Parasitology

Thick and thin blood films.

Thick and thin blood films are the most commonly used preparations for detection of blood parasites. ^{, ,} Blood can be obtained via venipuncture, or by finger-stick or earlobe puncture, and then spread onto clean, grease-free glass microscope slides. If the latter collection methods are used, then blood must be allowed to flow freely from the puncture site before collection is attempted to prevent contamination of the specimen with tissue fluid or the alcohol disinfectant. Ideally, the slides are prepared at the bedside immediately after obtaining the blood. However, if the blood must be transported to the laboratory prior to slide preparation, then it should be collected using ethylenediaminetetraacetic acid (EDTA) as the anticoagulant. It is important to prepare the slides as soon as possible after collection, since prolonged exposure to EDTA can result in distortion of the parasite morphology, particularly with Plasmodium spp.

Thin blood films are prepared in the same manner as for hematology evaluation, with creation of a thin layer of intact, mostly nonoverlapping blood cells. To make the thin film, one drop of blood (approximately 0.05 mL) is placed at one end of the slide, and a second slide is used to sweep the drop along the length of the slide. The well-prepared thin film contains a “feathered” edge of approximately 1.5 to 2 cm in length in which the erythrocyte and parasite morphology ( Plasmodium and Trypanosoma spp.) are ideal for microscopic examination ( Fig. 88.1 ). ^, The slide is fixed in methanol prior to staining, allowing the erythrocytes (and intraerythrocytic parasites, if present) to remain intact during staining. Therefore the thin film provides the ideal morphology for determining the species and percent parasitemia of intraerythrocytic parasites.

As the name implies, the thick film provides a greater concentration of blood than the thin film (16 to 30 times more blood per microscopic field than with the thin film) and therefore provides a more sensitive preparation for parasite screening. The reason that it is possible to examine this thicker film of blood is that the erythrocytes are lysed during the staining process so that only leukocyte nuclei, platelets, and parasites (if present) are visible. ^{, ,} The thick film is made by placing one to two drops of blood on the slide, and then using a second slide (or wooden stick) to spread the blood into a circular film. The ideal thick film should be 1.5 to 2 cm in diameter and of a thickness through which small typed print (e.g., newsprint) can just be read. The blood film must then be allowed to thoroughly dry so that it does not detach during the staining process. While older methods recommended thick film drying times of 8 to 12 hours, this prolonged period can be avoided by gently pushing down with the corner of the spreader slide while creating the film, thereby creating minute scratches on the carrier slide ( Fig. 88.2 ). This greatly improves adherence of the thick film to the carrier slide and allows for staining to take place as soon as the film is visibly dry (within 30 to 60 minutes). Regardless of the method used to distribute the blood on the slide, it is important not to fix the film in methanol since the erythrocytes will not effectively lyse during staining.

Giemsa is the preferred stain for both the thick and thin films since it provides the best contrast between the intraerythrocytic parasites and hemoglobin. It must be made fresh daily and is usually prepared by diluting Giemsa stock solution with phosphate-buffered distilled or deionized water. A small quantity of Triton X-100 (5 to 10%) can be added to the buffer to improve the contrast and staining of the organisms. The buffer pH should be at 7.0 to 7.2 to allow for optimal visualization of parasite-associated intracytoplasmic inclusions (i.e., Schüffner stippling and Maurer clefts). Wright-Giemsa and Wright stains may also be used, but the thick films must first be lysed in water since these stains commonly incorporate alcohol as a fixative. These stains generally have a pH of less than 6.8, and therefore stippling will usually not be seen. The Field stain may also be used as a rapid method for thick and thin blood film staining and is well suited for field use.

The entire thick and thin films should be examined using low magnification (i.e., 10× or 20× objective) to allow for detection of larger blood parasites such as microfilariae. At least 100 thick film fields or 200 thin film fields should also be examined using high magnification (100× oil immersion objective) before issuing a negative report ( Figs. 88.3 and 88.4 ). Malaria and babesiosis are potentially life-threatening conditions, and therefore testing should be performed on a STAT basis. ^, Some laboratories have opted to use rapid immunochromatographic tests for initial malaria testing due to their ease of use and interpretation (see the “Immunodiagnostic Methods” section later in this chapter). It is important to note that these methods are less sensitive than thick and thin blood film microscopy, particularly for non– Plasmodium falciparum infections and low levels of parasitemia. Therefore confirmatory testing using conventional blood films should still be performed, particularly in nonendemic settings. ^,

Concentration techniques.

Concentration techniques are recommended for more sensitive detection of the larger blood parasites (trypanosomes and microfilariae) and organisms that infect white blood cells (Leishmania). ^{, , ,} One relatively simple technique is the creation and examination of a buffy coat smear. In this method, anticoagulated blood is centrifuged to separate the denser erythrocytes from plasma and allow for formation of an intervening buffy coat layer. This layer, which primarily contains leukocytes and platelets, can be drawn off and used to prepare a wet mount (for identifying motile organisms) or Giemsa-stained blood films. The Knott’s concentration and membrane filtration techniques are also useful for detection of microfilariae, particularly in low-density infections. Like the buffy coat preparation, the Knott’s concentration method uses centrifugation to separate the larger microfilariae. However, the blood is first lysed with 2% formalin, allowing the microfilariae to concentrate in the sediment. The sediment can be examined as a wet or stained preparation. With the membrane filtration technique, approximately 1 mL of blood is lysed and passed through a filter with 3-to 5-μm pores to trap the microfilariae. The filter is then removed and placed on a microscope slide for examination.

Specimen collection and handling.

Proper collection and handling are essential for optimal recovery and identification of parasites in fecal specimens. Examination of poorly preserved specimens or those containing interfering substances (e.g., antibiotics, barium, bismuth, oily laxatives, and nonabsorbable antidiarrheal medications) provide little to no clinical value and should be avoided. ^{, ,} Ideally, specimens should be collected at least 7 days after completion of antibiotic therapy or product use. Fecal specimens should be collected in a clean container (not contaminated with urine, water, or soil) and may be submitted to the laboratory in a fresh or preserved state. Table 88.6 lists the most common preservatives and the types of preparations that can be made with each. The conventional two-vial preservative consists of 5 to 10% buffered formalin and polyvinyl alcohol (PVA). More recently, formalin- and mercury-free commercial single-vial proprietary fixatives have become increasingly popular due to their relative lack of toxicity and ease of use. ^{, ,} It is important to note that not all preservatives are amenable for use with enzyme immunoassay (EIA) and polymerase chain reaction (PCR) tests. ^, Fresh specimens must be examined as soon as possible (within 30 minutes for soft or liquid specimens or within 1 hour for semi-formed specimens) to avoid degradation of fragile trophozoites. If it is not possible to examine fresh specimens within this time frame, they should be placed immediately into an appropriate preservative. Vials of preservative should be given to patients for collection so that multiple specimens may be collected over a period of time. All specimens should be considered potentially infectious and handled using standard precautions. ^,

Ideally, three or more specimens (collected on consecutive days or every other day over a 10-day period) should be examined for optimal detection of most parasites; up to seven specimens must be examined for optimal detection of S. stercoralis.

Macroscopic and microscopic examination.

Fecal specimens should be examined macroscopically for consistency (e.g., watery, soft, formed) and the presence of blood, mucus, and whole worms and proglottids. The consistency of the specimen can provide important clues to the types of organisms that might be present; trophozoites are more frequently found in watery to soft specimens, while cysts are generally the only stages found in formed specimens.

A portion of the specimen can then be selected for microscopic examination, taking care to sample bloody or mucoid areas that may contain higher numbers of trophozoites. Three types of microscopic preparations may be used for parasite detection: the direct wet mount of fresh unpreserved stool, the wet mount of preserved concentrated stool, and the permanently stained smear.

Each preparation has specific limitations and advantages. The direct wet mount is made by placing a small amount of stool onto a slide and mixing it with a drop of 0.85% saline. The specimen is then coverslipped, and the entire specimen is examined using the 10× objective (i.e., 100× magnification). At least 1/3 of the coverslip should also be examined using the 40× objective (i.e., 400× magnification). A second (optional) preparation may be made in a similar fashion by using a drop of 1:5 dilution of Lugol iodine instead of saline in order to increase the contrast of protozoal trophozoites and cysts. Direct examination of the fresh, unpreserved specimen is used primarily for detection of motile parasites (larvae and trophozoites), and it is therefore essential to examine the specimen shortly after passage, before motile parasites die and begin to degrade. Since it is generally more important to preserve parasite morphology than motility, laboratories that cannot examine specimens within the narrow recommended time frame should forego the direct preparation in favor of placing the specimen immediately into a preservative.

Concentrated preparations can be made using either fresh or preserved feces (see Table 88.6 ). The two methods for specimen concentration are sedimentation and flotation. ^, Of these, sedimentation methods are most commonly used in human clinical parasitology laboratories. In these methods, parasites settle to the bottom of the specimen via centrifugation, whereas with flotation methods, parasites float to the top of a solution with a high specific gravity. ^{, ,} Most concentration methods also incorporate filtration/straining and defatting steps to remove extraneous fecal material. The classical method for concentrating human fecal specimens in the diagnostic laboratory is the formalin-ether sedimentation method, which uses formalin as a preservative and fixative and ether as a defatting agent. ^, This procedure is still commonly used worldwide, although ethyl acetate is often used in place of ether, given the high flammability of the latter. Prior to concentration, a filtration step is generally performed to remove large particulate matter from the stool. Filtration may occur using a simple mesh screen or employing a specially designed commercial filtration and concentration device. The concentrated wet mount is examined in the same manner as the direct wet mount, using low and high magnifications, with or without a second preparation using iodine. Examination of the concentrate allows for identification of protozoa and helminth eggs and larvae but is suboptimal for detection of small protozoa such as Dientamoeba fragilis . Fecal leukocytes, erythrocytes, and Charcot-Leyden crystals (slender crystals formed during eosinophil breakdown) can also be identified ( Fig. 88.5 ). The presence of fecal leukocytes is suggestive of an inflammatory cause of the patient’s symptoms such as invasive bacterial or parasitic infections, or inflammatory bowel disease, while the presence of eosinophils/Charcot-Leyden crystals may suggest a parasitic etiology or other cause of eosinophilia.

