of cyst material for the diagnosis of hydatid disease is a dangerous procedure and is normally performed only when open surgical techniques are used for cyst removal. Aspirated fluid usually contains hydatid sand (intact and degenerating protoscolices, hooklets, and calcareous corpuscles) (Fig. 6.8). Some older cysts contain material that resembles curded cottage cheese, and the hooklets may be very difficult to see. Some of this material can be diluted with saline or 10% KOH; usually, protoscolices or daughter cysts will have disintegrated. However, the diagnosis can be made by seeing the hooklets, which can be stained using the Ryan blue modified trichrome stain (see chapter 3) (Fig. 6.9).
Figure 6.8 Hydatid disease (Echinococcus granulosus). (Upper left) Hydatid cyst with protoscolices budding off from the germinal layer. (Upper right) Immature protoscolices, with the dark area being the hooklets. (Lower left) Higher magnification of the protoscolices taken from the hydatid cyst fluid. (Lower right) Hooklets from disintegrating protoscolices. Reprinted from reference 3. doi:10.1128/9781555819002.ch6.f8
Figure 6.9 Hydatid disease (Echinococcus granulosus). (Upper) Hydatid protoscolex revealed by Ryan modified trichrome stain. (Lower) Hydatid hooklets revealed by Ryan blue modified trichrome stain (reprinted from reference 30). doi:10.1128/9781555819002.ch6.f9
Examination of hydatid cyst material is carried out as follows.
1. If the cyst material is fluid, centrifuge at 500 × g for 3 min.
2. Carefully remove some of the sediment and prepare a wet mount.
3. Examine the material under low (100×) and high dry (430×) power (remember to use low light intensity, since some of the material may be very transparent).
4. If the cyst material is more viscous or solid, the material can be mixed with saline or 10% KOH and then centrifuged at 500× g for 3 min.
5. Some of the very viscous material can be placed on a glass slide (undiluted). Another glass slide can be placed on top of the first (material will now lie between the two slides). Rub the glass slides back and forth over each other and listen for a grating sound (like grains of sand being scratched). If this occurs, you may be hearing evidence of the presence of calcified hooklets. Place the material from the smears (may be diluted with saline) under the microscope to try to confirm the presence of hooklets (which may be extremely difficult to see with only low power).
6. A dried smear of the cyst aspirate can be stained using the Ryan blue modified trichrome stain (see chapter 3). The hooklets will stain reddish-purple in color; remember they are quite small, measuring 20 to 40 µm long.
Note Remember, the absence of protoscolices or hooklets does not rule out the possibility of hydatid disease, since some cysts are sterile and contain no protoscolices and/or daughter cysts. Review of the cyst wall from pathology (tissue sections) should be able to confirm the diagnosis.
Lymph Nodes, Spleen, Liver, Bone Marrow, Spinal Fluid, Eyes, and Nasopharynx
African Trypanosomiasis, Leishmaniasis, Chagas’ Disease, Primary Amebic Meningoencephalitis, Granulomatous Amebic Encephalitis, Amebic Keratitis, and Microsporidial Keratitis
Material from lymph nodes, spleen, liver, bone marrow, spinal fluid, eye specimens, or nasopharynx may be examined for the presence of parasites and should be processed as follows.
1. A portion of the fluid material can be examined under low (×100) and high dry (×400) power as a wet mount (diluted with saline) for the presence of motile organisms. Spinal fluid should not be diluted before examination.
2. Impression smears from tissues should be prepared and stained with Giemsa or another blood stain. The material is pressed between two slides, and a smear is formed when the slides are pulled apart (one across the other).
3. The smears are allowed to air dry and then processed like a thin blood film (fixed in absolute methanol and stained with Giemsa stain or one of the other blood stains).
4. Appropriate culture media should be inoculated with any remaining material (see chapter 8).
5. If microsporidia are suspected, modified trichrome stains (Ryan blue, Weber green) can be used; calcofluor white and immunoassay methods (currently under development) are also excellent options (36–43).
Primary amebic meningoencephalitis is rare, but the examination of spinal fluid may reveal the amebae, usually Naegleria fowleri (Fig. 6.10). Granulomatous amebic encephalitis is caused by Acanthamoeba spp., Balamuthia mandrillaris, or Sappinia diploidea. Uncentrifuged sedimented spinal fluid should be placed under a coverslip on a slide and observed for motile amebae; smears can also be stained with Wright’s, Giemsa, or one of the other blood stains (including the rapid stains). Spinal fluid, exudate, or tissue fragments can be examined by light microscopy or phase-contrast microscopy. Care must be taken not to confuse leukocytes with actual organisms and vice versa. The appearance of the spinal fluid may vary from cloudy to purulent (with or without red blood cells), with a cell count from a few hundred to over 20,000 white blood cells (primarily neutrophils) per ml. Failure to find bacteria in this type of spinal fluid should alert one to the possibility of primary amebic meningoencephalitis. These organisms can be isolated from tissues or soil with special media (see chapter 8).
Figure 6.10 Naegleria fowleri. Note the large karyosome and lobate pseudopodia. Spinal fluid should be examined on a slide, not in a counting chamber, where protozoan trophozoites could mimic white blood cells. (Armed Forces Institute of Pathology photograph.) doi:10.1128/9781555819002.ch6.f10 (Armed Forces Institute of Pathology photograph.)
Note When spinal fluid is placed in a counting chamber, any organisms that settle to the bottom of the chamber will tend to round up and look very much like white blood cells. For this reason, it is better to examine the spinal fluid on a slide directly under a coverslip, not in a counting chamber.
A rapid method for the diagnosis of Acanthamoeba or microsporidial keratitis involves the use of calcofluor white, which is a chemofluorescent dye with an affinity for the polysaccharide polymers of amebic cysts and microsporidial spores (Fig. 6.11 and 6.12). The following method has proven to be very successful for examination of corneal scrapings or biopsy material (42).
Figure 6.11 Acanthamoeba cyst. Note the fluorescence of the cyst wall. Trophozoites do not fluoresce when calcofluor white is used. doi:10.1128/9781555819002.ch6.f11
Figure 6.12 Microsporidial spores. (Upper left) Spores in a corneal scraping specimen; (upper right) spores in a fecal specimen (both images stained with silver stain).