhusiopathiae serovars 3, 7, 10, 14, 20, 22, and 23 (228). This species, which was first isolated from tonsils of healthy pigs, is considered to be less pathogenic than E. rhusiopathiae. Human infections attributed to this species have not been reported. In addition to differences in swine virulence and serovar grouping, E. tonsillarum can be differentiated from E. rhusiopathiae by production of acid from saccharose. E. rhusiopathiae ha s been found as a commensal or pathogen in a wide variety of mammals, reptiles, birds, and fish throughout the world. Domestic swine constitute an important natural reservoir for the organism. Symptomatic and nonsymptomatic animals may shed organisms through urination or defecation. Although this organism cannot persist in the environment indefinitely, soil that has been contaminated with E. rhusiopathiae can yield positive cultures for many weeks. Viable organisms have been recovered from a buried carcass after 9 months of interment. Low temperature, alkaline pH, and the presence of organic matter all favor its survival. Contaminated surface water, rodents, wild birds, and insects have been suggested as vectors for the dissemination of this organism in the environment (13 1). In swine, infections with E. rhusiopathiae may be endogenous or exogenous. Septicemia is followed by large numbers of bacteria being shed in the urine, feces, and saliva. Acute disease results in high fever, conjunctivitis, and vomiting. Purple lesions may occur on the skin, particularly the ears, abdomen, and legs. Lymphadenopathy and splenomegaly are common, as well as hemorrhagic lesions in the stomach, intestines, and kidneys. Chronic forms of the disease are usually associated with endocarditis and arthritis. In cattle arthritis is the most common malady. In domestic birds, septicemia and associated hemorrhages of the muscles in the legs, breast, and serous membranes occur. The liver, spleen, and alimentary tract may also be affected. Septicemia and systemic infections have also been reported in reptiles and fish (238). Mode of Transmission The most common mode of transmission of E. rhusiopathiae to humans is through direct contact with contam .inated organi c matter, particularly tissues and excrement of swine, fish, or poultry. Occupations

CUMITECH

37

associated with significant risk of infection include butchers, abattoir workers, fish handlers, and homemakers. Transmission of the agent to fish handlers and homemakers is particularly high due to abrasions caused by the handling of spines, fins, and fish bones. Laboratory workers and veterinary pathologists are also at higher risk of infection due to occupational exposure (198). Human Infections The most common clinical manifestation of E. rhusiopathiae infection in humans is a localized skin lesion, or erysipeloid. Within 2 to 7 days after infection (usually resulting from handling contaminated materials with lacerated or abraded hands or fingers), localized pain and swelling develop in the infected area. The resulting lesion is well-defined, slightly raised, violaceous in color, and nonsuppurative. This -type of i.nfection usually resolves without treatment within 3 or 4 weeks. Diffuse cutaneous and systemic infection with endocarditis are two other rarely encountered clinical syndromes of E. rhusiopathiae infection that have been described previously (199). The diffuse cutaneous manifestation is characterized by a proximal progression of the locali zed lesion from the original site of infection and by the appearance of other lesions at remote areas. Although fever and arthralgia are common, blood cultures are negative. Systemic infection with endocarditis represents a significant clinical challenge. A review of 60 cases by Gorby and Peacock reported a mortality rate of 3 8 %, with complications including congestive valve failure, myocardial abscesses, aortic valve perforation, meningitis, and glomerulonephritis ( 100). Diagnosis Skin biopsy samples or tissue aspirates are the specimens of choice for isolating E. rhusiopathiae from erysipeloid lesions. Successful cultivation requires sampling the entire thickness of the dermis, because the organisms tend to localize in the deeper parts of the skin (199). E. rhusiopatbiae grows on selective media for gram-positive bacteria, such as colistinnalidixic acid agar or phenylethanol agar (62). For systemic infections, the bacteria can successfully be cultivated from blood using standard techniques. Biochemical data for 81 strains identified as E. rhusiopathiae by the Special Bacteriology Reference Laboratory at the Centers for Disease Control and Prevention (CDC) in Atlanta, Ga., are given in Table 2. The most common source of these strains was human blood (47%), followed by human wounds (leg, toe, hand, and finger) (17%). Other human sources (chest tube, heart tissue, heart valve, liver, urine, and vaginal exudate) accounted for 14 %. Nonhuman sources (turkey liver, quail blood, mouse, fish,

CUMITECH

37

Table 2. Biochemical E. rhusiopathiae”

Diagnosis analysis

of 81 strains

of

Motlllty . . . . . . . . . 0 Acid from: . . . *.....,. 85 (15) Glucose . . . . . . . . . . ..a.........-. .*....... . . *......a.. Xylose . . . . . . . . . . .*.......... . . . ..***.**..........*....*.. ** *...,,,,* 0 ............ Mannrtol ............ .............. . ................. 0 Lactose ................. ....... .. ......*.. .. .. 85 (15) Sucrose . . . . . . . . . . . . . . . . . . , . , , . , , . . . . . *.........*......m......... . 0 Maltose . . .................. .... ....*. . ..**.,* IO (7) Nutrient broth, 0% NaCl . . . . . . . . . . . 53 (1) Nutrient broth, 6% NaCl . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . .. . . . . 9 Catalase . .............. . . . . . . . . . . . . . . . .. . . ...**.*..*......*....... 0 Oxldase . . . . . . . . . . . . . . . *..*..............*...*... . , . ..*.. . . . . 6 W Growth on MacConkey agar * ..*............ . .. . .. . . . . . * ..*..* 0 Esculln hydrolysis ....... . ......................... 0 0 Simmons citrate .............. ................................ .... Urea, Christensen’s ...................... ....................... 0 Nitrate reduction .... ........................... ............ 0 lndole . . . . . ..*....*.*.... . . . . . ..*.....*.....s......... . . . . . . . . . . ., 0 TSI slant, acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ...*..*. 98 TSI butt, acid. . . . . . . . . . . . . . . . . . . . . . . 98 H,S (TSI butt) . . ..a..... . . . : . . . * : ::.:..: . . . ..a.. . 99 Gelatin hydrolysis” . . . . . . . . . . . . .. . .. . . . . . . . . . . . . .:...:.I . . . . 0 Growth at: 25°C . . . . . . . . . . . . . . . . ...*... *.......................a. . . . . . . . . . . 82 35°C.. . . *. . . . . * . . . . , . . ,. .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ...* . 100 42°C . . . . . . . . . . . . . . . . . . . . . . . . ..*.........*........... .* 32 a Not all strains were tested In each test b Numbers In parentheses Indicate percentages 7 days of lncubatlon w, weak reaction ‘At 7 to 14 days of Incubation

of posltlve

Infections

5

growth pattern that is observed when stab cultures in gelatin medium are incubated at 25OC. % of tests posrtrve at 2 days of rncubatlor?

Characteristic

of Zoonotic

tests

at 3 to

pig joint, and bovine bladder) accounted for 16%. Biochemical testing was done using published methods (253). Carbohydrate reactions were performed in enteric fermentation base with Andrade’s indicator. Positive reactions include acid production from glucose and lactose, acid production in the slant and butt of the triple sugar iron (TSI) agar slant, and H,S production in the butt of the TSI agar slant. The production of H,S is a characteristic that differentiates E. rhusiopathiae from essentially all other grampositive aerobic or facultatively anaerobic bacterial species encountered in the clinical microbiology laboratory. The occasional H,S-positive Bacillus or Streptococcus isolate can be differentiated from E. rhusiopathiae by Gram stain and spore morphology. Acid is produced from lactose, but not from xylose, mannitol, and sucrose. Reactions are negative for catalase, Simmons citrate, urease, nitrate reduction, indole production, and esculin hydrolysis. Most of the strains studied were either negative for hemolysis or were alpha hemolytic; however, two strains were betahemolytic, and two strains lysed the erythrocytes without hemoglobin clearing. Although this species has been described as not able to grow in 6% NaCl (13 l), we have encountered seven strains that were positive for this characteristic. Another distinctive characteristic of this organism is the “bottle brush”

Antibiotic Susceptibility and Treatment Penicillin G (12 million to 20 million U per day) is the drug of choice for serious E. rhusiopathiae infections (199). Additional antibiotics to which this organism is susceptible include cephalosporins, clindamycin, imipenem, and ciprofloxacin (230, 245). Antibiotics to which this organism is resistant include sulfonamides, trimethoprim-sulfamethoxazole (SXT), novobiocin, teicoplanin, aminoglycosides, and vancomycin. Variable susceptibility has been reported for chloramphenicol, erythromycin, and tetracycline (230,245). Group

C Streptococci

Description, Natural Habitat, and Disease in Animals The organisms classified as group C streptococci include Streptococcus dysgalactiae subsp. equisimilis, S. dysgalactiae subsp. dysgalactiae, Streptococcus equi subsp. equi, and S. equi subsp. xooepidemicus. These organisms carry the Lancefield group C carbohydrate antigen in their cell walls and are commonly called large-colony-forming streptococci within the group. Another group of heterogeneous group C streptococci classified as the “Streptococcus anginosus or milleri” group also carries the group C antigen (as well as the groups A, F, and G antigens) but is known as smallcolony-forming Streptococcus. However, members of the ‘3. anginosus or miller? group are not zoonotic and are described here only due to their group antign similarity. Group C streptococci are beta-hemolytic, grampositive, catalase-negative, facultatively anaerobic organisms that are pyogenic by nature. Colonization or infection by the large-colony-forming group C streptococci is often associated with exposure to animals and animal products. S. dysgalactiae subsp. equisimilis, S. equi subsp. equi, and S. equi subsp. zooepidemicus primarily colonize the upper respiratory tract of horses. S. dysgalactiae subsp. equisimilis is considered the most common group C streptococcal species to colonize and cause infection in humans (75, 108, 121). S. equi subsp. xooepidemicus is reported to be a common pathogen of domestic animals (primarily horses, but also cattle, sheep, and pigs) but an uncommon human pathogen (16, 18, 222). S. equi subsp. equi is primarily a pathogen of horses, and S. dysgalactiae subsp. dysgalactiae is an uncommon human pathogen (30, 39, 193). Animal infections caused by these opportunistic pathogens include postviral respiratory infection, strangles (distemper), and abortions in horses; septicemia, arthritis, and endocarditis in pigs, and mastitis in cows (90).

6 Table

Bottone 3.

Differential

characteristics

of Lancefield

Colony size

Hemolysls Beta

Large

Alpha or none Alpha, beta, or none a Most strains b Varlable

CUMITECH

et al.

Large Small

group

C streptococci

Species S. S S. S S.

37

equi subsp equl equl subsp zooepldemicus dysgalactlae subsp equisimilis dysgalactlae subsp dysgalactlae anglnosus or miller/ group

Acetoln Voges-Proskauer +

Salicin + ha +

or acid formatlon Trehalose + + +

Sorbltol + -

Lactose Cb + +

are poslttve

Mode of Transmission Transmission may occur through the respiratory tract, skin contact with infected animals or tissues, or trauma (e.g., wound). Most human infections due to group C streptococci are community acquired and are secondary to underlying diseases such as cardiovascular diseases, malignancy, and immunosuppression (30). Human Infections In humans, S. dysg&ctide subsp. equisimilis causes various types of infections, including sepsis in neutropenic hosts, puerperal sepsis, cellulitis, postoperative wound infections, pneumonia, empyema, brain abscess, meningitis, osteomyelitis, septic arthritis, and endocarditis (17,30,173,268). It has been recovered from the pharynges of carriers and from individuals with exudative pharyngitis and tonsillitis as well as from intra-abdominal abscesses (6, 141). S. equi subsp. xooepidemicus has been implicated in outbreaks of human pharyngitis (18, 141). Outbreaks have been traced back to an animal source such as homemade cheeses (82) and unpasteurized cow milk (74,89) -the latter resulting in poststreptococcal glomerulonephritis in some patients (89). A case of a renal transplant recipient with S. equi subsp. zooepidemicus cellulitis has been reported (19, 72, SZ), as well as septicemia and meningitis in adult patients (86, 149, 173, 274) and endocarditis (159). S. equi subsp. equi has been implicated in bacteremia in a human in a single reported case secondary to a vulvectomy (73). A single case report of meningitis caused by S. dysgalactiae subsp. dysgalactiae in a premature infant has been reported (193). The “S. anginosus or miller? group can be isolated from pyogenic infections, notably abscesses. They reside in the pharynx as commensal bacteria and are probably not agents of pharyngitis (209). The members of the “S. anginosus or milleri” group are the most common group C beta-hemolytic streptococci isolated from the human throat and can also be a causative agent in bacteremia (48, 87). Diagnosis Group C streptococci are catalase-negative, spherical or ovoid, gram-positive cocci that are less than 2 Frn in diameter. They are facultatively anaerobic organisms that grow on conventional laboratory media

containing 5% sheep blood agar (BA). Optimal temperature for growth is 37OC. The colony size of largecolony-forming strains of group C streptococci is >0.5 mm in diameter, while those in the “S. anginosus or milleri” group are pinpoint, having diameters of ~0.5 mm. On BA, group C streptococci are beta hemolytic with the exception of S. dysgalactiae subsp. dysgalactiae, which produces alpha hemolysis or no hemolysis at all. The ‘?!3. anginosus or milleri” group can produce alpha, beta, or no hemolysis. Colonies of group C streptococci may vary in color, ranging from clear to gray to white with a glistening translucence. Highly encapsulated strains may be mucoid. Isolates of the small-colony-forming beta-hemolytic streptococci often produce a distinct odor described as buttery or carmel-like (209). Primary criteria for identifying these organisms are the presence of the Lancefield group C antigen. Several different identification products (i.e., Streptex [Murex Diagnostics, Norcross Ga.], Streptolex-OD [Orion Diagnostica, Somerset, N. J.], PathoDx [Diagnostic Products Corp., Los Angeles, Calif.], etc.) are commercially available for this purpose. The PathoDx test has been shown to rapidly detect group C streptococci in specimens from animals (124). Since the Lancefield group C antigen is not specific to a single type of Streptococcus, additional carbohydrate fermentation tests can be used to further identify the large-colony-forming organisms. Commercial systems such as Vitek (bioMerieuxVitek, Hazelwood, MO.), Microscan (Dade International, Miami, Fla.), and API 20 STREP (bioMerieuxVitek) have the capability to identify members of the group C streptococci to species level. However, the four subspecies can easily be differentiated presumptively by the fermentation pattern of four sugars: trehalose, sorbitol, lactose, and salicin (Table 3). The key test to differentiate the small-colony-forming beta-hemolytic “S. anginosus or milleri” group from the large-colony-forming pyogenic strains is the Voges-Proskauer (VP) test for acetoin production. Additional differential properties of group C streptococci are listed in Table 3 (181,209). Antibiotic Susceptibility and Treatment Penicillin is the drug of choice for treating streptococcal infections. Vancomycin is recommended for pa-