The third component of a complete O&P examination is the permanently stained smear. This preparation is used primarily to identify protozoa since it provides greater sensitivity for these parasites than concentrate and direct preparations alone. ^, However, it is not recommended for identification of helminth eggs and larvae, since the stain may obscure the features of these larger objects. The stained smear is useful for providing a permanent record of the patient’s specimen and allows for sharing of slides among laboratories. ^{, ,} It is made by spreading feces onto a slide in such a way that thick and thin regions are created, and then stained with a method that allows for good protozoal nuclear and cytoplasmic differentiation. While historically the permanent stained smear was made from unconcentrated preserved stool, it is becoming more commonplace to make the smear of a concentrated specimen, especially when using single-vial spin-column technology. The iron-hematoxylin stain was traditionally used due to its superior definition of key cytoplasmic and nuclear characteristics, but it is technically challenging and has largely been replaced with the Wheatley modification of the trichrome stain in the United States ( Fig. 88.6 ). Proprietary versions of the trichrome stain may be recommended for specimens preserved in commercial single-vial preservatives, so it is important to check with the manufacturer when selecting a stain for routine use.

Depending on the laboratory, the available resources, and the primary purpose of the stool parasite examination, any or all three of these preparations may be used. Many parts of the world, including those in which the primary concern is detection of helminth ova, will only examine a wet mount preparation of unconcentrated or concentrated stool. However, many laboratories in the United States, Australia, Canada, and parts of Europe will examine a permanent smear in addition to the concentrated wet preparation to provide additional sensitivity for detection of protozoa. Of note, both a concentrated wet preparation and permanent smear must be examined by laboratories accredited through the College of American Pathologists, including those outside of the United States.

Finally, additional special stains may be performed for sensitive detection of the coccidia and the microsporidia. ^, Modified acid-fast stains (modified Kinyoun method, modified acid-fast dimethyl sulfoxide, auramine-O) or modified safranin stains are recommended for detection of coccidian oocysts in stool preparations. ^{, ,} Instructions for performing staining procedures are available elsewhere. ^{, ,}

Additional morphologic methods for detection of parasites in feces.

A number of other techniques exist for the morphologic detection of parasites in fecal specimens. ^{, ,} A method that is uncommonly used in the United States and Europe, but may be used in endemic settings, is egg enumeration for estimation of intestinal worm burden. This may be requested to determine the need for treatment and to monitor therapeutic efficacy. Methods for enumerating eggs include the Stoll dilution egg count, direct smear method of Beaver, and the Kato-Katz technique. ^{, ,}

Finally, a number of concentration and culture techniques are used to increase the sensitivity of S. stercoralis detection, including the Baermann concentration, Harada-Mori filter paper test-tube culture, and stool agar culture methods. ^{, ,} These methods also allow for recovery of other nematodes including those shed as eggs in stool. With the Baermann technique, a portion of feces is placed on gauze that is suspended on a wire or nylon sieve over the top of a funnel. The funnel is held upright in a ring stand and attached to a section of rubber hose, which is clamped shut. The funnel is then filled with water up to the level of the gauze, so that the feces just barely touches the water. This allows nematode larvae that are present in the fecal specimen to migrate through the gauze into the water, from where they settle to the bottom of the funnel via gravity. Water can be sampled from the hose after 2 or more hours and centrifuged so that the sediment can be examined for the presence of larvae. ^,

The Harada-Mori filter paper test-tube method is similar to the Baermann concentration technique in that fecal material is placed on permeable substance (filter paper) and placed in contact with nonchlorinated water. ^, This method can also be used to detect nematodes in soil samples. With this method, the filter paper strip is placed into a test tube and the tube is allowed to stand upright at room temperature (25 to 28 °C) for 10 days. Larvae that are present in the sample will migrate into the water and drift to the bottom of the tube. A small amount of water is withdrawn from the bottom of the tube daily for 10 days and examined for the presence of live larvae. Hookworm and Trichostrongylus eggs in the specimen may also hatch during this time frame and release detectable larvae. Although the Baermann and filter paper test-tube culture methods provide greater sensitivity for detection of S. stercoralis larvae than conventional O&P testing, they are time-consuming and potentially hazardous (due to the infectivity of the larvae) and therefore rarely used in the clinical laboratory.

The stool agar culture method provides a safer, simpler, and more sensitive alternative to the Baermann technique and is therefore better suited for routine clinical use. ^{, , ,} Using this method, approximately 2 g of feces is placed in the center of an agar plate and incubated at room temperature. Larvae that are present in the specimen will migrate from the feces into the agar and carry fecal bacteria with them. The bacteria grow in the tracks of the migrating larvae and produce visual evidence of their presence ( Fig. 88.7 ), thus allowing for easier detection. The plates should be incubated at room temperature for 2 or more days (agar side up) and examined daily for evidence of larval tracks leading away from the stool. Migrating larvae from positive agar culture plates should be examined microscopically to confirm the identity of the infecting nematode. Any type of nutrient agar that can sustain bacterial growth is suitable for the stool agar culture, but clear agars are easier to macroscopically examine for larval migration tracks. Despite the increased sensitivity provided by the stool agar culture, testing multiple specimens may still be required for detection of latent S. stercoralis infection. ^, As noted earlier, the filariform larvae of S. stercoralis and hookworms are infectious, and therefore testing should be conducted wearing gloves. For additional protection of laboratory staff, stool agar culture plates should be sealed with cellulose tape and placed in a clear plastic bag for daily visual inspection. Feces should not be refrigerated since this will kill the larvae and cause false-negative results. ^,

Perianal skin sampling for pinworm.

Unlike other intestinal nematodes, stool is not the preferred specimen for detection of pinworm ( Enterobius vermicularis ) since eggs are deposited directly onto the perianal skin folds rather than in feces. Therefore sampling of the perianal skin using the cellulose tape test is preferred for detecting this helminth. Taenia spp. eggs may also be detected since proglottids containing eggs may migrate out of the anus. With this method, clear (non-frosted) adhesive cellulose tape (e.g., Scotch tape) is pressed onto the perianal skin, and then placed onto a glass slide for microscopic examination for detection of pinworm eggs, and occasionally adult females. Commercial adhesive paddle devices such as the Swube (Becton Dickinson, Franklin Lakes, New Jersey) and the Pinworm Collector (Scientific Device Laboratory, Des Plaines, Illinois) provide an alternative sampling device that is easier to use by parents and caregivers ( Fig. 88.8 ). Sampling should be performed early in the morning before the patient bathes or defecates. Gloves should be worn during specimen collection and examination, since E. vermicularis and Taenia solium eggs may be infectious. A single negative exam does not rule our pinworm infection, and up to six exams may be required for optimal sensitivity. Although pinworm eggs and/or worms may occasionally be detected in fecal specimens, the O&P is considered a suboptimal method for diagnosing pinworm infections.

Duodenal aspirates and the string test.

Direct duodenal sampling may allow for enhanced detection of S. stercoralis, G. duodenalis, Cryptosporidium spp., and Cystoisospora belli, and may, less commonly, recover Clonorchis sinensis, Opisthorchis spp., and Fasciola hepatica . ^{, ,} Specimens may be collected by upper endoscopy or through use of the “string test” (Entero-test, gelatin-capsule method). ^, With the string test, the patient swallows a weighted gelatin capsule that contains a spool of nylon string. The string also protrudes from one end of the capsule and is taped to the patient’s cheek or neck. Once swallowed, the gelatin capsule dissolves, and the string unwinds and is carried to the duodenum by peristalsis. Organisms that are present in the duodenum will adhere to the string and will then be removed when the string is pulled out of the patient approximately 4 hours later. Forceps are used to remove any adherent material (e.g., intestinal mucus) from the string for subsequent microscopic examination. With both duodenal aspirates and the string test, the material is examined by wet or stained preparation for the presence of parasites, in a similar manner to routine stool examinations. ^,

Genitourinary specimens.

Urine, urethral and vaginal discharge, and prostatic secretions may be examined for Trichomonas vaginalis, a sexually transmitted protozoal parasite. ^{, ,} This was traditionally done in the physician’s office using a wet mount of the specimen in saline, and could also be used to identify yeasts and pseudohyphae (suggestive of candidiasis) and clue cells (bacterial vaginosis) in vaginal secretions. T. vaginalis trophozoites are identified by their characteristic morphology and “jerky” motility. Unfortunately, this is a relatively insensitive method for diagnosis of trichomoniasis and is less commonly used in the United States due to regulatory/proficiency requirements for physician-provided microscopy. Instead, nucleic acid amplification tests (NAATs) are recommended for detection of T. vaginalis infection in men and women. ^, If imidazole drug resistance is suspected, culture is required for antimicrobial susceptibility testing (see the “Parasite Culture Techniques” section later in this chapter). ^,

Urine may also be examined for Schistosoma haematobium eggs and, less commonly, microfilariae. ^{, , ,} For detection of S. haematobium, several urine specimens should be examined over a period of several days to maximize the likelihood of detection. The optimal time for collection is between 10:00 am and 2:00 pm, since this is when peak egg shedding is thought to occur. Use of membrane filtration (as described earlier for microfilariae) ( Fig. 88.9 ) or collection of a 24-hour urine specimen may increase detection.

Respiratory specimens.

A variety of helminthic and protozoal parasites may be detected in respiratory specimens such as sputa, bronchial aspirates, and bronchoalveolar lavage specimens. The parasites most commonly seen in these specimens include S. stercoralis larvae, Paragonimus spp. eggs, Echinococcus spp. protoscoleces and free hooklets, and E. histolytica trophozoites. Specimens are usually examined as a wet mount, but a permanently stained slide is also appropriate for detection of protozoans. The type of stain used depends on the suspected organisms. Cryptosporidium spp. oocysts may rarely be seen and are best detected using a modified acid-fast stain.

Cerebrospinal fluid.

Cerebrospinal fluid (CSF) can be examined as a wet mount or stained preparation for trypanosomes ( T. brucei ) , free-living amebae (FLA) ( Naegleria fowleri ), and rarely T. solium (cysticercosis larval form), S. stercoralis, Echinococcus granulosus, Angiostrongylus cantonensis (L4 larvae), and various microfilariae. ^, Specimens should be centrifuged, and the pellet used for examination. It is important not to refrigerate the specimen if it will also be used for culture of N. fowleri.

Tissue aspirates, abscesses, cysts, and biopsies.

