Strongyloidiasis

Clinical Microbiology Newsletter Vol. 13, No. 5

March 1, 1991

Strongyloidiasis James W. Smith, M.D.

Professor of Pathology Director of Clinical Microbiology Indiana University Medical Center Indianapolis, IN 46202-5250

Case History A 25yr-old male truck driver from Kentucky received a cadaveric renal transplant. Because of allograft rejection he was given high doses of adrenal corticosteroids. Several weeks later he developed diffuse bilateral interstitial pneumonia with severe respiratory distress and Pneumocystis pneumonia was suspected. Impression smears of a lung biopsy were negative for infectious agents but a sputum cytology showed nematode larvae (Fig. 1). Subsequent fecal exam showed numerous rhabditiform larvae with short buccal cavities and prominent genital primordia diagnostic for Strongyloides stercoralis (Fig. 2). The patient was treated with thiabendazole (25 mg/kg twice daily for 5 d) and the pneumonia resolved (2). Several months later, interstitial pneumonia recurred and stools again were found to contain numerous rhabditiform larvae of Strongyloides stercoralis. The patient was treated again with thiabendazole and the chest infiltrates resolved. Follow-up stool examination at 4 mo showed no Strongyloides in a single specimen using the formalin ether concentration technique. He continued to be followed in the renal transplantation clinic and was doing well without pulmonary or gastrointestinal problems. On a clinic visit 10 yr after the second episode of pneumonia,

CMNEFJ 13(5)33-40,1991

the blood count showed an eosinophilia of 35%. Because of his past history, a fecal examination was requested which again showed numerous rhabditiform larvae of Strongyloides. He was treated again with thiabendazole and the eosinophilia disappeared. It is planned to check his stools at regular intervals for Strongyloides larvae using the Baermann technique (19) to determine if the infection recurs. This case raises a number of questions about strongyloidiasis including: (i). How is the infection acquired? (ii). How is infection able to persist? (iii). Why may Strongyloides cause serious, even fatal infections in immunocompromised hosts? (iv). What is the epidemiology of the infection? (v). What diagnostic tests can be used? This article will provide a brief overview of strongyloidiasis in normal and immunocompromised hosts and attempt to answer the above questions. For more detailed information recent reviews should be consulted (3, 4).

most never found in feces. Male worms are not found in the mucosa and reproduction may occur by parthenogenesis. In the external environment, the rhabditiform larvae may mature to the infective filariform stage in several days, depending on environmental conditions. If these filariform larvae come in contact with the skin of a human, they penetrate the skin, then reach small blood vessels or lymphatics and migrate to the lungs where they exit the circulatory system and undergo maturation in the pulmonary parenchyma. They then enter the trachea-bronchial tree where they are brought up with the pulmonary secretions and swallowed, thus reaching the intestinal tract where they invade the mucosa of the upper small intestine and begin to produce eggs. The period from penetration of skin to presence of larvae in feces is about 3 wk.

In This Issue

Life Cycle Strongyloides stercoralis is a small nematode parasite of humans and sometimes other animals. It has a complicated life cycle (Fig. 3) which may have both parasitic and free-living phases. Parasitic female worms live in the small intestinal mucosa producing embryonated eggs that hatch within the mucosa to release rhabditiform (feeding) larvae. These larvae migrate to the intestinal lumen and eventually are passed in the feces. Eggs are al-

ElSWkI

Strongyloidiasis . . . . . . . . . . . . . . . 33 A review of the clinical and laboratory aspects of a classic parasite and its newly appreciated importance as a cause of infections in immunocompromised hosts

Acute Suppurative Otitis Caused by Comamow acidovorans . . . . . 38 A case report

Letters to the Editors . . . . . . . . . . 39

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Clinical Infection There are two unique aspects of Strongyloides infection. First. there

Figure I. Strongyloides larva seen during cytologic examination of sputum (Papanicolaou stain X 500).