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tients with bacteremia who are allergic to penicillin (30). Sensitivity to tetracycline is variable, and resistance to erythromycin has been reported (15, 17). With the exception of SXT, to which the bacteria appear to be susceptible, resistance to sulfonamides is common. Some strains of S. dysgalactiae subsp. equisimilis are resistant to penicillin (6). Cephalosporins and erythromycin also serve as alternative choices of treatment. Listeria

monocytogenes

Description, Natural Habitat, and Disease in Animals L. monocytogenes is a short, gram-positive rod commonly found in the environment. Listeria spp. are facultative, non-spore-forming bacteria with peritrichous flagella that impart a characteristic tumbling motility to the microorganism. In young cultures, the organism appears as a rod, but as the culture ages, coccoid forms appear. It is not uncommon, therefore, for L. monocytogenes to appear as a mixture of rods and cocci in Gram-stained smears. Although the optimum growth temperature is between 30 and 37°C the organism will grow slowly at temperatures as low as 3OC. L. monocytogenes is common in soil, vegetation, and feces. It may also be isolated from the tonsils of swine and lymph nodes of healthy animals. Most animals are exposed to this bacterium and/or colonized at some time in their life. Disease in animals is usually associated with predisposing disease or stress. Animals are usually infected through the gastrointestinal tract, resulting in infection of Peyer’s patches and the liver, followed potentially by bacteremia. In pregnant animals abortion may occur. L. monocytogenes may infect a wide variety of animals; septicemia is the most common form of infection in most animals. Following septicemia death may occur suddenly or after several days’ illness, with necrosis of the liver, spleen, and lymph nodes. Meningoencephalitis is the most easily diagnosed form of listeriosis in cattle and sheep and has been called “circling disease” because the animal moves in circles in one direction (238). Mode of Transmission L. monocytogenes is widely distributed in nature and may be recovered from food products such as raw meats and vegetables, as well as from processed foods such as soft cheeses that may become contaminated after processing. However, an outbreak of listeriosis in Sweden was linked to the consumption of gravad, or cold-smoked rainbow trout (80). The most common portal of entry, for Listeria infections other than neonatal infections, is the gastrointestinal tract, with subsequent spread via the circulatory system. However, cutaneous listeriosis contracted by humans at

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the time of animal parturition is not uncommon, particularly among veterinarians. Although there are many reports of transmission of L. monocytogenes from animals or animal products to humans in the literature, for the majority of cases direct animal association has not been identified. Human Infections Of the eight species in the genus Listeria, L. monocytogenes is the only one commonly reported to cause infections in humans. Listeriosis is a potentially serious infection, which often affects pregnant women, newborns, and adults with malignancies or with defects in cell-mediated immunity; infections are frequently associated with eating food contaminated with L. monocytogenes. When contracted during pregnancy, L. monocytogenes shows a marked propensity to cross the placenta and infect the central nervous system of the fetus. In 1986 the CDC estimated that about 1,700 cases of listeriosis occurred annually, 27% of which occurred in pregnant women or the newborn, with an attack rate of 12.4 cases per 100,000 live births (93). Neonatal listeriosis may occur as early-onset disease, often presenting in premature infants as sepsis accompanied by disseminated microabscesses known as granulomatosis infantisepticum. Late-onset neonatal disease generally occurs in full-term infants, presenting as meningitis several days to weeks after birth. Among the remaining 73% of cases, the incidence of listeriosis was noted to increase with age, with 41% of nonperinatal cases occurring in patients over the age of 70 years (93). The disease spectrum, virulence factors, and mechanisms of pathogenicity of Listeria have recently been reviewed by Rouquette and Berche (206) as well as Low and Donachie (150). Diagnosis Listeria is not fastidious, and isolation from human clinical material can readily be accomplished by plating on 5% sheep BA or inoculation in commercial blood culture bottles containing any standard broth medium. A number of enrichment techniques to recover the organism from nonsterile sites have been described using selective and differential media, some of which are commercially available (226). The technique of cold enrichment, in which a nonselective broth medium is inoculated and held at 4OC for several weeks to even months, has been used to recover the organism from nonsterile sites. When cultivated at temperatures of 20 to 25”C, L. monocytogenes displays a characteristic tumbling motility, which when viewed in a wet preparation can be a useful feature for identification. On BA, colonies grow readily to a diameter of 0.5 to 1.5 mm and produce a small zone of subtle beta hemolysis, not unlike that of group B streptococci.

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Identification of L. monocytogenes using conventional biochemical tests can be made by observing positive reactions for catalase (useful in distinguishing the organism from streptococci), glucose, and rhamnose, accompanied by negative reactions for mannitol, xylose, H,S production, and hippurate hydrolysis. The CAMP test, in which I;. monocytogenes is streaked at a right angle to, but not in contact with, a streak of Staphylococcus aureus, reveals a zone of enhanced or synergistic beta hemolysis. In a similar fashion, a commercial disk containing S. aureus betalysin (Remel) that amplifies the beta-hemolysin of L. monocytogenes when placed on BA streaked with the organism is available. AccuProbe (GenProbe, San Diego, Calif.), which is a nucleic acid probe test for culture confirmation, and an API-Listeria test strip (bioMerieux-Vitek) of 10 biochemical substrates are also available. DNA hybridization and PCR techniques have been described a.nd may be useful for detecting L. monocytogenes, as well as other food-borne pathogens that may be transmitted via processed foods (e.g., meat, poultry, and dairy products). Such foods are often implicated in outbreaks of listeriosis (180). Antige nit variation (somatic and flagellar) among strains of L. monocytogenes has been described, and 13 serovars can be identified. While there is no documented relationship between serotype and virulence, the majority of human infections are caused by strains 1/2a, 1/2b, and 4b. Phage typing, multilocus enzyme electrophoresis, DNA fingerprinting, pulsed-field gel electrophoresis, and random amplified polymorphic DNA typing schemes are also available and have been reviewed elsewhere (226). Antibiotic Susceptibility and Treatment L. monocytogenes is uniformly susceptible to ampicillin, penicillin, erythromycin, tetracycline, chloramphenicol, rifampin, and gentamicin. In general, however, penicillin and ampicillin, which may be accompanied by gentamicin or another aminoglycoside, are considered the treatment of choice for most manifestations of listeriosis (132). SXT is the drug of choice for penicillin-allergic patients. Cephalosporins should not used for treatment of L. monocytogenes infections regardless of the results of in vitro susceptibility testing. Rhodococcus

equi

Description, Natural Habitat, and Disease in Animals The genus Rhodococcus is composed of a heterogeneous group of microorganisms that are members of the aerobic actinomycetes. They are quite variable in morphology, growth patterns, and biochemical characteristics. Rhodococci are gram positive, nonmotile, nonsporulating, and partially acid-fast. Microscopic

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morphology varies from coccoid to coccobacillary and bacillary forms. Bergey’s Manual of Systematic Bacteriology lists 20 different Rhodococcus species (144). Among them, R. equi causes most of the reported human and animal infections. Other species of Rhodococcus that have been reported to cause human infections include R. (Gordona) bronchialis, R. (Tsukamurella) aurantiacus, R. luteus, R. erythopolis, R. rhodochrous, and R. rubropertinctus (35). R. equi is widely distributed in the environment and has been isolated from 54% of soil samples and from intestinal contents, feces, and dung of all grazing herbivorous species examined (2 1). R. equi has been known to cause animal disease since it was first isolated from lung lesions of foals in 1923, and at that time it was described as Corynebacterium equi (153). Clinical presentations vary from the asymptomatic carrier state to pneumonia in foals. On the majority of horse farms, it is unrecognized as a pathogen. R. equi may become endemic on some horse farms or cause sporadic disease on other farms (204). Less commonly, R. equi may cause submaxillary lymphadenitis (197) and pneumonia (263) in pigs. Although rarely, R. equi also causes infections in a wide variety of other animals, such as lymphadenitis and pneumonia in cattle ( 166,174); pneumonia, liver, and spleen abscesses in goats (50, 255); lung abscess in deer (47); lymphadenitis, pyogranuloma, and abscess in cats (78, 113, 129); and skin lesions in dogs (191). For an excellent review of R. equi, readers are referred to reference 192. Mode of Transmission Human R. equi infections are frequently associated with a history of contact with farm animals, soil, or both (49, 99, 133,243). Although commonly associated with disease in animals, infections have occurred in patients who have denied any close contact with animals. Infections in both animals and humans are thought to occur through the respiratory tract (35). Human Infections R. equi infections in human have been rare and almost always associated with a state of immune suppression. In 1967, R. equi pneumonia was reported in a patient receiving corticosteroid therapy for chronic hepatitis (99). Depending on the underlying immune status or age of the host, the site of inoculation, and the virulence of the organism, the infection may have variable clinical presentations. R. equi has been reported to cause posttraumatic cutaneous infections (5 1, 177) and foot mycetoma in an HIV-infected patient (5). Lymphadenitis due to R. equi in a healthy person has also been reported (144). R. equi bacteremia is usually secondary to pulmonary or cutaneous infection (51, 164,220). R. equi has been reported to be responsible for catheter-associated sepsis (41; C. Hinnebusch, D.

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Glenn and l? Colonna, Abstr. 92nd Gen. Meet. Am. 1992, abstr. C-461, p. 497, 1992) as well as pericarditis in a patient with a failed renal transplant (145). In severely immunocompromised patients, especially patients with AIDS, primary pulmonary R. equi infections (pneumonia and lung abscess) have been reported most frequently (27, 109, 142, 156, 212, 251; L. A. Haglund, J. A. Trotter, L. N. Slater, S. L. Harris, and I?. J. Retting, Letter, N. Engl. J. Med. 321:395,1989; D. C. Sane and D. T. Durack, Letter, N. Engl. J. Med. 314:56-57, 1986). However, fatal R. equi pneumonia was reported in an U-year-old woman without AIDS (247).

Sot. Microbial.

Diagnosis In humans with respiratory disease, sputum is the least-desirable specimen for diagnosis. A more reliable diagnosis can be made by bronchial brushing, percutaneous thoracic aspiration, or open lung biopsy. In Gram-stained smears of clinical specimens (purulent material and tissue), the organisms frequently appear as gram-positive cocci and bacillary forms. Some strains are partially acid-fast. Acid-fastness is more prominent on direct smears and on fresh isolates using the Kinyoun technique. Subcultures are rarely acid-fast. R. equi grows aerobically on most conventional media used in clinical microbiology laboratories: 5% sheep BA, chocolate agar, brain heart infusion agar, aerobic blood culture medium, thioglycolate broth, Sabouraud’s medium, and LowensteinJensen’s medium. The organism grows over a wide range of temperatures: from 10 to 40°C. Cultures grow equally well at room temperature or 37OC. The colony morphology of R. equi is diverse. The most frequently observed colony type is moist and pale pink when growing on BA aerobically at 35OC for 2 to 4 days. The other colony type is salmon-pink and nonslimy. Occasionally, colonies are pale yellow and nonslimy. The microscopic morphology of R. equi in culture varies in form from coccoid to bacillary, depending on incubation time and growth conditions. At 6 h on heart infusion agar incubated at 35°C the organism is completely bacillary; at 24 h it converts to a coccoid morphology (152). In nutrient broth, the organism is predominately bacillary. R. equi is non-spore forming, nonmotile, oxidase negative, catalase positive, indole negative, weakly urease, positive, CAMP positive, and noncarbohydrate fermenting in fermentative basal medium with Andrade’s indicator (62). In Gordon’s basal medium, it produces acid via oxidation of glucose in 14 days (24). It is lipase, phosphatase, and nitrate reduction positive and negative for hippurate hydrolysis, esculin hydrolysis, and gelatin hydrolysis. A commercial miniaturized differentiation system,

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the Rapid CORYNE (bioMerieux-Vitek), offers a 24to 48-h identification test for coryneform bacteria, including R. equi. The kit does not, however, discriminate among species of Rhodococcus; all Rhodococcus and Gordonae spp. key out as R. equi. Preliminary evaluations showed the system to have limited sensitivity, with only 65 % of R. equi being identified (22). Definitive identification of R. equi may require 107 to 129 biochemical tests. A less extensive panel of tests for routine identification of Rhodococcus spp. that is used at the Actinomycete Laboratory at the CDC is more practical but may not identify all species of Rhodococcus (143). Clinical laboratories capable of performing molecular biology procedures may use PCR and restriction fragment length polymorphism (RFLP) techniques to identify and further differentiate rhodococcal species. A PCR-RFLP method to amplify a 439bp segment of the 65-kDa heat shock protein gene of actinomycetes has been developed to rapidly identify clinically significant species of aerobic actinomycetes, including Rhodococcus (224). A ribotyping technique to analyze RFLP in rRNA genes of Rhodococcus species (41) can be used for further differentiation of rhodococcal isolates. Antibiotic Susceptibility and Treatment In vitro, R. equi isolates are usually susceptible to erythromycin, rifampin, vancomycin, SXT, tetracycline, ampicillin-sulbactam, amoxicillin-clavulanic acid, aminoglycosides, and imipenem. Broad-spectrum cephalosporins and fluoroquinolones may have good activity against R. equi; however, susceptibility may be variable. R. equi is moderately susceptible or resistant to penicillins, penicillinase-resistant penicillins, and the narrow- and extended-spectrum cephalosporins (63). Emergence of rifampin-resistant R. equi in human and foal isolates has been reported (23 1). Successful treatment of R. equi infection depends on the use of lipophilic antimicrobials that may penetrate the macrophage and neutrophils in which the organisms often survive. A combination of erythromycin and rifampin (both penetrate the macrophage cell membrane) may have synergistic activity against R. equi and is the therapy of choice for foals. However, disappointing results have been reported with this combination for patients with AIDS. Gentamicin, tobramycin, and ciprofloxacin have good activity in vitro as well as good penetration into macrophages. Vancomycin is effective for treating R. equi infections in monotherapy as well as in combination therapy. Nontuberculous

Mycobacferium

spp.

The zoonotic potentia 1 of Mycobacterium tuberculosis and M. bovis has been extensively documented. Except for Myco bacterium marinum, however, it is

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not as clear that the nontuberculous mycobacteria (NTM) can be transmitted directly from infected animals to humans. Most NTM are noninvasive and opportunistic in nature. These infections are most often acquired from environmental sources rather than from human-to-human contact (45). Direct infection from animals to humans has not yet been demonstrated (236). Nevertheless, instances of animals infected with the NTM known to cause disease in humans have been documented, and therefore there is zoonotic potential. Patients who are immunocompromised are particularly susceptible to these organisms. Due to the number of species in this group, they are described individually below. Description, Natural Habitat, Mode of Transmission, and Animal and Human Infections by Species MAC

Mycobacterium avium-Mycobacterium intracellulare complex (MAC) organisms are the most common disseminated bacteria in patients with AIDS. MAC organisms are not common agents of infections among pet birds (116), but 211.avium is the cause of tuberculosis in chickens, pigeons, and other wild birds. It may also occasionally cause diseasein pigs, cattle, sheep, dogs, cats, and humans. Although birds are the natural reservoir for M. avium, direct transmission of MAC from birds to humans is not likely. The more probable source of infection is soil and water contamination (97). Nonetheless, it is possible that birds function in transmitting MAC to previously uncontaminated sources of water and soil, although this has not been proven. Human infections and detection of the MAC organisms are well described elsewhere (110) and are not described further here. Mycobacterium

fort&urn

complex

Members of the rapid-growing (recovered in 3 to 5 days) M. fortuitum complex (M. abscessus,111.chelonae, M. fortuitum, and M. peregrinum) have been reported to infect a variety of animals, including fish (66), shrimp (172), and dogs (88; D. S. McKinsey, M. Dykstra, and D. L. Smith, Letter, N. Engl. J. Med. 332:338,1995). Direct transmission from the affected animal to humans has not been demonstrated, but indirect transmission and the potential to infect humans bear some scrutiny. This group of mycobacteria consists of opportunistic pathogens. 111.fortuitum has been reported to causemultiple cutaneous abscessesin a dog following a bite wound (88). Mycobacterium chelonae, which is the slowest growing of this group and prefers 28 to 30°C for primary isolation, has been isolated from the soft tissue of the heel of a woman who would rub her heel against her Scottish terrier (McKinsey et al., letter).