A number of parasites may be found in tissue aspirates, abscesses, cysts, and biopsies, and the type of preparation and stains should be selected based on the clinical presentation and suspected parasite. ^{, ,} In addition to the wet mount preparation, Giemsa-stained aspirate smears and tissue “touch preps” (impression smears) are particularly useful for identification of the protozoa, including the hemoflagellates, T. brucei and Leishmania spp., and FLA. Leishmania spp. amastigotes are most commonly seen within macrophages in bone marrow, liver, lymph node, skin biopsies, and splenic aspirates. Culture of the aspirated or biopsied material should be performed to increase the sensitivity of detection and to determine the infecting species if required for treatment purposes; ^{, , ,} see the “Parasite Culture Techniques” section later in this chapter. T. brucei trypomastigotes can also be seen in lymph node aspirates, whereas E. histolytica causes abscesses in the liver, and less commonly, the lung, brain, and other organs, and the FLA ( N. fowleri, Acanthamoeba spp., Balamuthia mandrillaris ) are found primarily in brain tissue. All can be detected using wet mounts and Giemsa-stained preparations, although the trichrome stain is preferred for detection of E. histolytica. ^{, ,} It is important to note that E. histolytica trophozoites are often few in number and may be difficult to identify among necrotic host cells; therefore culture, antigen-detection methods, or PCR of aspirated or biopsied material may be useful. Culture or PCR is also recommended for enhanced detection of the FLA, N. fowleri , and Acanthamoeba spp.; see “Parasite Culture Techniques” later in this chapter. ^{, ,}

The helminths most commonly found in tissue aspirates, cysts, and biopsies are Echinococcus spp., T. solium (cysticercosis larval form), and S. stercoralis filariform larvae (in hyperinfection syndrome). These can usually be identified using wet mounts, but can also be highlighted with a variety of stains including Giemsa and trichrome.

Skin biopsies (e.g., bloodless skin snips) submitted for detection of Onchocerca volvulus or Mansonella streptocerca should be placed in 1 mL saline for incubation at room temperature for up to 4 hours. ^, The skin snip and the saline are then examined for detection of motile microfilariae.

Lastly, human muscle biopsies (or animal meat) sent for suspected trichinosis are generally examined by compressing the fresh specimen between two glass slides (“squash prep”) or using a trichinoscope that compresses the muscle in a similar manner. ^, Bladder and rectal biopsies may be examined in a similar fashion for detection of Schistosoma spp. eggs. Specimens can also be submitted for routine histologic identification of Schistosoma eggs or Trichinella larvae.

Skin scrapings for scabies diagnosis.

Skin scrapings are the preferred specimen for detection of Sarcoptes scabiei mites, eggs, and fecal pellets (scybala). Specimens should be obtained from the infected area using a scalpel blade. A drop of mineral oil should be applied to the blade and allowed to flow onto the region of skin being sampled since this will facilitate adherence of the mites to the blade. Because the mites live within the epidermis, the area should be vigorously scraped, such that flecks of blood are seen in the oil. The material is then transferred to a glass slide using a wooden applicator stick. Another drop of oil is added, and the material is mixed together. The slide can then either be coverslipped for immediate examination, or covered with another slide and rubber-banded together for transportation to the laboratory. ^, This method can also be used for the detection of Demodex mites. Gloves should be worn during collection and examination of skin scrapings due to the highly infectious nature of the S. scabiei mites.

Corneal scrapings.

Corneal scrapings are used for detecting Acanthamoeba spp. cysts and trophozoites. ^{, ,} Giemsa is sufficient for detection of both Acanthamoeba cysts and trophozoites, but Calcofluor white will detect the cysts only. Smears are generally insensitive for detection of amebic keratitis, and therefore culture and/or PCR should also be performed on corneal specimens. Swabs are suboptimal for collecting ocular material for testing.

Enzyme immunoassays.

EIAs are commonly used in the parasitology laboratory and generally have the benefit of established performance and utility for many other tests in the laboratories. They also usually demonstrate good sensitivity and specificity for detection of fecal protozoa when compared to the O&P examination. Most EIA methods require 2 to 3 hours to perform and can be read manually or using a spectrophotometer. EIAs commonly use microtiter plates containing immobilized monoclonal antibodies (capture antibodies) to the parasite antigens of interest. The test sample is mixed with conjugate antibodies (e.g., polyclonal antibodies against the parasite antigens) and added to the microtiter plate containing the monoclonal antibodies. If the antigen is present, an immune complex will form and remain bound when washed to remove unbound antibodies. A substrate is then added and, if antigen/antibody complexes are present, a color will develop as a result of an enzymatic reaction (e.g., peroxidase).

Lateral-flow immunochromatographic assays.

LFICAs are available for the detection of parasite antigens in stool, blood, and vaginal/cervical samples (see Table 88.7 ). In general, LFICAs are designed to be used as point-of-care tests and are relatively simple to perform. Results are usually available within a short time frame (15 to 30 minutes). It is important to note, however, that analytical sensitivity and specificity may be lower than reported for EIAs. The typical assay design consists of a nitrocellulose membrane test strip that has been coated with mouse or rabbit “capture” antibodies to the parasite antigen of interest on the test line portion of the strip. Another portion of the strip is coated with goat anti-mouse or anti-rabbit Ig to serve as a control for the test mechanics. To perform the test, straight or diluted samples are added to a loading chamber, where the specimen migrates to a conjugate pad containing immobilized antibodies. These antibodies have been conjugated with colloidal gold or colored latex particles and are specific to the targeted parasite antigen. If parasite antigen is present in the specimen, then antigen/antibody complexes are formed, and these migrate along the nitrocellulose test strip via capillary action until reaching the control and test lines. An immune complex will form if the sample is positive for the targeted antigen and is visible as a colored line or dot ( Fig. 88.10 ). A positive control line reaction should always be detected to confirm that the test was performed correctly.

LFICAs are available for Cryptosporidium spp., E. histolytica, E. histolytica/Entamoeba dispar, Giardia , Plasmodium spp., and W. bancrofti (see Table 88.7 ). The sensitivity and specificity of these assays vary and are highest in regions with a higher prevalence of the disease. Also, not all assays are capable of species-specific identification; for example, some Entamoeba spp. antigen tests detect both E. histolytica and E. dispar, and do not differentiate between them. Due to the limited sensitivity and specificity of some tests, additional laboratory testing may be recommended for confirmation of negative results. It should be noted that some of these (e.g., those for W. bancrofti ) are not FDA approved for routine clinical diagnosis in the United States.

Epidemiology.

Malaria, from the Italian “mal aria,” meaning “bad air,” is one of the most important infections worldwide. It is a disease of antiquity, and several mutations have evolved over the course of human evolution that provide some protective benefit against severe malaria. Among the most well known are sickle cell trait, hemoglobin C, hemoglobin E, hereditary spherocytosis, <alpha>-and <beta>-thalassemia, and glucose-6-phosphate dehydrogenase (G6PD) deficiency. See Chapter 78 for further information.

According to the World Health Organization (WHO) World Malaria Report 2015, 3.2 billion individuals are at risk for malaria ( Fig. 88.11 ), and millions do not have access to reliable services and medications to prevent and treat the disease. There were an estimated 228 million cases of malaria in 2018 (95% confidence interval: 206 to 258 million) and 405,000 deaths. Of the malaria-related deaths, 94% occurred in the WHO African Region, with Nigeria, the Democratic Republic of the Congo, the United Republic of Tanzania, Angola, Mozambique, and Niger accounting for the most deaths. An estimated 67% of all deaths worldwide (272,000) were in children younger than 5 years of age. Malaria is found primarily in tropical and subtropical regions of the world today, although P. vivax is also found in some temperate regions, and used to be widespread in the United States. P. falciparum and P. vivax are the two most widely encountered species worldwide, followed by P. malariae and P. ovale . A list of countries in which malaria is endemic can be found in the CDC Yellow Book, which is an important source of information for travelers and physicians that care for international travelers, as well as the CDC Red Pages. ^, In nonendemic countries, malaria is seen primarily in immigrants and travelers. The majority of imported malaria is detected in patients who have traveled to an endemic region to visit friends and relatives (VFRs), as these individuals do not usually seek medical advice prior to traveling, tend to visit more rural settings where there is an increased exposure to mosquito bites, and tend to stay for longer periods of time. In 2016, there were 2088 cases of imported malaria reported to the CDC, of whom 69.4% were VFRs.

Despite these sobering figures, there have been significant reductions in malaria incidence and mortality rates due to extensive global malaria campaigns. Between 2010 and 2018, incidence rates fell from 71 to 57 cases per 1000 population at risk. Furthermore, malaria has fallen from number 9 in all causes of cause-specific disability-adjusted life year (DALYs) in 2000 to number 12 in 2012, and is no longer in the top 20 causes as of 2016. Malaria was estimated to cause 37,369,000 DALYs in 2016. These successes have been due to extensive efforts in controlling the Anopheles mosquito vector, providing highly effective artemisinin-based combination therapies (ACTs), and providing intermittent preventive treatment during pregnancy.

Life cycle.

Human Plasmodium spp. have a complex life cycle involving sexual reproduction in an Anopheles mosquito vector and asexual reproduction in the human host ( Fig. 88.12 ). It is important to understand that an initial asymptomatic liver stage precedes erythrocytic infection by 8 to 40 days (depending on the species) and that dormant hypnozoite stages remain in the liver with P. vivax and P. ovale infection, which may reactivate weeks, months, or years later. Other, less common ways that humans acquire infection are transplacental spread (congenital infection) and through blood transfusion. ^, The asexual forms of infection are seen in humans, but rarely exflagellated microgametes and ookinetes (sexual life cycle stages) may be seen in human blood only if the blood specimen is allowed to sit at room temperature for a prolonged period of time prior to examination. ^,

Clinical disease.

