CHAPTER 120: LABORATORY DIAGNOSIS OF PARASITIC INFECTIONS

The cornerstone for the diagnosis of parasitic infections is a thorough history of the patient’s illness. Epidemiologic aspects of the illness are especially important because the risks of acquiring many parasites are closely related to occupation, recreation, or travel to areas of high endemicity. Without a basic knowledge of the epidemiology and life cycles of the major parasites, it is difficult to approach the diagnosis of parasitic infections systematically. Accordingly, the medical classification of important human parasites in this chapter emphasizes their geographic distribution, their transmission, and the anatomic location and stages of their life cycle in humans. The text and tables are intended to serve as a guide to the correct diagnostic procedures for the major parasitic infections; in addition, the reader is referred to other chapters that contain more comprehensive information about each infection (Chaps. 121, 122, 123, 124, 126, 127, 128, 129, 131, 132, 133, 134, 135). Tables 120-1, 120-2, and 120-3 summarize the geographic distributions, the anatomic locations, and the methods employed for the diagnosis of flatworm, roundworm, and protozoal infections, respectively.

In addition to selecting the correct diagnostic procedures, physicians must counsel their patients to ensure that specimens are collected properly and arrive at the laboratory promptly. For example, the diagnosis of bancroftian filariasis is unlikely to be confirmed by the laboratory unless blood is drawn near midnight, when the nocturnal microfilariae are active. Laboratory personnel and surgical pathologists should be notified in advance when a parasitic infection is suspected. Continuing interaction with the laboratory staff and the surgical pathologists increases the likelihood that parasites in body fluids or biopsy specimens will be examined carefully by the most capable individuals.

Most helminths and protozoa exit the body in the fecal stream. Many laboratories now use a stool collection kit with instructions for patients to transfer portions of the sample directly into bacterial carrier medium and fixative, which increases yield. If kits are not available, the patient should be instructed to collect feces in a clean waxed or cardboard container and to record the time of collection on the container. Refrigeration will preserve trophozoites for a few hours and protozoal cysts and helminthic ova for several days. Contamination with water (which could contain free-living protozoa) or with urine (which can damage trophozoites) should be avoided. Fecal samples should be collected before ingestion of barium or other contrast agents for radiologic procedures and before treatment with antidiarrheal agents and antacids; these substances change the consistency of the feces and interfere with microscopic detection of parasites. Because of the cyclic shedding of most parasites in the feces, a minimum of three samples collected on alternate days should be examined. Examination of a single sample can be up to 50% less sensitive.

Analysis of fecal samples entails both macroscopic and microscopic examination. Watery or loose stools are more likely to contain protozoal trophozoites, but protozoal cysts and all stages of helminths may be found in formed feces. If adult worms or tapeworm segments are observed, they should be transported promptly to the laboratory or washed and preserved in fixative for later examination. The only tapeworm with motile segments is Taenia saginata, the beef tapeworm, which patients sometimes bring to the physician. Because the ova of T. saginata are morphologically indistinguishable from those of Taenia solium (the cause of cysticercosis), motility is an important distinguishing characteristic.

Microscopic examination of feces is not complete until direct wet mounts have been evaluated and concentration techniques as well as permanent stains have been applied. Before accepting a report of negativity for ova and parasites as final, the physician should insist that the laboratory undertake each of these procedures. Some intestinal parasites are more readily detected in material other than feces. For example, examination of duodenal contents is sometimes necessary to detect Giardia lamblia, Cryptosporidium, and Strongyloides larvae. Use of the “cellophane tape” technique to detect pinworm ova on the perianal skin sometimes also reveals ova of T. saginata deposited perianally when the motile segments disintegrate (Table 120-4).

Two routine solutions are used to make wet mounts for identification of the various life stages of helminths and protozoa: physiologic saline for trophozoites, cysts, ova, and larvae and dilute iodine solution for protozoal cysts and ova. Iodine solution must never be used to examine specimens for trophozoites because it kills the parasites and thus eliminates their characteristic motility.