Mycobacterium

37

genavense

First described as a causative agent of disseminated infection in AIDS patients, M. genavense was previously recognized as a fastidious, nonculturable, acidfast organism. Supplementation of the growth medium with mycobactin enabled the organism to grow in vitro. Subsequent analysis of the 16s rRNA gene identified this mycobacterium as a new species (28). Investigation of nonculturable mycobacteria affecting various species of birds indicated that 111.genavense was the most common cause of mycobacteriosis in psittacine birds. An examination of mycobacteria isolated from pet birds presenting with clinical symptoms over a 9-year span indicated that 71% of the mycobacteria isolated were M. genavense (116 ). Water was suspected as the source of infection, but this could not be confirmed. A similar study of birds kept at the Antwerp Zoo (Antwerp, Belgium) was conducted over an 1 l-year period, and M. genavense was found in the tissues of seven birds examined at necropsy. Most of the birds in this study had a predominance of organisms in the lungs, liver, and spleen, as well as the intestine (190). Severe hind limb weakness with infection of the cervical lymph node due to 211. genavense has also been reported in a dog (137). M. genavense rarely invades the lungs in human patients. Rather, it more commonly causessepticemia and disseminated infections. M.

marinum

111.marinum is found in both fresh- and saltwater environments and can cause granulomas and death in fish. Humans can become infected following contact with infected fish or their waste, usually secondary to some minor trauma to the skin (97). A common source of 211.marinum infections to humans is fish aquariums (120). Reported casesinclude nodular skin lesions on the hands and arm (94, 1l2), papular lesions on the knuckle (2O), and dissemination in AIDS patients (97; K. M. Ries, G. L. White, Jr., and R. T. Murdock, Letter, N. Engl. J. Med. 322:633, 1990). Although all caseswere a result of contact with aquarium debris or water, no evidence of direct fishto-human transfer was demonstrated. One report suggested that water fleas used to make commercial fish food could be a likely source that infected the fish and then the aquarium environment (155). Mycobacterium

thermoresistibile

M. thermoresistibile is a rapidly growing, chromogenic mycobacterium, originally thought to be nonpathogenic. However, both pulmonary and nonpulmonary clinical caseshave been reported (4.5). Cases of extrapulmonary involvement in humans have involved primarily the skin and lymph nodes (179,260). The first reported case of an animal to exhibit extrapulmonary lesions of M. thermoresistibile involved

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a short-haired lO.S-year-old female domestic cat (257). The cat presented with a number of lumbrosacral-area skin lesions, which had been present over a lo-month period but grew larger prior to examination. There was no history of trauma to the site, and it could not be determined how the animal contracted the organism. Fluid was aspirated from the lesions, and acid-fast bacilli were observed and subsequently identified as 211. thermoresistibile. Treatment with penicillin had no effect, and the cat was euthanized. Diagnosis An acid-fast stain can be employed directly on specimens to determine if mycobacteria may be present. For isolation of NTM from blood and disseminated infections, BACTEC 12B broth bottles have been useful. In addition to standard mycobacterial agar medium, some NTM (such as M. fortuitum) grow on BA or chocolate agar. For most NTM species, conventional biochemical methods remain the standard for identification ( 136). Information from biochemical tests may be limited for recently identified or rarely encountered NTM, and alternative methods should be utilized. The analysis of high-molecular-weight mycolic acids by high-performance liquid chromatography has been shown to be a useful and rapid tool for identification to the species level (44, 239). Genetic methods, such as analysis of the hypervariable region of the 16s rRNA gene, have been used to define the species taxonomically (28). To date, however, genetic methods are not standardized and are available primarily in research laboratories Antibiotic Susceptibility Currently, standardized methods for susceptibility testing of the NTM do not exist, and therefore correlation between in vitro results and clinical response may be unclear (45). Furthermore, many mycobacteria classified as rapid growers (visible growth of bacteria on solid media within 5 to 7 days) do not respond to conventional antituberculosis drugs. Because of the wide variety of responses by the different species, and even by strains of the same species, an individualized approach to therapy is advocated. Susceptibility results must be interpreted cautiously and used as adjuncts to published therapeutic regimens if available (45). The M. fort&urn complex may be susceptible to amikacin, doxycycline, cefoxitin, clarithromycin, minocycline, SXT, and sulfisoxazole. Infections due to 211.genavense have been successfully treated with clarithromycin, ethambutol, and enrofloxacin ( 13 7). Treatment of M. mdrinum infections has varied (20, 135). Although in some cases treatment with antituberculosis drugs has failed, skin lesions have responded to extended treatment with isoniazid and streptomycin ( 112). An HIV-infected individual with lesions on his chest, extremities, and face responded to

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6 months of treatment with isoniazid, rifampin, and ethambutol (Ries et al., letter). SXT has been used successfully in cases where antituberculosis drugs failed (20, 135). Bruce//a

spp.

Description, Natural Habitat, and Diseasein Animals Brucellae are agents of brucellosis of domestic and wild animals that are zoonotic to humans. Despite the recognition of brucellae as a single genospeciesbased on DNA-DNA hybridization studies, they are systematically classified in the genus Brucella as B. abortus, B. melitensis, B. suis, B. canis, B. ovis, and B. neotomae (123, 168). Brucellae have also been isolated from marine mammals ( 128, 246). They are small gram-negative coccobacilli, nonmotile, aerobic, and slow growers. All brucellae except B. ovis and B. neotomae produce catalase and oxidase, are urease variable, and reduce nitrate to nitrite. Infection due to brucellae is ubiquitous. However, the diseaseis endemic in the Mediterranean basin, the Arabian peninsula, the Indian subcontinent, and in parts of Mexico and Central and South America (271). Brucellosis is mainly a diseaseof farmers, shepherds, veterinarians, microbiologists, butchers, and slaughterhouse workers (40, 269). B. abortus is mainly found in cattle, but other animals, such as the camel, yak, and buffalo, may also be infected by this species. B. melitensis primarily infects goats and sheep; non.etheless, camels can be a source of this agent in some countries. B. suis biovar 4 and B. canis biovars 1 to 3 are common to reindeer, caribou, and feral swine, respectively. B. canis, the least-common cause of human infections, is found in dogs (67, 79). However, there has been a single case report of a laboratory infection by a brucella isolate from a marine mammal (S. D. Brew, L. L. Perrett, J. A. Stach, A. I?. MacMillan, and N. J. Staunton, Letter, Vet. Rec. 144:483,1999). B. ovis causesinfections in sheepand B. neotomae causesinfections in rats, but these species have not been reported to cause infections in humans. Brucellosis in animals is a chronic, lifelong infection with localization of the organism in reproductive organs. The clinical manifestations of infection include primarily abortion and sterility (79). Brucellae are also considered class B bioterrorism agents by the CDC. Mode of Transmission Human Brucella infections are acquired by consumption of unpasteurized milk and dairy products or by direct exposure to animals and their carcasses.Contaminated fresh goat’s cheeseis the major source of Brucella bacteria in the general population (8, 58, 232, 235, 273). Human-to-human transmission is

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highly unlikely, although sexual transmission of brucellosis is possible, as the organisms have been isolated from human semen (208, 242; J. I? Veenstra et al., Letter,]. Infect. Dis. 166:209-210, 1992). Veterinarians have acquired the infection as a result of self-inoculation with the attenuated B. abortus strain 19 vaccine previously used for cattle immunization (211). However, the current attenuated vaccine strain (RB51) is safer and has not been reported to cause disease in humans or animals. Wild animals may play a role in the epidemiology of brucella infections. For example, wild hare, the reservoir for B. suis biovar 2 in Europe, may transmit the agent to domestic and feral swine. Also, many bison in Yellowstone National Park are infected with B. abortus, which may serve as a reservoir for the disease (67,271). It should be emphasized that laboratory-acquired infections with Brucella are relatively common, with over 100 such infections reported ( 176). Human Infections Brucellae are the cause of abortion in animals and can cause systemic infections involving any organ in the human body (271). The smooth lipopolysaccharide is an important virulence factor of brucellae, and B. melitensis is the most virulent biovar. The incubation period of human brucellosis is 2 to 3 weeks, and its initial symptoms are insidious. However, fever, sweats, malaise, weight loss, arthralgia, splenomegaly, and hepatomegaly are common clinical presentations of the infection (175, 272). Localized and chronic infections usually involve organs important in nonspecific defense mechanisms, such as liver, bone, spleen, and kidneys ( 158, 249). Brucellae can rarely cause meningoencephalitis, which may be misdiagnosed as tuberculosis (167). Diagnosis The isolation of brucellae from blood, bone marrow, or other tissues is the definitive means of diagnosis of human brucellosis. Conventional and commercial blood culture systems with extended incubation periods are appropriate for the detection of brucellae (259). Inoculated biphasic media and vented blood culture bottles are incubated at 35OC under 5% CO;?, with subcultures performed every 4 to 5 days for 4 weeks (176, 259). Bone marrow cultures are preferred, as they are usually positive for acute, subacute, and chronic brucellosis. In contrast, blood cultures are positive only for patients with acute infections ( 101). The lysis-centrifugation technique has also been found useful for the recovery of brucellae from blood, bone marrow, and spinal fluid specimens. The mean recovery time is 3 days for lysis-centrifugation versus 14 days for the older BACTEC NR system (140, 178). However, the BACTEC 9240 system has been shown to detect Brucella spp. within 5 days of

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37

incubation (176). The BacT/Alert system has been reported to require blind subculture for isolation of Brucella spp. (176). The ESP system may also be capable of isolating Brucella spp. but has not been adequately studied. Due to the lack of adequate investigation of the performance of automated blood culture systems to isolate and identify Brucella spp., it would be wise to perform standard blind subcultures with bottles in which Brucella is part of the differential diagnosis and to hold all bottles 21 days (176). Exudates and tissues can be cultured on tryptose agar with 5% bovine serum with or without antibiotics. Skirrow agar, a selective medium, has been used in veterinary microbiology laboratories to isolate brucellae from contaminated bovine semen, milk, and vaginal secretions (234). Brucellae also grow on Thayer-Martin or unsupplemented chocolate agar (36). Biovars of brucellae are identified by means of colony characteristics, biochemical reactions, phage typing, dye inhibition assay, molecular diagnostic techniques, and serology. Brucellae produce nonhemolytic, oxidase-positive colonies that agglutinate in unabsorbed smooth phase (encapsulated) antiserum. Production of H,S, growth requirement for increased CO;?, production of urease, absence of fermentative capability, and growth on solid media containing basic fuchsin and thionin have been shown to be useful characteristics for identification of brucellae (176). Phage typing is used to differentiate certain biotypes of B. abortus from B. melitensis. DNA probe assay systems are used to identify most common biovars of B. abortus, B. melitensis, B. suis, and B. ovis (102). PCR h as b een used successfully to differentiate between B. abortus biovars 1,2, and 4; B. melitensis; B. suis biovar 1; and B. ovis (34, 160). Commercially available identification kits, however, have misidentified B. melitensis as Moraxella phenylpyruuica or as Haemophilus infhenxae (14). Brucellosis can also be diagnosed serologically by the demonstration of high or rising specific serum antibody titers. The serum agglutination test uses B. abortus strain 19 and detects both immunoglobulin G (IgG) and IgM to B. abortus, B. melitensis, and B. suis, but not B. canis, because B. canis lacks smooth lipopolysaccharide (271). Titers of IgG against bruh.igh following cellae in serum remain Pe rsistently an increase in both IgG and infection. Nevertheless, IgM titers by enzyme-linked immunosorbent assay (ELISA) is a strong indication for the relapse of a chronic infection (7). Antibiotic Susceptibility and Treatment The antibiotic susceptibility of brucellae in vitro does not correspond with therapeutic efficacy in clinical trials of new antibiotics. Doxycycline is considered the most effective single drug for the treatment of uncomplicated brucellosis. Combination therapy with

CUMITECH

Diagnosis

37

tetracycline and streptomycin or doxycycline plus rifampin is recommended for 6 weeks due to a high rate of relapse when a single drug is used (1, 106). For children under 8 years of age and pregnant women, SXT plus gentamicin is equally effective. However, children older than 8 years can be treated with combinations of doxycycline and gentamicin (151, 270). Meningitis and endocarditis due to Brucella have been successfully treated with doxycycline in combination with SXT and rifampin (127, 167). Laboratory Safety Brucellosis is the most common laboratory-associated infection reported. A wide variety of fluid specimens may be infected (blood, semen, spinal fluid, and urine), and therefore aerosol exposure is possible. However, most laboratory infections are the result of working with large volumes of pure cultures. Biosafety level 2 (BSL-2) laboratory conditions are recommended for work with clinical specimens, but BSL-3 laboratory conditions should be employed when manipulation of pure cultures is required. Suspected cultures may also be sent to the CDC Special Bacteriology Reference Laboratory if facilities are not available to safely handle these organisms. Transport of cultures should follow current CDC-National Institutes of Health (NIH) recommendations (201). Burkholderia mallei

pseudomallei

and Burkholderia

Description, Natural Habitat, and Disease in Animals B. pseudomallei is the etiologic agent of melioidosis, a disease of humans and animals that resembles the equine disease glanders, which is caused by B. mall& Melioidosis (pseudoglanders, or Whitmore’s disease) and B. pseudomdllei were first described by Whitmore and Krishnaswami in 1912 (256). Interest in this disease, which is endemic in Southeast Asia and Australia, has increased in the United States as a result of infections occurring among U.S. Armed Forces personnel stationed in Vietnam (119). Glanders (farcy, equine nasal phthisis, or maliasmus) is a more severe zoonotic disease of solipeds (horses, mules, or donkeys). However, infections of humans and other animals can occur, although other species are less susceptible than solipeds (223). B. pseudomallei is a motile, gram-negative rod with a cellular diameter of 0.8 pm and a cell length of 1.5 km. Individual cells usually have a bipolar appearance on microscopic examination. Motility is achieved by flagella arranged in polar tufts of three or more individual organelles. This organism grows vigorously on most commonly used bacteriologic media and forms colonies that range in appearance from rough to mucoid and range in color from cream to

of Zoonotic

Infections

13

bright orange. Cultures tend to have a musty aromatic odor (although the sniffing of open plates is not recommended). The optimal growth temperature is 37°C. B. mallei is very similar to B. pseudomallei except that B. mallei lacks flagella (183). B. pseudomallei utilizes some substrates that B. mallei does not, and B. pseudomallei grows much more quickly than B. mallei. If specimens containing B. pse~doma~~ei are contaminated, the selective Ashdown’s medium, which contains crystal violet, neutral red, and gentamicin, is recommended; colonies are typically dry, wrinkled, and purple (264). Following culture, the organism may have an odor of putrification or a sweet, earthy smell (96). Since its original description in 1912, the nomenclature of B. pseudomallei and B. mallei has often been revised. It has been variously classified with the genera Actinomyces, Pfeifferella, Actinobacillus, Loeflerella, Malleomyces, and, most recently, Pseudomonas (96,119). As a member of the Pseudomonas rRNA homology group II of Palleroni, it was proposed that these two species, along with Burkholderia cepacia, Burkholderia gladioli, and Burkholderia pickettii, be included in the new genus Burkholderia by Yabuuchi in 1992 (184,265). B. pseudomallei may be cultured from soil and freshwater sources in temperate zones of the Far East, Africa, and middle and South America ranging 20’ latitude on either side of the equator (119). B. pseudomallei has been recovered from soil samples stored at room temperature for 1onger than 1 year (104) and from moist clay soil stored at room temperature for 30 months (238). All known areas of endemic disease are located within this zone, and human infections detected outside of this zone can usually be traced to travel through an area of endemic disease. Serologic surveys in areas of endemicity indicate that the level of exposure and subclinical infection is much higher than overt disease (170). Epizootic outbreaks of B. pseudomallei involving swine, sheep, goats, calves, marine mammals, and other anim .als have been reported in locations both inside and outside areas of endemic human disease within the temperate zone (170). Infections can be acute, ending with death, but more often they are subclinical or chronic. Clinical signs vary with animal species bu t often include n asal an .d ocular discharge, lameness, septicemia, and lesions of internal organs. Glanders mav also present w -ith acute-to-chronic disease, with the chronic form bei ng more comma n in horses and the acute form more common in mules and donkeys. The acute form of glanders presents with septicemia, bloody nasal discharge, and ulceration of lymph nodes. Death may occur within 1 week. In horses the disease usually begins with pulmonary infection, begins and progresses slowly over weeks to