Malaria can range from asymptomatic to severe, life-threatening disease ( Table 88.9 ). The classic symptom is the malaria fever paroxysm, which begins abruptly when parasite destruction of erythrocytes and subsequent release of parasite antigens into the peripheral blood results in host cytokine production. With long-standing infection, erythrocytes may begin to rupture in synchrony, resulting in the classic (but rarely seen) fever cycles (see Table 88.9 ). Signs and symptoms of severe disease include generalized convulsions, hypoglycemia, respiratory distress, renal failure, circulatory collapse, and unarousable coma. Nonimmune individuals, pregnant women (and their fetuses), and young children are at greatest risk for severe disease. P. falciparum is associated with the highest morbidity and mortality due in part to its ability to infect erythrocytes of all ages, leading to high levels of parasitemia, anemia, jaundice, and splenomegaly. ^, In comparison, other human Plasmodium spp. preferentially infect old ( P. malariae ) or young ( P. vivax and P. ovale ) erythrocytes, and cause significantly less destruction. Erythrocytes infected with P. falciparum also sequester in small blood vessels due to receptor-mediated cytoadherence to endothelial cells and other erythrocytes, resulting in accumulation of infected cells in the microvasculature. This phenomenon, called sequestration, is associated with local tissue hypoxia, cytokine release, and metabolic acidosis, particularly in the brain, kidney, and lung. ^, Sequestration also occurs in the placenta due to cytoadhesion of infected erythrocytes to syncytiotrophoblasts, resulting in poor fetal outcomes. Other Plasmodium spp. do not sequester, and cause significantly lower mortality, although rare fatal infections have been reported due to P. vivax and P. knowlesi infection. ^{, ,} Given the broad range of clinical symptoms associated with malaria, and the potential for life-threatening infections, malaria should be considered in all patients who travel to endemic areas and have unexplained fever.

A number of drugs are available for preventing and treating malaria, including ACTs. At this time, the only ACT that is readily available in the United States is artemether-lumefantrine (Coartem, Novartis, East Hanover, NJ), although intravenous (IV) artesunate is available through the CDC malaria hotline under an expanded-access investigational new drug (IND) protocol for patients in the United States with severe malaria and those who are unable to tolerate oral medications. Selection of therapy is complex due to the widespread resistance of P. falciparum, and to a lesser extent, P. vivax to chloroquine and other antimalarials. It is therefore important to know the causative species, travel history, degree of parasitemia, clinical status of the patient, and patient medications and drug allergies, since this will help direct treatment. Patients with P. vivax or P. ovale infection must be given primaquine, in addition to a primary therapeutic antimalarial, in order to eradicate the hepatic hypnozoite stage. ^, Primaquine can cause hemolytic anemia in patients with G6PD deficiency, and therefore screening for this disorder is recommended before initiating therapy (for further details, see Chapter 78 for a discussion on enzymes of the red blood cell).

In addition to antimalarial therapy, supportive care may be necessary in patients with respiratory or renal failure, lactic acidosis, and hypoxia. Exchange transfusion may be used for cases in which the parasite load exceeds 10% of circulating red blood cells. While this method is no longer recommended by the CDC, the American Society for Apheresis guidelines support its use for patients with severe malaria and high parasitemia.

Laboratory diagnosis.

Malaria is a potentially life-threatening disease, and testing for malaria should be performed immediately and be available 7 days a week on a 24-hour basis. ^{, ,} Examination of thick and thin blood films is the gold standard for diagnosing malaria but requires considerable expertise for accurate screening and interpretation. If this expertise or resources are not available, then laboratories should consider using a rapid malaria LFICA or transport the specimen immediately to a nearby laboratory. ^, In the past decade, malaria molecular tests (primarily PCR) have become available at reference and public health laboratories. While these methods usually offer equal or greater sensitivity and specificity than conventional blood films, they are rarely performed rapidly enough to be used for primary diagnosis. Finally, malaria serologic tests may be useful in some settings, primarily for epidemiologic studies, but play little to no role for detection of acute disease.

Testing a single blood specimen may be insufficient for ruling out a diagnosis of malaria. The Clinical and Laboratory Standards Institute (CLSI) recommends examining repeat blood films every 6 to 8 hours for up to 3 days before excluding the diagnosis of malaria. There are no similar guidelines for repeat testing with LFICAs or PCR. Of note, DNA and/or antigen may remain detectable for days to weeks after successful treatment, and therefore NAATs and LFICAs cannot be used as a “test of cure.”

Microscopy.

As mentioned earlier in the section on microscopic examination of blood specimens, microscopic examination of Giemsa-stained thick and thin blood films remains the gold standard for diagnosis of malaria. At least two thin and two thick blood films should be prepared as described earlier in this chapter. The thick film is up to 20 times more sensitive than the thin film for detection of Plasmodium parasites and is therefore the preferred method for screening (see Fig. 88.3 ). The reported detection threshold is approximately 10 to 50 parasites/μL blood, although this may be different in field conditions (e.g., 500 parasites/μL blood). The thin film, in contrast, is ideal for determining the species of the infecting Plasmodium spp., since the erythrocytes remain intact with this method (see Fig. 88.4 ). Species determination is made based on a number of morphologic features as shown in Figs. 88.3 , 88.4 , and 88.13–88.20 . Among the most important features are the size of the infected erythrocyte, presence or absence of cytoplasmic inclusions (when stain pH is 7.0 to 7.2), and the stages of parasites present ( Table 88.10 , Figs. 88.13–88.20 ). Importantly, all stages (early and late trophozoites, schizonts, and gametocytes) are generally seen in peripheral blood smears for all species except P. falciparum, in which only early trophozoites (ring forms) and gametocytes are usually seen. This is because late-stage asexual forms (late-stage trophozoites and schizonts) of P. falciparum are contained in erythrocytes that are sequestered in the microvasculature. While rare, the presence of P. falciparum late-stage asexual forms in the peripheral blood is indicative of severe, overwhelming infection. When examining blood films for malaria, it is important to also look for other blood parasites (trypanosomes, microfilariae), bacteria (intraleukocytic morulae of Ehrlichia spp./ Anaplasma phagocytophilum and relapsing fever Borrelia spirochetes), and yeasts (e.g., Histoplasma capsulatum ). The presence of extracellular bacteria and yeast could be indicators of infection or contaminants, the latter of which might require investigative actions. Mixed infections with multiple Plasmodium spp. may also be observed, with rates of co-infection varying widely by region and methods used for Plasmodium species detection. In malaria cases reported to the CDC with onset of illness in 2016, co-infection was noted in 1.1% of cases. Finally, it is essential to differentiate Plasmodium spp. from the similar-appearing Babesia spp. parasites ( Table 88.11 ).

TABLE 88.10

Comparison of Plasmodium Species Causing Human Malaria in Giemsa-Stained Thin Films

Modified from Fritsche TR, Pritt BS. Medical parasitology. McPherson RA, Pincus MR, eds. Henry’s clinical diagnosis and management by laboratory methods. 24th ed. Philadelphia: Elsevier; 2022.

Species a	Appearance of the Infected RBC	Intracellular Inclusions b	Degree of RBC Infectivity	Characteristic Features
Plasmodium falciparum	Normal size, multiple infections of RBCs common	Rarely Maurer clefts	High	Usually only rings and gametocytes seen. Delicate rings, <1/3 of RBC diameter, appliqué and “headphone” forms; “banana-shaped” gametocytes; delicate brown-black pigment, often in one mass. Mature schizonts (rarely seen, except in very heavy infections) have 8–24 merozoites. Percent parasitemia may exceed 2%.
Plasmodium malariae	Small to normal size	Rarely Ziemann stippling	Low	All stages seen; small coarse rings, ≤1/3 of RBC diameter, growing rings form basket, band and bird’s eye forms. Schizont with 6–12 merozoites, occasionally in a “rosette” configuration; round-oval gametocytes. Coarse brown pigment. Percent parasitemia <2%.
Plasmodium vivax	Enlarged size, multiple infections of RBCs not uncommon	Schüffner dots (stippling)	Low	All stages seen; ring forms >1/3 of RBC diameter, ameboid forms; schizonts with 12–24 merozoites; round-oval gametocytes. Delicate yellow-brown pigment. Percent parasitemia <2%.
Plasmodium ovale	Enlarged size, round-to-oval, occasionally fimbriated	Schüffner dots (stippling)	Low	All stages seen; ring forms >1/3 of RBC diameter, more compact than P. vivax; schizonts with 6–14 merozoites, round-oval gametocytes. Coarse brown pigment. Percent parasitemia <2%.

RBC, Red blood cell.

a Zoonotic species, such as Plasmodium knowlesi, are not included in this table. P. knowlesi is geographically restricted to parts of Southeast Asia.

b Inclusions are best visualized using a Giemsa stain with a pH of 7.0 to 7.2.

TABLE 88.11

Comparative Morphology of Plasmodium falciparum and Babesia Species

Morphologic Feature	Plasmodium falciparum	Babesia spp.
Size of infected RBC	Normal	Normal
Parasite stages in peripheral blood	Ring forms; less commonly “banana-shaped” gametocytes. High degree of RBC infectivity possible.	Ring forms only. High degree of RBC infectivity possible.
Ring form characteristics	Small size, <1/3 diameter of RBC, appliqué and “headphone” forms. Multiply infected RBCs common.	Small size, 1/3–1/6 diameter of RBC; irregular, spindle, and ameboid forms; single or multiple chromatin dots. Rarely, “Maltese cross” formation. Multiply infected RBCs common.
Presence of intracellular inclusions	Rarely Maurer clefts	None
Extracellular forms	Rare	Common
Pigment	Brown-black	None

RBC, Red blood cell.

Determining the percentage of infected erythrocytes can be performed using the thick or thin blood film, and is useful for predicting prognosis, determining treatment approaches, and monitoring response to therapy. Formulas are widely available for this purpose. ^{, ,}

A number of other microscopic methods have also been described for malaria detection, including stains for hemozoin and nucleic acid. One of the commonly used methods is the QBC Malaria Test (Drucker Diagnostics, Port Matilda, Pennsylvania), which uses the fluorescent acridine orange stain to highlight parasites in centrifuged blood. This and other fluorescent stain methods may decrease screening time and provide increased sensitivity but requires fluorescence microscopy and may be difficult to interpret due to nonspecific staining patterns.

Other methods.

Other laboratory methods are rarely used for diagnosis. Histopathology is not usually useful for detection of acute disease but may be important in autopsy studies. In tissues, evidence of malaria is seen primarily as hemozoin pigment (parasite-derived hemoglobin breakdown product) within erythrocytes in small blood vessels. “Ring hemorrhages” are commonly seen in cerebral malaria ( Fig. 88.21 ).

Some newer, promising laboratory techniques include detection of hemozoin-generated nanobubbles across intact skin and flow cytometry for malaria screening and diagnosis. ^,

Finally, the use of digital image analysis and AI continues to be investigated for detection of malaria parasites and hemozoin using whole blood and stained blood films. While these approaches currently lack sensitivity compared with conventional light microscopic analysis, they hold promise for simplifying malaria detection and may even be adaptable for use with a cell phone.

