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Haemophilus ducreyi and chancroid: Practical aspects for the clinical microbiology laboratory
Clinical Microbiology Newsletter Vol. 12, No. 18
September 15, 1990
Haemophilus ducreyi and Chancroid: Practical Aspects for the Clinical Microbiology Laboratory J. E. Clarridge, Ph.D. R. Shawar, Ph.D. B. Simon, M.T. Departmem of Microbiology Veterans Affairs Medical Center and Baylor College of Medicine Houston, TX 77030 Recent reviews of Haemophilus ducreyi have highlighted the increased interest in this organism and in chancroid, the sexually transmitted disease (STD) it causes (1, 2). This article reviews important aspects of the disease and the organism and provides practical guidelines for culturing and identifying H. ducreyi in geographic areas where its incidence is low. Historical Background Chancroid is an STD that is being reported with increasing frequency in North America (3), and is often related to epidemic drug use or prostitution. Approximately 5,000 cases are reported annually to the Centers for Disease Control. Most of these case reports are based on clinical diagnosis rather than confirmed by culture and the actual number is probably much higher (4). In some developing countries, particularly Africa and Southeast Asia, the disease is endemic and occurs as frequently as syphilis. It is difficult to determine whether chancroid existed in ancient times because the infection can be confused with other genital ulcer diseases such as
CMNEEI 12(18)137-144,1990
syphilis and lymphogranuloma venereum. Bassereau f'n-st described the disease in 1852 and subsequently, Auguste Ducrey in 1889. Both workers induced lesions in the forearms of patients who had been inoculated with exudate from their own genital ulcers. They characterized the organism by Gram stain, allowing more reliable data collection on disease occurrence. However, the problem of underreporting of cases continues today because the organism is so difficult to culture. During the 1920s and 1930s, many publications on the pathology, physiology, and epidemiology of H. ducreyi appeared. In the 1940s, the availability of penicillin to treat certain STDs (without the perceived need to determine an etiology) led to a decline in interest in H. ducreyi which continued until the mid- 1970s. The biology of H. ducreyi has been reviewed recently (1). Although the organism is classified in the genus Haemophilus on the basis of its hemin requirement, H. ducreyi has little genetic relationship to this genus or even to other members of the Pasteurellaceae family (5). Because of strain variability and difficulties in growing the organism, even its basic physiology and structure are not well known. There are reports that the organism needs albumin, selenium, and L-glutamine for growth, but no successful defined medium has been developed.
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Although the organism is thought to penetrate normal skin at the site of a minor lesion, the pathogenetic factors allowing adherence and tissue necrosis are unknown. Electron microscope studies have shown a standard gramnegative cell wall and intercellular structures that may be responsible for the ordered cell-to-cell adhesions seen under some culture conditions. Clinical Manifestations The incubation period of chancroid following contact with an infected partner ranges from 1 to 2 wk but most initial lesions appear within 4 to 7 d. The first pathological change, a small inflammatory papule surrounded by a
In T h i s I s s u e
Haemophilus ducreyi and Chancroid . . . . . . . . . . . . . . . . . . . 137 A review including practical diagnostic aspects for the clinical microbiology laboratory Tularemia . . . . . . . . . . . . . . . . . . . . 141 A case of typhoidal tularemia: laboratory diagnosis and review of the disease
Cryptococcus neoformans grant forms on direct microscopic examination . . . . . . . . . . . . . . . . . . A case report
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narrow erythematous zone, occurs around a break in the epithelium. The papule then becomes pustular and erodes to form a painful, nonindurated, ulcer with ragged edges (2, 4, 6). Individual ulcers vary from l to 20 mm in diameter (4). Adjacent lesions may coalesce to form giant ulcerations. Males usually have single ulcers that occur most often on the preputial oririce, internal surface of the prepuce, or the frenulum. In women, ulcers are usually multiple, very irregular, and frequently occur on the fourchette, labia, vestibule, or clitoris. Chancroid lesions are of several varieties. One of these, transient chancroid, is an ulceration that resolves rapidly (4 to 6 d) and is followed by acute, painful, inguinal lymphadenitis. If left untreated, the infection results in formation of a bubo within 10 to 20 d. This form of chancroid is difficult to distinguish from lymphogranuloma venereum. Classic chancroid refers to infection with multiple ulcers, each measuring 0.3 to 2 cm; these may be formed by autoinoculation. Multiple ulcers occur in 40% of infected men and 80% of infected women. Other characteristic lesions have been called follicular, papular, giant, and phagedenic (2, 4, 6). Because the ulcerative lesions in chancroid appear similar to those of syphilis, herpes, and lymphogranuloma venereum, chancroid may be confused with these diseases. Several studies have compared the sensitivity and specificity of clinical diagnosis to culture. In endemic areas, clinical diagnosis has a relatively high positive predictive value whereas in areas of low endemicity, the clinical diagnosis has a lower predictive value (3, 7-9). In part because of strain differences (2, 10, 11), serologic tests are not effective for diagnosing the disease (12). Rather, culture is the gold standard for diagnosing chancroid. H. ducreyi has not been shown to
cause systemic disease, but extragenital lesions on the oropharynx, lip, and breast have been reported (13). There has been some suggestion that chancroid can enhance the spread of human immunodeficiency virus because the lesion provides access to the bloodstream (4).
