Medical Parasitology

Thick and Thin Blood Films

Examination of permanently stained blood films is required to identify most blood parasites ( ). Thin films are prepared in the same manner as for hematologic differential evaluation; blood is spread over the slide in a thin layer, yielding intact, nonoverlapping cellular elements ( Fig. 65.1 ). Integrity of the blood cell membranes is important for determining the intracellular or extracellular nature of the infection and the size of the infected erythrocyte. In the thick film, blood is concentrated in a small area that is many cell layers deep. During staining, erythrocytes are dehemoglobinized and only leukocyte nuclei, platelets, and parasites (if present) are visible. The thick film is preferred for parasite detection because it contains 15 to 30 times more blood per microscopic field than does the thin film, thus increasing the chances of detecting light parasitemia and decreasing the time needed for reliable examination ( ). Although examination of thick films increases the likelihood of detecting an infection, species identification is usually performed by examination of thin films because morphology is often more definitive, especially for malarial parasites. For routine detection of malaria and babesiosis, both thick and thin films should be prepared ( ).

Preparation of Slides

Blood for examination may be obtained by fingerstick, earlobe puncture, or venipuncture. Fingerstick blood should flow freely to prevent dilution with tissue fluid, and it should not be contaminated with the alcohol disinfectant, which should be allowed to dry first. If obtained by venipuncture, the first drop of blood (anticoagulant-free) from the needle is used to prepare the films at the bedside ( ). Use of anticoagulants is discouraged when malaria is suspected because they may cause distortion of the parasites and interfere with staining. In practice, however, blood usually is submitted to the laboratory in an anticoagulant, which may be the only practical method to ensure that high-quality smears can be prepared. Ethylenediaminetetraacetic acid (EDTA)-anticoagulated blood is preferred in such cases; the specimen should be transported to the laboratory within the hour to prevent deterioration of organism morphology ( ). Anticoagulants do not interfere with the staining of microfilariae.

Both thin and thick films should be prepared on clean, grease-free slides. Thick films are prepared by placing 1 to 2 small drops of blood onto a slide and spreading them into an area the size of a dime (1.5 cm) with the edge of a second slide ( ; ). The blood film is then allowed to dry flat at room temperature. Drying time may be decreased by placing the slides in a laminar flow hood. A proper thick film should be thin enough that newspaper print may be faintly readable through it ( ). If it is too thick, the film may peel from the slide. Adherence can be greatly improved by gently pushing (grinding) down with the corner of the second slide while spreading the droplet, creating minute scratches on the carrier slide that provide additional surface area for the blood film (see Fig. 65.1 ) ( ).

This method does not affect the microscopic morphology and allows the film to be stained as soon as it is dry (within 30–60 minutes). Heating the slides to decrease drying time is discouraged since excess heat may fix erythrocytes and prevent them from lysing when placed in the stain reagents ( ).

Staining

Blood begins to lose its affinity for stain in about 3 days, and the erythrocytes in older thick films do not lyse well. Therefore, slides stored for archival purposes should first be stained. Best staining results are achieved when using Giemsa stain, because host cell and parasite chromatin stains vividly while the hemoglobin in the erythrocytes is only a pale purple-red. When used at a neutral pH (7.0–7.2), this method also allows for optimal visualization of erythrocyte inclusions (e.g., Schüffner stippling, Maurer clefts) that occur with infection by certain malarial parasites ( ). Wright and Wright-Giemsa stains may also be used for thin films (as is common in the hematology lab), but they stain parasites less well than Giemsa and do not allow for visualization of erythrocyte inclusions due to the lower pH (<6.8). Because these stains incorporate alcohol as their fixative, thick films must be lysed in water before staining ( ; ).

The Giemsa staining procedure requires somewhat more attention to preparation of reagents and staining protocol than does the Wright staining procedures, which are often automated. Generally, fresh Giemsa stain must be made each day of use by diluting stock solution into phosphate-buffered water ( ). Each new lot of stock Giemsa stain must be checked to determine optimal staining time and dilution because some variation is seen from lot to lot ( ).

Examination of Smears

Both thick and thin smears are examined in their entirety under the low-power (10×) objective to detect microfilariae, which rarely occur in large numbers. In particular, the feathered edge of thin smears should be examined, as microfilariae are often carried there during preparation of the smear ( ). Examination using a 50× oil immersion objective may subsequently be used to screen blood films for protozoa, although thorough examination using the 100× oil immersion objectives still is necessary to detect the smallest parasites, such as Plasmodium and Babesia spp. The optimal location for examining the thin film is the region of the feathered edge where there is minimal overlap of cells and the erythrocytes maintain their central pallor ( ). A common mistake is to examine regions of the thin film where the blood is too thick or too thin and the parasite morphology is distorted. An experienced microscopist should examine at least 100 oil immersion fields on both the thick and thin blood film and up to at least 300 fields for immunologically naïve patients who might present with more severe symptoms at a lower parasitemia ( ).

Blood Concentration Techniques

A variety of special techniques have been described for the concentration of blood parasites—specifically, leishmaniasis, trypanosomes, and microfilariae—details of which may be found elsewhere ( ; ; ; , ).