Epidemiology.

Unlike malaria, babesiosis is primarily a disease of temperate climates. It is zoonotic and in nature cycles between rodents and Ixodes ticks. Highly endemic regions in the United States are the northeastern and upper Midwestern states, as well as areas of the South-Central and Atlantic states, where B. microti is the predominant pathogen and ticks in the I. scapularis -complex are the tick vectors. Babesia duncani is responsible for a smaller number of cases in the Pacific Northwest and is transmitted by Ixodes pacificus, while B. divergens –like organisms (sometimes referred to as Babesia MO-1) have been reported from patients in the states of Missouri, Kentucky, and Washington. From 2011 to 2015, there were 7612 cases of babesiosis reported to the CDC from 27 states, with most cases coming from 7 states (Connecticut, Massachusetts, Minnesota, New Jersey, New York, Rhode Island, and Wisconsin). The number of annual cases mostly increased each year over this time period, with 1126 cases in 2011 and 2074 in 2015. Most patients were 60 years of age or older.

Substantially fewer cases of babesiosis occur in Europe, with approximately 39 published cases to date. ^{, ,} Of these, Babesia divergens is responsible for the majority of cases, while Babesia microti and B. venatorum cause a smaller number of cases. ^, Ixodes ricinus is the primary tick vector in Europe. Elsewhere, rare cases of human babesiosis due to unnamed species have been reported from Taiwan, Japan, South Korea, China, India, South Africa, Egypt, Mozambique, Mexico, and Colombia.

Life cycle.

As with malaria, babesiosis is acquired primarily through the bite of an infected vector ( Fig. 88.22 ). Unlike malaria, there is no liver stage, and parasites immediately invade and multiply in erythrocytes. Also, the trophozoite, merozoite, and gamete stages are morphologically similar, appearing as round, oval, and pleomorphic ring stages. In the United States, infections occur primarily during the spring, summer, and fall when the tick vectors are actively seeking blood meals. Less commonly, humans acquire infection transplacentally and through infected blood transfusion (whole blood or blood-derived platelets). ^, The FDA considers transfusion-transmitted babesiosis to be a relevant threat and now requires blood collection centers in specific high-risk states to test each donation using a licensed B. microti NAAT. There are two FDA-approved NAATs for blood-donor babesiosis screening, as well as an arrayed fluorescent immunoassay for detection of B. microti antibodies (AFIA). The FDA has also approved pathogen reduction devices, capable of reducing B. microti in plasma and platelet components.

Clinical disease.

Most patients with babesiosis due to B. microti are asymptomatic. When present, symptoms commonly include fever, headache, and myalgias. ^, Elderly, immunocompromised, and asplenic individuals are at risk for severe life-threatening disease; complications may include hemolytic anemia, hepatosplenomegaly, hematuria, jaundice, disseminated intravascular coagulation (DIC), respiratory failure, acute renal failure, congestive heart failure, coma, and death (see Table 88.5 ). ^, Most patients with B. divergens infection have been immunocompromised and presented with severe disease, and most patients with B. divergens –like infections in the United States have similar characteristics. Only a few cases of B. venatorum infection have been described to date, but disease appears to be mild with this agent. ^,

Babesiosis is generally treated with combination therapy using atovaquone and azithromycin or clindamycin and quinine. ^{, ,} Supportive care including vasopressors, mechanical ventilation, dialysis, and blood transfusions may be necessary. As with malaria, high levels of parasitemia (≥10%) should prompt consideration of exchange transfusion therapy. ^{, ,}

Laboratory diagnosis.

Testing for acute babesiosis is performed in a similar manner as for malaria, with collection and examination of thick and thin blood films. Due to the potentially life-threatening nature of babesiosis, particularly in immunocompromised and asplenic patients, testing should be performed on an emergent basis. Serology plays little to no role in the diagnosis of acute disease but may be useful for detecting cases of chronic infection, for screening blood donor units, and for epidemiologic studies. There are currently no FDA-approved antigen or molecular detection methods for clinical diagnosis, although PCR is offered by select reference and public health facilities. In vivo culture may also be performed in rare instances at public health laboratories such as the CDC. It is important to note that other infectious agents (e.g., Borrelia burgdorferi and B. mayonii, A. phagocytophilum, Borrelia miyamotoi, Powassan virus) may also be transmitted by Ixodes spp. ticks, and therefore testing for these agents may also be indicated.

Microscopy.

Blood films are examined in the same manner as described earlier for malaria, including calculation of percent parasitemia, although clinically relevant breakpoints may be different. ^, An important consideration is the differentiation of Babesia spp. parasites from the morphologically similar-appearing Plasmodium spp. (see Table 88.11 ), especially when a patient’s travel history is not available. In particular, P. falciparum shares several morphologic features with Babesia spp.: they are both present primarily as thin intraerythrocytic ring forms, may cause high parasitemia levels, and can produce multiply infected cells. However, Babesia parasites usually demonstrate greater pleomorphism in size and shape than P. falciparum , are more likely to have extracellular forms, and can exhibit the characteristic (although uncommon) tetrad (“Maltese cross”) form ( Fig. 88.23 ). Babesia spp. also do not produce hemozoin pigment, and they do not have morphologically distinct gametocytes like Plasmodium spp. ^{, , ,} Of note, it is not possible to differentiate the human-infecting Babesia spp. by morphology alone, but the causative agent can usually be inferred by the patient’s travel history.

Epidemiology.

T. brucei is found primarily in sub-Saharan Africa. T. brucei gambiense is endemic in 24 countries of west and central Africa, and is responsible for the majority (>97%) of human cases. In contrast, T. brucei rhodesiense is endemic in 13 countries in eastern and southeastern Africa and is considered to be primarily a zoonosis, causing only ~3% of reported human cases. Numbers of reported cases have dropped significantly due to the success of multiple control efforts; between 1999 and 2014, new cases of West African trypanosomiasis fell from 27,862 to 3679 (77%), and new cases of East African trypanosomiasis fell from 619 to 117 (71%). There were only 977 cases detected in 2018, which is a record low for this historic disease.

Life cycle.

The life cycle in humans is relatively simple. Infective (metacyclic) trypomastigotes are injected into skin when an infected tse tse fly takes a blood meal. These enter the circulatory system, transform into bloodstream trypomastigotes (see Fig. 88.24 ), and are carried throughout the body. They eventually reach the lymphatics, and later, the CSF. The motile trypomastigote is the only stage seen in humans (compared to T. cruzi, which also has a nonmotile amastigote stage), and these forms replicate in the blood and other body fluids through binary fission. The tse tse fly becomes infected when it ingests trypomastigotes with its blood meal from an infected host. The parasite then undergoes further replication to form procyclic trypomastigotes, which migrate from the gut and transform into epimastigotes. These multiply in the tse tse fly salivary gland and transform into metacyclic trypomastigotes, which can be injected into the next host.

Clinical disease.

HAT generally occurs in three stages, beginning with a trypanosomal chancre at the site of inoculation. ^, This is followed by the hemolymphatic stage during which trypomastigotes may be found in the blood and lymph nodes. During this time period, patients may present with fever, pruritus, and lymphadenopathy. In the third (late) stage of infection (meningoencephalitic stage), trypomastigotes enter the CSF and cause headache, mental status changes, somnolence, loss of consciousness, and ultimately, coma and death. ^, These three stages are most pronounced in West African trypanosomiasis, which has a more chronic course. Posterior cervical lymphadenopathy, referred to as Winterbottom sign, may be pronounced. In comparison, East African trypanosomiasis is an acute and rapidly progressive febrile illness, in which patients may die before CNS involvement is conspicuous. ^, The diagnosis is usually suspected based on the clinical presentation and travel/exposure history. Antitrypanosomal therapy is indicated for all cases of infection. The choice of therapy varies with both the infecting subspecies and the stage of disease. Pentamidine and suramin are used for treatment of T. b. gambiense and T. b. rhodesiense early-stage infection, respectively, while eflornithine and melarsoprol (an arsenic derivative) are used for late-stage (meningoencephalitic) infection with T. b. gambiense and T. b. rhodesiense, respectively. ^, Potential side effects and toxicities occur with all of these medications, but are most severe for melarsoprol, which causes a life-threatening encephalopathy in 5 to 10% of patients who receive this drug. ^, Death occurs in approximately one-half of the patients with melarsoprol-induced encephalopathy. In 2019, the WHO published new guidelines for the treatment of HAT, which includes the use of the new medication, fexinidazole, for T. b. gambiense infection.

Laboratory diagnosis.

The primary means of diagnosis is examination of peripheral blood and, in late-stage disease, CSF, for the presence of trypomastigotes. ^, Trypomastigotes can also be seen in aspirated chancre fluid, lymph nodes, and bone marrow. High total IgM levels are usually detected in the blood and CSF, while pleocytosis (50 to 500 mononuclear cells/μL) is found in the CSF. ^, In endemic regions, detection of antigen or antibodies is also commonly used, and provides increased sensitivity over microscopic examination. Molecular and culture methods are not widely available and are used primarily in research settings. ^,

Microscopy.

Extracellular flagellated trypomastigotes can be identified on routine thick and thin blood films, but are often present in low numbers, and therefore concentration techniques such as buffy coat preparations and mini–anion-exchange centrifugation are commonly used to increase diagnostic sensitivity. ^{, ,}

The two subspecies of T. brucei are morphologically indistinguishable. They are slender extracellular organisms ranging from 14 to 33 μm in length with a small kinetoplast at the posterior end and a centrally located nucleus ( Figs. 88.24 and 88.25 A). A single flagellum arises from the kinetoplast and runs along the length of the organism as an undulating membrane, leaving the body at the anterior end as a free flagellum. Travel history is usually sufficient for differentiating T. brucei from T. cruzi and T. rangeli since the latter organisms are only found in the Americas.

Epidemiology.