The two most common concentration procedures for detecting small numbers of cysts and ova are formalin-ether sedimentation and zinc sulfate flotation. The formalin-ether technique is preferable because all parasites sediment but not all float. Slides permanently stained for trophozoites should be prepared before concentration. Additional slides stained for cysts and ova may be made from the concentrate.

In many instances, especially in the differentiation of Entamoeba histolytica from other amebas, identification of parasites from wet mounts or concentrates must be considered tentative. Permanently stained smears allow study of the cellular detail necessary for definitive identification. The iron-hematoxylin stain is excellent for critical work, but trichrome staining, which can be completed in 1 h, is a satisfactory alternative that also reveals parasites in specimens preserved in polyvinyl alcohol fixative. Modified acid-fast staining and fluorescent auramine microscopy are useful adjuncts for detection and identification of several intestinal protozoa, including Cryptosporidium and Cyclospora, while direct fluorescent antibody testing is in common use for Cryptosporidium and Giardia. Microsporidia, which cause chronic diarrhea in HIV-infected patients, may be missed unless a special modified trichrome stain or fluorescent screening with calcofluor white is requested (Table 120-3).

Invasion of tissue by protozoa and helminths renders the choice of diagnostic techniques more difficult. For example, physicians must understand that aspiration of an amebic liver abscess rarely reveals E. histolytica because the trophozoites are located primarily in the abscess wall. They must remember that the urine sediment offers the best opportunity to detect Schistosoma haematobium in the young Ethiopian immigrant or the American traveler who returns from Africa with hematuria. Tables 120-1, 120-2, and 120-3, which offer a quick guide to the geographic distribution and anatomic locations of the major tissue parasites, should help the physician to select the appropriate body fluid or biopsy site for microscopic examination. Tables 120-5 and 120-6 provide additional information about the identification of parasites in samples from specific anatomic locations. The laboratory procedures for detection of parasites in other body fluids are similar to those used in the examination of feces. The physician should insist on wet mounts, concentration techniques, and permanent stains for all body fluids. The trichrome or iron-hematoxylin stain is satisfactory for all tissue helminths in body fluids other than blood, but microfilarial worms and blood protozoa are more easily visualized with Giemsa or Wright’s staining.

The parasites most commonly detected in Giemsa-stained blood smears are the plasmodia, microfilariae, and African trypanosomes (Table 120-5). Most patients with Chagas disease present in the chronic phase, when Trypanosoma cruzi is no longer microscopically detectable in blood smears. Wet mounts are sometimes more sensitive than stained smears for the detection of microfilariae and African trypanosomes because these active parasites cause noticeable movement of the erythrocytes in the microscopic field. Filtration of blood through a polycarbonate filter (pore size, 3–5 μm) facilitates the detection of microfilariae. The intracellular amastigote forms of Leishmania species and T. cruzi can sometimes be visualized in stained smears of peripheral blood, but aspirates of the bone marrow, liver, and spleen are the best sources for microscopic detection and culture of Leishmania in kala-azar and of T. cruzi in chronic Chagas disease. The diagnosis of malaria and the critical distinction among the various Plasmodium species are made by microscopic examination of stained thick and thin blood films (Chap. 123). Because the lab infrastructure and technical expertise may not be available in many rural areas with high levels of malaria, rapid detection tests (RDTs) are increasingly being used to fill this gap. These are immunochromatographic capture assays with monoclonal antibodies to species-specific antigens (histidine-rich protein 2 [PfHRP2] or aldolase of Plasmodium falciparum) or conserved Plasmodium antigens (lactate dehydrogenase). The World Health Organization sponsored a major testing program evaluating the different RDTs. Lower performance rates have been reported in a variety of field sites, especially in areas where deletions of the pfhrp2 gene have been detected. Subpatent infection and identification of Plasmodium species can also be confirmed by polymerase chain reaction (PCR), but for this purpose PCR is primarily a research tool. P. knowlesi, a simian parasite, has been identified as the cause of an increasing number of infections in Malaysian Borneo and other areas of Southeast Asia; PCR or another molecular method is required to differentiate P. knowlesi from P. malariae.