-

--._ .HABDITIFORM

STAGE

FILARIFORM

STAGE

can be a free-living cycle in which the rhabditiform larvae mature into freeliving adult male and female worms which mate; then, the females lay eggs that hatch to release larvae that can again develop into adult, free-living stages or can develop into infective filarifonn larvae. Second. autoinfections with Strongyloides occur often so that the infection may persist for many years. There are three possible ways in which autoinfection may occur. First, rhabditiform larvae in fecal material left in the perianal area may mature to the filariform stage and penetrate the skin. Second, rhabditiform larvae may mature to the infective stage during transit through the intestinal tract and penetrate through the mucosa into the circulation. Third, larval stages hatching from eggs in the mucosa of the small

-1

kOlP: )

45

9p Microns

Genital primoridum

Hookworm

StrongYlo~des

Hookworm

J

Figure 2. Identification of hookworm and Strongyloides larvae (I). Rhabditiform larvae of Strongyloides have a short buccal cavity and prominent genital primordium whereas those of hookworm have a long buccal cavity and inconspicuous genital primordium. Filariform larvae of Strongyloides have a long esophagus (VII to % length of body) and notched tip of the tail whereas those of hookworm have a short esophagus (% length of body) and a pointed tail.

NOTE. No responsibihty is assumed by the Publisher for any injury and/or damage lo persons or property as a matter of products liatnhty, negligence or otherwise, or from any use or operation of any methods, products, insttuctions 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 mdependent veritication 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 Clinical Microbiology Newsletfer (ISSN Ol%-4399) IS issued twce monthly in one indexed volume by Elsevier Scwx~ce Publishing Co., 655 Avenue of the Americas. New York. NY 10010. Subscription prices per year: $1 IO.00 in&ding postage and handling in the United States. Canada, and Mexico. Add $48.00 for postage in the rest of the world. Second-&s\ postage paid at New York, NY, and at additional mailing offices. Postmaster: send address changes to CIinirol Microhiolo~y Newsletrer. Elsev~er Science Publishing Co., Inc 655 Avenue of the Americas, New York. NY 10010.

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Figure 3. Life cycle of Strongyloides stercoralis (see text) lfrom ref. 5).

intestine may mature, invade the vessels and complete the life cycle without reaching the intestinal lumen. In the normal host there is usually a very low grade autoinfection which is generally asymptomatic or is associated with skin eruptions, eosinophilia, or mild nonspecific gastrointestinal complaints such as epigastric discomfort (6). The host’s immune system appears to play an important role in controlling the infection (7) by allowing only a few of the parasites to complete the cycle and survive to adulthood. Low-grade infection may persist for over 40 years. However, patients who are immunocompromised may develop hyperinfection syndrome because this control is removed. In this syndrome

Clinical Microbiology Newsletter 13:5,1991

there may be great numbers of larvae and adult female worms in the intestine, lungs, and sometimes other organs. It appears that many larvae mature to a stage that allows them to invade the intestinal mucosa before they are passed and are able to survive migration to become adult females in tissue. Hyperinfection is seen in immunocompromised patients such as transplant recipients, patients receiving steroids for conditions such as chronic obstructive pulmonary disease or rheumatoid arthritis, patients being treated with antineoplastic therapy, or occasionally patients with AIDS (4, 8). For reasons that are not completely clear, hyperinfection is uncommon in patients with AIDS, even in highly endemic

0 1991 Elsevier Science Publishing Co., Inc.

areas (9). Patients with hyperinfection often present with gastrointestinal or pulmonary symptoms resulting from the severe infection and host reaction to it. Eosinophilia is sometimes noted but often is not seen due to the immune suppression. The patient described above is an example of one with hyperinfection syndrome. In some patients with hyperinfection, adults and larvae may be found in sites in addition to intestine and lungs including brain, liver, and kidneys, and thus have disseminated strongyloidiasis. Hyperinfection is a life-threatening condition which, if not diagnosed promptly, may be fatal. Patients with hyperinfection may have recurrent episodes of gram-nega-

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tive sepsis caused by bacteria that are brought into the circulatory system by the larval stages when they penetrate the colonic mucosa. In some patients, peritonitis or gram-negative meningitis ( 10, 11) may develop. Strongyloidiasis with hyperinfection should always be considered in an immunocompromised patient who develops sepsis, meningitis, or peritonitis due to gram-negative organisms without an obvious cause (3).