As many as 8 million individuals are estimated to be infected worldwide, with most infected individuals living in poverty in Latin America. ^, Common triatomine vector species belong to the genera Rhodnius, Triatoma, and Panstrongylus, and these are found throughout much of the Americas, including parts of the United States. The genera and species of triatomine vector vary from country to country and even among ecologic niches. While some triatomine bugs maintain infection in a sylvatic (i.e., wild) cycle involving animal reservoirs, others infest poorly constructed houses in rural areas and have adapted to a domiciliary life, feeding primarily on humans. ^,

Based on the estimated prevalence data in Latin America and immigration rates, the CDC estimates that 300,000 individuals with Chagas disease live in the United States. Most infected individuals in the United States are thought to have acquired infection in Latin America; however, autochthonous (locally acquired) cases have been reported from Texas and Louisiana, and up to 50% of triatomines in these regions were found to be infected with T. cruzi. It should be understood the risk for vector-borne human infection in the United States is low, due to the feeding habits of Nearctic triatomines (with a delay between feeding and defecation) and better housing and infrastructure in the United States.

Life cycle.

In comparison to T. brucei, the life cycle of T. cruzi is relatively complex. Infective (metacyclic) trypomastigotes are present in the triatomine bug feces, and these may be inoculated into the bite site or nearby mucosal membranes after the infected triatomine takes a blood meal ( Fig. 88.26 ). ^, This form of transmission differs from other vector-borne parasitic infections, in which the infective form is usually introduced into the host through the vector’s saliva. Less commonly, humans become infected with T. cruzi through ingestion of food or beverages contaminated with infected triatomine bug feces, as well as through receipt of an infected blood transfusion or organ transplant. Congenital infection is also possible. Once in the body, the parasite alternates between intracellular, nonmotile amastigotes and extracellular, motile trypomastigotes. ^,

Clinical disease.

There are two main stages of disease: acute and chronic. ^, The earliest signs of infection include local swelling and erythema at the infection site. This may include formation of a chagoma (subcutaneous swelling), and unilateral periorbital edema (Romaña sign) when the conjunctiva is the site of inoculation. Localized lymphadenopathy may also occur. ^, Acute systemic signs and symptoms may be seen 2 to 3 weeks after infection, but only occur in a small subset (~1%) of infected individuals. ^, Manifestations include high fevers, myalgias, diffuse rash, hepatosplenomegaly, lymphadenopathy, keratitis, subcutaneous edema of the extremities and face, acute myocarditis, and meningoencephalitis. The myocarditis is directly related to replication of amastigotes within the cardiac myocytes and the associated host response, resulting in conduction abnormalities, loss of contractility, and death. ^, Children younger than 5 years of age are most likely to have severe acute disease. The acute stage lasts for approximately 8 to 12 weeks, after which the parasitemia decreases to undetectable levels and patients enter the chronic stage of infection. ^, The chronic stage may be asymptomatic, with patients having normal physical examination, electrocardiogram, and radiographic findings. This is sometimes referred to as an indeterminate form of disease, and it can last for years to decades. It is important to note that patients can still transmit infection to biting triatomine bugs during this time, as well as to other humans through donated blood or organs. ^,

Approximately 20 to 30% of individuals with indeterminate disease eventually progress to chronic symptomatic Chagas disease and experience one or more of the three “mega” syndromes: megacardia (cardiomyopathy), megaesophagus, and megacolon. ^, The associated pathology is thought to be due to complex parasite, host, social, and environmental factors and cannot be tied solely to destruction by the replicating amastigotes. Cardiomegaly and cardiac conduction abnormalities are the most common manifestations of chronic Chagas disease. ^, Nifurtimox and benznidazole are the most commonly used therapies for acute Chagas disease, but they have little benefit in chronic disease. ^{, ,} Long-term supportive therapies may be necessary for patients with end-stage cardiac or gastrointestinal manifestations of Chagas disease, including dietary modifications and pacemaker use.

Laboratory diagnosis.

The recommended method for laboratory diagnosis is dependent upon the stage of infection. Blood film microscopy is most useful during the acute form of disease since trypomastigotes are present in high numbers and are detectable as early as 10 days after infection. Antigen and molecular-based detection methods may also be performed using peripheral blood. ^{, ,} The level of parasitemia decreases significantly within the first 90 days of infection, and trypomastigotes are rare or absent during the indeterminate and chronic stages, thus requiring the use of alternate diagnostic modalities. Antibody detection is most commonly used for detection of chronic infections, while NAAT is used for acute cases or cases of reactivation and while monitoring seropositive patients who have undergone recent organ transplantation. ^,

Other.

In endemic areas, xenodiagnosis is sometimes used to detect low-grade and chronic infections. With this method, trypanosome-free triatomine bugs are allowed to feed on patients under evaluation. The feces of the bugs are then examined monthly for 3 months for the presence of epimastigotes.

On occasion, tissue biopsy and histopathologic examination is used to identify amastigotes in infected tissues. Parasites are most commonly seen in the heart, and rarely in other tissues. Tissue sections generally show collections of amastigotes within “pseudocysts” in association with lymphoplasmacytic inflammation and fibrosis ( Fig. 88.27 A). The amastigotes of T. cruzi are morphologically indistinguishable from those of Leishmania spp., and clinical presentation, site of amastigotes, and travel history may aid in distinguishing the two diseases. Amastigotes of T. cruzi may also resemble small yeasts such as H. capsulatum. This can usually be accomplished by the type of infected cell/tissue, size of the collection of organisms, and stain reaction using fungal stains.

Epidemiology.

Leishmaniasis is considered an NTD by the WHO, with an estimated 700,000 to 1 million new cases (1 million CL cases, 300,000 VL cases) and 26,000 to 65,000 attributable deaths occurring annually. ^, There are more than 1 billion people at risk of infection worldwide. Most (95%) cases of CL occur in the Americas, the Middle East, the Mediterranean basin (including parts of Spain and Italy), and Central Asia, with greater than 95% of cases occurring in Afghanistan, Algeria, Brazil, Colombia, Rian, Iraq, and the Syrian Arab Republic. Endemic foci of CL are also found in Texas and Oklahoma. Causative species include the Old World species, Leishmania major, L. tropica, and L. aethiopica, and the New World species, L. mexicana, L. venezuelensis, L. amazonensis, and L. peruviana. Local geographic names for cutaneous disease include Oriental sore, Delhi boil, Baghdad boil, Aleppo evil, and chiclero ulcer. In comparison, most cases of VL occur in Brazil, East Africa, and Southeast Asia, with L. donovani and L. infantum (Old World) and L. chagasi (New World) being the primary etiologic agents. Of note, L. infantum and L. chagasi have many genetic similarities and are considered by many to be the same organism, or subspecies of the same organism. Ongoing control and elimination programs have made significant progress in reducing VL in India, Bangladesh, and Nepal. Finally, MCL is found only in select parts of the world. Classical MCL occurs in the Americas due to L. braziliensis and related species in the Viannia subgenus. Additionally, a mucosal form of disease is seen with L. aethiopica infection in Ethiopia, and rare cases of mucosal involvement with L. donovani have also been reported. Nearly 90% of cases of MCL occur in Brazil, Bolivia, Ethiopia, and Peru.

Life cycle.

Humans acquire Leishmania infection primarily through the bite of an infected phlebotomine sand fly ( Fig. 88.28 ). Like T. cruzi, the motile parasites become intracellular amastigotes shortly after inoculation; in this case, they are phagocytosed by macrophages and other mononuclear phagocytic cells and proceed to multiply within these cells by binary fission. Promastigotes are only seen in skin specimens if there is a delay in processing the tissue, and no circulating motile blood form is present.

Clinical disease.

Most infected patients are asymptomatic, with only a small fraction of patients developing clinical disease. ^, CL, the most common form of infection, manifests as skin lesions on exposed parts of the body. The primary skin lesion usually appears 2 to 8 weeks after the sand fly bite, and commonly takes the form of a firm, painless papule at the bite site. ^{, ,} Over time, the lesion generally progresses to an ulcer with an exudative necrotic center and raised border. The lesions usually heal over time, but satellite lesions may appear and merge with the initial ulcer, and sporotrichoid lymphatic spread can occur. Other subtle differences have been noted among diseases due to different Leishmania spp. CL is caused by members of the L. major, L. tropica, L. mexicana, L. braziliensis, L. guyanensis, and L. enrietii complexes. L. tropica may also cause VL, as was seen in military personnel who served in Operation Desert Storm. Leishmania aethiopica, an Old World species in the L. tropica complex, causes an aggressive form of CL, which may metastasize to other sites to cause diffuse CL, as well as mucosal and nodular cutaneous lesions resembling lepromatous leprosy. The forms of New World CL are very similar to those seen with Old World CL, with the important exception that some species are capable of metastasizing to the nasal, buccal, and pharyngeal mucosa and causing MCL, even after resolution of the initial cutaneous lesions. New World CL due to L. mexicana generally causes single ulcers that heal over a relatively short (i.e., 6-month) period. However, this species also involves the earlobe in up to 40% of cases and may cause destructive chronic lesions (chiclero ulcer). Cutaneous lesions due to L. braziliensis commonly take longer to heal than lesions due to other New World species and may be disseminated to the mucosa of the head and neck in approximately 1 to 3% of cases, resulting in MCL.

MCL, also known as espundia, may occur concomitantly with CL, or occur several months to years after the initial cutaneous infection. Mucosal involvement usually begins with edema, nasal stuffiness, and erythema and occasional bleeding. This presentation later progresses to erosion, ulceration, and eventually disfiguring destruction of the soft tissues and cartilage. Death is rare but can occur due to secondary bacterial infection or aspiration pneumonia.

VL is the most serious form of disease and may be fatal if untreated. Early infection is commonly subclinical but may progress as amastigotes within macrophages multiply in the liver, spleen, and bone marrow. In immunologically naïve patients, symptoms may be more acute, with high fever, weight loss, and diarrhea. The disease is called kala-azar in India due to the associated skin darkening that is commonly observed. Post–kala-azar dermal leishmaniasis (PKDL) is a special form of VL in which widespread cutaneous lesions develop 6 months to several years after treatment. The cutaneous lesions contain multiple parasites, and therefore individuals with PKDL are an important reservoir for disease.

VL may be contained, or even cured, by the cell-mediated immune response, but hosts with an increased risk for developing severe, symptomatic disease include young children, nonimmune individuals (e.g., travelers, soldiers), malnourished individuals, and patients with AIDS. Unfortunately, the response to treatment is generally poor in patients with concurrent AIDS, unless cell-mediated immunity can be restored.