Culture Techniques and Specimen Gram S t a i n After cleaning the ulcer with sterile gauze and saline, exudate can be collected on a premoistened swab and transported immediately to the laboratory or placed in Amie's or Stuart's transport media. Aspirate from an inguinal lymph node should be retained within the syringe which should be handled carefully to prevent needle sticks and contamination. Specimens should reach the laboratory within 1 h. Some workers have recommended inoculating culture plates at the patient's bedside, but we have not found this procedure necessary. Several media have been recommended for culturing H. ducreyi. Early recommendations described use of heat-inactivated, clotted blood from humans or rabbits (14) followed by identification by Gram stain morphology. More recently recommended media include: (i) a gonococcal agar medium supplemented with hemoglobin, fetal calf serum, and either CVA (Gibco, Madison, Wis.) or IsoVitaleX (Becton-Dickinson Microbiology Systems, Cockeysville, Md.) enrichment; (ii) a Mueller-Hinton agar supplemented with chocolatized horse blood and enrichment (15-19); or (iii) other more exotic media (18, 21-23) such as Bieling or Sheffield media (Table 1). Most workers recommend adding 3 to 5 mg of vancomycin/L to inhibit certain gram-positive flora that can be associated with genital ulcers. Isolation rates of 70 to 90% have been achieved on these media; however, they are not usually readily available in the
average clinical microbiology laboratory where the need for diagnosing chancroid is low. In addition, strains of H. ducreyi vary in their vancomycin susceptibility with some having low vancomycin MICs (24). Overall recovery has been reported to be better on media without antibiotics (25, 26). We have achieved a 90% isolation rate on commercially prepared media (27). On both enriched chocolate gonococcal agar and enriched chocolate Mueller-Hinton agar, H. ducreyi colonies usually grew within 48 h. Growth characteristics vary on media of the same formulation from different manufacturers (28). Plates from Becton Dickinson Microbiology Systems (Cockeysville, Md.) and Regional Media Laboratories (REMEL, Lenexa, Kans.) were most satisfactory. The appropriate incubation conditions are essential for achieving isolation on these agar media. High humidity (>90%) and 3 to 9% CO 2 are necessary and can be attained either in a candle extinction jar (about 3% CO2) humidified by insertion of four water-saturated, crumpled paper towels or a CO 2 incubator set at 8 to 8.5% CO2 and 90 to 95% humidity. When the plates are inoculated, the exudate material on the swab is carefully transferred to the agar surface; the plate is streaked for isolation and immediately placed in the proper environment. A slide for Gram stain examination is made by pressing the swab to a small area of the slide. Do not spread the material around. Of the specimens positive for H. ducreyi by culture, we found that all bubo and most ulcer specimens contained numerous inflammatory cells. Often, only a few gram-negative bacilli were seen, and these were always as single organisms. We did not observe chains or the characteristic "school of fish" arrangement. Some Gram stained smears of exudate from bubos were initially thought to be negative; but
NOTE. No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. No suggested test or procedure should be carried out unless, in the reader's judgment, its risk is justified. Because of rapid advances in the medical sciences, we recommend that the independent verification of diagnoses and drug dosages should be made. Discussions, views and recommendations as to medical procedures, choice of drugs and drug dosages are the responsibility of the authors.
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Clinical Microbiology Newsletter 12:18,1990
TABLE 1. Haemophilusducreyi culture media
about the organism's enzymatic activities have appeared in the literature. Most prominent among these are differences in the oxidase reaction, which depends on the reagent used (1, 2); inability of commercial systems to detect a positive nitrate test, which is demonstrable by conventional tests (29); and differences in the catalase reaction (1, 2). A porphyrin test for X-factor (hemin) requirement and a test for nitrate reductase using an enriched nitrate broth (30, 31) should also be performed. We found that the sodium polyanetholsulfonate (SPS) disk susceptibility test is useful for differentiating H . d u c r e y i from similar organisms because only H . d u c r e y i ,
References indicating media was Media base Mueller Hinton Gonococcal Columbia (Bieling) Proteose #3 Trypticase soy
Common additives
Good
Horse blood, IsoVitaleX, CVA Hemoglobin, fetal calf serum, IsoVitaleX, CVA Horse blood, yeast dialysate Hemin, starch Sheep blood
15,16,19,22
more careful review after the culture became positive revealed a few gramnegative rods.
Colony Morphology and O r g a n i s m Identification H . d u c r e y i colonies are usually not visible after 24 h incubation. After 48 h, they are 0.1 to 0.5 mm in diameter and either smooth, white, and domed (on blood agar) or grey and domed or slightly flattened on chocolate agar. The colonies are cohesive and can be pushed intact across the agar plate. Sometimes the colonies are so adhesive, particularly when grown on blood agar, that subculture is difficult. In this case, the whole colony can be mashed with a sterile wooden stick in a test tube containing a small amount of
Poor
15,16,17,18, 19,24 18,34 21,23 23,25
broth. Gram stain of carefully separated portions of colonies reveals a very structured morphology. Gram-negative organisms of 0.5 I~m by 1.0 to 1.5 i~m are aligned in regular rows. This arrangement is the so-called "finger print," "railroad track," or "school of fish" arrangement. We have not observed bacilli with rounded ends as has been reported by others (1). The morphology of individual organisms resembles that of H a e m o p h i l u s
Gardnerella
vaginalis,
and Capnocyto-
spp. show a zone of inhibition of /> 14 mm (29). Several commercially available kit systems including the RapiD NH (25) and RaplD-ANA (29) (Innovative Diagnostics Systems, Atlanta, Ga.) have been evaluated as aids in identification. We found that the RaplD-ANA was more useful for detecting a positive alkaline phosphatase test, and the number of positive peptidase reactions allowed better differentiation of the isolate. A standard alkaline phosphatase test could be performed instead. Table 2 illustrates important characteristics that distinguish
phaga
parainfluenzae.
Colonies can be identified presumptively as H . d u c r e y i by their adherent colony characteristics, the Gram stain results, and a positive oxidase and negative catalase test. Few biochemical tests are reliable because H . d u c r e y i is asaccharolytic. Conflicting reports
TABLE 2. Biochemical tests differentiating Haemophilus ducreyi from other fastidious gram-negative organisms Organism
Ox
Cat
NO2
PO4
IND
Biochemical Tests a Urea ORN
X
V
Glu
Suc
SPS
Haemophilus
ducreyi
+
-
+
++_
-
-
-
+
-
-
-
+
Haemophilus
influenzae
+
+
+
+
V
V
V
+
+
+
-
-
Haemophilus
parainfluenzae
+
V
+
+
-
V
V
-
+
+
+
-
Haemophilus
segnis
-
V
+
+
.
+
w
w
ND
Haemophilus
aegyptius
+
+
+
+
-
+
+
-
ND
Haemophilus
aphrophilus
V
-
+
+
.
+
+
-
Haemophilus
paraphrophilus
+
-
+
+
.
+
+
+
ND
Haemophilus
parahaemolyticus
+
V
+
+
-
+
V
-
+
+
+
ND
Haemophilus
haemolyticus
+
+
+
+
V
+
-
+
+
+
-
-
Haemophilus
actinomycetemcomitans
V
+
+
+
.
+
-
-
+
-
-
-
+
.
+
+
-
Pasteurella multocida
+
+
+
+
+
-
+
+
-
Kingella denitrificans
+
-
+
.
+
-
ND
+
-
ND
+
+
ND
V
+
Cardiobacterium
hominis
+
-
-
+
.
+
-
-
+
+
Eikenella corrodens
+
Gardnerella vaginalis
.
+ .
.
.
.
.
. .
-
. .
.
.
+
.
. .
.
+ .
-
.
.
.
.
.
. .
.
.
.
.
+
.