Epidemiology Strongyloidiasis is found worldwide, particularly in underdeveloped countries where a large percentage of the population may be infected (9). There appear to be differences in strains of Strongyloides from different geographic areas. For example, the strains found in IndoChina are infective for dogs whereas those found in India are not. Dogs may thus serve as a reservoir host in some areas, although most infections are probably of human origin (3). Infection is usually acquired by environmental exposure rather than by direct person to person spread; for example. wives of chronically infected persons are usually not infected ( 12). In normal individuals, there are usually few or mild symptoms from strongyloidiasis such as bloating and epigastric discomfort; however, malnourished individuals may develop serious diarrhea. During World War Il. diarrhea was a major problem in the prisoners of war mentioned below In addition, people who have had gastric surgery, who take antacids, or who have blind intestinal loops may have more severe infections. The prevalence and epidemiology of strongyloidiasis have recently been reviewed by Genta (9). People in the United States most likely to have strongyloidiasis come from developing countries such as Africa, Asia, and Latin America or Southern, Central, or Eastern Europe. The infection is particularly common in native Americans born in the southeastern Appalachian region, who work in custodial institutions, or who have traveled or served in the armed forces in the areas mentioned above. Infection may be seen in Vietnam veterans but is not common

(13). In the Appalachian region, the prevalence in children may be 2 to 4%. Chronic infection with Strongyloides has been clearly described in former prisoners of the Japanese in World War II from England ( 14), Australia ( 15), and the United States ( 16) who worked on the Burma-Thailand railroad (the railroad depicted in the movie Bridge on the River Kwai). Many of these people still have evidence of Strongy loides infection after over 45 years. A common manifestation in these former prisoners and others who have acquired strongyloidiasis in Southeast Asia is a skin condition known as larva currens (17, 18). These people have pro gressing linear skin eruptions that are felt to represent filariform larvae migrating through the skin with subsequent reaction to larval products. U&aria may be seen in some infected patients worldwide.

Laboratory Diagnosis Diagnosis is traditionally establish& by wet mount examination of concentrated fecal specimens to detect the rhabditiform larvae that must be differentiated from those of‘ hookworm (Fig. 2). Shedding of parasites may vary from day to day and the number of larvae may be small. Examination of a single specimen allows diagnosis in only 30 to 60% of patients. Even patients with hyperinfection syndrome may not always have larval stages detectable in feces. .A total of three specimens should be examined and even then some intections, especially chronic infections. will be missed. The most common concentration procedure used in the i !nited States is the formalin-ethyl acctatc centrifugal sedimentation. .A more sensitive method. Baermann c.oncentration ( 19). is recommended for screening for

GAUZE SCREEN

RUBBER TUBING - i-’( CLAMP ----+ LI

Figure 4. Baermann concentration apparatus (adapted from ref. 20). Wuter level should be such that the gauze and bottom qf the,fecal specimen are \b’et.