14

Bottone

et al

CUMITECH

months, and may progress to upper respiratory and cutaneous gl anders. Animals tha t are subclinically infected may be a more significant source of infection because they may be shedding bacilli in respiratory secretions for months before they are clinically symptomatic. Lesions and a bloody, green-yellow nasal discharge appear first, with ulcers developing along the respiratory tract. Lymphadenitis develops with ulceration through the skin, resulting in the cutaneous form of the disease (farcy). Both B. m&i and B. pseudomallei are considered class B agents of bioterrorism by the CDC because of their virulence and potential transmission by aerosol. During World War I German spies infected horses with B. mdllei at ports in the United States. Since in nature it is a zoonotic disease, infection of horses with 13. m&i could be another way by which the agent could be spread. Furthermore, infection of horses would be a form of agroterrorism. Mode of Transmission Although B. pseudomdllei causes disease in both animals and humans, human infection does not involve direct contact with an infected animal. Since this organism survives as a free-living soil or water microbe within the temperate zone, human infection typically results from direct skin contact with contaminated soil or water. However, animals may serve as a source of the infectious agent. Pulmonary melioidosis also occurs as a result of inhalation of contaminated dust. This mode of transmission was observed among U.S. helicopter crewmen stationed in Vietnam (119). A few cases of human-to-human transmission have been reported (104). Glanders is spread by aerosol and direct contact with infective material from horses and other solipeds. The route of infection may be through the digestive tract (ingestion of contaminated food or water), through the respiratory tract, or by exposure of wounds to fluids or tissues of infected animals. However, under field conditions the risk of transmission to humans is low, as only a few human handlers of infected animals have developed the disease (223). Human-to-human transmission has also occurred from patients to health care workers (104). Human Infections Melioidosis in humans can present as various clinica manifestations, including inapparent infection, tran sient bacteremia, asymptomatic pulmonary infiltra tion, acute localized suppurative infection, acute pul monary intection disseminated septicemic intection, nondisseminated septicemic infection, and chronic suppurative infection (214). The organism may produce abscesses in a wide variety of tissues, including the lungs, liver, spleen, lymph nodes, skin, soft tissue, joints, and bones (96). The incubation period between l

r

.

1’

.

1

.

l

l

r

.

37

exposure and the onset of symptoms can range from 2 or 3 days to as long as 26 years (214). The incidence of infection is higher in males than females due to occupational differences in exposure. Glanders in humans may present as a nasal-pulmonary form (glanders) or a cutaneous form (farcy). Both forms can be present simultaneously and have systemic involvement. Glanders may also be acute or chronic, depending on the route of infection, dose, and virulence of the strain; the acute form is most common in humans (104, 223). However, glandersassociated nodules have been identified at autopsy in large numbers of people with equine contact, suggesting that mild or subclinical disease may be more common than previously thought ( 104). In acute cases clinical signs appear within 14 days of exposure, and the course of the disease lasts 2 to 4 weeks. Initially, painful nodules and swellings appear on the face and extremities that spread and give rise to pustular eruptions over most of the body. The nasal mucosa may ulcerate with a purulent discharge. Pneumonia with abscesses, pleural effusions, and empyema often develops. Later, suppurating pustules cover the body, and internal abscesses develop, leading to death. The chronic form of the disease is similar, but symptoms are milder and may last from months to up to 25 years (104,223). Diagnosis B. pseudomdlEei may be isolated from abscess drainage, sputum, tissue biopsy specimens, or blood specimens. The organism grows well on most commonly used media in the clinical laboratory, including gramnegative selective media such as MacConkey agar. Colonies can be observed within 24 to 48 h of incubation at 3.5 to 37OC and usually show a characteristic white opaque growth with a sheen by 48 h. At first colonies are smooth and convex, but slowly they become umbonate, with an uneven wrinkled surface. In a young culture at 35°C most cells will show bipolar staining; after 48 h most cells are oval to round, and only the periphery of the cell stains; such cells may be mistaken for endospores. The biochemical reactions for 70 strains identified as B. pseudomalEei by the CDC Special Bacteriology Reference Laboratory are given in Table 4 (253). Oxidation of carbohydrates was performed in King’s oxidation-fermentation (OF) medium. Sources of strains include sputum (21%), wound (14%), animal isolate (13%), blood (lo%), abscess or drainage (6%), lung (4%), skin (3%), other (including one prostatic fluid) (15%), and unknown (14%). This species is very active, producing positive reactions for oxidase and arginine dihydrolase. Glucose, xylose, mannitol, lactose, maltose, and som .etimes sucrose are oxidized, and nitrate is reduced, usually with formation of gas. With some strains detection of gas pro-

CUMITECH

37

Table 4. Biochemical 6. pseudomallei

Diagnosis analysis

of 81

of

Motrlrty . . . . . . . . . . . . . . . . . . . . . . . **..* -.... . . . . . . . . . . . . . . . . . . . 100 Acid from: . . . . . . . . . . . . . . . . . . . . ..*...*... . . . . . . . . . . . .*. . . . .. . . . .. 100 D-Glucose D-Xylose .*..*..... . ..*a............ *.... . . . . .. . . .. . .. . . . . .. . .. . . . . . 86 D-Mannrtol . . ..*.. ..a....... . . . . . . . . . . . . . . . . ...*.... .. . . .. . .*... 94 Lactose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ‘.,,.. . . . . . 99 Sucrose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ..s.... 66 Maltose . . . . . . . . . . . . . . . . . . . . . . . ....” . . .. ’. ...*....... *“’ . . . . . . . . . . . . 99 Catalase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Oxrdase . . . . . . . . . . . . . . . . . . . . . . . ...*.... . . . . . . Growth on: MacConkey agar. . . . . . .. . . . 100 Salmonella-shrgella agar ........ . . .................... 8 Cetnmrde agar ................ 7 Simmons citrate . . .. . . .. . .. . . .. . .. . . . . . . . 77 Urea, Christensen’s a............... . . . ..**....*............ . . . . . . . . . 13 Nitrate reduction. . . . .. . . . . . ............... .... ..... . *IO0 Gas from nrtrateb ............................ ... .* ................ 100 Indole ............................................ ...... ................. 0 TSI slant, acid ...................... ........................ ... 72 TSI butt, acid ........................................................... 0 H,S (TSI butt) ............................. . ............... 0 H,S (Pb ac paper). ............................... .... . . 26 Gelatin hydrolysisC.. . . .. .. . . . .......... .. ..* . . . . 79 96 Litmus milk peptonrzatron ....... ........................ Growth at: 25°C .............................................. . ..................... 100 35OC ............................................ . ........................... 100 42OC ..... ................... . . . ............................. 100 Esculrn hydrolysis.. ............................... ... ..... 59 Lysrne decarboxylase ............. ...... ... .................. ...... 0 Arginine dihydrolase . . . . . . . . . . . . . . . . . . . ..s..... *. . . . . . . .. . . . . .. . . .. . 100 Ornrthrne decarboxylase .. .. ... .. ............ ... .. . 0 Nutrient broth, 0% NaCl . . . ..*......................... . .. .I00 Nutrient broth, 6% NaCl . . . . . . . . 12 a Numbers In parentheses Indicate 7 days of lncubatlon ‘The volume of gas may be small cubatlon IS done at 25°C ‘At 7 to 14 days of lncubatlon

percentages

of posltwe

Gas may

not be detected

tests

(14)

6) (I)

(4) (1)

(31) (3) (4) (8)

at 3 to

unless

In-

duction from nitrate may be difficult using standard methods. In these cases, incubation of the nitrate test at 25OC or utilization of the semiaerobic method of Stanier (253) can be used to enhance gas formation. The lysine decarboxylase test is usually negative. B. pseudomallei may be confused with Pseudomonas stutxeri. However, P. stutzeri colonies usually develop a slightly yellow pigment, and this bacterium does not oxidize lactose, is rarely proteolytic, and is motile by a single polar flagellum. Also, I?. cepacia may be confused with B. pseudomallei when gas is not formed readily from nitrate; however, B. cepacia is lysine decarboxylase positive and arginine dihydrolase negative. The API 20NE kit has misidentified B. pseudomallei as pigment-negative Chromobacterium violaceum (122). Therefore, if commercial kits are used to identify this agent, results should be interpreted with caution. A diagnosis of glanders can often be made based on

of Zoonotic

Infections

15

clinical signs alone, as most are pathognomonic. However, glanders is so rare that most physicians have never encountered it. Specimens for culture can be taken from lymph nodes or abscess, inoculated to media supplemented with glycerol, and cultured at 37OC. Intraperitoneal inoculation of a male guinea pig produces an ulcerative orchitis, referred to as the Strauss reaction, and is considered a definitive test for B. mallei (104, 223). However, inoculation with B. pseudomallei will give the same result, so biochemical testing of cultured bacteria is usually required for a definitive diagnosis. B. mallei can be distinguished from B. pseudomallei by the former’s lack of motility, utilization of xylose, and failure to utilize azelate and levulinate (183). Colonies of B. mallei are smooth, viscid, and moist. They are initially white to cream in color and become dark brown with continued incubation (104,223). Diagnosis and screening of equines for glanders is most commonly done by intrapalpebral mallein skin testing. Mallein is a filtered extract from killed cells of B. mallei, and the test is performed by intradermal injection into the animal’s lower eyelid. In animals infected with B. mallei a swelling, purulent conjunctivitis with pain, photophobia, and depression will develop within 1 to 2 days and persist for several days. Subcutaneous mallein testing is not recommended, because it will induce false-positive serologic reactions. Serologic tests for B. mallei, which are required of any equine that is imported from a region of endemicity, consist of complement fixation (CF), indirect hemaglutination (IH), CF-IH, ELISA, and fluorescent antibody tests. Tests that utilize complement cannot be used with sera from donkeys or mules, because sera from these animals are anticomplementary. A major drawback to serologic testing is false-positive reactions due to exposure to B. pseudomallei, which is antigenically very similar to B. mallei (104, 223). A similar skin test for detecting reactivity of animals to B. pseudomallei (melioidin skin test) is available but is subject to the same limitations and crossreactivity as the glanders skin test. In other words, an animal infected with B. mallei would test positive for B. pseudomallei in a melioidin skin test. IH and CF are more useful and commonly used serologic tests to diagnose melioidosis in animals ( 104,237). Antibiotic Susceptibility and Treatment Treatment approaches to melioidosis vary depending upon the clinical manifestation of the disease. The in vitro susceptibility data and treatment recommendations that have recently been summarized by Sanford are presented here (214). B. pseudomallei is usually sensitive in vitro to the tetracyclines, chloramphenicol, novobiocin, kanamycin, amoxicillin-clavulanate, ticarcillin-clavulanate, piperacillin, imipenem, many of the broad-spectrum cephalosporins, sulfadiazine or

16

Bottone

CUMITECH

et al

sulfisoxazole, and SXT (9, 219). Resistance has been reported for penicillin, erythromycin, ampicillin, streptomycin, colistimethate, ciprofloxacin, aztreonam, ticarcillin, and amdinocillin (95). Increased resistance to SXT has been reported in strains from Thailand (219). Patients with infections caused by SXT-sensitive strains have been successfully treated with either sulfisoxazole or SXT. Infections involving SXT-resistant strains have been successfully treated with amoxicillin-clavulanate, tetracycline, or chloramphenicol(57,214). A combination therapy of SXT and ceftazidime is recommended for the treatment of septicemic infections (2 18,254). Successful treatment regimens can range in duration from 60 to 150 days for localized nontoxic infections to as long as 1 year for patients with systemic infections (214). There has been little opportunity to evaluate treatment options with human glanders because the disease is so rare. Isolates of B. mallei from infected horses have been shown to be susceptible to sulfamethizole, gentamicin, tetracycline, sulfathiazole, kanamycin, tobramycin, streptomycin, and SXT. All equine isolates tested have been shown to be resistant to the narrow-spectrum cephalosporins and penicillins and to nitrofurantoin; extended- and broad-spectrum cephalosporins were not tested. Treatment of experimental glanders infections in laboratory animals has been successful with sodium sulfadiazine and various other sulfonamides ( 104). Laboratory Safety and Precautions Laboratory-acquired infections with B. pseudomallei have been reported (216). The current edition of the CDC-NIH guide Biosafety in Microbiological and Biomedical Laboratories recommends BSL-3 procedures for activities with a high potential for aerosol or droplet production and for activities involving production-level quantities or concentrations of this organism (201). Although BSL-2 safety preca utions are recommended for work with suspected clinical specimens, pure cultures of B. mallei or B. pseudomallei should be handled with caution under BSL-3 laboratory conditions. In particular, generation of aerosols should be avoided. The risk of laboratory-acquired infection due to these bacteria is much greater than that of acquiring the infection in the field. B. mallei has been described as “the most dangerous [bacterium] to work with in a laboratory” (65). Suspected cultures should be sent to the CDC Special Bacteriology Reference Laboratory or the Bacteriology Division of the U.S. Army Medical Research Institute of Infectious Diseases, Fort Detrick, Md. Details on how to package, label, and ship human pathogens are described in the CDC-NIH guide Biosafety in Microbiological and Biomedical Laboratories.