.
.
.
+
.
Kingella kingae
-
.
.
.
Kingella indologenes
.
.
. .
.
.
. +
.
° OX, oxidase; CAT, catalase; NO2, nitrate reduction; PO,), alkaline phosphatase; IND, indole production; UREA, urea hydrolysis;ORN, ornithine decarboxylase; X, requires X factor; V, requires V factor;GLU, produces acid from glucose; SUC, produces acid from sucrose; SPS, susceptibleto SPS. OX, CAT, NO2, PO4, X, and V tests performed by standard methods; data from refs. 15, 30, 31, 35. PO(, IN-D,UREA, ORN, GLU, and SUC tests performed by the Rapid NH system; data compiled from Rapid NH charts and refs. 31 and 35. SPS data compiled from ref. 29.
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H. ducreyi from other gram-negative
Summary
bacilli. For reference laboratories and STD clinics, rapid processing of specimens is often not possible because the laboratory is located a great distance from the patient. In this setting, specific H. ducreyi probes, with or without amplification by the polymerase chain reaction (32), may be useful because they can detect even nonviable organisms.
Most clinical microbiology laboratories can isolate H. ducreyi from clinical specimens. The essential factors include collecting the specimen properly in clinically appropriate situations; delivering the specimen to the laboratory quickly; rapidly inoculating it onto appropriate media such as enriched chocolate agar; incubation in a very high humidity with added CO2; and recognizing the distinctive colony morphology after 48"h or more of incubation. Subsequent identification can be made readily by SPS disk susceptibility, the RaplD-ANA system, and other routine tests.
Antimicrobial Susceptibility No standard method is available for performing antimicrobial susceptibility tests for H. ducreyi. The broth dilution method is seldom used because the organism grows poorly in broth. The disk diffusion method is not acceptable because this organism does not grow rapidly nor sufficiently on standard media such as Mueller-Hinton agar with or without supplementation. The agar dilution method is performed most commonly but many different media have been recommended and used. Care must be taken in interpreting resuits among laboratories because small differences in media composition can significantly affect results (27). In vitro, H. ducreyi is usually susceptible to rifampin, erythromycin, ceftriaxone, cefotaxime, trimethoprimsulfamethoxazole (2), and the newer quinolones such as ciprofloxacin (33). These drugs have been used successfully for treatment (3). Tetracycline and the aminoglycosides have also been used for therapy but susceptibility studies have shown that resistance patterns vary widely with geographic source of the strain. Penicillin and ampicillin have been useful in the past, but most strains isolated in the past 20 years produce a 13-1actamase that confers resistance to these drugs. Amoxicillin, in combination with clavulanic acid, a [3-1actamase inhibitor, has been effective for treatment (2). In addition to the penicillins, resistance to trimethoprim, chloramphenicol, sulfonamide, tetracycline, streptomycin, kanamycin, and gentamicin has been reported. The plasmids conferring resistance to many of these antimicrobial agents have been isolated and characterized (2).
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References i. Albritton, W. L. 1989. Biology of Haemophilus ducreyi. Microbiol. Rev. 53:377-389. 2. Morse, S. A. 1989. Chancroid and Haemophilus ducreyi. Clin. Microbiol. Rev. 2:137-157. 3. Schmid, G. P., et al. 1987. Chancroid in the United States, reestablishment of an old disease. J. Am. Med. Assoc. 258:3265-3268. 4. Hand, W. L. 1989. Haemophilus species, p. 1729-1733. In G. L. Mandell, R. G. Douglas, and J. E. Bennett (ed.), Principles and practice of infectious diseases, 3rd ed. Churchill Livingston, New York. 5. Casin, I., et al. 1985. Lack of deoxyribonucleic acid relatedness between Haemophilus ducreyi and other Haemophilus species. Int. J. Syst. Bacteriol. 35:22-25. 6. McCarley, M. E., P. D. Cruz, and R. D. Sontheimer. 1988. Chancroid: clinical variants and other findings from an epidemic in Dallas county, 1986-1987. J. Am. Acad. Dermatol. 19:330-337. 7. Chapel, T. A., et al. 1977. How reliable is the morphological diagnosis of penile ulcerations? Sex. Transm. Dis. 4:150-152. 8. Fast, M. V., et al. 1984. The clinical diagnosis of genital ulcer disease in men in the tropics. Sex. Transm. Dis. 11:72-76. 9. Salzman, R. S., et al. 1984. Chancroidal ulcers that are not chancroid. Arch. Dermatol. 120:636-639. 10. Taylor, D. N., et al. 1985. Antimicrobial susceptibility and characterization of outer membrane proteins of Haemophilus ducreyi isolated in Thailand. J. Clin. Microbiol. 21:442-444. 11. Van Dyck, E. and P. Piot. 1987. En-
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zyme profile of Haemophilus ducreyi strains isolated on different continents. Eur. J. Clin. Microbiol. 4:40-43. 12. Museyi, K., et al. 1988. Use of an enzyme immunoassay to detect serum lgG antibodies to Haemophilus ducreyi. J. Infect. Dis. 157:1039- 1048. 13. Kinghorn, G. R., S. Hafiz, and M. G. McEntegart. 1983. Oropharyngeal Haemophilus ducreyi infection. Lancet ii:656. 14. Borchardt, K. A. and W. A. Hoke. 1970. Simplified laboratory technique for diagnosis of chancroid. Arch. Dermatol. 102:188-192. 15. Albritton, W. L., et al. 1985. Haemophilus ducreyi and Calymmatobacterium granulomatis, p. 869-873. In E. H. Lennette et al. (ed.), Manual of clinical microbiology, 4th ed. American Society for Microbiology, Washington, D.C. 16. Dylewski, J., et al. 1986. Laboratory diagnosis of Haemophilus ducreyi: sensitivity of culture media. Diagn. Microbiol. Infect. Dis. 4:241-245. 17. Hammond, G. W., et al. 1978. Comparison of specimen collection and laboratory techniques for isolation of Haemophilus ducreyi. J. Clin. Microbiol. 7:39-43. 18. Kunimoto, D. Y., et al. 1986. Field testing of modified Bieling media for the isolation of Haemophilus ducreyi in Kenya. Eur. J. Clin. Microbiol. 