Although most tissue parasites stain with the traditional hematoxylin and eosin, appropriate special stains should also be applied to surgical biopsy specimens. The surgical pathologist who is accustomed to applying silver stains for Pneumocystis to induced sputum and transbronchial biopsies may need to be reminded to examine wet mounts and iron-hematoxylin–stained preparations of pulmonary specimens for helminthic ova and E. histolytica. The clinician should also be able to advise the surgeon and pathologist about optimal techniques for the identification of parasites in specimens obtained by certain specialized minor procedures (Table 120-6). For example, the excision of skin snips for the diagnosis of onchocerciasis, the collection of rectal snips for the diagnosis of schistosomiasis, and punch biopsy of skin lesions for the identification and culture of cutaneous and mucocutaneous species of Leishmania are simple procedures, but the diagnosis can be missed if the specimens are improperly obtained or processed.

Eosinophilia (>500/μL) commonly accompanies infections with most of the tissue helminths; absolute numbers of eosinophils may be high in trichinellosis and the migratory phases of filariasis (Table 120-7). Intestinal helminths provoke eosinophilia only during pulmonary migration of the larval stages. Eosinophilia is not a manifestation of protozoal infections. Parasitic causes of eosinophilia in cerebrospinal fluid include nematodes (e.g., Angiostrongylus, Gnathostoma, Toxocara, and Baylisascaris species) as well as flatworms (e.g., T. solium and Schistosoma species).

Like the hypochromic microcytic anemia of heavy hookworm infections, other nonspecific laboratory abnormalities may suggest parasitic infection in patients with appropriate geographic and/or environmental exposures. Biochemical evidence of cirrhosis or an abnormal urine sediment in an African immigrant certainly raises the possibility of schistosomiasis, and anemia and thrombocytopenia in a febrile traveler or immigrant are among the hallmarks of malaria. CT and MRI also contribute to the diagnosis of infections with many tissue parasites and have become invaluable adjuncts in the diagnosis of neurocysticercosis and cerebral toxoplasmosis.

Useful antibody assays for many of the important tissue parasites are available; most of those listed in Table 120-8 can be obtained from the Centers for Disease Control and Prevention (CDC) in Atlanta. The results of serologic tests not listed in the tables should be interpreted with caution.

The value of antibody assays is limited by several factors. For example, the preparation of thick and thin blood smears remains the procedure of choice for the diagnosis of malaria in individual patients because diagnostic titers to plasmodia develop slowly and do not differentiate species—a critical step in patient management. Filarial antigens cross-react with those from other nematodes; as in assays for antibody to most parasites, the presence of antibody in the filarial assay fails to distinguish between past and current infection. Despite these specific limitations, the restricted geographic distribution of many tropical parasites increases the diagnostic usefulness of both the presence and the absence of antibody in travelers from industrialized countries. In contrast, a large proportion of the world’s population has been exposed to Toxoplasma gondii, and the presence of IgG antibody to T. gondii does not constitute proof of active disease.

Fewer antibody assays are available for the diagnosis of infection with intestinal parasites. E. histolytica is the major exception. Sensitive, specific serologic tests are invaluable in the diagnosis of amebiasis. Commercial kits for the detection of antigen by enzyme-linked immunosorbent assay or of whole organisms by fluorescent antibody assay are now available for several protozoan parasites (Table 120-8).

DNA hybridization with probes that are repeated many times in the genome of a specific parasite and amplification of a specific DNA fragment by PCR have now been established as useful techniques for the diagnosis of several protozoan infections (Table 120-8). Although PCR is very sensitive, it is an adjunct to conventional techniques for parasite detection and should be requested only when microscopic and immunodiagnostic procedures fail to establish the probable diagnosis. For example, only multiple negative blood smears or the failure to identify the infecting species justifies PCR for the diagnosis or proper management of malaria. In addition to PCR of anticoagulated blood, the CDC (www.cdc.gov/dpdx/) and several commercial laboratories now perform PCRs for detection of certain specific parasites in stool samples, biopsy specimens, and bronchoalveolar lavage fluid (Table 120-8). Although PCRs are now used primarily for the detection of protozoans, active research efforts are likely to establish their feasibility for the detection of several helminths.