Francisella

37

tularensis

Description, Natural Habitat, and Disease in Animals F. tularensis is the etiologic agent of tularemia. The disease cycle is maintained in nature between wild animals, biting vectors, and the contaminated environment. Humans become at risk for acquiring tularemia by participating in activities that expose them to the natural cycle of tularemia. Tularemia is distributed over the northern hemisphere and not found in the southern hemisphere. Infection is widespread, but since international reporting of the disease is not required, the relative incidence is difficult to determine in many areas. Reported human cases in the United States between 1990 and 2000 ranged from 86 to 193 cases per year, for a total of 1,368 cases from 44 states (Fig. 1) (56). F. tularensis is a tiny, pleomorphic, gram-negative coccobacillus known to be involved in epizootics but is not considered highly transmissible. F. tularensis has been known to survive for long periods of time in the environment ( 117). Its ability to remain viable in the environment is in contrast to its fastidious nature in the laboratory, where it requires nutritional supplementation for growth and propagation. Tularemia in both animals and humans is treatable with antibiotics, and resistance to the recommended drugs for treatment is uncommon (70,215). F. tularensis cultures and specimens should be handled carefully since tularemia is one of the most frequently acquired laboratory infections (42, 182). The etiologic agent of tularemia was first described in 1912 as the cause of a plague-like disease of California ground squirrels (162). The disease remains widely enzootic in North America, Europe, and northeastern Asia. There are four subspecies of F. tularensis: F. tularensis subsp. tularensis, known as type A, is found in North America, while F. tularensis subsp. holarctica (formerly known as F. palaearctica), or type B, is more wide1 y distributed and is isolated from regions withi n the entire northern hemisphere; the other two subspecies, F. tularensis subsp. novicida and F. tularensis subsp. mediasiatica, are sporadically recovered and limited in their geographic distribution. Tularemia has a broad host distribution. F. tularensis infection has been reported in more than 100 species of both domestic and wild animals (117). The disease may be continually present in the animal population but at an unperceivable level or may appear in seasonal peaks depending on the dynamics of the epizootic. Disease affecting cottontails (Sylvilagus spp.) and jackrabbits (Lepus californicus), beavers (Castor canadensis), muskrats (Ondatra xibethicus), meadow voles (Microtus spp.), and sheep are most frequently reported in the United States. Within the United States, the character of the natural cycle differs. For instance, a deerfly-tick-animal cycle seems to be im-

CUMITECH

FIGURE patient’s

Diagnosis

37

1. Reported cases of tularemla In the United States between r-es ldence rn the continental United States Ten cases In four

portant in maintaining tularemia in the western United States. In contrast, in the southwestern region of the United States, the wild rabbit-tick cycle predominates. In Europe and Russia, mosquitoes are acknowledged as important vectors, whereas the mosquito’s role in the tularemia cycle in North America is not defined. F. tularensis has a very broad host range but is primarily associated with disease in wild animals. The most common species affected are hares, rabbits, voles, and muskrats. Tularemia has not been transmitted from domesticated rabbits to humans. The most common domestic animals infected are sheep, although stress usually plays a role in developing disease. However, birds and rarely fish, amphibians, and reptiles have also become infected. Dogs are resistant to infection, but cats are susceptible and present with a clinical syndrome similar to that in humans: depression, generalized lymphadenopathy, and hepatosplenomegaly. Necrotic lesions may also be present in the spleen, liver, and lungs. Enterocolitis and ulceration of Peyer’s patches may also be present. Such infections most commonly occur in outdoor cats in areas of endemicity. A similar disease presentation has been noted in prairie dogs (117). F. tularensis is considered a class A bioterrorism agent (like Yersinia pestis and Bacillus anthracis) by the CDC, because it is highly infectious, can survive in water and soil for weeks, and can potentially be transmitted by aerosol. During World War II, the potential use of F. tularensis as a biological weapon was studied by Japan as well as by the United States and its allies. F. tularensis was one of several biological weapons that were stockpiled by the U.S. military in the late 196Os, all of which were destroyed by 1973. The Soviet Union continued weapons production of

1990 and 2000, based counties were reported

on 1,347 In Alaska

of Zoonotic

cases during

Infections

reporting the county this period

17

of the

antibiotic- and vaccine-resistant strains into the early 1990s (Johns Hopkins University Center for Civilian Biodefense Strategies [http://www.hopkins-biodefense .org/index.html]). Mode of Transmission Dozens of biting and blood-sucking insects have been implicated as mechanical vectors of tularemia. Biting flies, ticks, mosquitoes, mites, and fleas have been involved in transmitting tularemia to animals and humans ( 117). Aside from being acquired from the bite of infected arthropods, the infection can be acquired by ingestion of infected tissues or contaminated water, entry of bacteria through broken skin when in contact with contaminated hay, mud, or carcasses; and by inhalation of aerosolized particles that contain F. tularensis. Urine or carcasses from infected voles have been known to serve as a source of epizootics in water-living animals (beavers and muskrats). Transmission from person to person has been reported in the autopsy suite but not from living persons via the respiratory route (250). Human Infection (Clinical Disease) Human cases of tularemia typically are sporadic, but outbreaks do occur if there is a contaminated common source of exposure (200). Tularemia is frequently misdiagnosed early in infection since its symptoms are not unique: sudden onset of chills, fever, headache, and generalized malaise. The incubation period is from 2 to 10 days after infection. Tularemia presents in humans primarily as an ulceroglandular disease in which the bacteria replicate in the skin at the localized site of penetration, leading to formation of ulcers; as a glandular infection, typically resulting from ingestion of contaminated food or water; and less frequently, as oculoglandular (eye

18

Bottone

Table

5.

Laboratory

ldentrfrcatron category

CUMITECH criteria

for

identification

Presumptive

Confirmed

tests

37

of F. fularensis Appropriate laboratory procedures

Cntena

Suspect

Supplemental

et al

1. Clrnrcal symptoms and exposure history compatrble with tularemra and 2. A smear or Isolate that has these charactenstrcs. a Weakly starnrng, tiny, gram-negative coccobacrllr b. Weakly catalase pos rtrve, and negative for oxrdase, beta-lactamase, XV factor or satellite test, and urease 1. The specimen or recovered Isolate IS posrtrve by any one of four tests: a DFA b SA c DNA detection (by PCR or other format) d. Fatty acid profile (MIDI system) or 2 A single serum specimen that has a mrnrmum MA titer of 21 128 or a TA titer of ~I:160 1. The recovered Isolate IS posrtrve by DFA or SA test, DNA detection, or fatty acid profile and has charactenstrc growth on agar or 2. Paired serum specrmens, taken at least 14 days apart, have at least a fourfold rise or fall In titer by MA or TA. One of the pair of serum specimens must have a titer of ~1 :I28 by MA or 21.160 by TA. Nonconfirmatory, for charactenzatron

involvement), septic (becoming blood borne), oropharyngeal (ingestion and swallowing), pneumonic (inhalation or dissemination to the lung), and typhoidal (no obvious portal of entry) forms. Most cases are associated with outdoor activities such as rabbit hunting, muskrat or beaver trapping with handling of diseased animals or carcasses, and being bitten by an arthropod that is carrying the bacteria from another host. Without treatment, nonspecific symptoms usually persist for several weeks, with sweats, chills, progressive weakness, and weight loss characterizing the illness. Any of the principal forms of tularemia may be complicated by bacteremic spread, resulting in pneumonia (common), sepsis (uncommon), and meningitis (rare). The severity of F. tularensis infections varies with the type or strain. Type A isolates are associated with more virulent forms of the disease, while infections with type B manifest in a less virulent course of infection. Both F. tularensis subsp. nouicida and F. tularensis subsp. mediaasiatica result in milder forms of tularemia. Diagnosis Appropriate specimens should be taken for examination, and tests should be performed with proper controls. The levels of laboratory diagnosis are suspect, presumptive, and confirmed (Table 5). Tularemia should be suspected if there are a known exposure to risks associated with tularemia; suggestive clinical symptoms; and the presence of poorly staining, gramnegative, tiny (0.2 to 0.7 km by 0.2 Frn) coccobacilli. F. tularensis is weakly catalase positive and may be differentiated from other small gram-negative bacteria by oxidase, urease, beta-lactamase, and XV factor

Gram stain, catalase, oxrdase, beta-lactamase, XV factor or satellite, urease DFA, SA, DNA, fatty acid profile

Serum

agglutination

DFA and culture, SA and culture, DNA and culture, fatty acid and culture Serum agglutination

Brochemical profile, susceptrbrlrty

antibrotrc

or satellite tests. A presumptive diagnosis is based on a positive result in any one of the following tests performed on specimens taken from ulcer or wound swab, tissues, and cultures: (i) direct fluorescent antibody (DFA), (ii) slide agglutination (SA), (iii) forensic DNA detection by PCR or by Southern blotting, or (iv) fatty acid profile. In addition, a positive serological test result on a single specimen would meet the presumptive criteria. New molecular tests and antigen-IgM capture assays can provide presumptive early diagnostic results, although these tests are not yet fully validated for the clinical laboratory. Confirmation is by the recovery and identification of the causative agent, F. tularensis, by a positive DFA or SA result, DNA detection test, or fatty acid profile and characteristic colonial morphology on agar. However, tularemia is most frequently confirmed by serology, where a change in titer of fourfold or more occurs between two (acute and convalescent) serum specimens taken at least 14 days apart, with one of the pair having a positive titer. It is up to the laboratory to determine which test(s) will be performed based on time, cost, availability of reagents, and biosafety considerations. The DFA test can be performed and read within an hour from the time of receipt of the specimen in the laboratory. Specimens that would likely yield F. tularensis are direct swabs of skin ulcers, affected lymph nodes, liver or spleen, and swabs from the eye or throat as clinical presentation would dictate. The specimen is applied to a heat-fixed smear, and F. tularensis is identified by specific fluorescence following staining with fluorescein isothiocyanate-labeled

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37

hyperimmune anti-F. tularensis antibody. Blood cultures are usually not ideal since few organisms are found in the blood, but prolonged incubation may increase chances of recovery. A positive DFA test is a presumptive result and should be reported to the physician or submitting laboratory. A presumptive positive result also mandates that all further work with the specimen and cultures should be carried out, at a minimum, in a BSL-2 laminar-flow cabinet. However, a false-positive DFA test for Legionella due to F. tularensis has been reported (207). For culture recovery, direct plating on chocolate, Thayer-Martin, Legionella (buffered charcoal yeast extract), and cysteine-supplemented media is recommended. Cysteine heart agar with 9% sheep blood is the optimum medium for growth at 37OC either aerobically or in 5% CO2 and incubated for up to 7 days. Recovery is enhanced if a saline solution of the specimen or triturate of pooled arthropods can be inoculated into susceptible laboratory mice subcutaneously. F. tularensis grows rapidly in spleen and liver, and these tissues serve as a good source of the bacteria. After 48 h of incubation on a plate of cysteine heart agar with 9% sheep blood the colonial morphology of F. tularensis is unique: the raised, round colonies are 2 to 4 mm in diameter; have a smooth, entire surface and a dense, butyrous consistency; and are greenish-white in color, with a distinctive opalescent sheen. Once a culture is recovered, further characterization and supportive biochemical profiling and antimicrobial susceptibility tests may be undertaken. The cellular fatty acid content of Francisella is unique and has distinctive features (115). F. tularensis, is a nonmotile, oxidase-negative organism that is relatively inert, using glucose as its main carbohydrate source. Glycerol fermentation differentiates type A (glycerol-positive) from type B (glycerol-negati ve) isolates. Commercial identification kits should not be relied on for identification, because there are numerous reports of misidentification of F. tularensis (25, 61). In our own experience the Sensititre Ap80 System (Trek) misidentified F. tularensis as Moraxella osloensis. The GN MicroPlate (Biolog) failed to identify the agent to species level, but the only Francisella sp. in its database is F. philomiragia (T. J. Inzana and D. Jones, Abstr. 4Sth Annu. Meet. Am. Sot. Vet. Lab. Diagn., p. 78, 2002). Once identification of F. tularensis is confirmed, a report should be made to the attending physician and to the public health authorities so that proper and timely treatment and control efforts may be undertaken if needed. Serological testing is done either by tube agglutination (TA) or microagglutination (MA) (3 8). A TA titer of 1:160 or more or a MA titer of 1:128 or more denotes a positive serology result. Cross-reactivity with Brucella antigens has been reported, but such

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reactivity is usually below the positive serum titer threshold. ELISAs have been developed, but although ELISA formats are more sensitive and specific than agglutination, they have not been widely adopted by laboratories. For most laboratories, requests for tularemia antibody testing are infrequent. Thus, it is easier to use the agglutination test than the more-laborintensive ELISA format. Antibiotic Susceptibility and Treatment Most laboratories are reluctant to carry out further testing of F. tularensis because of (i) the inherent danger of laboratory-acquired infections, (ii) the rarity of antimicrobial resistance to drugs used for treatment, and (iii) lack of special medium needed to perform antimicrobial susceptibility testing. Typically once an isolate is obtained, it is sent to a reference laboratory where such tests can be performed. F. tularensis infections are treatable with antibiotics, and appropriate and early treatment is effective (182, 215). Streptomycin is the drug of choice. It is given to adults in a dosage of 0.5 to 1.0 g intramuscularly (i.m.) every 12 h for 7 to 10 days. Gentamicin is an alternative drug, which can be given at 3 to 5 mg/kg of body weight/day i.m. in equal divided doses at S-h intervals for 10 days. Tetracyclines (doxycycline) or chloramphenicol may be administered to patients with less severe disease, but treatment failures using these bacteriostatic agents may result in relapses. A dosage schedule of 14 to 21 days may be necessary to prevent relapses. A small number of patients have been successfully treated with ciprofloxacin, suggesting that fluoroquinolones may be useful in treating tularemia. Penicillins and cephalosporins are not effective and should not be used to treat tularemia. Prior to the advent of antibiotics, the overall mortality from infections with the more severe type A strains was in the range of 5 to 10%. However, fatality rates as high as 40 to 60% have been reported for untreated typhoidal and pneumonic forms of disease. Untreated, type B strain infections have been associated with a fatality rate of only 1 to 3%. In the United States, the fatality rate for all forms of tularemia in recent years has been less than 2%. Precautions and Decontamination Standard, universal precautions should be applied for hospitalized patients since person-to-person transmission has not been proven. However, respiratory droplet precautions may be advisable in dealing with cases of overwhelming tularemia pneumonia. Due to the potential for aerosolization, invasive treatments and examination of infected materials should be performed under BSL-3 conditions. Clothing or linens contaminated with body fluids or tissues infected with

Bottone

CUMITECH

et al

FIGURE 2 Scanning electron mlcrograph cells bound to a 0.1.pm-pore-size fllter x3.500 Source CDC Pubk Health Image

of L mterrogans strain RGA Magnlflcatlon, approximately Library (http://phll cdc.gov)

F. tularensis should be disinfected as per routine hospital protocol. The greatest risk of laboratory-acquired tularemia is by aerosol inhalation when working with infected materials and cultures (182). The organism should be handled by trained and, whenever possible, vaccinatCd personnel. F. tularensis may survive for long periods in water, mud, and animal carcasses, and its survival is thought to be enhanced by low ambient temperatures. The bacteria are readily killed by incubation at >6O”C for 2 h or with exposure to common laboratory disinfectants. Laboratory benches should be cleaned with a combination of 10% hypochlorite followed by 70% alcohol. Leptospira

spp.

Description, Natural Habitat, and Disease in Animals Leptospirosis is a zoonosis of worldwide distribution. Although tropical areas have a higher incidence of human disease, cases of leptospirosis have been reported from every continent except Antarctica (52). This disease is caused by infection with Leptospira species, bacteria that are carried and excreted by domestic and feral mammals, birds, and reptiles. Leptospira species are members of the family Leptospiraceae, which are flexible, helical rods, 0.1 pm in diameter and 6 to >12 pm in length. The helical conformation is right-handed, with more than 18 coils per cell. Usually one or both ends of the cells are hooked. An electron micrograph of Leptospira cells is shown in Fig. 2. Individual cells stain faintly gram negative but are best viewed by dark-field or phasecontrast microscopy. Leptospiraceae are motile, with two subterminal periplasmic flagella, and are able to pass through 0.2-km-pore-size filters. These organisms are aerobic and utilize long-chain fatty alcohols as carbon and energy sources. Amino acids and car-

37

bohydrates are not utilized. Purines but not pyrimidines are utilized. These bacteria may be free-living or closely associated with human or animal hosts. There are currently two recognized taxonomic systems for classification of the Leptospiraceae. In the classical system, which is described in Bergey’s Manual of Systematic Bacteriology, the basic taxon is the serovar. Pathogenicity is the key criterion for differentiating between species (130). For example, pathogenic serovars are included in the species Leptospira interrogans and free-living nonpathogenic serovars are included in the species Leptospira biflexa. Failure to grow at 13°C and inhibition of growth by S-azaguanine at 225 &ml are other phenotypic characteristics that differentiate L. interrogans from L. bifiexa. The most recent edition of the revised list of Leptospira serovars contains 212 L. interrogans serovars which are organized into 23 different serogroups and 63 L. biflexa serovars organized into 38 serogroups, based on shared major antigens (138). An example of the nomenclature of the classical taxonomic system would be strain Ictero #l, the type strain of L. interrogans. Strain Ictero #l is in the serovar icterohaemorrhagiae of the serogroup Icterohaemorrhagiae of

L. interrogans. Molecular taxonomic studies of the family Leptospiraceae have shown a high degree of genetic heterogeneity within L. interrogans and L. biflexa (3 1,32, 118, 196, 267). Based on this foundation, a phylogenetic system was developed. In this system, serovars formerly considered to be L. interrogans have been assigned to seven named and five unnamed Leptospira specie&L. interrogans, L. noguchi, L. weilii, L. san-

tarosai, L. borgpetersenii,

L. kirschneri,

L. inadai,

and Leptospira genomospecies 1, 2, 3, 4, and 5. L. bifiexa serovars have been assigned to one of five species in two genera: L. biflexa, L. meyeri, L. wolbachii, L. puma, and Leptonema illini. Leptospires have a worldwide distribution. These organisms are carried, either symptomatically or asymptomatically, by wild or domestic mammals. Birds and reptiles have also been identified as carriers (84). The natural reservoir of the pathogenic leptospires is the lumen of nephritic tubules (238). Organisms are excreted in the urine of infected animals and, in tropical areas, may survive for an extended time in freshwater, soil, or mud provided the conditions are suitable. Leptospires do not survive well in saltwater and conditions that include low pH, low humidity, and low temperature. In addition to being transmitted through urine, leptospires may be transmitted through milk, placenta, and aborted fetuses. Infections in animals vary depending on whether the disease is acute or chronic and may involve renal disease, abortion and infertility, hepatitis, mastitis, arthritis, and focal lesions in other internal organs.