Preparation of buffy coat smears, which most clinical laboratories can perform with existing resources, is helpful in the detection of L. donovani , trypanosomes, and microfilariae ( ). Following centrifugation of an anticoagulated blood sample, the layer of cells between plasma and packed erythrocytes is drawn off and used to prepare blood films for staining or for preparation of a wet mount to detect motile organisms ( ).

For detection of microfilariae, the Knott concentration technique or membrane filtration is helpful, particularly when the density of microfilariae in peripheral blood is very low. With the Knott concentration technique, anticoagulated blood is lysed with 2% formalin and centrifuged to concentrate the microfilariae in the sediment, which then may be examined as a wet preparation or stained with Giemsa or hematoxylin stain. In the membrane filtration procedure, blood is lysed and passed through a 5-μm membrane filter, which is subsequently stained with hematoxylin to reveal any microfilariae ( ; ; ; ).

Finally, use of the fluorochrome acridine orange in a microhematocrit centrifuge format (QBC Malaria test; Drucker Diagnostics, Port Matilda, PA) allows detection of Plasmodium spp. and other blood parasites. While it appears to be more sensitive than traditional thick and thin smears, it is not widely used due to the need for a fluorescent microscope ( ). Fluorescent attachments to traditional light microscopes may make this test more widely accessible. In positive cases of malaria or babesiosis, traditional blood films must be examined for Plasmodium spp. identification (if relevant) and calculation of parasite burden.

Specimen Collection, Handling, and Preservation

Recovery and subsequent identification of parasites in fecal specimens requires proper collection and handling. Old, poorly preserved, or contaminated specimens are of little value. Additionally, specimens should not be collected for 1 week after the patient has ingested any materials that leave a crystalline residue, such as nonabsorbable antidiarrheal compounds, antacids, bismuth, barium, or antimalarial agents. Oily laxatives such as mineral oil may also interfere with examination ( ). Use of antibiotics or contrast media may decrease the numbers of organisms, especially protozoa, in the intestinal tract for several weeks ( ; , ; ).

Specimens may be submitted to the laboratory fresh or in appropriate preservatives. Fresh specimens should ideally be examined shortly after passage, with liquid specimens being examined within 30 minutes of passage, whereas semi-formed specimens should be examined within 1 hour, and solid specimens within 24 hours of passage ( ). Given that most labs cannot accommodate these time frames, it is best to place the specimen immediately into a suitable preservative to maintain parasite morphology and ensure that fragile protozoal trophozoites are not inadvertently destroyed ( ). Unpreserved specimens should be refrigerated if they cannot be examined immediately. Specimens in preservative can be stored at room temperature.

Specimens may be passed directly into clean, dry containers, or onto a specially designed wax or plastic collection sheet that is placed over the toilet bowl ( ; ). Diarrheic specimens may also be collected in clean bedpans. Containers should have tight-fitting lids and should be placed in plastic bags before transport to the laboratory. Inadvertent introduction of urine or toilet water with the specimen may readily destroy protozoal trophozoites and should be avoided. Also, contamination with water or soil may accidentally introduce free-living organisms that may prove difficult to differentiate from parasitic ones ( ).

Kits consisting of vials of preservatives appropriate for performing direct examinations, concentration procedures, and preparation of stained smears are available from a number of commercial sources at relatively low cost. Aliquots of freshly passed stool should be immediately placed into these vials and mixed thoroughly. These kits are especially helpful for those patients who are unable to bring in a fresh sample in timely fashion or for those who will be collecting several specimens over the course of several days. With the classic two-vial technique, one portion of specimen is fixed in three parts of 5% to 10% buffered formalin and another portion in three parts of polyvinyl alcohol (PVA) fixative. Other available preservation systems include merthiolate-iodine-formalin (MIF) and sodium acetate–formalin (SAF; Table 65.3 ). SAF has an advantage in that it can be used for permanent stains as well as for direct mounts and concentration procedures and it does not contain mercury, which is present in Schaudinn and PVA fixatives. In addition to being poisonous, mercury presents disposal problems in an increasing number of states. However, the quality of permanent stains when SAF is used is not as good as when Schaudinn or PVA fixative is used. Zinc sulfate–based PVA and other newer commercial products such as single-vial multipurpose fixatives are gaining popularity, and their use may be indicated when mercury chloride–based compounds cannot be used ( ; ).

Examination of three specimens collected on different days over a maximum 10-day period is considered the minimum necessary to perform an adequate O&P evaluation ( ; ; ). This procedure ensures an optimum interval for recovery of those parasites known to shed diagnostic forms intermittently. However, for certain parasites, such Strongyloides stercoralis , up to seven O&P examinations may be necessary for optimal detection. Additional sensitivity may be achieved in detecting these parasites, as well as E. histolytica , using antigen detection methods, nucleic acid amplification tests (NAATs) and concentration techniques (discussed later). In general, it is not useful to examine specimens from hospitalized patients who develop diarrhea after the third hospital day since parasites are an unlikely cause of their symptoms. Once the specimens are received in the laboratory, they should undergo macroscopic and microscopic examination as detailed next.

Macroscopic Examination

Fecal specimens should be examined grossly for consistency (formed, soft, loose, or watery) and for the presence of mucus, blood, adult worms, and proglottids. Protozoan trophozoites are more likely to be found in watery or loose specimens, whereas cysts predominate in formed or soft specimens. Helminths or their eggs may be found in any type of fecal specimen. Most parasites are uniformly distributed in the stool as a result of the mixing action of the cecum, although some eggs (especially schistosomes) may enter the fecal stream in the lower colon and rectum and may be unevenly distributed, as may pinworm and Taenia spp. eggs. Protozoan trophozoites may be more numerous in the last portion of stool evacuated and should be specifically sought in mucus ( ).