5:673-675. 19. Lubwama, S. W., et al. 1986. Isolation and identification of Haemophilus ducreyi in a clinical laboratory. J. Med. Microbiol. 22:175-178. 20. Hafiz, D., M. G. McEntegart, and G. R. Kinghom. 1984. Sheffield medium for cultivation of Haemophilus ducreyi. Br. J. Vener. Dis. 60:196198. 21. MacDonald, K., et al. 1989. Comparison of Sheffield media with standard media for the isolation of Haemophilus ducreyi. Sex. Transm. Dis. 16:88-90. 22. Sottnek, F. O., et al. 1980. Isolation and identification of Haemophilus ducreyi in a clinical study. J. Clin. Microbiol. 12:170-174. 23. Sturm, A. W. and H. C. Sanen. 1984. Characteristics of Haemophilus ducreyi in culture. J. Clin. Microbiol. 29:672674. 24. Hannah, P. and J. R. Greenwood. 1982. Isolation and identification of Haemophilus ducreyi. J. Clin. Microbiol. 16:861-864. 25. Oberhofer, T. R. and A. E. Black. 1982. Isolation and cultivation of Haemophilus ducreyi. J. Clin. Microbiol. 15:625-629. 26. Sanson-Le Pors, M. J., et al. 1983. In
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vitro susceptibility of thirty strains of Haemophilus ducreyi to several antibiotics including six cephalosporins. J. Antimicrob. Chemother. 11:271-280. 27. Clarridge, J. E., et al. 1989. The routine isolation and recognition of Haemophilus ducreyi. Progr. Abstr. 29th Intersci. Conf. Antimicrob. Agents Chemother. no. 1055. 28. Clarridge, J. E. 1989. The growth of Haemophilus ducreyi on similar media from different manufacturers and various media from the same source. Diagnosis of sexually transmitted diseases. Texas Department of Health Bureau of Laboratories. Section IX. 29. Shawar, R., J. Sepulveda, and J. E. Clarridge. 1990. Use of the RapiD-
ANA system and sodium polyanetholesulfonate disk susceptibility testing in identifying Haemophilus ducreyi. J. Clin. Microbiol. 28:108-111. 30. Kilian, M. 1985. Haemophilus, p. 387-393. In E. H. Lennette et al. (ed.), Manual of clinical microbiology, 4th ed. American Society for Microbiology, Washington, D.C. 31. Kilian M. and E. L. Biberstein. 1984. Genus II. Haemophilus, p. 558-569. In N. R. Kreig and J. G. Holt. Bergey's manual of systematic bacteriology, vol. 1. Williams and Wilkins Co., Baltimore. 32. Parson, L. M., et al. 1990. Construction of DNA probes for the identification of Haemophilus ducreyi, p.
69-94. Gene probes for bacteria. Academic Press, New York. 33. Bodhidatta, L., et al. 1988. Evaluation of 500-mg and 1,000-mg doses of ciprofloxacin for the treatment of chancroid. Antimicrob. Agents Chemother. 32:723-725. 34. Nobre, G. N. 1982. Identification of Haemophilus ducreyi in the clinical laboratory. J. Med. Microbiol. 15:343-245. 35. Weaver, R. E., et al. 1982. Revised tables from "The identification of unusual pathogenic gram-negative bacteria (Elizabeth O. King)." Division of Bacterial Diseases, Center for Infectious Diseases, Centers for Disease Control, Altanta, Georgia.
The purpose of this report is to describe how this organism was recovered from the blood of a patient with fever of unknown origin using the Isolator microbial detection system (E. I. DuPont de Nemours, Wilmington, Del.). An initial diagnosis of tularemia had not been considered in this patient and was made by subsequent recovery of the organism from culture. A four-fold rise in the patient's antibody titers further confirmed the diagnosis.
showed a total white blood cell count of 15 x 106/p,l with 80% segmented neutrophils, 9% band forms, 5% lymphocytes, 5% monocytes, and 1% basophils. The hemoglobin was 13.1 gm/dl, hematocrit, 38.2%, urinalysis and Gram stain of urine were negative, and routine chemistry tests were normal. Blood cultures were drawn and cefoxitin was started. The patient was thought to have diverticulitis with abscess formation. However, an extensive evaluation of her abdomen revealed only three adenomatous polyps and an abdominal aortic aneurysm. The patient remained febrile and on the fourth hospital day, her chest X ray showed a left lower lobe infiltrate. CT scan of her chest also showed bilateral pleural effusions and infiltrates, but followup bronchoscopy was negative. Cefoxitin was discontinued and erythromycin therapy begun. An echocardiogram that was obtained to exclude bacterial endocarditis was normal except for moderate mitral valve regurgitation. The patient continued to receive an extensive evaluation for fever of unknown origin which included a bone marrow biopsy, MRI of the head, and a complete serologic evaluation for rickettsial, mycoplasmal, and viral infection. The bone marrow biopsy revealed small areas of necrosis which, in retrospect, contained minute gram-negative bacteria (Fig. 1). A tiny, gram-
Case Reports Tularemia: A Present Day Problem Denise Rule, B.A., M(ASCP) Laboratory Associate Robin Miller-Catchpole, M.D. Associate Director, Microbiology Harold G. Wedell, M.D.
Department of Medicine Eileen L. Randall, Ph.D. Director, Microbiology Evanston Hospital Evanston, IL 60201 Francisella tularensis is a fastidious, small, gram-negative coccobacillus. It was first described in 1911 by McCoy who wrote about a plaguelike disease of rodents occurring in Tulare County, California (1). McCoy and Chapin discovered the causative organism of the disease and named it Bacterium tularense. In 1925, Francis wrote extensively about the history of tularemia in humans, elaborating on the epidemiology, pathology, and laboratory diagnosis of the disease (2). Previously, the disease had been given many different names depending on the conditions encountered by various workers; for example, plaguelike disease of rodents, deerfly fever, Bacillus tularensis infections of the eye, rabbit fever, and glandular type of tick fever. Francis named the disease tularemia. In 1959, he was awarded the Nobel Prize for his work and was honored by having the name of the bacterium changed to Francisella tularensis.