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37

The distribution of Leptospira serovars varies by region and host species. In the United States serovars isolated from humans include icterohaemorrhagiae, bataviae, canicola, autumnalis, intrav, and mini. Serovars commonly isolated from feral animals include ballum, zanoni, and australis. Knowledge of the predominant serovars of the test area is important for effective leptospirosis diagnosis. The most significant infections occur in cattle, swine, and dogs, and these animals are a major source of leptospires transmitted to humans. Mode of Transmission The most common mode of transmission of leptospires to humans is by contact of exposed skin with water or mud that has been contaminated with urine from an infected animal. Organisms may also gain access through mucous membranes of the oropharynx, conjunctiva, or genital tract (85). Recent outbreaks involving large numbers of cases have occurred after periods of heavier-than-normal rainfall and flooding (275). Occupations involving contact with infected animals or contaminated water, such as farming, military service, or animal husbandry, are associated with increased risk of infection. Human Infections Leptospirosis in humans may be manifested in a variety of clinical symptoms, ranging from a mild self-limiting febrile illness (most cases) to a fulminating fatal illness associated with hepatorenal failure (Weil’s disease). The incubation period of this disease is usually 7 to 14 days but can range from 2 to 21 days (84). Mild cases of disease are characterized by the sudden onset of low-grade fever, headache, and muscle pain, which may last between 1 day and 2 weeks. Due to the nonspecific nature of these symptoms, mild cases of leptospirosis may be confused with a viral illness. Severe cases of leptospirosis are characterized by a biphasic manifestation, which includes a septicemic phase lasting 4 to 7 days and an icteric phase that begins 1 to 3 days after the septicemic phase resolves. The septicemic phase begins with the abrupt onset of rapidly rising fever, chills, severe headache, and myalgia, which is often most pronounced in the thighs and calves. The icteric phase is characterized by jaundice associated with hepatic and renal failure. The mortality rate in untreated cases of severe disease may be as high as 15 to 40% (84). Diagnosis Blood, cerebrospinal fluid (CSF), and urine are the specimens of choice for the recovery of leptospires from patients with leptospirosis. The most appropriate samples to culture during the first 10 days of illness are blood and CSF. These specimens should be collected prior to antibiotic treatment and while the patient is febrile. Specimens should be collected using

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21

aseptic techniques and immediately inoculated into tubes containing 5 ml of a semisolid leptospira growth medium, such as Fletcher’s, Ellinghausen’s, or PLM (polysorbate liquid medium) (84,225). For blood, the inoculum volume should be kept to a minimum due to the presence of inhibitory substances. No more than 3 drops (approximately 50 ~1 per drop) should be added to 5 ml of medium. Satisfactory results may be obtained with an inoculum of 1 drop of blood/5 ml of medium. If culture media are not immediately available, blood should be collected into tubes containing heparin or sodium oxalate for transport. Tubes containing citrate solutions should be avoided because they may be inhibitory (2). Specimens should be stored and transported at room temperature and inoculated within 1 week of collection. For CSF, inocula as high as 0.5 ml of undiluted specimen per 5 ml of medium can be used. After the first 10 days of illness, the optimal specimen for isolation of leptospires is urine. Extreme care should be taken to collect an uncontaminated specimen, since most urine contaminants will overgrow leptospires in culture. Cultures should be inoculated as soon as possible, especially if the pH of the urine is acidic. The specimen should be prediluted l:lO, l:lOO, and l:l,OOO in either culture medium or phosphate-buffered saline (PBS), pH 7.2 to 7.8, and 2 to 3 drops of each dilution should be inoculated per 5 ml of culture medium. If culture media are not immediately available, the urine can be diluted 1:10 in 1% bovine serum albumin and stored at room temperature (64). For specimens obtained from field work, media can be made selective by the addition of 5-fluorouracil(lO0 to 300 pg/ml) or neomycin (6 kg/ml). In fatal cases of leptospirosis, viable leptospires can be isolated from multiple tissues, particularly the liver, kidney, and brain (2,64). Samples should be collected and processed within 4 h of death since leptospires will not survive in autolytic tissues. However, some serovars of leptospira may be very difficult to grow in vitro, even if present. For epidemiological investigations, freshwater ponds represent potential sources of pathogenic leptospires. Water should be collected in sterile containers and transported at room temperature to the laboratory within 72 h of collection (126). A presumptive identification of leptospires can be made by direct microscopic methods. Blood, CSF, and tissues may be screened microscopically for the presence of leptospires by either dark-field microscopy or DFA assay. Blood is prepared for analysis by mixing 5 ml of freshly drawn whole blood with either 0.5 ml of 1% sodium oxalate or 1.0 ml of 1% heparin. The mixture is then centrifuged at 500 X g for 15 min, and the supernatant is transferred to another tube and centrifuged at 1,500 X g for 30 min. The supernatant

22

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FIGURE Source:

et al.

3 Histopatholoav of kldnev showlna leptomres CDC Public Health’lmage Library (http~//phil.cdc.gov).

CUMITECH

(arrows)

is discarded, and a wet mount is prepared from the sediment. For urine and CSF, the sample is centrifuged at 1,500 X g for 30 min, the supernatant is discarded, and a wet mount is prepared from the sediment. Tissues may be prepared for analysis by disruption in a sterile Ten Broeck tissue grinder or ground with mortar and pestle. The suspension is mixed with 9 parts sterile PBS (pH 7.2 to 7.8), and the resulting suspension is centrifuged at 500 X g for 15 min. The sediment is discarded, and the supernatant is poured into another tube and centrifuged at 1,500 X g for 30 min. The supernatant is discarded, and a wet mount is prepared from the sediment (134). Although a specific transport medium is usually not required, if there is a lengthy delay in culture, specimens (particularly tissues) can be transported in Stuart’s or EllinghausenMcCullough/Johnson-Harris (EMJH) medium or 1% bovine serum albumin containing fluorouracil (200 pg/ml) or neomycin sulfate (300 p&ml). Examination of specimens processed as indicated above by dark-field microscopy can be useful in establishing a rapid, presumptive diagnosis. However, this technique should not be relied upon as the sole diagnostic criterion. In specimens in which the numbers of leptospira are few, observing the organisms may be difficult, even after centrifugation of the specimen. It is also important to differentiate leptospires from artifacts, such as fibrils or cellular extrusions. If a fluorescence microscope is not available, several silver staining techniques can be used to visualize leptospires in tissues (225). A silver-stained preparation of human kidney tissue containing leptospires is shown in Fig. 3. As with the other microscopic techniques described in

Dleterle

sliver

stain

was

used

Magnlflcatlon,

approximately

37

xl80

this section, the sensitivity of the technique is poor in samples in which the number of leptospires is small. A slightly more sensitive technique for visualizing leptospires in specimens involves staining them with fluorescein-conjugated antibodies and observing with UV microscopy. The National Veterinary Service Laboratory (Animal and Plant Health Inspection Service) of the U.S. Department of Agriculture, Ames, Iowa, produces multivalent conjugate reagents for the veterinary profession. These conjugates are available through local Animal and Plant Health Inspection Service representatives. The fastidious nature of the leptospires makes them good candidates for the development of rapid and sensitive nucleic acid hybridization-based assays. In recent years PCR-based methods have been developed for the rapid detection of Leptospira DNA in human specimens (37, 169). Two of these molecular methods have been evaluated in comparison to culture and serology with specimens from patients with active disease, and both appear to be more sensitive than culture in patients with serologically confirmed disease (37, 169). PCR analysis is most useful within the first 10 to 14 days after the onset of clinical symptoms, when leptospiral DNA may be detected prior to the development of the humoral immune response. Specimens in which leptospiral DNA has been detected by PCR include serum, urine, aqueous humor, and CSF. When leptospires are cultivated in semisolid media, leptospiral growth will initially appear as a diffuse zone near the top of the tube. As the culture matures the leptospires will condense into a well-differentiated

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ring located at the level of the tube corresponding to the optimum oxygen tension for the organisms (Dinger’s Ring). In general, the saprophytic serovars will form rings closer to the top of the tube than the pathogenic serovars. Saprophytic serovars can also be differentiated from pathogenic serovars by their ability to grow at 13°C and in the presence of 225 pg of 8-azaguanine per ml (2). Leptospiral growth can be confirmed by examining a sample of the growth ring by dark-field microscopy. Cultures in semisolid media should remain viable for at least 8 weeks at room temperature. For permanent storage, cultures in semisolid medium can be frozen in liquid nitrogen with the addition of either glycerol or dimethyl sulfoxide (2). Once grown in culture, leptospires can be identified to the serogroup level by the microscopic agglutination test using reference rabbit sera prepared against the type strains of serovars known to cause disease in the local area (225). The microscopic agglutination test is performed on isolates that have been serially passed at least three times in liquid medium, such as PLM liquid. This effectively dilutes agar granules that may produce artifacts and reduces nonspecific agglutination. Reference sera are diluted with PBS in twofold steps from 1:25 through 1:3,200. Subsequently, 50 ~1 of each serum dilution is mixed with 50 ~1 of an antigen suspension standardized at 100 to 200 organisms per high-power field. The antigen-antiserum mixtures are incubated at room temperature for 2 to 4 h in a 96-well flat-bottom microtiter plate and read by dark-field microscopy. At the Leptospirosis Reference Laboratory of the CDC, isolates from the United States are serogrouped using a battery of hyperimmune rabbit sera made against serovars from the following serogroups: Ballum, Canicola, Icterohaemorrhagiae, Bataviae, Grippotyphosa, Pyrogenes, Autumnalis, Pomona, Sejroe, Australis, Tarassovi, Mini, Cynopteri, Celledoni, Diasiman, Hebdomadis, Shermani, Javanica, and Andamana. A battery this extensive might not be necessary if the common local serovars are known. Final identification of an isolate to the serovar level requires the use of the agglutinin adsorption technique (225). This technique involves raising isolatespecific hyperimmune rabbit antiserum and requires the maintenance of many reference serovars and antisera. Final identification to the species level using the phylogenetic taxonomic approach requires nucleic acid hybridization studies. The complexity of these techniques limits their use to reference laboratories. The reference laboratory for human leptospirosis for the United States is the WHO/FAO Collaborating Center for Reference and Research on Leptospirosis at the CDC, Atlanta, Ga. Some other international reference laboratories for leptospirosis include the WHO/FAO Collaborating Centers in Amsterdam,

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23

The Netherlands, and Brisbane, Australia. Additional information regarding leptospirosis is available on the World Wide Web at the International Leptospirosis Society website (http://www.med.monash.edu.au /microbiology/staff/adler/ilspage.htm). Less complex alternatives for identification are currently under development. Whole-chromosome restriction endonuclease patterns and ribotyping are used for serovar identification in some reference laboratories (157), and PCR-RFLP methods that differentiate between Leptonema illini and Leptospira species (261) and between L. interrogans and L. biflexa (262) have been described. Random amplified polymorphic DNA fingerprinting and arbitrarily primed PCR represent two extremely promising approaches for identification. Within the past 4 years, investigators, using primarily reference strains, have successfully applied these methods to identify serovars (194, 195). As these methods are applied to more clinical isolates, the utility of these approaches will become more apparent. Antibiotic Susceptibility and Treatment Due to the fastidious nature of leptospires, standard methods for determining in vitro antimicrobial susceptibility have not been established. However, clinical and animal studies indicate that intravenous penicillin G and ampicillin are useful in the treatment of severe cases. In less severe cases where oral administration is possible, doxycycline, ampicillin, and amoxicillin are recommended ( 85). Doxycycline has also been shown to be effective as a prophylactic agent (227). Laboratory Safety and Precautions Laboratory-acquired leptospirosis has been well documented, particularly in cases of handling experimentally infected animals or tissues (171, 188). Infected animals, specimens, and cultures should be handled under BSL-2 laboratory conditions. Gloves should be used when handling infected animals, their tissues, or their body fluids. Yersinia

pestis

Description, Natural Habitat, and Disease in Animals The plague, known as the Black Death in the Middle Ages, is a feared and devastating disease with a remarkable place in history. For centuries, plague represented disaster for those living in Asia, Africa, and Europe, where, it has been said, populations were so affected that sometimes there were not enough people left alive to bury the dead. The fundamental but separate works by Yersin and Kitasato in 1894 on the discovery of the etiologic agent of plague in Hong Kong opened the way for investigating the disease and its mode of transmission. Kitasato and Yersin de-

24

Bottone

et al.

scribed, within days of each other’s findings, the presence of bipolar staining organisms in the swollen lymph node (bubo), blood, lungs, liver, and spleen of patients who had died (91). Cultures isolated from patient specimens were inoculated into a variety of laboratory animals, including mice. These animals died within days after injection, and the same type of bacilli as those found in patient specimens were present in the animal organs. Yersin recorded that black or roof rats (Rattus rdttus) were not only affected by plague during epidemics but often were found dying of Y. pestis infection preceding such epidemics in humans. The transmission of plague organisms was first described by Simond in 1898 (91). He demonstrated that the rat flea (Xenopsylla cheopis) transmitted the agent in a now classic experiment in which a healthy rat, separated from direct contact with a rat that had recently died of plague, also died of plague after the infected fleas jumped from the first rat to the second. So imprinted in our minds is the fear of plague that, even now in the 21st century, a suspected plague outbreak can cause mass panic and bring enormous burden to much of the medical and public health services, as it did in 1994 in India (60). Plague in the 21st century has changed from its historical classic urban, or rat-borne form to an enzootic form affecting sylvatic rodents in rural regions. Although the number of human plague infections is low, plague still invokes an intense, irrational fear, disproportionate to its transmission potential in the postantibiotic and postvaccination era. Unfortunately the fear of plague has again become a public health concern because of its potential for use in bioterrorism. Plague is an ancient disease that is not likely to disappear, and continued outbreaks throughout the world attest to its tenacious presence. Plague is established in rodent populations in widely scattered rural areas throughout the world except for Australia. The organism cycles naturally in its enzootic foci, circulating between small mammals and fleas without human involvement. The quiescent periods, during which few or no human cases are detected, may last for years, leading to mistaken declarations of plague eradication. There have been three pandemics of plague, and with each one, the range of Y. pestis has expanded. In some of the range expansion, a region might have been only transiently affected, while in other regions, the expansion led to the establishment of new foci of Y. pestis among local species of rodents and their fleas. The modern pandemic began in 1886 in the Yunnan Province of China, reached Hong Kong by 1896, and from there infected rats (both R. rattus and Rattus norvegicus) were transported aboard ships to North and South America, Southern Africa, Southea st Asia, and i .sland nations such as Madagascar (91).