Microscopic Examination

Specimens may be examined microscopically by direct wet mounts of fresh or preserved material, wet mounts of concentrated feces, or permanent stains. Each procedure has specific advantages and limitations, and not all may be routinely used in the clinical laboratory. Direct saline wet mounts of fresh feces allow detection and observation of motile protozoan trophozoites and helminth larvae. Direct mounts of preserved feces may allow detection of parasites that do not concentrate well. Concentration procedures increase the examiner’s ability to detect protozoan cysts and helminth eggs and larvae but are unsatisfactory for detecting protozoan trophozoites. Permanent stains are useful for detection and morphologic examination of protozoan trophozoites and cysts.

The circumstances under which each procedure is performed vary depending on the workflow of the laboratory. As mentioned earlier, examination of unpreserved specimens allows for enhanced detection of motile parasites. Many protozoa can also be differentiated by their characteristic motility patterns. However, examination of unpreserved stool is useful only when specimens can be examined shortly after being passed. The direct wet mount may be omitted if the specimen is submitted in a preservative. At a minimum, fixed specimens should be examined by a concentration procedure, which provides increased sensitivity for detection of parasites. However, improved yield has been demonstrated when a permanent stain is also examined, as the two preparations are complementary. Therefore, the optimal stool examination includes both a concentrated wet prep and a permanently stained preparation ( , , ; ). The following provides detailed instructions for performing the direct wet mount, concentration techniques, and permanently stained preparations.

Additional Techniques for Examination of Enteric Parasites

Cellulose Tape and Paddle Techniques for Pinworm

The female pinworm, Enterobius vermicularis , migrates from the cecum to the perianal skin, where she deposits typical eggs that are partially embryonated. The eggs or, occasionally, adult worms may be detected on examination of clear, adhesive cellophane tape or commercial collection kits that have been pressed on to the perianal skin ( Fig. 65.2 ).

Eggs or adults are not commonly found in stool concentrates, which is considered to be an inappropriate specimen for detection of this parasite. Specimens should be collected first thing in the morning before bathing or defecation. Several specimens taken on different days should be examined before infection is ruled out. Commercial devices such as the SWUBE (Becton Dickinson, Franklin Lakes, NJ) use an adhesive paddle for collection and greatly simplify specimen collection and examination.

Nematode Culture and Recovery Techniques

Several culture techniques (coproculture) assist in the detection and identification of certain nematode infections, including the Harada-Mori filter paper strip culture, filter paper/slant culture, and charcoal culture ( ; , ; ; ). Differentiation of hookworms and trichostrongyles on the basis of egg morphology is difficult, whereas infective-stage larvae are more readily identified. Such culture techniques may also prove useful in recovery of Strongyloides larvae, which may be few in number, and in differentiating them from those of hookworms. With all culture methods, feces are incubated in a humid environment to encourage egg hatching. With the Harada-Mori and filter paper/slant techniques, larvae migrate from the feces into a water phase, where they may be readily detected. In the charcoal culture, larvae first migrate into a dampened gauze pad, which is then placed in water, allowing the larvae to settle out. These methods are most commonly used in clinical laboratories in endemic settings.

The Baermann funnel technique and agar culture methods are sensitive and reliable methods for recovery of Strongyloides and other nematode larvae from a stool specimen. In the Baermann assay, feces are placed on several layers of gauze on top of a wire screen that is suspended in a funnel. The bottom of the funnel is clamped off, and water is added to the level of the gauze. Larvae actively migrate through the gauze and settle to the bottom of the funnel, where they may be drawn off for examination. Although this method provides increased sensitivity over the traditional O&P exam, it is labor-intensive and used infrequently in the clinical laboratory. The agar culture technique provides a simpler and more sensitive means for detecting S. stercoralis in feces. With this method, feces are plated on a nutrient agar and incubated at room temperature for several days. Over time, the larvae will migrate out of the feces into the agar and carry fecal bacteria with them. Growth of the bacteria in the larval tracks facilitates identification of larvae in the specimen ( Fig. 65.3 ).

In latent Strongyloides infection, in which few larvae are being shed, several examinations over 1 week using a concentration technique may be required to detect the infection ( , ; , ). It is important to note that filariform larvae of S. stercoralis and hookworms are highly infectious; therefore, testing must be performed using universal precautions.

Objects Resembling Enteric Parasites

A large variety of objects that closely resemble various parasite life cycle stages may be seen in feces and other specimens sent for O&P examination. Careful differentiation of these objects from real parasites is necessary to prevent inappropriate or unnecessary treatment. White blood cells, macrophages, and squamous and columnar epithelial cells may resemble amebae; yeasts and starch granules may resemble protozoal cysts or oocysts; pollen and fungal conidia may resemble helminth eggs; plant fibers may resemble nematode larvae; and pieces of vegetables or vegetable skins may resemble adult worms or proglottids. Examples of artifacts and pseudoparasites have been reviewed elsewhere ( ; ; ).