The choice of therapy depends on multiple factors, including the type and severity of disease (CL, VL, MCL) and anatomic location, and must be individualized for the infected patient, ideally with expert consultation. ^, All patients with visceral and mucocutaneous infection should be treated. Furthermore, treatment is indicated for patients with CL acquired in Latin America, when the causative species is capable of causing MCL. In these cases, antimony-containing compounds including pentavalent antimonial (SbV; not licensed in the United States) and sodium stibogluconate (Pentostam, GlaxoSmithKline, Philadelphia, Pennsylvania) have been the mainstay of treatment, although liposomal amphotericin B deoxycholate and miltefosine are also FDA approved for VL treatment. Patients with CL due to species that do not cause MCL may be treated in a variety of manners. Options include doing nothing (waiting for self-healing), local therapy (cryotherapy, thermotherapy, intralesional administration of SbV, topical paromomycin), and systemic therapy (Sbv, amphotericin B deoxycholate, ketoconazole, fluconazole, miltefosine).

Laboratory diagnosis.

In many endemic areas, diagnosis is made primarily on clinical findings and epidemiologic factors. ^, However, the clinical differential diagnosis for leishmaniasis may be broad, and confirmation of infection is preferred whenever possible. The most common means of diagnosis is by visualizing amastigotes in aspirated or biopsied material. In CL, the most active lesion should be sampled, taking an aspirate or biopsy from the border of the ulcer. Giemsa is the most commonly used stain. When VL is suspected, blood (for buffy coat preparations), bone marrow, lymph node or splenic aspirates, and liver biopsies may be obtained. Splenic aspirate provides the highest rate of detection (>95%) but carries the risk for splenic laceration and is therefore not commonly performed in nonendemic settings. Antibody detection is commonly used for indirect detection of VL but reported sensitivities vary widely and it is not useful for CL or MCL (see below).

In addition to light microscopic examination and serology, culture and molecular testing may be performed in specialized centers. Culture offers increased sensitivity over microscopy alone, and provides ample material for isoenzyme and molecular studies. In one study, the sensitivity of hematoxylin and eosin (H&E)–stained histologic sections was only 14% for diagnosis of CL, while examination of imprints was 19% and culture of biopsies/aspirated material was 58%. Combining the methods yielded higher sensitivity (67%). Finally, molecular studies may be performed on culture material or directly on clinical specimens and offers the highest detection rates of all direct methods.

Epidemiology.

T. gondii has a global distribution and can infect humans, in addition to a wide array of wild and domestic animals. Approximately 25 to 30% of the world’s population is estimated to be infected, with more than 40 million infected children and adults in the United States alone. ^, According to the CDC, toxoplasmosis is the second leading cause of death from a food-borne illness in the United States. Seroprevalence rates vary significantly from country to country and among socioeconomic groups, and some countries such as El Salvador and France have rates as high as 85%. ^, In the United States, it is estimated that 11% of individuals ages 6 or older have been infected with T. gondii.

Life cycle.

Humans can become infected with T. gondii in a number of ways, and the predominant means of exposure may vary by country. The two most common means of acquiring infection are ingestion of meat containing infectious cysts, and infection of oocysts in food, water, and other environmental sources contaminated with cat feces ( Fig. 88.29 ). Transmission may also occur via receipt of an infected blood or organ donation and congenitally across the placenta.

Clinical disease.

Infection is entirely asymptomatic in the vast majority (~90%) of patients, while the remaining 10% experience a self-limited mononucleosis-like syndrome with fever, malaise, and lymphadenitis. ^, Following acute infection, the parasite becomes dormant and patients remain asymptomatic unless they become immunocompromised later in life. Types of immunocompromised states commonly associated with toxoplasmosis are AIDS (CD4+ T lymphocytes <100 cells/mm ³ ) and receipt of immunosuppressive therapies after organ transplant or for cancer therapy. ^, In solid organ transplant recipients, the highest risk for infection is in seronegative recipients from a seropositive donor (D+/R−).

The most common form of toxoplasmosis in the immunocompromised host is encephalitis (see Table 88.5 ). Affected patients commonly present with motor or sensory deficits, mental status changes, lethargy, fever, seizures, and chorea. Other forms of disease seen in immunocompromised individuals are pneumonitis, myocarditis, chorioretinitis, cystitis, hepatitis, and multiorgan failure. ^, Congenital infection may also result in severe disease in the neonate, with blindness, epilepsy, developmental delays, and fetal demise. The risk and severity of disease is unrelated to the presence or absence of maternal symptoms, but correlates strongly with the gestational age at the time of maternal infection. The incidence of transmission to the fetus is low (9%) when maternal infection occurs during the first trimester, but severe fetal toxoplasmosis occurs in 78% of cases. ^{, ,}

Conversely, the incidence of transmission to the fetus is highest when maternal infection occurs during the third trimester (59%), but 90% of neonates are asymptomatic at birth. Maternal infection acquired prior to pregnancy does not usually pose a risk to the fetus, but infection may reactivate in significantly immunocompromised mothers during gestation. Infected infants may be asymptomatic at birth, but commonly develop symptoms later in life. Therefore infected infants should be treated regardless of symptoms. ^{, ,}

The preferred treatment varies by the type and severity of infection and the immune status of the host. ^, Immunocompetent patients with mild illness do not generally require treatment, whereas immunocompromised patients with visceral disease are usually given pyrimethamine and sulfadiazine plus folinic acid (leucovorin). Long-term treatment and/or prophylaxis is recommended for as long as a host remains severely immunocompromised. ^, When possible, immune restoration is an important part of therapy. In pregnant women, spiramycin is the recommended treatment for infection during the early stages of pregnancy (first and early second trimesters), while pyrimethamine and sulfadiazine with leucovorin are given during the later stages (late second and third trimesters). ^{, ,} The prognosis depends on the type and extent of infection. Most congenital abnormalities are irreversible. ^,

Laboratory diagnosis.

Serology, NAATs, microscopy, and rarely, in vivo culture, are used for diagnosis of toxoplasmosis. The preferred approach is based primarily on the patient’s immune status. ^, Serology is widely available and used as a first-line test for immunocompetent individuals, including pregnant women and neonates, who are likely to mount a detectable immune response. Serologic testing also allows for differentiation of other clinically similar infections such as acute HIV and mononucleosis. However, there are multiple caveats of serologic testing for toxoplasmosis that must be considered, which are described in more detail in the section on Antibody Testing below. NAAT and/or microscopic examination of involved tissues are generally recommended for evaluation of profoundly immunocompromised patients, particularly in the setting of negative or inconclusive serologic test results. NAAT of amniotic fluid is also recommended for prenatal diagnosis of congenital toxoplasmosis. Finally, T. gondii culture isolation may be attempted at specialized reference laboratories in cases of suspected prenatal congenital infection by inoculating mice or cell culture with amniotic fluid. In vivo and in vitro cultures of CSF, fresh tissue, and other specimens may also be helpful in detecting cases of toxoplasmosis in immunocompromised patients.

Test interpretation can be challenging, particularly for T. gondii serology. Therefore consultation and testing at a T. gondii reference lab may be advised. In the United States, the CDC recommends consulting with the Dr. Jack S. Remington Laboratory for Specialty Diagnostics (formerly the Palo Alto Medical Foundation Toxoplasma Serology Laboratory; https://www.sutterhealth.org/pamf/services/lab-pathology/toxoplasma-serology-laboratory ) for additional testing options. Further information about the main testing modalities is presented below, and full information about testing in different patient populations is available through published reviews and guidelines. ^{, ,} The Geneva Foundation for Medical Education and Research maintains a list of country-specific toxoplasmosis guidelines, programs, and publications at https://www.gfmer.ch/Guidelines/Maternal_neonatal_infections/Toxoplasmosis.htm .

Microscopy.

T. gondii parasites may be seen as either dormant cysts or actively replicating tachyzoites ( Fig. 88.30 ). Tachyzoites are arc-shaped organisms measuring 4 to 5 μm in length that may be seen extracellularly or within infected cells. ^, Tissue cysts, on the other hand, range from 10 to 100 μm in diameter and contain hundreds of smaller bradyzoites, measuring 3 to 4 μm in length. The types of specimens that should be examined depend on the host and the clinical presentation. Parasites can be seen in amniotic fluid, CSF, blood, broncho-alveolar lavage (BAL) fluid, and numerous tissue types, although detection sensitivity depends on the parasite load, and a negative result may not reliably exclude infection. Giemsa-stained impression smears are preferred for demonstrating the parasite morphology, but organisms can also be seen using routine H&E and parasite-specific immunohistochemical stains. Identification of only cysts in the absence of an associated tissue inflammatory response may indicate latent rather than active infection.

Epidemiology.

The FLA have a worldwide distribution. Acanthamoeba spp. are the most prevalent and widespread of the FLA, having been found in water (fresh and brackish), chlorinated water, soil, sewage, air-conditioning units, heating systems, dental and medical equipment, and even in the nose and throats of healthy individuals. Several genotypes have been described (T1–T20), with species in the T4 genotype ( A. castellanii, A. lugdunensis, A. polyphaga, A. royreba, A. mauritaniensis, A. rhysodes, and A. triangularis ) being most commonly associated with human infection. As a result of the broad environmental distribution, humans are commonly exposed to Acanthamoeba spp., and this is supported by the approximate 80% seropositivity rate detected in otherwise healthy individuals. Less is known about the environmental niche of B. mandrillaris compared with Acanthamoeba; it is most commonly found in soil and less often in water sources. Most cases of B. mandrillaris infection in the United States have been reported from the Southwest, with Hispanic Americans having a relatively high incidence of infection. GAE due to Acanthamoeba spp. is primarily seen in immunocompromised individuals, whereas GAE due to B. mandrillaris may be seen in both immunocompromised and immunocompetent patients (see Table 88.5 ). ^, AK is seen primarily in contact lens wearers who improperly store, handle, or disinfect their lenses. Specific risk factors for infection include rinsing contact lenses with tap water, and wearing lenses while swimming, showering, or using a hot tub. ^, A conservative estimate of the incidence of disease in the United States is 1 to 2 cases per 1 million contact lens users.

N. fowleri is commonly found in warm freshwater sources, including lakes, rivers, geothermal springs, and untreated domestic water sources. ^, Historically, most cases in the United States occurred in southern states, but cases as far north as Minnesota have now been reported. Surveillance from the CDC detected 142 cases of PAM in the United States from 1937 to 2013, with 76% of the patients being male, and 83% being children (median 12 years, range 8 months to 66 years). Most cases are associated with recreational water use, but cases linked to nasal irrigation with inadequately treated domestic water have also been reported. ^,

Life cycle.