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In the United States plague is endemic in the western states, and its presence in the counties of these states is monitored by a combination of the presence of specific antibodies in carnivores and by the isolation of plague bacteria from animals and humans (Fig. 4). Plague does not exist in the eastern states of the United States. Sylvatic plague has been associated with 76 species of mammals in the United States, but most of these are likely to be “accidental” hosts of Y. pestis and play only a minor role in the ecology of plague (91). The primary amplification of Y. pestis is limited to a few host species of rodents and their fleas: (i) ground squirrels of the genus Spermophilus (S. variegatus, S. beecheyi, S. elegans, S. beldingi, S. townsendi, S. armatus, and S. lateralis) and their fleas (Oropsylla montana, Oropsylla labis, Oropsylla idahonesis, Oropsylla tuberculata, Hoplopsyllus anomalus, Thrassis bacchi, Thrassis francisi, Thrassis pandorae, and Thrassis petiolatus); (ii) the chipmunk family (Tamias spp.) and their fleas (Eumolpianus eumolphi and re lated species and Ceratophyllus cilia tus); (iii) prairie dogs (Cynomys gunnisoni, Cynomys ludovicianus, and Cynomys leucurus) and their fleas (Oropsylla hirsuta and 0. tuberculata); and (iv) woodrats (Neotoma spp.) and their fleas (Orchopeas, Anomiopsyllus, Stenistomera, and Megarthroglossus spp.). Maintenance, or enzootic, plague is believed to be carried by mice (Peromyscus maniculatus) and voles (Microtus californicus) and their flea s (92). Epizootic plague is present in diverse habitats and is more likely to occur when high densities of multiple susceptible hosts and fleas coexist. Such conditions are found i n the moun tainous and plateau regions of the southwestern United States. Rodents are highly susceptible to infection by Y. pestis. Experimental infection of California ground squirrels by McCoy (161) indicated that the disease could take one of three forms: (i) an acute infection, with the animals dying within 3 to 5 days (hemorrhagic buboes and splenomegaly were present, but there were no lesions in the liver or spleen); (ii) a subacute form in which the animals died 6 or more days after infection and had necrotic buboes and necrotic lesions in the liver and spleen; and (iii) a convalescing form, with only lymphadenopathy and necrotic foci present. Other studies have shown that there is considerable variation in the susceptibility of individual squirrels within a population ( 189). Carnivores -particularly dogs, but also coyotes, raccoons, and skunks-appear to be much more resistant to Y. pestis infection than rodents. Infection appears to be the result of ingestion of plague-infected animals rather than by flea bites. Seroconversion occurs, but the animals either do not become ill or have only moderate illness with a mild to moderate febrile response, and Y. pestis cannot be isolated from the blood or feces (210). In contrast,

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FIGURE reports

Diagnosis

37

4.

Reported

cases

of plague

m the western

United

States

cats appear to be almost as susceptible to Y. pestis as rodents. Following oral infection cats become bacteremit and may die. Lesions are present in the liver and spleen, and a suppurative pneumonia may occur; buboes may also be present (189,210). In a CDC study of 16 infected cats, 88% became ill, 75% became bacteremic, and 38% died. Furthermore, natural plague pneumonia in cats has been observed during epizootics of rodent plague (189). Mode of Transmission Plague is a bacterial infection of small mammals transmitted from animal to animal by the bite of infected fleas. Rodent hosts become bacteremic when infected with Y. pestis, and they serve as the infective blood meal for fleas. Blood is the sole source of nutrition for fleas. The bacteria multiply in the flea proventriculus and midgut, where they grow and coalesce to form sticky masses that eventually occlude the proventriculus and gut tract. Because blood cannot pass beyond the blocked proventriculus, the flea begins to starve and will attempt to feed repeatedly on almost any available host. The repeated feeding leads to regurgitation of infective Y. pestis, thereby transmitting the bacteria to the host. Rodent-consuming carnivores

between

1970

and 2001.

Shaded

of Zoonotic

areas

show

Infections

counties

with

25

oositlve

may then become infected through ingestion of infected prey or being bitten by the infected fleas. Depending on the relative resistance of the carnivore, the infection may be of short duration, or if the animal is susceptible, it may become an amplifying host source for further transmission of the agent. As mentioned above, the Cmis genus (dogs and coyotes) and foxes (Vulpes and Urocyon spp.) seroconvert if exposed to plague but rarely succumb to infection. Thus, they become good sentinel animals for plague. In contrast, the felids (cats and bobcats) are very susceptible to disease, and exposure via close contact or handling of infected cats and bobcats has led to human cases. The greatest risk to humans for acquiring plague is during plague epizootics. Humans acquire plague from the bite of infected fleas, from direct contact with contaminated tissue, and by inhalation of bacterium-laden droplets (91). The ingestion of infected animals has also been documented to result in transmission of plague to humans, but this mode of transmission is very rare (59). Bubonic plague is the most common form of infection, comprising nearly 8.5% of reported plague cases. Bubonic plague results from the bacteria being taken up by host macrophages in

26

Bottone

Table

6.

Laboratory

et al

CUMITECH

criteria

for

identification

of Y. pestis

ldentrfrcatron category

Presumptive

Confirmed

Supplemental

Approprtate laboratory procedures

Cntena

Suspect

tests

I. Clinrcal symptoms and exposure history compatrble with and 2 A smear or Isolate from affected trssues-lymph node (septrcemic), tracheal or lung aspirate (pneumonrc)-with charactenstrcs: a. Small, gram-negative and/or bipolar-starnrng, single or short chains b. Catalase posrtrve, oxrdase and urease negative 1. The specimen or recovered Isolate IS posrtrve by either a DFA stain b DNA detection or 2 A single serum specimen that has a mrnrmum PHA/Hlb or a posItIvea ELISA titer 1 The recovered Isolate IS posrtrve by DFA or DNA detection by specrfrc bacteriophage at both ambient temperature and 37°C or 2. Paired serum specrmens, taken at least 14 days apart, a fourfold rise or fall In titer by PHA/HI or ELISA Nonconfrrmatory, for charactenzatron of the Isolate

a Positwe ELISA toters vary with each format Known rum samples must be tested at different drlutrons b PHA/HI, passive hemagglutrnatron and rnhrbrtron

37

(I

negatrve and posrtrve serum samples e , 1 100, 1 1,000) and In duplicate

the lymph node closest to the site of the flea bite. The affected lymph node becomes inflamed (bubo), enlarged, and painful as the bacteria replicate. From multiplication within the infected lymph node, bacteria sometimes become blood borne (septicemic) and occasionally lodge in the lungs (pneumonic). When plague infection becomes pneumonic, direct personto-person transmission of the bacteria via respiratory droplets becomes possible (46). Progression of pneumonic plague is rapid, and if untreated, it may lead to death in a few days. However, pneumonic plague is rare and requires close contact for transmission to occur. Early diagnosis with prompt antibiotic treatment is effective against all forms of plague infection, and antibiotic resistance to plague is rare (69). Human Infections The most common naturally occurring clinical presentation in humans is bubonic plague, which occurs following the bite of an infected flea. Following an incubation period of 2 to 8 days patients have a sudden onset of fever, chills, lethargy, and headache, and shortly thereafter develop painful lymphadenopathy (bubo), usually in the groin, axilla, or neck. The buboes are very painful, 1 to 10 cm in length, and raised under the intact skin. Hepatosplenomegaly is often present. Skin lesions may occur but are not common. Overwhelming infection may result in bacteremia, which may be common even in acute bubonic plague. However, moribund patients often have such a high density of bacteria in the blood that a direct micro-

plague

Infection

(bubonic), blood the following plump

rods,

of two

tests:

titer

Gram stain, differential stain, culture, catalase, oxrdase, urease

DFA

of 1 :I 0

or DNA

Serological

and IS lysed (22 to 25°C)

Bacteriophage

have

Serological

at least

detection

assays lysls

assays

Brochemrcal profile, braI susceptrbrlrty must

be run together

with

test

samples

Antrmrcroprofile

for each

test

Se-

scopic exam of the blood may demonstrate characteristic bipolar staining bacilli. Disease in which patients die with bacteremia without development of buboes is referred to as septicemic plague. Pneumonic plague can result from hematogenous spread of bacteria from the bubo, by transmission from another patient with pneumonic plague by coughing, or even by domestic cats with pneumonia or submandibular abscesses (54). Plague pneumonia is highly contagious, and patients often present with fever, buboes, cough, chest pain, and hemoptysis. Plague pneumonia is uniformly fatal if antibiotics are delayed more than 1 day. Therefore, aerosol transmission of Y. pestis in a bioterrorism attack is of grave concern. Rare complications of plague are meningitis, pharyngitis, or gastrointestinal symptoms (43). Diagnosis Y. pestis, the etiologic agent of plague, is a pleomorphic, non-spore-forming, gram-negative coccobacillus (0.5 to 0.8 pm by 1 to 2 km) that is catalase positive and negative for urease and oxidase. The laboratory criteria for diagnosis of plague have been categorized as suspect, presumptive, and confirmed (60) (Table 6). Appropriate specimens should be examined for evidence of plague if an infected animal or person resides in or has a recent history of travel to plague-infected areas and presents with symptoms suggestive of plague (fever, sepsis syndrome, lymphadenopathy, and/or acute pneumonitis). The preferred specimen for microscopic examination and isolation from a patient with bubonic plague

CUMITECH

37

is aspirated fluid from the affected bubo, which should contain numerous organisms. Blood for culture should be taken whenever possible. Organisms may be seen in blood smears if the patient is septicemic, while blood smears taken from suspected bubonic plague patients are usually negative for bacteria. Bacteria may be intermittently released from affected lymph nodes into the bloodstream; therefore, a series of blood specimens taken about 30 min apart may be productive in the isolation of Y. pestis. Sputum or throat smears taken from pneumonic plague patients may contain too many other organisms to be of diagnostic value when only Gram’s or a differential stain is used. These smears should be examined with the more specific DFA test to obtain a presumptive diagnosis. Preferably, a bronchial or tracheal washing specimen should be taken from a suspected pneumonic plague patient for culture instead of sputum. In biopsy specimens or in specimens taken postmortem, where recovery of live organisms is compromised, lymphoid tissues, lung, and bone marrow samples may yield evidence of plague infection by the DFA test or by detection of Y. pestis DNA. In the initial diagnostic workup, specimens intended for culture should be taken before initiation of antibiotic treatment. Specimens are inoculated onto general laboratory media and, if available, into laboratory mice, for isolation. A thin smear is made from the remaining materials for examination by fluorescence microscopy. If a specimen is suspected to contain mixed flora, passage of the material through laboratory mice will increase the likelihood of recovery of pure Y. pestis. Plague bacilli express a unique diagnostic envelope glycoprotein called the fraction 1 (Fl) antigen or capsular antigen at ~33°C; this unique envelope antigen is the primary target antigen used for plague DFA tests and other antibody-based tests. Plague bacilli are susceptible to lysis by a specific bacteriophage at both 2.5 and 37OC. A positive bacteriophage lysis test result is confirmatory for Y. pestis. Plague bacilli are relatively inactive by standard enteric biochemical reactions. Therefore, biochemical profiles should be used as a supplemental diagnostic test. If a patient has been treated with a bacteriostatic antibiotic (e.g., tetracycline) for more than 4 days, bacterial cultures should be incubated for more than 5 days to enable the organisms to recover. Serologic testing is advised in the event cultures yield negative results. One serum specimen should be taken as early in the illness as possible, to be followed by a second sample 4 weeks or more after onset of disease and after antibiotic therapy has ceased. Bone marrow from desiccated animal carcasses may yield positive results when other tissues are not available or are heavily contami nated. In addition, serum and blood specim .ens may be taken for detec-

Diagnosis

of Zoonotic

Infections

27

tion of specific antibody. Fleas should be identified and placed in pools by species. A flea pool is then triturated and inoculated into laboratory mice for isolation of plague bacteria. The serum from inoculated laboratory mice may be examined for the presence of antibody to Fl after an appropriate incubation time. For serosurveillance of plague in animal populations, filter paper strips may be soaked with blood, dried, and sent to the laboratory for the detection of antibody to Fl. Lastly, as with human specimens, in cases where no specimens or sera are available for testing, both animal and flea material may be tested by PCR to determine if plague DNA is present in the specimens. Y. pestis grows well, but slowly, on all general media. Standard sheep BA is used for culture; day-old to 4%h-old growth in broth can be described as suspended flocculent or crumbly clumps ( “stalactites”). These clumps are visible at the side and bottom of the tube; the rest of the medium remains clear. Longer incubation times will result in the clumps of cells falling to the bottom of the tube and loss of the characteristic formation, but the medium above will remain clear. Yersinia pseudotuberculosis and Streptococcus pneumoniae may exhibit the same type of clumping. Therefore, this characteristic growth formation in broth is not diagnostic for Y. pestis. Subculture onto sheep BA, identification by the DFA test, and bacteriophage lysis should be performed to validate the observation. Colonies of Y. pestis are gray-white, translucent, and usually too small to be seen as individual colonies after 24 h of incubation at 37OC. After 48 h, colonies are about 1 to 2 mm in diameter, gray-white to slightly yellow in color, and more opaque, with a raised, irregular, “fried egg” morphology, which becomes more prominent as the culture ages. Colonies also can be described as having a “hammered-copper,” shiny surface. There is little or no hemolysis of the sheep red blood cells. Characteristic colonies grown on BA should also be examined by the DFA test, bacteriophage lysis, and supportive biochemical reactions to distinguish the isolate from Y. pseudotuberculosis or other bacteria that have similar colony morphology and minimal biochemical metabolism. Y. pestis has been misidentified as Y. pseudotuberculosis, other Yersinia spp., Shigella spp., H,S-negative Salmonella spp., and Acinetobacter sp. (258). Antibiotic Susceptibility and Treatment Prompt diagnosis combined with appropriate antimicrobial treatment are important for successful resolution of plague infection. All attempts must be made to recover live bacteria so that an in vitro antimicrobial susceptibility profile may be ascertained. Although a multidrug-resistant isolate from Madagascar has been recovered from a patient with bubonic plague, antibi-

28

Bottone

et al

otic resistance has not been problematic in treating plague. Naturally occurring isolates of Y. pestis resistant to the antibiotics of choice are rare (69). The potential for the spread and public health implication of multidrug-resistant Y. pestis needs to be monitored by research and heightened surveillance. Untreated plague is fatal in more than 50% of patients with the bubonic form of the disease and in nearly all persons with septicemic, pneumonic, or meningeal plague. The recent overall mortality in plague cases in the United States is nearly 1.5%, and fatalities almost always arise from delays in seeking treatment or in making the correct diagnosis. The drugs of choice for treatment of plague are streptomycin and gentamicin (68). Tetracyclines, sulfonamides, and chloramphenicol are also very effective alternative drugs for treatment. Penicillins and cephalosporins are not recommended. Penicillins appear effective in inhibiting Y. pestis in vitro. However, this antibiotic is considered ineffective against resolution of human disease. Therapy should ensue for 10 days, and for at least 3 days beyond apparent clinical recovery to ensure complete resolution of the infection. Streptomycin should be given i.m. at 2 g/day in two doses per day for adults and at 30 mg/kg/day in two to three doses for children. Gentamicin should be given at a dosage of 3 mg/kg/day by i.m. or intravenously divided into three doses per day and at 6.0 to 7.5 mg/kg/day for children. Tetracycline, doxycycline, and chloramphenicol may be taken for prophylaxis orally (68). Ciprofloxacin and its fluoroquinolone derivatives have been used for treatment, but their efficacies have not been verified by clinical trials. Precautions and Decontamination Prevention of plague is attained by reducing risk of exposure and in seeking prompt treatment. All patients with suspected plague should be isolated for at least 48 h after treatment begins to reduce the risk of person-to-person transmission (68). Patients with pneumonic plague should be isolated until sputum or tracheal cultures are negative for Y. pestis. Individuals who have had significant and close contact with pneumonic plague patients should be advised of their risks, monitored for illness, and offered prophylactic antibiotic therapy. Persons who have had more-distant contact (2 m or further) are unlikely to become infected but should be informed of their risks and monitored closely during the week following the potential exposure. Epidemiologists and surveillance personnel should act quickly to identify the likely exposure sites, assess the potential risks, and institute control measures to limit the risk of further infections. Vaccination is recommended only for persons at high risk, such as research laboratory workers, fi.eld investiga tors, or others who may be repeatedly exposed to source s of Y. pestis. All suspected plague cases should be re-