Antigen Detection

Antigen detection methods are commercially available for several parasitic diseases, including amebiasis, cryptosporidiosis, giardiasis, malaria, and trichomoniasis ( Table 65.4 ). These methods may be useful for initial testing or in instances in which traditional tests are negative, yet a high index of clinical suspicion remains. These tests offer the advantage of detecting current infection and can often be performed by someone other than an experienced morphologist ( ; Shimizu & Garcia, 2018).

Antigen detection in stool samples is usually performed using fecal immunoassays. A number of published studies have suggested that these assays have good or superior sensitivity and specificity when compared with routine ova and parasite examination ( ; ). These immunoassays are easy to use and rapid, permit batch processing, and do not require experienced microscopists. Given the current shortage of medical technologists and individuals with specialized training in parasitology, use of immunoassays appears to be an attractive alternative. However, laboratories that use rapid cartridge-based immunoassays should be aware of potential problems with false-positive results and should closely monitor test performance. Currently, fecal immunoassays are marketed for G. duodenalis , C. parvum / C. hominis , the E. histolytica / E. dispar group, and E. histolytica . Antigen detection tests using blood or serum are also available for Plasmodium spp. and W. bancrofti . A latex agglutination test for T. vaginalis antigen detection in vaginal swabs has also been introduced. Immunoassays are usually available in three formats: EIA, DFA, and lateral flow (immunochromatography) cartridges. Fresh or preserved stool samples are appropriate for most antigen detection kits ( ). Although each kit has unique operating characteristics, most are generally comparable in performance ( ; ).

Rapid antigen detection tests (RDTs) developed for malaria may detect histidine-rich protein II (HRP-II), parasite lactate dehydrogenase (pLDH), parasite aldolase, or a combination of these antigens in peripheral blood. HRP-II tests are specific for Plasmodium falciparum , and pLDH and aldolase tests detect all four human Plasmodium spp. These assays have highly variable performance characteristics, but in general are adequate for detecting moderate to heavy infections with Plasmodium falciparum . They are often significantly less sensitive for detecting lower levels of P. falciparum infection and infections with other Plasmodium spp. At the time of this writing, only the BinaxNow (Abbott Diagnostics, Chicago, IL) is cleared by the U.S. Food and Drug Administration (FDA) for clinical diagnosis of malaria in the United States ( ).

Trichomonas vaginalis antigens from vaginal samples may also be detected using rapid antigen tests such as the OSOM Trichomonas Rapid Test (Sekisui Diagnostics, Burlington, MA), which is an FDA-cleared, Clinical Laboratory Improvement Amendments (CLIA)–waived test for detecting T. vaginalis antigen in vaginal swabs. These tests can be used for rapid detection of T. vaginalis infection in the clinical setting and may replace wet mount examinations, which generally have lower sensitivities (75%–96%) compared with NAATs ( ).

Most EIAs are available in microwell format ( ). Antigens from frozen, fresh, or 10% formalin–preserved stool samples are suitable for testing by this method. Concentrated or PVA samples are not suitable for testing with EIA kits. Parasite antigen is captured by immobilized antibodies coated on microwells and is detected by an enzyme-conjugated secondary antibody that is capable of producing a colored reaction following the addition of substrate. Although the colored wells can be read visually or with the use of a spectrometer, the latter seems to be the preferred option because of occasional ambiguous results obtained with some kits ( ). In general, EIA tests have good sensitivities and specificities. Garcia and colleagues evaluated nine immunoassay kits for detection of G. duodenalis and Cryptosporidium spp. in comparison with a reference DFA test that visualizes the parasite directly in the sample. Investigators found that all kits had high sensitivities, ranging from 94% to 99%, and 100% specificities ( , ). This is in contrast to first-generation EIAs, which had been previously reported to produce false-positive results, resulting in their recall ( ). Hence, a strong quality control (QC) pro and participation in proficiency test pros are required to ensure high-quality test results. Local epidemiology of the parasitic infection can help to determine whether additional confirmatory testing is required; consultation with local public health authorities may prove useful in characterizing which infections are being seen locally. Additionally, for some diseases such as giardiasis, examination of two specimens by EIA or microscopy may be necessary to achieve diagnostic sensitivity greater than 90% ( ).

Lateral flow cartridges are a popular format of immunoassay because of their ease of use and the minimal performance time required. These kits can be stored conveniently at room temperature and may be used in single or batch processing. The parasite antigen in the sample migrates through the membrane and binds to specific capture antibodies. Use of a secondary reagent results in development of a colored reaction. These kits also have an internal control to ensure that the colloidal dye conjugates used in the assay are intact and that proper capillary flow has occurred. To ensure complete migration of the specimen, only the supernatant of a well-mixed stool sample is used, and some samples may be diluted to a liquid state before testing. Any color visible at the reagent test zone (usually a band) is interpreted as positive. Some studies have demonstrated that cartridge assays are somewhat less sensitive than a microwell EIA plate assay ( ; ). When lack of sensitivity is a concern, it may be necessary to perform alternative O&P or NAAT studies if the patient’s symptoms persist.