chronic strongyloidiasis or for evaluating success of therapy; even with this method, multiple specimens may need to be examined to detect infection in some patients. Baermann concentration is performed by placing a fecal specimen on a wire mesh covered with several layers of gauze in a wide funnel and filling the funnel with water such that the water just touches the specimen (Fig. 4). Larval stages migrate into the water. After several hours or overnight, a specimen is collected by releasing the rubber clamp at the tip of the funnel, removing a small amount of fluid and examining the fluid microscopically for larvae. Filter paper cultures for larval maturation can also be performed but are not widely used (19, 20). Examination of duodenal aspirate or of material obtained by the string test may be helpful in diagnosing some patients. The string test (21) is performed by having the patient swallow a portion of string and taping the proximal end to the cheek. If the patient takes small sips of water over a period of several hours, the distal string reaches the upper small intestine. The string is then retrieved and the bile-stained material from the distal portion of the string is examined microscopically to detect the wiggling larvae. Larvae may be detected by sputum examination as in the patient described here; however, it must be noted that we have also seen larval nematodes in cytology slides as a result of free-living nematodes contaminating the water in an automatic stainer. Sputum examination may be helpful in monitoring the success of therapy in patients with hyperinfection. Occasional infections have been diagnosed by alert technologists in the bacteriology laboratory who noted “tracks” of bacteria inoculated across agar plates by wandering larvae (22). These cultures have usually been of fecal or sputum specimens. Serologic tests for antibodies may be helpful in diagnosing strongyloidiasis in both epidemiologic and patient care situiitions (23, 24, 25). These tests genetJly have both sensitivity and specificity of over 90%, but unfortunately, they are not widely available. Thiabendazole is the treatment of

Clinical Microbiology Newsletter 13:5,1991

choice in acute severe infections and response is rather prompt; however, there may be failure to cure in up to 30% of patients (3, 26, 27) and infections in immunocompromised patients may be particularly difficult to eradicate. Such patients should be monitored to determine cure or to detect recurrence of strongyloidiasis as the case presented illustrates. Infection has been transmitted through transplanted organs (28) but most infections in immunocompromised patients probably arise from activation of chronic subclinical infection. In the case presented above, the origin is not certain as the patient was from an endemic area and had received a kidney transplant. Before being immunosuppressed, people with a history of strongyloidiasis, who are from an endemic area, or who have had significant potential exposure in such an area should be evaluated for strongyloidiasis, preferably with the Baermann method or serologic testing. In summary, strongyloidiasis should be considered as a cause of intestinal complaints in healthy individuals or as a cause of interstitial pneumonia or gastrointestinal symptoms, including epigastic discomfort and diarrhea in immunocompromised patients. Strongyloidiasis should also be considered when immunocompromised patients present with recurrent episodes of gram-negative sepsis or with meningitis caused by gram-negative bacteria. Diagnosis is established by demonstrating larvae in feces, sputa, or intestinal aspirates.

8.

11.

12. 13.

14.

15.

References

Brooke, M. M. and D. M. Melvin. 1984. Morphology of diagnostic stages of intestinal parasites of humans, 2nd ed. HHS Pub. No. (CDC) 848116. U.S. Dept. of Health and Human Services, Centers for Disease Control, Atlanta, Ga. Leapman, S. B. et al. 1980. Strongyloides stercoralis in chronic renal failure. Safe therapy with tbiabendazole. South. Med. J. 73:1400-1402. Genta, R. M. and P. D. WaJzer. 1989. Strongyloidiasis, p. 463-525. In P. D. Walzer and R. M. Genta (ed.), Parasitic infections in the compromised host. Marcel Dekker, Inc., New York. lgra-Siegman, Y. et al. 1981. Syn-

0 1991 Ekvier Science Publishing Co.,

6.

Inc.

16.

17.