The life cycle of the FLA in humans is shown in Fig. 88.31 . N. fowleri enters the body through the olfactory mucosa and migrates along the olfactory nerve through the cribriform plate to the olfactory bulb into the adjacent meninges and cortex ( Fig. 88.32 A). From there, it spreads throughout the CNS, causing edema, necrosis, and ultimately brain herniation and death. In comparison, B. mandrillaris and Acanthamoeba enter the body through the respiratory tract or a break in the skin and disseminate hematogenously to the CNS. Acanthamoeba spp. can also enter the eye through contaminated contact lens solution or corneal trauma.

Clinical disease.

PAM usually begins with meningeal signs such as severe headache, stiff neck, fever, and vomiting and rapidly progresses over a period of several days to mental status changes, coma, and death. Several approaches have been used to treat patients with PAM, including administration of systemic and intrathecal amphotericin B, miconazole, rifampin, and miltefosine, and whole-body cooling (therapeutic hypothermia). Miltefosine was previously only available in the United States through the CDC for single-patient emergency treatment of FLA infections, but is now commercially available, as it has received FDA approval for treatment of cutaneous and muco CL. Despite aggressive interventions, fatality rates exceed 97%. ^,

In contrast to PAM, GAE is a subacute to chronic infection presenting with headache, mental status changes, focal neurologic deficits, and eventually death, if untreated. ^, Acanthamoeba spp. and B. mandrillaris also cause granulomatous skin lesions, which may precede CNS involvement and serve as the source of subsequent hematogenous dissemination. The incubation period is unknown, but probably spans weeks to months. Likewise, the clinical course can span days to months. No specific treatment is recommended for GAE, although a number of agents including amphotericin B, paromomycin, azoles, metronidazole, macrolides, sulfadiazine, and miltefosine have been used with limited success. ^, Skin lesions can be successfully treated if the organism has not yet spread to the CNS.

Patients with AK commonly present with unilateral severe eye pain (out of proportion to the clinical findings), erythema, blurred vision, foreign body sensation, photophobia, and excessive tearing. ^, Less commonly, disease is present in both eyes. Physician exam findings include paracentral stromal ring infiltrate and nonhealing corneal ulcers. Amebic keratitis is a potentially sight-threatening infection and needs to be aggressively treated. Topical agents such as chlorhexidine and polyhexamethylene biguanide are usually prescribed for prolonged periods (6 months to 1 year or more), but corneal transplantation may be required for severe or refractory cases. The presence of residual cysts can lead to relapsed disease.

Laboratory diagnosis.

The first step in diagnosis is to have a high index of suspicion based on the clinical findings. Unfortunately, diagnosis of PAM and GAE is challenging and commonly made at autopsy. Microscopy of CSF, brain tissue, corneal scrapings, and corneal biopsies, in conjunction with culture, are the most commonly used methods for diagnosis. More recently, laboratory-developed PCR tests have been implemented at specialized reference and public health laboratories for testing of these same specimen types and provide more rapid diagnosis than culture. Culture and/or PCR are highly recommended, as they are the most sensitive methods for detection of the FLA from clinical specimens. Serologic testing is not routinely available and not clinically useful due to the high seroprevalence of antibodies to the FLA.

Epidemiology.

E. histolytica infection is found worldwide, but is most commonly reported from resource-poor countries with poor sanitation, especially in the tropics and subtropics in Latin America, Africa, and Asia. Unfortunately, the accuracy of available prevalence data is dependent on the methods used for detection, with rates calculated using microscopy-based studies inflated due to morphologic similarities among E. histolytica and morphologically identical Entamoeba spp. ( E. dispar, E. moshkovskii, and E. bangladeshi ) (see Table 88.12 ). ^, E. dispar is the more common of these Entamoeba spp., and may be more than ten times more common than E. histolytica in endemic regions. Newer studies using methods capable of identifying and differentiating the various Entamoeba spp. have estimated that there are 28 million illnesses each year, with as many as 1470 associated deaths.

The actual prevalence rates vary significantly by geographic location. Studies from South Africa, Egypt, and Cote d’Ivoire using methods capable of specifically identifying E. histolytica show prevalence rates ranging from 0% to greater than 21% for asymptomatic carriage. In Northern India and China, seroprevalence and PCR data showed overall prevalence rates of approximately 11.1%, ^, while Northwestern Saudi Arabia reported rates of 16.5%. E. histolytica infection is much less common in developed countries. The reported prevalence rates range from 0.2 to 12.5%, with the higher rates being from outbreak settings. In these countries, cases of E. histolytica are more commonly detected in patients with travel history to developing countries, recent immigrants, and immunocompromised patients, particularly those with HIV infection (see Table 88.5 ). Data from the International GeoSentinel-associated clinics showed that most returned travelers diagnosed with E. histolytica infection from 2007 to 2011 had traveled to India, Indonesia, Mexico, and Thailand. Tourism was the most common reason cited for travel in this group.

Life cycle.

As with the other intestinal amebae, E. histolytica infection is acquired following ingestion of food or water contaminated with mature cysts ( Fig. 88.39 ). In the intestinal lumen, the cyst matures into a trophozoite through a process known as excystation. Colonized individuals may shed mature and immature cysts that can be detected in stool by microscopy. Noninvasive, asymptomatic infections occur when the trophozoites attach to the surface of the cells lining the lumen. Trophozoites may also be shed in stool, but unlike the cysts, are susceptible to environmental factors and therefore are not always detected using the O&P examination. Rarely, trophozoites can invade the intestinal epithelium and cause gastroenteritis. Extraintestinal disease occurs when invading trophozoites enter the portal bloodstream from the bowel and migrate to the liver (primarily) and other organs such as the lung and brain. ^{, ,}

Clinical disease.

Most individuals with confirmed E. histolytica infection are asymptomatic, and only a minority goes on to develop symptomatic disease. Longitudinal studies revealed that approximately 4 to 10% of infected patients develop symptomatic intestinal disease within 1 year of acquiring the infection if left untreated. ^, When present, symptoms range from self-limited diarrhea to amebic dysentery, with the latter causing profuse bloody diarrhea and abdominal pain. In severe cases, complications such as extensive ulceration ( Fig. 88.40 A), toxic megacolon, ameboma (formation of an inflammatory mass in the intestine), and intestinal perforation may ensue.

ALA is the most commonly reported extraintestinal form of amebiasis caused by E. histolytica, occurring in approximately 5% of patients with a previous history of intestinal amebiasis ( Fig. 88.40 B). ^, Patients with ALA commonly present with fever, right upper-quadrant abdominal pain, and acute liver tenderness. Diarrhea may be present, although stool O&P examination is negative for E. histolytica in most cases. ^, Laboratory findings include leukocytosis, increased alkaline phosphatase, and increased liver transaminases. Disease often progresses relatively quickly and can be fatal if untreated. One of the most important complications of ALA is rupture and extension into the peritoneum or across the diaphragm to the lung and heart. Although rare, hematogenous dissemination to other organs may also occur, and cerebral amebiasis must be differentiated from infection with the FLA. ^,

To prevent progression to symptomatic disease and contamination of the environment, asymptomatic carriers of E. histolytica are routinely treated with drugs that effectively concentrate in the intestinal lumen such as paromomycin or diloxanide furoate. ^, Amebic colitis and extraintestinal disease are treated with a course of a nitroimidazole drug such as metronidazole followed by a luminal drug as prescribed for asymptomatic carriage. Percutaneous drainage of liver abscesses is not generally recommended unless patients do not respond to therapy within 5 to 7 days and there is a risk for abscess rupture. Antibacterial therapy is warranted in cases of ALA with bacterial superinfection.

Laboratory diagnosis.

The most common methods of E. histolytica laboratory detection are microscopy, stool antigen detection, and more recently, NAATs. ^, Serology is most useful for detection of extraintestinal and invasive disease. Culture is not routinely used for clinical diagnosis but may be performed in specialized research and public health laboratories. ^,

Other.

Culture is no longer routinely used for clinical diagnosis and is now only available in limited research and public health settings. Histopathology is not generally used for primary diagnosis but may be useful for detecting infection in atypical cases. On intestinal biopsy, mucosal erosion and ulceration are commonly observed, and characteristic “flask”-shaped ulcers may be seen ( Fig. 88.41 A). ^{, ,} Trophozoites are best identified at the leading edge of the ulcer in the submucosa ( Fig. 88.41 B) and demonstrate morphologic features that are similar to those seen in stool preparations, including ingested erythrocytes in the cytoplasm. Trophozoites may also be seen at the edge of extraintestinal abscesses but are seldom detected in aspirated abscess fluid. Cysts are not seen in tissue.

Epidemiology.

Giardiasis is found in both resource-rich and resource-poor settings and causes epidemic and endemic patterns of infection. Infections are commonly endemic in tropical regions with poor sanitation, where infection is transmitted largely person-to-person and clinical immunity develops over time. Children are frequently infected and shed large numbers of organisms into their stool. In contrast, sporadic epidemics can occur in resource-rich settings with good sanitation and are usually transmitted through consumption of Giardia cysts in contaminated food, municipal water supplies, or untreated groundwater. There is no preexisting immunity in the population, and therefore symptomatic disease occurs in both children and adults. ^, Immunocompromised patients, including those with HIV/AIDS and selective IgA deficiency (sIgAD) disorder, are at risk for persistent and chronic infections.

Global epidemiology data show a prevalence rate as low as 0.4% in developed countries and as high as 30% in developing countries. Giardiasis is a nationally reportable disease in the United States, and therefore good data exist for its incidence. During the 2011 to 2012 reporting period, the rates of giardiasis were 6.4 and 5.8 per 100,000 populations in 2011 and 2012, respectively, with higher rates reported in the northwestern states compared to the southwestern states (4.6 to 4.8 versus 8.5 to 9.4 per 100,000 population). In Canada, an average rate of 19.6 per 100,000 population per year (1999–2002) has been reported. The rate of infections in North America peaks in the summer months, which reflects its association with ingestion of contaminated surface water by campers and hikers. Other populations with a high incidence of infection are travelers, children attending day care (and their guardians), and men who have sex with men. A number of large water-borne outbreaks have been associated with contaminated municipal water supplies.