DFA testing allows direct visualization of the parasites in stool specimen using antibodies conjugated to fluorescent dyes. These assays are easy to perform and to interpret, permitting rapid screening of slides when compared with some of the traditional stains used in parasitology ( ). A fluorescence microscope is necessary for this procedure, which is a limiting factor in some laboratories. Currently, kits are available for detection of cysts of G. duodenalis and oocysts of Cryptosporidium spp. Fixed stool specimens may be used for this procedure (10% formalin, SAF, or one of the mercury- or formalin-free products) ( ). Although fresh stool samples can be tested directly, the sensitivity of the assay can be improved by performing the test on centrifuged stool (500 g for 10 minutes). Occasionally, fluorescing bacteria and yeasts may be seen, but these are readily distinguished from Giardia and Cryptosporidium on the basis of their size and shape. The edges of the wells should be carefully examined to avoid missing the rare parasite in light infections. Given the relatively recent recognition of additional Cryptosporidium spp. that may infect humans, commercial assays currently available may not be adequate for detection of all infections.

Antibody Detection

Tests that are available from public health, hospital, or commercial laboratories to detect immunologic reactivity to parasitic diseases are summarized in Table 65.5 . Historically, serologic procedures for parasitic diseases have been plagued by low sensitivity and specificity, primarily owing to the complex antigenic nature of parasites and the possibilities for cross-reactions from related species. However, newer test methods combined with the use of more highly defined antigenic components has provided more accurate results with greater predictive values. Many of the newer tests use the EIA or immunoblot (Western blot) format, although IFA, indirect hemagglutination (IHA), and complement fixation (CF) are still in use ( ).

In general, serologic diagnosis of parasitic infection is used as an adjunct to the usual diagnostic modalities or in special situations in which identification of the parasite itself or its antigen or nucleic acid from host tissue or excreta is not possible ( ). For example, parasitic infections such as toxoplasmosis and toxocariasis reside in deep tissues and cannot be readily diagnosed by morphologic means. Others such as cysticercosis and echinococcosis develop in organs, where invasive studies that may be required are not recommended in the initial patient evaluation. Additional conditions—such as filariasis, schistosomiasis, and strongyloidiasis—may remain subclinical because of light infections or because the clinical evaluation occurred during the prepatent period (interval from infection to demonstration of symptoms or recovery of organism). In these settings, serologic testing may be invaluable for making a presumptive diagnosis in correlation with the accompanying clinical and radiographic features. Other circumstances in which serologic evaluation may prove useful include diagnosis of extraintestinal amebiasis (e.g., amebic liver abscess), trichinellosis, and chronic stages of trypanosomiasis. Lastly, serologic studies serve as a powerful tool in enhancing our understanding of the epidemiology of diseases such as schistosomiasis, toxoplasmosis, amebiasis, Chagas disease, malaria, and babesiosis, and for screening blood donors for select infections (e.g., malaria, Chagas disease) ( ).

Interpretation of serologic testing may be challenging. Detection of antibodies, especially immunoglobulin (Ig) G, provides evidence of infection but may not be able to differentiate active from past exposure. High antibody levels are useful for diagnostic purposes if they occur in a patient with no previous exposure to the parasite and no recent history of travel to an endemic area. Unfortunately, positive antibody levels in persons living in endemic areas often do not help in the clinical diagnosis. In some parasitic diseases, levels of antibodies may decline slowly following successful therapy or self-cure and thus may be useful for monitoring response to treatment ( ).

Serologic tests for parasitic diseases generally evaluate IgG levels with the exception of toxoplasmosis and babesiosis, in which IgM- and IgA-specific antibodies may be helpful for determining the age of infection ( ). Unfortunately, IgM and IgA may persist for as long as 2 years after the primary infection, thus limiting their utility for differentiating acute from past infection. When Toxoplasma gondii IgG antibodies are detected, avidity testing may also be useful to distinguish recent from past infection, particularly during pregnancy ( ; ; ). This testing is based on the principle that the initial host response results in the production of low-avidity antibodies. Over time, the antibodies gain higher avidity as the host immune response is enhanced.

Because serologic tests for most parasitic diseases are requested infrequently, specimens generally are submitted to public (CDC) or private reference laboratories. Some of the more commonly requested tests are available as commercial kits. However, many of these assays are developed in-house and, hence, lack correlation with universal standards. Interpretive criteria are established by reagent manufacturers or by the center performing the test; these criteria often vary from institution to institution. Individuals requesting such tests should inquire about the performance characteristics, including sensitivity and specificity, and should be aware that cross-reactions may occur. For example, antibody tests for Chagas disease are known to cross-react with antibodies produced in response to Leishmania infections. However, reactivity to homologous antigen is greater, and this test is useful in diagnosing chronic stages of the disease when parasitemia is generally low. Usually, serology for chronic Chagas disease correlates well with molecular diagnostic methods ( ). Helminthic parasites are well known to cross-react in serologic assays that use crude antigen preparations because of phylogenetic, hence antigenic, similarities.