Clinical Microbiology Newsletter 12:18,1990
Case History A 75-year-old white female presented to Glenbrook emergency room with a two-day history of fever and right lower quadrant abdominal pain. She denied any nausea, vomiting, diarrhea, or dysuria. She had a past history of hypertension and an appendectomy. At the time of admission she was receiving propranolol hydrochloride and lovastatin. On physical examination, the patient was a well-developed female in no apparent distress. Her vital signs were: blood pressure, 110/60; pulse rate, 108; respiratory rate, 24; and temperature, 103.5°F. Pertinent physical findings included no rash or skin ulceration, no adenopathy, clear lungs, decreased bowel sounds, no abdominal distension and no masses. She had mild discomfort on palpation of the right lower quadrant. Admission laboratory results included a complete blood count that
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Haemophilus ducreyi and Chancroid: Practical Aspects for the Clinical Microbiology Laboratory J. E. Clarridge, Ph.D. R. Shawar, Ph.D. B. Simon, M.T. Departmem of Microbiology Veterans Affairs Medical Center and Baylor College of Medicine Houston, TX 77030 Recent reviews of Haemophilus ducreyi have highlighted the increased interest in this organism and in chancroid, the sexually transmitted disease (STD) it causes (1, 2). This article reviews important aspects of the disease and the organism and provides practical guidelines for culturing and identifying H. ducreyi in geographic areas where its incidence is low. Historical Background Chancroid is an STD that is being reported with increasing frequency in North America (3), and is often related to epidemic drug use or prostitution. Approximately 5,000 cases are reported annually to the Centers for Disease Control. Most of these case reports are based on clinical diagnosis rather than confirmed by culture and the actual number is probably much higher (4). In some developing countries, particularly Africa and Southeast Asia, the disease is endemic and occurs as frequently as syphilis. It is difficult to determine whether chancroid existed in ancient times because the infection can be confused with other genital ulcer diseases such as
CMNEEI 12(18)137-144,1990
syphilis and lymphogranuloma venereum. Bassereau f'n-st described the disease in 1852 and subsequently, Auguste Ducrey in 1889. Both workers induced lesions in the forearms of patients who had been inoculated with exudate from their own genital ulcers. They characterized the organism by Gram stain, allowing more reliable data collection on disease occurrence. However, the problem of underreporting of cases continues today because the organism is so difficult to culture. During the 1920s and 1930s, many publications on the pathology, physiology, and epidemiology of H. ducreyi appeared. In the 1940s, the availability of penicillin to treat certain STDs (without the perceived need to determine an etiology) led to a decline in interest in H. ducreyi which continued until the mid- 1970s. The biology of H. ducreyi has been reviewed recently (1). Although the organism is classified in the genus Haemophilus on the basis of its hemin requirement, H. ducreyi has little genetic relationship to this genus or even to other members of the Pasteurellaceae family (5). Because of strain variability and difficulties in growing the organism, even its basic physiology and structure are not well known. There are reports that the organism needs albumin, selenium, and L-glutamine for growth, but no successful defined medium has been developed.
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Although the organism is thought to penetrate normal skin at the site of a minor lesion, the pathogenetic factors allowing adherence and tissue necrosis are unknown. Electron microscope studies have shown a standard gramnegative cell wall and intercellular structures that may be responsible for the ordered cell-to-cell adhesions seen under some culture conditions. Clinical Manifestations The incubation period of chancroid following contact with an infected partner ranges from 1 to 2 wk but most initial lesions appear within 4 to 7 d. The first pathological change, a small inflammatory papule surrounded by a
In T h i s I s s u e
Haemophilus ducreyi and Chancroid . . . . . . . . . . . . . . . . . . . 137 A review including practical diagnostic aspects for the clinical microbiology laboratory Tularemia . . . . . . . . . . . . . . . . . . . . 141 A case of typhoidal tularemia: laboratory diagnosis and review of the disease
Cryptococcus neoformans grant forms on direct microscopic examination . . . . . . . . . . . . . . . . . . A case report
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narrow erythematous zone, occurs around a break in the epithelium. The papule then becomes pustular and erodes to form a painful, nonindurated, ulcer with ragged edges (2, 4, 6). Individual ulcers vary from l to 20 mm in diameter (4). Adjacent lesions may coalesce to form giant ulcerations. Males usually have single ulcers that occur most often on the preputial oririce, internal surface of the prepuce, or the frenulum. In women, ulcers are usually multiple, very irregular, and frequently occur on the fourchette, labia, vestibule, or clitoris. Chancroid lesions are of several varieties. One of these, transient chancroid, is an ulceration that resolves rapidly (4 to 6 d) and is followed by acute, painful, inguinal lymphadenitis. If left untreated, the infection results in formation of a bubo within 10 to 20 d. This form of chancroid is difficult to distinguish from lymphogranuloma venereum. Classic chancroid refers to infection with multiple ulcers, each measuring 0.3 to 2 cm; these may be formed by autoinoculation. Multiple ulcers occur in 40% of infected men and 80% of infected women. Other characteristic lesions have been called follicular, papular, giant, and phagedenic (2, 4, 6). Because the ulcerative lesions in chancroid appear similar to those of syphilis, herpes, and lymphogranuloma venereum, chancroid may be confused with these diseases. Several studies have compared the sensitivity and specificity of clinical diagnosis to culture. In endemic areas, clinical diagnosis has a relatively high positive predictive value whereas in areas of low endemicity, the clinical diagnosis has a lower predictive value (3, 7-9). In part because of strain differences (2, 10, 11), serologic tests are not effective for diagnosing the disease (12). Rather, culture is the gold standard for diagnosing chancroid. H. ducreyi has not been shown to
cause systemic disease, but extragenital lesions on the oropharynx, lip, and breast have been reported (13). There has been some suggestion that chancroid can enhance the spread of human immunodeficiency virus because the lesion provides access to the bloodstream (4).