18.

drome of hyperinfection with Strongyloides stercoralis. Rev. Infect. Dis. 3:397-407. Melvin, M. D., M. M. Brooke, and E. H. Sadun. 1980. Common intestinal helminths of man, life cycle charts (HHS Pub. No. [CDC] 80-8286). U.S. Dept. of Health and Human Services, Centers for Disease Control, Atlanta, Ga. Milder, J. E. et al. 1981. Clinical features of Strongyloides stercoralis infection in an endemic area of the United States. Gastroenterology 80:14811488. Genta, R. M. 1986. Strangyloides stercoralis: imrnunobiological consideration on an unusual worm. Parasitol. Today 2:241-246. Maayon, S. et al. 1987. Strongyloides stercoralis hyperinfection in a patient with the acquired immune deficiency syndrome. Am. J. Med. 83:945-948. Genta, R. M. 1989. Global prevalence of strongyloidiasis: critical review with epidemiologic insights into the prevention of disseminated disease. Rev. lnfeet. Dis. 11:755-767. Vishwanath, S., R. A. Baker, and B. J. Mansheim. 1982. Strongyloides infection and meningitis in an immunocompromised host. Am. J. Trop. Med. Hyg. 31:857-858. Smallman, L. A. et al. 1986. Strongyloides stercoralis hyperinfestation syndrome and E. coli meningitis: report of two cases. J. Clin. Pathol. 39:366-370. Grove, D. I. 1982. Strongyloidiasis: is it transmitted from husband to wife? Br. J. Vener. Dis. 58:271-272. Genta, R. M. et al. 1987. Strongyloidiasis in United States veterans of the Vietnam and other wars. J. Am. Med. Assoc. 258~49-52. Gill, G. V. and D. R. Bell. 1979. Strongyloides stercoralis infection in former Far East prisoners of war. Br. Med. J. 21572-574. Grove, D. I. 1980. Strongyloides in Allied ex-prisoners of war in SouthEast Asia. Br. Med. J. 1:598-601. Pelletier, L. L. 1984. Chronic strongyloidiasis in World War II Far East exprisoners of war. Am. J. Trop. Med. Hyg. 33:55-61. Smith, J. D., D. K. Goette, and R. B. Odom. 1976. Larva currens. Cutaneous strongyloidiasis. Arch. Dermatol. 112:1161-1163. VonKuster, L. C. and R. M. Genta. 1988. Cutaneous manifestations of strongyloidiasis. Arch. Dermatol. 124:1826-1830. Ash, L. R. and T. C. Orihel. 1987. Parasites: A Guide to Laboratory Procedures and Identification. American

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Society of Clinical Pathologists Press, Chicago. 20 Melvin, D. M. and M. M. Brooke. 1982. Laboratory procedures of the diagnosis of intestinal parasites, 3rd. ed. HHS Publication No. (CDC) 82-8282 U.S. Dept. of Health and Human Services, Centers for Disease Control, Atlanta, Ga. 21 Beal, C. B. et al. 1970. A new technique for sampling duodenal contents. Am. J. Trop. Med. Hyg. 19:349-352. 33Panosian, K. J., P. Ma, and S. G. Edberg. 1986. Elucidation of Srrongyloides stercoralis by bacterial colony II.

displacement. J. Clin. Microbial.

24:86-88. 23. Genta, R. M. 1986. Strongyloidiasis, p. 183-199. In K. Walls and P. Schantz (ed.), Immunodiagnosis of parasitic diseases. Academic Press, New York.

26. Pelletier, L. L. Jr. et al. 1988. Diagnosis and evaluation of treatment of chronic strongyloidiasis in ex-prisoner5 of war. J. Infect. Dis. 157:573- 576.

21. Shelhamer, J. H., F. A. Neva. and D. R. Finn. 1982. Persistent strongyloidiasis in an immunodeficient patient. Am. J. Trop. Med. Hyg. 31:746-751.

24. Genta, R. M. 1988. Predictive value of an enzyme-linked immunosorbent assay (ELISA) for the serodiagnosis of strongyloidiasis. Am. J. Clin. Pathol. 89:391-394. 25. Neva, F. A. 1986. Biology and immunology of human strongyloidiasis. J. Infect. Dis. 153:397-406.