Several factors that may influence the test performance of serologic assays include disease manifestation, test format, reagents used, and parasite viability, to name a few. The sensitivity of the test is increased in patients with invasive amebiasis but may be weak in intestinal amebiasis with minimal tissue invasion and absent for asymptomatic carriers. The type of serologic assay format may also determine the sensitivity, as in the diagnosis of toxoplasmosis ( ). The double-sandwich IgM enzyme-linked immunosorbent assay (ELISA) is known to be more sensitive and specific than IgM immunofluorescence for detecting recently acquired and congenital toxoplasmosis. IHA has been the primary test for serodiagnosis of amebiasis. The sensitivity of the assay is also dependent on the type or stage of parasite antigen used. For example, the sensitivity of cutaneous leishmaniasis can be improved by using amastigote antigens in place of promastigote antigens in the IFA test. Finally, serologic assays are also affected by parasite viability; hydatid cysts occurring in the lung and dead or calcified cysts are less frequently detected than active cysts in the liver. This also holds true for neurocysticercosis (larval infection with the cestode Taenia solium ), in which sensitivity is low when only a single parenchymal cyst or calcified lesions are present ( ).

Life Cycle

Plasmodium spp. have a complex life cycle involving a mosquito definitive host and a vertebrate intermediate host (see Fig. 65.4 ). Human infection is initiated when a female Anopheles mosquito takes a blood meal and injects infectious sporozoites into the bloodstream. Sporozoites are carried to the liver and invade hepatic parenchymal cells, initiating a proliferative phase known as extraerythrocytic sporogony . Eventually, these hepatic schizonts rupture, releasing merozoites into the bloodstream, which infect erythrocytes. Developing parasites are known as trophozoites . Early trophozoites are usually characterized by a thin or thick vacuolated cytoplasm and one or two chromatin dots and are referred to as rings or ring-form trophozoites . Hemozoin pigment, a breakdown product of hemoglobin, is characteristic of erythrocytes containing mature stages of malarial parasites but is not usually evident in ring forms. As the trophozoites mature, they take one of two pathways.

In one pathway, trophozoites become erythrocytic schizonts that produce merozoites and perpetuate the erythrocytic cycle. The erythrocytic cycle takes approximately 48 hours (tertian periodicity) for P. falciparum , P. ovale , and P. vivax infections, and 72 hours (quartan periodicity) for P. malariae infection. Clinical symptoms of fever spikes are ties to the rupturing of erythrocytic schizonts. In the second pathway, trophozoites become macro- (female) or microgametocytes (male). Gametocytes are a dead-end stage in the human host; their sole purpose is to be picked up by a mosquito host to initiate the sexual cycle. In the mosquito host, microgametocytes exflagellate and the released microgametes fertilize a macrogamete, resulting in the formation of a motile ookinete . Ookinetes migrate outside the gut wall of the mosquito and become an oocyst. Within the oocyst, sporogony results in the formation of infectious sporozoites. The mature oocyst ruptures into the body cavity, releasing sporozoites, which then migrate through the tissues to the salivary glands, from which they are injected into the vertebrate host as the mosquito feeds. The time required for development in the mosquito ranges from 8 to 21 days.

Plasmodium vivax and P. ovale are unique in that true disease relapses may occur weeks to months following subsidence of previous attacks. This occurs as a result of renewed extraerythrocytic and, eventually, erythrocytic schizogony from latent hepatic parasite forms known as hypnozoites . Recurrences of disease due to P. falciparum or P. malariae , called recrudescences , arise from increased numbers of persisting blood stage forms to clinically detectable levels and not from persisting liver stage forms. Liver cells are infected only by sporozoites from the mosquito; thus, transfusion-acquired P. vivax or P. ovale infection does not relapse ( ).

Epidemiology

Endemic transmission of malaria requires a reservoir of infection, an appropriate mosquito vector, and a susceptible host. Control of malaria is directed at elimination of mosquito hosts, treatment of active cases, and prophylaxis of susceptible persons. However, the emergence of mosquitoes resistant to insecticides, the development of resistance to prophylaxis and therapy by P. falciparum and P. vivax ( ) and lack of adequate funding have made control difficult in endemic areas. Fortunately, renewed global efforts have dramatically decreased the morbidity and mortality of malaria worldwide, although further efforts are needed to achieve full control and elimination ( ).

A number of hereditary erythrocyte genotypes offer some degree of protection against severe P. falciparum malaria, including sickle cell trait, glucose-6-phosphate dehydrogenase (G6PD) deficiency, hereditary ovalocytosis, and thalassemia ( ; ). Alterations of erythrocyte cell membrane surface proteins may interfere with parasite invasion and confer a protective effect against malaria infection and severe disease. Individuals who lack the Duffy blood group antigen, for example, have a decreased likelihood of P. vivax infection since this is the primary protein used by P. vivax to bind and invade erythrocytes ( ). Alternate binding pathways have recently been identified for P. vivax , providing an explanation for the numerous reports of P. vivax malaria in Duffy blood group–negative individuals ( ).

Transfusion-acquired malaria may occur when blood donors have subclinical malaria and may prove fatal for the recipient. Similarly, congenital malaria may occur in infants born to mothers from endemic areas. The infant acquires the infection at birth as a result of rupture of placental blood vessels leading to maternal-fetal transfusion. Neither transfusion nor congenital malaria is expected to relapse because exoerythrocytic schizogony does not occur. The number of cases of malaria reported in the United States averages about 1700 cases per year ( ). The most recent surveillance summary from the CDC reported 1517 total cases in 2015 ( ). Species causing infection were P. falciparum (67.4%), P. vivax (11.7%), P. ovale (4.1%), P. malariae (3.1%), and undetermined (12.9%), with less than 1% having infection with two species. Of reported cases, 17.1% were associated with severe disease, including the 11 persons who died. Only 4.7% of patients with malaria reported adhering to a CDC-recommended chemoprophylaxis malaria regimen. Patients acquired the infection in Africa (84.7%), Asia (8.6%), the Western Hemisphere (4.6%), or Oceania (2 cases). In patients for whom the reason for travel was known, the majority (68.4%) were visiting friends and relatives (VFRs). These individuals have long been known to be at higher risk for acquiring malaria than other travelers since they often do not seek pre-travel counseling, do not take malaria chemoprophylaxis, travel to more rural areas, and often stay for longer periods of time.