Culture Techniques and Specimen Gram S t a i n After cleaning the ulcer with sterile gauze and saline, exudate can be collected on a premoistened swab and transported immediately to the laboratory or placed in Amie's or Stuart's transport media. Aspirate from an inguinal lymph node should be retained within the syringe which should be handled carefully to prevent needle sticks and contamination. Specimens should reach the laboratory within 1 h. Some workers have recommended inoculating culture plates at the patient's bedside, but we have not found this procedure necessary. Several media have been recommended for culturing H. ducreyi. Early recommendations described use of heat-inactivated, clotted blood from humans or rabbits (14) followed by identification by Gram stain morphology. More recently recommended media include: (i) a gonococcal agar medium supplemented with hemoglobin, fetal calf serum, and either CVA (Gibco, Madison, Wis.) or IsoVitaleX (Becton-Dickinson Microbiology Systems, Cockeysville, Md.) enrichment; (ii) a Mueller-Hinton agar supplemented with chocolatized horse blood and enrichment (15-19); or (iii) other more exotic media (18, 21-23) such as Bieling or Sheffield media (Table 1). Most workers recommend adding 3 to 5 mg of vancomycin/L to inhibit certain gram-positive flora that can be associated with genital ulcers. Isolation rates of 70 to 90% have been achieved on these media; however, they are not usually readily available in the
average clinical microbiology laboratory where the need for diagnosing chancroid is low. In addition, strains of H. ducreyi vary in their vancomycin susceptibility with some having low vancomycin MICs (24). Overall recovery has been reported to be better on media without antibiotics (25, 26). We have achieved a 90% isolation rate on commercially prepared media (27). On both enriched chocolate gonococcal agar and enriched chocolate Mueller-Hinton agar, H. ducreyi colonies usually grew within 48 h. Growth characteristics vary on media of the same formulation from different manufacturers (28). Plates from Becton Dickinson Microbiology Systems (Cockeysville, Md.) and Regional Media Laboratories (REMEL, Lenexa, Kans.) were most satisfactory. The appropriate incubation conditions are essential for achieving isolation on these agar media. High humidity (>90%) and 3 to 9% CO 2 are necessary and can be attained either in a candle extinction jar (about 3% CO2) humidified by insertion of four water-saturated, crumpled paper towels or a CO 2 incubator set at 8 to 8.5% CO2 and 90 to 95% humidity. When the plates are inoculated, the exudate material on the swab is carefully transferred to the agar surface; the plate is streaked for isolation and immediately placed in the proper environment. A slide for Gram stain examination is made by pressing the swab to a small area of the slide. Do not spread the material around. Of the specimens positive for H. ducreyi by culture, we found that all bubo and most ulcer specimens contained numerous inflammatory cells. Often, only a few gram-negative bacilli were seen, and these were always as single organisms. We did not observe chains or the characteristic "school of fish" arrangement. Some Gram stained smears of exudate from bubos were initially thought to be negative; but
NOTE. No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. No suggested test or procedure should be carried out unless, in the reader's judgment, its risk is justified. Because of rapid advances in the medical sciences, we recommend that the independent verification of diagnoses and drug dosages should be made. Discussions, views and recommendations as to medical procedures, choice of drugs and drug dosages are the responsibility of the authors.
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Clinical Microbiology Newsletter 12:18,1990
TABLE 1. Haemophilusducreyi culture media
about the organism's enzymatic activities have appeared in the literature. Most prominent among these are differences in the oxidase reaction, which depends on the reagent used (1, 2); inability of commercial systems to detect a positive nitrate test, which is demonstrable by conventional tests (29); and differences in the catalase reaction (1, 2). A porphyrin test for X-factor (hemin) requirement and a test for nitrate reductase using an enriched nitrate broth (30, 31) should also be performed. We found that the sodium polyanetholsulfonate (SPS) disk susceptibility test is useful for differentiating H . d u c r e y i from similar organisms because only H . d u c r e y i ,
References indicating media was Media base Mueller Hinton Gonococcal Columbia (Bieling) Proteose #3 Trypticase soy
Common additives
Good
Horse blood, IsoVitaleX, CVA Hemoglobin, fetal calf serum, IsoVitaleX, CVA Horse blood, yeast dialysate Hemin, starch Sheep blood
15,16,19,22
more careful review after the culture became positive revealed a few gramnegative rods.
Colony Morphology and O r g a n i s m Identification H . d u c r e y i colonies are usually not visible after 24 h incubation. After 48 h, they are 0.1 to 0.5 mm in diameter and either smooth, white, and domed (on blood agar) or grey and domed or slightly flattened on chocolate agar. The colonies are cohesive and can be pushed intact across the agar plate. Sometimes the colonies are so adhesive, particularly when grown on blood agar, that subculture is difficult. In this case, the whole colony can be mashed with a sterile wooden stick in a test tube containing a small amount of
Poor
15,16,17,18, 19,24 18,34 21,23 23,25
broth. Gram stain of carefully separated portions of colonies reveals a very structured morphology. Gram-negative organisms of 0.5 I~m by 1.0 to 1.5 i~m are aligned in regular rows. This arrangement is the so-called "finger print," "railroad track," or "school of fish" arrangement. We have not observed bacilli with rounded ends as has been reported by others (1). The morphology of individual organisms resembles that of H a e m o p h i l u s
Gardnerella
vaginalis,
and Capnocyto-
spp. show a zone of inhibition of /> 14 mm (29). Several commercially available kit systems including the RapiD NH (25) and RaplD-ANA (29) (Innovative Diagnostics Systems, Atlanta, Ga.) have been evaluated as aids in identification. We found that the RaplD-ANA was more useful for detecting a positive alkaline phosphatase test, and the number of positive peptidase reactions allowed better differentiation of the isolate. A standard alkaline phosphatase test could be performed instead. Table 2 illustrates important characteristics that distinguish
phaga
parainfluenzae.
Colonies can be identified presumptively as H . d u c r e y i by their adherent colony characteristics, the Gram stain results, and a positive oxidase and negative catalase test. Few biochemical tests are reliable because H . d u c r e y i is asaccharolytic. Conflicting reports
TABLE 2. Biochemical tests differentiating Haemophilus ducreyi from other fastidious gram-negative organisms Organism
Ox
Cat
NO2
PO4
IND
Biochemical Tests a Urea ORN
X
V
Glu
Suc
SPS
Haemophilus
ducreyi
+
-
+
++_
-
-
-
+
-
-
-
+
Haemophilus
influenzae
+
+
+
+
V
V
V
+
+
+
-
-
Haemophilus
parainfluenzae
+
V
+
+
-
V
V
-
+
+
+
-
Haemophilus
segnis
-
V
+
+
.
+
w
w
ND
Haemophilus
aegyptius
+
+
+
+
-
+
+
-
ND
Haemophilus
aphrophilus
V
-
+
+
.
+
+
-
Haemophilus
paraphrophilus
+
-
+
+
.
+
+
+
ND
Haemophilus
parahaemolyticus
+
V
+
+
-
+
V
-
+
+
+
ND
Haemophilus
haemolyticus
+
+
+
+
V
+
-
+
+
+
-
-
Haemophilus
actinomycetemcomitans
V
+
+
+
.
+
-
-
+
-
-
-
+
.
+
+
-
Pasteurella multocida
+
+
+
+
+
-
+
+
-
Kingella denitrificans
+
-
+
.
+
-
ND
+
-
ND
+
+
ND
V
+
Cardiobacterium
hominis
+
-
-
+
.
+
-
-
+
+
Eikenella corrodens
+
Gardnerella vaginalis
.
+ .
.
.
.
.
. .
-
. .
.
.
+
.
. .
.
+ .
-
.
.
.
.
.
. .
.
.
.
.
+
.
.
.
.
+
.
Kingella kingae
-
.
.
.
Kingella indologenes
.
.
. .
.
.