28. Hoy, W. E. et al. 198 I. Transmlssion of strongyloidiasis by kidney transplant’? Disseminated strongyloidiasis in both recipients of kidney allografts from a single cadaver donor. J. Am. Med. Assoc. 246:1937-- 1939.

which gave the bionumber 14102000010, and positive reactions for the hydrolysis of acetamide, maltose, and mannitol. The definitive identification was obtained with the biochemical and physiological tests recommended by Tamaoka et al. (1) Antibiotic susceptibility studies, performed with the GNS-BH and GNS-BI cards (Vitek Systems Co.), showed the microorganism to be susceptible with the respective MICs (pg/mL), cefotaxime (<4), ceftazidime (<8), ceftriaxone (<8), aztreonam (<8), imipenem (<4), chloramphenicol (4), tobramycin (2), amikacin (16), ciprofloxacin (0.5), trimethoprim-sulfamethoxazole (lo), and resistant with the respective MICs to ampicillin (>32), cefazolin (>32), cefuroxime (>32), carbenicillin (128). and gentamicin (>16). The species included previously in the acidovorans group of the genus Pseudomonas were taxonomically reclassified by Tamaoka et al. (1) on the basis of their phenotypic characteristics, DNA-DNA relatedness, and chemotaxonomic characteristics. They were finally included in the genus Comamonas as Comamonas acidovorans and Comamonas testosteroni. These two species are nonfermenting, motile (polar tuft of flagella), oxidase-positive, gram-negative rods, that are found in soil, water, and in animals such as the rabbit, turtle, frog, and cobra. C. acidovorans is distinguished from C. testosteroni by its ability to produce acid from fructose

and mannitol and to hydrolyze acetamide. C. acidovorans has been isolated as a contaminant of hospital equipment (intravenous tubing, urinary catheter, ultrasonic nebulizer), as well as from some clinical sources, such as urine, sputum, feces, oropharynx, CSF, and wounds (2). These isolates have, however, almost always been considered nonpathogenic and insignificant. Cases of nosocomial infection presenting as bacteremia associated with the use of an indwelling pressure-monitoring device (3) and an infection of a comeal ulcer (4) have also been described. Recently, Horowitz et al. (5) have reported the first case of endocarditis caused by this microorganism in an intravenous drug abuser. Human infections by C. ucidovorans appear to be infrequent. The origin of C. ucidovorans in this case was probably from the river water in which the patient had bathed a few days before the first signs of infection appeared, particularly because fresh waters of rivers and lakes are the natural habitat for this microorganism (6). Since there are few reports of infection by C. acidovorans in the literature, little information is available regarding its antibiotic susceptibility. A study of nonfermenting gram-negative rods, performed by Gilardi (7), shows this microorganism is resistant to the majority of penicillins and aminoglycosides, and susceptible to third generation cephalosporins, trimethoprim-sulfamethoxazole. nalidixic acid,

Case Report

Acute Suppurative Otitis Caused by Comamonas

acidovorans

Jordi Reina, M.D., Ph.D. Isabel Llompart, Phar.D. , Pedro Alomar, M.D., Ph.D.

Servicio de Microbiologia, Hospital Son Dureta 07014-Palma de Mallorca, Spain

A 7-yr-old male, in previously good health, was evaluated in an emergency department for pain, inflammation, edema, and purulent suppuration of the right ear. The infection had started 3 d after he bathed in a river near his rural home and had worsened progressively until his arrival at the hospital. A sample of the ear drainage was obtained for microbiological culture and parenteral ceftazidime therapy was initiated. The Gram stain of the sample showed many polymorphonuclear cells, epithelial cells, and gram-negative rods. The sample was plated onto blood, chocolate agar, MacConkey agars, and incubated at 37°C for 18 to 24 h. At the same time the sample was plated onto Sabouraud dextrose agar and incubated at 30°C for 18 to 24 h. All plates grew an abundance of a highly motile, oxidase- and catalasepositive gram-negative rod. The initial identification was performed using the AutoMicrobic System (Vitek System Co., St. Louis, MO), using a GNI card

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