Clinical Disease

Most patients who develop P. falciparum infection become symptomatic within 1 month of exposure, whereas a delay of up to 6 months or more may be seen with the other Plasmodium species. Common presenting symptoms of malaria include chills and fever, which often are associated with splenomegaly. In the early stages of the disease, febrile episodes occur irregularly but eventually become more synchronous, assuming the usual tertian ( P. vivax , P. falciparum , and P. ovale ) or quartan ( P. malariae ) periodicity. Patients with malaria may develop anemia and may have other manifestations, including diarrhea, abdominal pain, headache, and muscle aches and pains. Plasmodium falciparum malaria can result in high rates (50%) of parasitemia, which can lead to severe hemolysis with hemoglobinuria and profound anemia. Erythrocytes infected with growing trophozoites and schizonts of P. falciparum become sequestered in small vessels of the body, which may lead to occlusion of these vessels, causing symptoms related to capillary obstruction and tissue anoxia. Involvement of the brain is known as cerebral malaria, in which the patient becomes disoriented, progressing to delirium, coma, and often death.

The course of untreated malaria depends on the species. Most fatal cases of malaria are due to P. falciparum , although P. knowlesi can also cause fatalities. In nonfatal cases, the febrile paroxysms become less severe with time and the disease gradually subsides. Patients with P. vivax or P. ovale infection may have relapses after many months or, occasionally, years. Persons with P. falciparum and P. malariae infection may have symptom-free periods but suffer from sporadic recrudescences owing to persisting low-grade parasitemia. Relapses and recrudescences may be associated with changes in the host’s defense mechanisms or possibly with antigenic changes in the infecting organisms.

Peripheral smears may show leukocytes that contain malaria pigment (hemozoin). Increased reticulocyte counts occur commonly and are associated with rapid erythrocyte turnover. The presence of greatly enlarged platelets may be noted on peripheral blood films and may occur as a result of their rapid turnover secondary to splenic sequestration. Malarial infection may interfere with certain serologic tests, producing false-positive results, especially those for syphilis.

Therapy and prophylaxis of malaria have become highly complex topics because of the widespread appearance of resistance by P. falciparum to chloroquine and other antimalarials and, to a lesser extent, resistance by P. vivax to chloroquine. Artemisinin combination therapy is now recognized as the preferred treatment by the WHO for treatment of P. falciparum infection and chloroquine-resistant P. vivax infection ( ). Persons with P. vivax or P. ovale malaria should receive treatment with primaquine phosphate or tafenoquine in addition to standard primary treatment in order to eradicate hepatic hypnozoites and to prevent relapse. Use of primaquine or tafenoquine may be dangerous in patients who have G6PD deficiency. Screening of at-risk patients before therapy is initiated may be necessary ( ; ). Exchange transfusion may decrease morbidity and mortality in severe cases of P. falciparum infection in which the parasitemia is ≥10% ( ). While exchange transfusion is no longer recommended in this setting by the CDC ( ), it is still supported in cases of severe malaria with high parasitemia by the American Society for Apheresis ( ).

Diagnosis

Malaria should always be included in the differential diagnosis of fever in patients who have a history of travel to or residence in endemic areas ( ). Given the potentially life-threatening nature of infection, testing must be performed on a STAT basis. Diagnosis usually is established by demonstrating parasites in thick and thin blood films. Blood specimens ideally are collected just before the next anticipated fever spike or at the onset of fever. Specimens drawn several hours apart sometimes may be required to demonstrate infection or to diagnose the species because the number and morphologic stage of parasites vary during the cycle. Careful examination of thick films should reveal the presence of the parasites in almost all patients with clinically apparent malaria.

Identification of malarial parasites on thin blood films requires a systematic approach. Three major factors should be considered: appearance of infected erythrocytes, appearance of parasites, and stages found. Table 65.7 summarizes diagnostic characteristics of the species, which are illustrated in Figures 65.5 and 65.6 . The size of the infected erythrocyte is a particularly useful feature for determining the infecting species; erythrocytes infected by P. vivax or P. ovale parasites often appear enlarged compared with adjacent uninfected cells, whereas P. malariae and P. falciparum parasites are usually found in erythrocytes of normal size. Erythrocytes infected with P. ovale are often oval or fimbriated (having irregular, usually polar or bipolar, projections of the cell margins), a feature rarely observed with P. vivax . Schüffner stippling, noted as numerous small uniform pink granules in the erythrocyte, is usually seen in cells infected with P. vivax and P. ovale , although it may not be evident in cells infected with early ring forms or on slides that have not been stained at the appropriate pH, such as the Wright-Giemsa stain, used widely in hematology laboratories (see the Laboratory Methods section earlier in the chapter). The presence of Schüffner stippling is helpful because it is not seen in P. malariae or P. falciparum infection.