. +
.
° OX, oxidase; CAT, catalase; NO2, nitrate reduction; PO,), alkaline phosphatase; IND, indole production; UREA, urea hydrolysis;ORN, ornithine decarboxylase; X, requires X factor; V, requires V factor;GLU, produces acid from glucose; SUC, produces acid from sucrose; SPS, susceptibleto SPS. OX, CAT, NO2, PO4, X, and V tests performed by standard methods; data from refs. 15, 30, 31, 35. PO(, IN-D,UREA, ORN, GLU, and SUC tests performed by the Rapid NH system; data compiled from Rapid NH charts and refs. 31 and 35. SPS data compiled from ref. 29.
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H. ducreyi from other gram-negative
Summary
bacilli. For reference laboratories and STD clinics, rapid processing of specimens is often not possible because the laboratory is located a great distance from the patient. In this setting, specific H. ducreyi probes, with or without amplification by the polymerase chain reaction (32), may be useful because they can detect even nonviable organisms.
Most clinical microbiology laboratories can isolate H. ducreyi from clinical specimens. The essential factors include collecting the specimen properly in clinically appropriate situations; delivering the specimen to the laboratory quickly; rapidly inoculating it onto appropriate media such as enriched chocolate agar; incubation in a very high humidity with added CO2; and recognizing the distinctive colony morphology after 48"h or more of incubation. Subsequent identification can be made readily by SPS disk susceptibility, the RaplD-ANA system, and other routine tests.
Antimicrobial Susceptibility No standard method is available for performing antimicrobial susceptibility tests for H. ducreyi. The broth dilution method is seldom used because the organism grows poorly in broth. The disk diffusion method is not acceptable because this organism does not grow rapidly nor sufficiently on standard media such as Mueller-Hinton agar with or without supplementation. The agar dilution method is performed most commonly but many different media have been recommended and used. Care must be taken in interpreting resuits among laboratories because small differences in media composition can significantly affect results (27). In vitro, H. ducreyi is usually susceptible to rifampin, erythromycin, ceftriaxone, cefotaxime, trimethoprimsulfamethoxazole (2), and the newer quinolones such as ciprofloxacin (33). These drugs have been used successfully for treatment (3). Tetracycline and the aminoglycosides have also been used for therapy but susceptibility studies have shown that resistance patterns vary widely with geographic source of the strain. Penicillin and ampicillin have been useful in the past, but most strains isolated in the past 20 years produce a 13-1actamase that confers resistance to these drugs. Amoxicillin, in combination with clavulanic acid, a [3-1actamase inhibitor, has been effective for treatment (2). In addition to the penicillins, resistance to trimethoprim, chloramphenicol, sulfonamide, tetracycline, streptomycin, kanamycin, and gentamicin has been reported. The plasmids conferring resistance to many of these antimicrobial agents have been isolated and characterized (2).
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References i. Albritton, W. L. 1989. Biology of Haemophilus ducreyi. Microbiol. Rev. 53:377-389. 2. Morse, S. A. 1989. Chancroid and Haemophilus ducreyi. Clin. Microbiol. Rev. 2:137-157. 3. Schmid, G. P., et al. 1987. Chancroid in the United States, reestablishment of an old disease. J. Am. Med. Assoc. 258:3265-3268. 4. Hand, W. L. 1989. Haemophilus species, p. 1729-1733. In G. L. Mandell, R. G. Douglas, and J. E. Bennett (ed.), Principles and practice of infectious diseases, 3rd ed. Churchill Livingston, New York. 5. Casin, I., et al. 1985. Lack of deoxyribonucleic acid relatedness between Haemophilus ducreyi and other Haemophilus species. Int. J. Syst. Bacteriol. 35:22-25. 6. McCarley, M. E., P. D. Cruz, and R. D. Sontheimer. 1988. Chancroid: clinical variants and other findings from an epidemic in Dallas county, 1986-1987. J. Am. Acad. Dermatol. 19:330-337. 7. Chapel, T. A., et al. 1977. How reliable is the morphological diagnosis of penile ulcerations? Sex. Transm. Dis. 4:150-152. 8. Fast, M. V., et al. 1984. The clinical diagnosis of genital ulcer disease in men in the tropics. Sex. Transm. Dis. 11:72-76. 9. Salzman, R. S., et al. 1984. Chancroidal ulcers that are not chancroid. Arch. Dermatol. 120:636-639. 10. Taylor, D. N., et al. 1985. Antimicrobial susceptibility and characterization of outer membrane proteins of Haemophilus ducreyi isolated in Thailand. J. Clin. Microbiol. 21:442-444. 11. Van Dyck, E. and P. Piot. 1987. En-
© 1990ElsevierSciencePublishingCo., Inc.