CUMITECH

37

ported to national health authorities, and all laboratory-confirmed cases must be reported to the World Health Organization, Geneva, Switzerland. Plague is one of three international diseases subject to quarantine. Universal precautions with protective clothing, gloves, and eye protection should be used, and cultures or infectious materials should be handled in a class II or class III biological safety cabinet in a BSL-3 laboratory. Y. pestis is readily killed by incubation at temperatures of >6O”C for 2 h or by exposure to common laboratory disinfectants. Laboratory benches should be cleaned with a combination of 10% hypochlorite followed by 70% alcohol. Rare, Miscellaneous

Bacterial

Agents

Rarely, bacterial agents not normally associated with zoonotic infections are isolated from human clinical specimens. Often these infections result from direct contact, such as from bite wounds. However, in immunocompromised patients, direct contact can often not be documented. Below is a brief description of bacterial agents that are part of the normal flora of animals and that have rarely caused infections in humans. Of members of the Pasteurellaceae, Pasteurella multocida is the most common cause of human infections (reviewed in Cumitech 2 7 [29]). However, other species may, less often, also cause infections in humans. Such infections are usually obtained from bite wounds from dogs or cats. Pasteurella caballi, which may be part of the normal upper respiratory tract flora of horses, has been isolated from a wound of a veterinary surgeon (26) and from the hand wound of a 56-year-old man following a horse bite (81). Other Pasteurellaceae species from food animals and horses that have rarely caused human infections include ” Pasteurella aerogenes” (bite wounds) (146), and Mannheimia (Pasteurella) haemolytica (endocarditis) (266). Five cases of human infections due to the Pasteurella SP group, which are normally isolated from guinea pigs and rabbits, have also been reported (147). In addition, Actinobacillus lignieresii, A. equuli, and A. suis have been responsible for infections due to horse or sheep bites (23, 71, 186). Most members of the Pasteurellaceae are difficult to identify to the species level. Commercial kits and automated systems generally are inadequate for species identification. However, most species of Pasteurellaceae are oxidase positive and weakly fermentative. Thus, a gram-negative coccobacillus that is oxidase positive and is a weak lactose or sucrose fermenter (uniform orange color change in a TSI agar tube) should be considered a possible member of the Pasteurellaceae. Conventional biochemical tube tests should be used for further identification, as described

CUMITECH

37

by Bergey’s Manual of Systematic Bacteriology (154). A. lignieresii may require incubation on media containing blood or serum for initial isolation. A. equuli and A. suis are hemolytic on BA and may also grow on MacConkey agar as small, red (lactose-positive) colonies. Staphylococcus caprae is a coagulase-negative, DNase-positive member of the genus Staphylococcus that is associated with goats. However, since 1991 this species has occasionally been isolated from human clinical specimens. Shuttleworth et al. (217) described 14 strains isolated from human specimens. The majority of these strains were from bone and joint infections. A close association with goats was not a risk factor, but wearing an orthopedic prosthesis was. Commercial identification systems may misidentify this organism as other Staphylococcus spp., such as S. carnosus, S. haemolyticus, S. aureus, S. warneri, S. simulans, or S. epidermidis, because it is not adequately represented in their databases. However, the species may be presumptively identified based on positive results for DNase, pyrrolidonyl aminopeptidase, and acid production from mannitol and maltose; it is negative for ornithine decarboxylase and tube coagulase (217). Streptococcus suis is primarily a pathogen of swine, causing arthritis meningitis, pneumonia, septicemia, endocarditis, bol y serositis, , abortions, and abscesses (221). Infections in humans most often occur in abattoir workers and swine handlers (203). There are 3.5 serotypes of S. suis, but type 2 is the most common serotype to infect humans; types 4 and 14 have also been reported (240). Type 2 strains have been reported to cause meningitis and/or septicemia with deafness, or diplopia; arthriti .s, acute gastroenteritis, endophthalmitis, and death have also been reported (221). Four cases of human endocarditis have been attributed to S. suis type 2 (233). S. suis type 2 has been isolated from the tonsils of people working in slaughterhouses, and antibodies to S. suis type 2 may occur in 10% of meat inspectors and 21% of pig farmers (203). S. suis type 2 colonies are alpha-hemolytic on sheep BA and beta-hemolytic on horse BA. Biochemical typing is highly variable and requires a large battery of biochemical tests. Some commercial systems are capable of identifying this species, such as the Rapid ID 32 Strep (bioMerieux-Vitek, Marcy-1’Etoile France) (233). Confirmation is best done by serotyping at a reference laboratory, such as the National Veterinary Services Laboratory (Ames, Iowa) Strep tococcus iniae, a pathogen of aquatic animals, has recently been shown to cause serious infections in humans as well. S. iniae has been reported to cause skin lesions in freshwater dolphins and meningitis and mortality in rainbow trout and tilapia (76, 77, 187).

Diagnosis

of Zoonotic

Infections

29

Infections in humans are associated with handling and cleaning of freshwater fish, most commonly tilapia. Infections have included cellulitis, meningitis, endocarditis, and possibly septic arthritis (5.5, 252). On initial isolation S. iniae is alpha-hemolytic and may be misidentified as a viridans group streptococcus. Under anaerobic conditions it may appear beta-hemolytic and could be confused with Streptococcus pyogenes because it is pyrrolidonyl arylamidase positive and may be susceptible to bacitracin. However, it is not typeable into any of the established Lancefield groups. S. iniae is negative for growth on bile esculin agar and does not grow in 6.5% NaCl within 24 h. Esculin and arginine are hydrolyzed, and the CAMP test is positive. PCR amplification of the universal chaperonin 60 gene followed by reverse checkerboard hybridization has been shown to identify this species (98). S. iniae should be considered in cases of wounds of patients with a history of handling freshwater fish. Confirmatory identification can be done at reference laboratories such as the CDC. Miscellaneous bacteria that may be transmitted from fish and cause infections in humans include Vibrio carchariae (MS), Vibrio uulnificus (3; J. Veenstra et al., Letter, J. Infect. Dis. 166:209-210, 1992), and Edwardsiella tarda (10, 13, 241), among others that may be potential pathogens. Detailed procedures for the identification of bacteria from fish are described in references 11 and 12.

FUNGAL

INFECTIONS

Dermatophytes Description, Natural Habitat, and Disease in Animals Most fungi that cause systemic diseases in animals are identical to those that cause similar diseases in humans. The identification of dermatophytes from food animal and equine specimens is emphasized here because dermatophytes are the only zoonotic fungi that, under normal circumstances, are transmitted directly from animals to humans. Although not absolutely host specific, zoophilic dermatophyte species do predominate in certain animal hosts, such as Microsporum nanum in swine, Trichophyton equinum in horses, Trichophyton verrucosum in cattle, and Trichophyton gallinae in poultry. The most common geophilic dermatophyte pathogen of animals is Microsporum gypseum. Trichophyton mentagrophytes is the most common dermatophyte pathogen of all animals, and the varieties of this agent that infect animals can readily infect humans. Dermatophyte infections in animals are similar to those in humans, characterized by alopecia, erythema, and crusting of the skin. Lesions may or may not be pruritic and raised. Infections of the nails can occur

30

Bottone

CUMITECH

et al.

FIGURE 5. Human dermatophyte infection due to M. canis. of inflammatory keratosis surrounding an area of healing.

Note

the ring

but are rarer than such infections in humans. T. gallinae infections in birds usually manifest as white patches on the comb, which can result in a white, moldy layer several millimeters thick. The infection is usually self-limiting, unless lesions extend to the feathers, in which case the bird may become seriously ill and die (238). The spores of dermatophytes can survive in the environment for years. Lesions usually occur around the head or neck or in regions where irritation may occur, such as where the harness or straps on horses rub the skin. Adverse environmental conditions and stress also contribute to the occurrence of disease. Thus, infections are more common in fall and winter, when humidity is high, and when animals are in stables rather than on pasture (238).

Mode of Transmission to Humans, Human Infections, and Treatment The most common zoophilic species that infect humans are Microsporum canis, T. verrucosum, and T. mentagrophytes. Zoophilic dermatophyte infections in humans are, for the most part, indistinguishable from anthrophilic or geophilic dermatophyte infections. Most of the lesions are restricted to the skin and may be localized to a single lesion or area. The inflammatory response may be more intense, and a hyperkeratosis may occur more commonly than from the anthrophilic dermatophytes (Fig. 5). Transmission is through direct or indirect contact of spores with a break in the stratum corneum of the skin. Documented zoophilic onychomycosis is rare, if it occurs at all. The animal itself, being the natural host, may not show any lesions or clinical signs. In the absence of lesions, it is not practical to take specimens for culture from the animal. Fine, septate hyphae germinate and spread uniformly in all directions, resulting in a ringlike lesion. The greatest inflammatory response is at the leading edge, with a centralized area of healing.

37

Diagnosis A zoophilic dermatophyte infection should be considered when a patient presents with a history of animal contact followed by an acute, intensely inflammatory, circular skin lesion. A rapid, presumptive diagnosis can sometimes be made by mixing a skin scraping from the edge of the lesion with 10% KOH, covering with a coverslip for a few minutes, and examining the specimen under high-dry (40X) magnification. The fine hyphae of the dermatophytes must be distinguished from the junction between epithelial cells. Although this examination may be negative with chronic, anthrophilic dermatophyte infections, acute, zoophilic lesions may have high concentrations of arthrospores and hyphae (Fig. 6). Confirmation and identification of the etiologic agent require culture, usually on a selective medium containing chloramphenicol and cycloheximide. Dermatophyte test medium (DTM) is a selective and differential medium that will suppress the growth of contaminants. Dermatophytes usually grow as a white-to-tan, powdery or fluffy mold in 1 to 2 weeks and turn DTM red as they are growing. A tease preparation of the growth needs to be done to confirm and identify the. agent, because contaminating fungi may eventually grow on DTM and turn it red. Identification is based primarily on recognition of specific spores and hyphae or nutritional requirements. Macroscopically, many of the dermatophytes look identical. Most dermatophytes of the genus Microsporum can be identified by distinctive macroconidia. Some species of Trichophyton, which rarely produce macroconidia, look very similar microscopically, with abundant microconidia of similar shape and size (163). An exception is T. verrucosum, which has a more tan, glabrous appearance with few aerial hyphae, and grows best at 37°C. Microscopically, the hyphae have large numbers of chlamydoconidia in chains, making them distinctive. For some of the

FIGURE 6. Skin scraping, the arm of a patlent with

suspended T. verrucosum

In 10% potassium hydroxide, Infectjon Magnlflcatlon.

from x40.

CUMITECH

37

zoophilic Trichophyton spp., the species can be determined by nutritional requirements using Trichophyton media (Remel and BD Biosciences). These media consist of seven different agar’ preparations that are nutritionally distinct. Three zoophilic agents that can be identified using these media are T. ueyrucosum, which requires inositol and/or thiamine-HCl for growth; T. equinum, which requires nicotinic acid for growth; and T. gallinae, which requires ammonium nitrate for growth (according to the 10th edition of the Difco Manual [BD Biosciences, Sparks, Md.]). Treatment

Diagnosis

of Zoonotic

Infections

31

7. Ariza, J., T. Pellicer, R. Pallares, A. Fez, and F. Gudio. 1992. Specific antibody profile in human brucellosis. Clin. Infect. Dis. 14:131-140. 8. Arnow, I? M., M. Smaron, and V. Ormiste. 1984. Brucellosis in a group of travelers to Spain. JAMA 251:.50.5-507. 9. Ashdown, L. R., and R. J. Frettingham. 1984. In vitro activity of various cephalosporins against Pseudomonas pseudomallei. J. Infect. Dis. 150:779 -780. 10. Ashford, R. U., I?. D. Sargeant, and G. D. Lum. 1998. Septic arthritis of the knee caused by Edwardsielza tarda after a catfish puncture wound. Med. J. Aust. 168:443-444.

Zoophilic dermatophyte infections are usually selfhealing, but recovery can be hastened by application of topical treatments. Appropriate therapy includes Whitfield’s ointment (salicylic and benzoic acids), chlorphenesin, undecylenate, and tolnaftate. The broad group of azole drugs are very active against these fungi and include miconazole, clotrimazole, ketoconazole, oxiconazole, and others. Other, newer agents include cyclopiroxolamine, terbinafine, and naftifine. Topical therapy twice a day will normally clear the infection in 2 to 4 weeks; the response to terbinafine may be more rapid. Zoophilic dermatophyte infections should not require oral therapy (111). For additional information on zoophilic dermatophyte infections, the reader is referred to Cumitech 28 (105).

11. Austin, B., and D. A. Austin. 1999. Bacterial Fish Pathogens: Disease of Farmed and Wild Fish, 3rd ed. Praxis Publications, London, United Kingdom.

1. Acocella, G., A. Bertrand, J. Beytout, J. B. Durrande, J. A. Garcia Rodriguez, J. Kosmidis, M. Micoud, M. Rey, M. Rodriguez Zapata, J. Roux, et al. 1989. Comparison of three different regimens in the treatment of acute brucellosis: a multicenter multinational study. J. Antimicrob. Chemother. 23:433-439.

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12. Austin, B., and D. A. Austin. 1989. Methods for the Microbiological Examination of Fish and Shellfish Ellis Horwood, New York, N. Y. 13. Banks, A. S. 1992. A puncture wound complicated by infection with Edwardsiella tarda. J. Am. Podiatr. Med. Assoc. 82:.529 -53 1. 14. Barham, W. B., P. Church, J. E. Brown, and S. Paparello. 1993. Misidentification of Brucella species with use of rapid bacterial identification systems. Clin. Infect. Dis. 17:1069-1070. 15. Barnham, M., G. Cole, A. Efstratiou, J. R. Tagg, and S. A. Skjold. 1987. Characterization of Streptococcus zooepidemicus (Lancefield group C) from human and selected animal infections. Epidemiol. Infect. 98:171-182.

17. Barnham, M., J. Kerby, R. S. Chandler, and M. R. Millar. 1989. Group C streptococci in human infection: a study of 308 isolates with clinical correlations. Epidemiol. Infect. 102:379-390. 18. Barnham, M., A. Ljunggren, and M. McIntyre. 1987. Human infection with Streptococcus zooepidemicus (Lancefield group C): three case reports. Epidemiol. Infect. 98:183-190. 19. Barnham, M., T. J. Thorton, and K. Lange. 1983. Nephritis caused by Streptococcus zooepidemicus (Lancefield group C). Lancet i:945-948. 20. Barrow, G. I., and M. Hewitt. with Mycobacterium marinum tank. Br. Med. J. 760:505-506.

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