As trophozoites grow in the infected cells, the amount of hemoglobin in the erythrocyte decreases and hemozoin pigment accumulates. The amount and appearance of the pigment vary among species. Ring forms of all parasites may have a similar appearance, and if only occasional ring forms are found, the species may not be identifiable. Young rings of P. falciparum tend to be smaller than those of the other species (one-sixth the diameter of the red blood cell compared with one-third the diameter of the red blood cell for the other species). Rings of P. falciparum that have grown are similar in size to those of the other species. Trophozoites that appear to be lying on the surface of the erythrocyte or protruding from it are called appliqué or accolé forms, most often seen in P. falciparum infection. Doubly infected cells and double chromatin dots in ring trophozoites occur most commonly in P. falciparum infection but can occur with the other species as well.

Growing trophozoites of P. vivax have irregular shapes and are termed ameboid . Those of P. malariae and P. ovale tend to remain compact, although vacuolated forms can occur (with P. malariae , these can take on a characteristic shape called the “basket-form” trophozoite). Mature trophozoites and schizonts of P. falciparum are usually sequestered in capillary beds secondary to cytoadherence to endothelial cells and are not seen in the peripheral blood except in very severe cases of infection. When schizonts are identified in the peripheral blood, determining the number of merozoites is helpful in identifying the various species. Gametocytes of P. falciparum are readily identified by their characteristic sausage shape. Gametocytes of P. vivax , P. malariae , and P. ovale have a similar shape. Thus, they are difficult to differentiate, although characteristics of infected red blood cells can aid in identification.

The varieties of developmental stages in the peripheral blood aid in diagnosis. In P. falciparum infection, ring forms predominate; finding numerous ring forms without more mature stages is highly suggestive for P. falciparum infection. In P. vivax , P. malariae , and P. ovale infections, various stages of parasites are found with some predominance of one stage depending on the phase of the cycle.

Thick films are preferred for detecting malarial infections because a greater quantity of blood is examined (see the Laboratory Methods section earlier in the chapter). Ring forms often have the appearance of punctuation marks rather than complete rings, and the presence of red chromatin and blue cytoplasm should be required to identify them as parasites (see Fig. 65.6A ). Schüffner stippling still may be a helpful identifying characteristic, which may be recognized around growing trophozoites as a pink halo rather than as distinct granules seen in thin films. The ameboid character of P. vivax trophozoites is not as evident in thick films, but the number of merozoites in mature schizonts is helpful. Gametocytes usually cannot be differentiated. The distinctive sausage shape of P. falciparum gametocytes is still evident, although they may appear stubbier than in thin films. Gametocytes of the other species can be detected and are easily differentiated from host cell nuclei by the presence of refractile hemozoin pigment.

Mixed infections occur occasionally, but caution should be used in making such a diagnosis unless definite evidence reveals two separate populations of parasites. The most common mixed infections are P. falciparum and P. vivax . Finding gametocytes of P. falciparum in a person obviously infected with P. vivax is diagnostic, for example.

Multiple artifacts may be confused with malarial parasites on thick and thin films. The most common artifacts on thin films are blood platelets superimposed on red blood cells. These platelets should be readily identified because they do not have a true ring form, do not show differentiation of the chromatin and cytoplasm, and do not contain pigment. Clumps of bacteria or platelets may be confused with schizonts. At times, masses of fused platelets may resemble gametocytes of P. falciparum but do not show the differential staining or the pigment. Precipitated stain and contaminating bacteria, fungi, or spores may also be confused with these parasites.

Species-specific serologic tests for malaria are not useful for diagnosis of acute infection but may be useful for epidemiologic surveys and detection of infected blood donors ( ). Such tests do not reliably differentiate current from past infection, however. Sensitive and specific IFA tests using antigens from the four human species are available from the CDC. Assays for the direct detection of malarial antigens in blood are especially useful (see the Laboratory Methods section earlier in the chapter) ( ).

Trypanosoma spp. (African and American Trypanosomiasis)

Infections with trypanosomes include those caused by Trypanosoma brucei (African trypanosomiasis) and T. cruzi (American trypanosomiasis, or Chagas disease). Both are of great importance in endemic areas. A third species, Trypanosoma rangeli , has been described in humans in the Americas but does not cause clinical illness. Bloodstream trypomastigotes of the T. brucei group (see Fig. 65.6D ) are up to 30 μm long with graceful curves and a small kinetoplast. Those of T. cruzi are somewhat shorter (20 μm) and display a larger kinetoplast. Also, dividing forms may be seen in blood films with T. brucei but not T. cruzi , as the latter replicates only in the amastigote stage in the human host.

Leishmania spp. (leishmaniasis)

Leishmaniasis is a disease of the reticuloendothelial system caused by kinetoplastid protozoa of the genus Leishmania . All species that infect humans have animal reservoirs and are transmitted by sand flies belonging to the genera Phlebotomus in the Old World and Lutzomyia in the New World. The parasites assume the amastigote form in mammalian hosts and the promastigote form in insect vectors. Species of Leishmania cannot be differentiated by examination of amastigotes or promastigotes. Leishmaniasis may assume many different clinical forms; cutaneous, mucocutaneous, and visceral diseases are best known. The form and severity of disease vary with the infecting species, the particular host’s immune status, and prior exposure ( ).