zyme profile of Haemophilus ducreyi strains isolated on different continents. Eur. J. Clin. Microbiol. 4:40-43. 12. Museyi, K., et al. 1988. Use of an enzyme immunoassay to detect serum lgG antibodies to Haemophilus ducreyi. J. Infect. Dis. 157:1039- 1048. 13. Kinghorn, G. R., S. Hafiz, and M. G. McEntegart. 1983. Oropharyngeal Haemophilus ducreyi infection. Lancet ii:656. 14. Borchardt, K. A. and W. A. Hoke. 1970. Simplified laboratory technique for diagnosis of chancroid. Arch. Dermatol. 102:188-192. 15. Albritton, W. L., et al. 1985. Haemophilus ducreyi and Calymmatobacterium granulomatis, p. 869-873. In E. H. Lennette et al. (ed.), Manual of clinical microbiology, 4th ed. American Society for Microbiology, Washington, D.C. 16. Dylewski, J., et al. 1986. Laboratory diagnosis of Haemophilus ducreyi: sensitivity of culture media. Diagn. Microbiol. Infect. Dis. 4:241-245. 17. Hammond, G. W., et al. 1978. Comparison of specimen collection and laboratory techniques for isolation of Haemophilus ducreyi. J. Clin. Microbiol. 7:39-43. 18. Kunimoto, D. Y., et al. 1986. Field testing of modified Bieling media for the isolation of Haemophilus ducreyi in Kenya. Eur. J. Clin. Microbiol. 5:673-675. 19. Lubwama, S. W., et al. 1986. Isolation and identification of Haemophilus ducreyi in a clinical laboratory. J. Med. Microbiol. 22:175-178. 20. Hafiz, D., M. G. McEntegart, and G. R. Kinghom. 1984. Sheffield medium for cultivation of Haemophilus ducreyi. Br. J. Vener. Dis. 60:196198. 21. MacDonald, K., et al. 1989. Comparison of Sheffield media with standard media for the isolation of Haemophilus ducreyi. Sex. Transm. Dis. 16:88-90. 22. Sottnek, F. O., et al. 1980. Isolation and identification of Haemophilus ducreyi in a clinical study. J. Clin. Microbiol. 12:170-174. 23. Sturm, A. W. and H. C. Sanen. 1984. Characteristics of Haemophilus ducreyi in culture. J. Clin. Microbiol. 29:672674. 24. Hannah, P. and J. R. Greenwood. 1982. Isolation and identification of Haemophilus ducreyi. J. Clin. Microbiol. 16:861-864. 25. Oberhofer, T. R. and A. E. Black. 1982. Isolation and cultivation of Haemophilus ducreyi. J. Clin. Microbiol. 15:625-629. 26. Sanson-Le Pors, M. J., et al. 1983. In
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vitro susceptibility of thirty strains of Haemophilus ducreyi to several antibiotics including six cephalosporins. J. Antimicrob. Chemother. 11:271-280. 27. Clarridge, J. E., et al. 1989. The routine isolation and recognition of Haemophilus ducreyi. Progr. Abstr. 29th Intersci. Conf. Antimicrob. Agents Chemother. no. 1055. 28. Clarridge, J. E. 1989. The growth of Haemophilus ducreyi on similar media from different manufacturers and various media from the same source. Diagnosis of sexually transmitted diseases. Texas Department of Health Bureau of Laboratories. Section IX. 29. Shawar, R., J. Sepulveda, and J. E. Clarridge. 1990. Use of the RapiD-
ANA system and sodium polyanetholesulfonate disk susceptibility testing in identifying Haemophilus ducreyi. J. Clin. Microbiol. 28:108-111. 30. Kilian, M. 1985. Haemophilus, p. 387-393. In E. H. Lennette et al. (ed.), Manual of clinical microbiology, 4th ed. American Society for Microbiology, Washington, D.C. 31. Kilian M. and E. L. Biberstein. 1984. Genus II. Haemophilus, p. 558-569. In N. R. Kreig and J. G. Holt. Bergey's manual of systematic bacteriology, vol. 1. Williams and Wilkins Co., Baltimore. 32. Parson, L. M., et al. 1990. Construction of DNA probes for the identification of Haemophilus ducreyi, p.
69-94. Gene probes for bacteria. Academic Press, New York. 33. Bodhidatta, L., et al. 1988. Evaluation of 500-mg and 1,000-mg doses of ciprofloxacin for the treatment of chancroid. Antimicrob. Agents Chemother. 32:723-725. 34. Nobre, G. N. 1982. Identification of Haemophilus ducreyi in the clinical laboratory. J. Med. Microbiol. 15:343-245. 35. Weaver, R. E., et al. 1982. Revised tables from "The identification of unusual pathogenic gram-negative bacteria (Elizabeth O. King)." Division of Bacterial Diseases, Center for Infectious Diseases, Centers for Disease Control, Altanta, Georgia.
The purpose of this report is to describe how this organism was recovered from the blood of a patient with fever of unknown origin using the Isolator microbial detection system (E. I. DuPont de Nemours, Wilmington, Del.). An initial diagnosis of tularemia had not been considered in this patient and was made by subsequent recovery of the organism from culture. A four-fold rise in the patient's antibody titers further confirmed the diagnosis.
showed a total white blood cell count of 15 x 106/p,l with 80% segmented neutrophils, 9% band forms, 5% lymphocytes, 5% monocytes, and 1% basophils. The hemoglobin was 13.1 gm/dl, hematocrit, 38.2%, urinalysis and Gram stain of urine were negative, and routine chemistry tests were normal. Blood cultures were drawn and cefoxitin was started. The patient was thought to have diverticulitis with abscess formation. However, an extensive evaluation of her abdomen revealed only three adenomatous polyps and an abdominal aortic aneurysm. The patient remained febrile and on the fourth hospital day, her chest X ray showed a left lower lobe infiltrate. CT scan of her chest also showed bilateral pleural effusions and infiltrates, but followup bronchoscopy was negative. Cefoxitin was discontinued and erythromycin therapy begun. An echocardiogram that was obtained to exclude bacterial endocarditis was normal except for moderate mitral valve regurgitation. The patient continued to receive an extensive evaluation for fever of unknown origin which included a bone marrow biopsy, MRI of the head, and a complete serologic evaluation for rickettsial, mycoplasmal, and viral infection. The bone marrow biopsy revealed small areas of necrosis which, in retrospect, contained minute gram-negative bacteria (Fig. 1). A tiny, gram-
Case Reports Tularemia: A Present Day Problem Denise Rule, B.A., M(ASCP) Laboratory Associate Robin Miller-Catchpole, M.D. Associate Director, Microbiology Harold G. Wedell, M.D.
Department of Medicine Eileen L. Randall, Ph.D. Director, Microbiology Evanston Hospital Evanston, IL 60201 Francisella tularensis is a fastidious, small, gram-negative coccobacillus. It was first described in 1911 by McCoy who wrote about a plaguelike disease of rodents occurring in Tulare County, California (1). McCoy and Chapin discovered the causative organism of the disease and named it Bacterium tularense. In 1925, Francis wrote extensively about the history of tularemia in humans, elaborating on the epidemiology, pathology, and laboratory diagnosis of the disease (2). Previously, the disease had been given many different names depending on the conditions encountered by various workers; for example, plaguelike disease of rodents, deerfly fever, Bacillus tularensis infections of the eye, rabbit fever, and glandular type of tick fever. Francis named the disease tularemia. In 1959, he was awarded the Nobel Prize for his work and was honored by having the name of the bacterium changed to Francisella tularensis.
Clinical Microbiology Newsletter 12:18,1990
Case History A 75-year-old white female presented to Glenbrook emergency room with a two-day history of fever and right lower quadrant abdominal pain. She denied any nausea, vomiting, diarrhea, or dysuria. She had a past history of hypertension and an appendectomy. At the time of admission she was receiving propranolol hydrochloride and lovastatin. On physical examination, the patient was a well-developed female in no apparent distress. Her vital signs were: blood pressure, 110/60; pulse rate, 108; respiratory rate, 24; and temperature, 103.5°F. Pertinent physical findings included no rash or skin ulceration, no adenopathy, clear lungs, decreased bowel sounds, no abdominal distension and no masses. She had mild discomfort on palpation of the right lower quadrant. Admission laboratory results included a complete blood count that
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