Diagnostic Medical Parasitology. Lynne Shore Garcia
of organisms. However, depending on the level of infection, examination of the fecal material as a direct smear may or may not reveal organisms. The direct wet smear is prepared by mixing a small amount of stool (about 2 mg) with a drop of 0.85% NaCl; this mixture provides a uniform suspension under a 22- by 22-mm coverslip. Some workers prefer a 1.5- by 3-in. (1 in. = 2.54 cm) slide for the wet preparations rather than the standard 1- by 3-in. slide, which is routinely used for the permanent stained smear. A 2-mg sample of stool forms a low cone on the end of a wooden applicator stick. If more material is used for the direct mount, the suspension is usually too thick for an accurate examination; any sample of less than 2 mg results in the examination of too thin a suspension, thus decreasing the chances of finding organisms. If present, blood or mucus should always be examined as a direct mount. The entire 22- by 22-mm coverslip should be systematically examined with the low-power objective (10×) and low light intensity (Fig. 3.1); any suspicious objects may then be examined with the high dry objective (40×). At least one-third to one-half of the coverslip should be examined under high dry power (total magnification, ×400), even if nothing suspicious has been seen. Use of an oil immersion objective (100×) on mounts of this kind is not routinely recommended unless the coverslip is sealed to the slide (a no. 1 thickness coverslip is recommended for oil immersion). For a temporary seal, a cotton-tipped applicator stick dipped in equal parts of heated paraffin and petroleum jelly should be used. Nail polish can also be used to seal the coverslip. Many workers think that the use of the oil immersion objective on this type of preparation is impractical, especially since morphological detail is more readily seen by oil immersion examination of the permanent stained smear. This is particularly true in a busy clinical laboratory situation.
Figure 3.1 Method of scanning direct wet film preparation with a 10× objective. Note that the entire coverslip preparation should be examined before indicating the examination is negative. (Illustration by Nobuko Kitamura.) Note: All methods contained in the figures can be found in reference 48. doi:10.1128/9781555819002.ch3.f1
The direct wet mount is used primarily to detect motile protozoan trophozoites. These organisms are very pale and transparent, two characteristics that require the use of low light intensity. Protozoan organisms in a saline preparation usually appear as refractile objects. If suspicious objects are seen on high dry power, at least 15 s should be allowed to detect motility of slowly moving protozoa. Application of heat by placing a hot penny on the edge of a slide may enhance the motility of trophic protozoa. Tapping on the coverslip can also stimulate the fluid to move; objects will roll over, thus providing a better view of the parasite or artifact. Helminth eggs and/or larvae, protozoan cysts, and coccidian oocysts may also be seen on the wet film, although these forms are more likely to be detected after fecal concentration procedures (Fig. 3.2).
Figure 3.2 Direct wet smear with saline. (Top row) Giardia lamblia (G. duodenalis, G. intestinalis) trophozoite (left), G. lamblia cyst (right); (second row) Entamoeba sp. (probably E. coli) (left), Blastocystis spp. central body form (right); (third row) Entamoeba hartmanni trophozoite (left), E. hartmanni cyst (right); (fourth row) Cystoisospora belli immature oocyst (left), Iodamoeba bütschlii cyst (right); (bottom row) Balantidium coli cyst (left), Chilomastix mesnili cyst (right). doi:10.1128/9781555819002.ch3.f2
After the wet preparation has been thoroughly checked for trophic amebae, a drop of iodine can be placed at the edge of the coverslip or a new wet mount can be prepared with iodine alone (Fig. 3.3). A weak iodine solution is recommended; too strong a solution may obscure the organisms. Several types of iodine are available; Lugol’s and D’Antoni’s are discussed here. Gram’s iodine, used in bacterial work, is not recommended for staining parasitic organisms.
Figure 3.3 Direct wet smear with saline and iodine. (Top) Entamoeba coli cyst with saline (left), E. coli cyst with iodine (note chromatoidal bars with sharp ends) (right); (next row) Trichuris trichiura egg in saline (left), T. trichiura egg with iodine added (right); (next row) Iodamoeba bütschlii cyst with saline (left), I. bütschlii cyst with iodine (right); (bottom row) Blastocystis spp. in saline (left), Blastocystis spp. in iodine (right). Note that more detail can be seen once the iodine is added to the wet mount. Also, when iodine is used, the glycogen vacuole stains dark (brownish gold to brown) in the Iodamoeba cysts and is clearly visible. doi:10.1128/9781555819002.ch3.f3
If preserved specimens are submitted to the laboratory, it is more cost-effective and clinically relevant to omit the direct smear and begin the stool examination with the concentration procedure, particularly since motile protozoa are not viable because of the prior addition of preservative. Even if parasites are seen on a direct mount of preserved stool, they would almost certainly be seen on the concentration examination as well as on the permanent stained smear (protozoa in particular). With few exceptions, intestinal protozoa should not be identified on the basis of a wet mount alone; permanent stained smears should be examined to confirm the specific identification of suspected organisms.
Saline (0.85% NaCl)
1. Dissolve the NaCl in distilled water in a flask or bottle, using a magnetic stirrer.
2. Distribute 10 ml into each of 10 screw-cap tubes.
3. Label as 0.85% NaCl with an expiration date of 1 year.
4. Sterilize by autoclaving at 121°C for 15 min.
5. When cool, store at 4°C.
D’Antoni’s Iodine
1. Using a magnetic stirrer, dissolve the potassium iodide and iodine crystals in distilled water in a flask or bottle.
2. The potassium iodide solution should be saturated with iodine, with some excess crystals left on the bottom of the bottle.
3. Store in a brown, glass-stoppered bottle at room temperature and in the dark.
4. This stock solution is ready for immediate use. Label as D’Antoni’s iodine with an expiration date of 1 year (the stock solution remains good as long as an excess of iodine crystals remains on the bottom of the bottle).
5. Aliquot some of the iodine into a brown dropper bottle. The working solution should have a strong-tea color and should be discarded when the color lightens (usually within 10 to 14 days).
Note The stock and working solution formulas are identical, but the stock solution is held in the dark and will retain the strong-tea color while the working solution will fade and have to be periodically replaced (Fig. 3.4).
Figure 3.4 Commercially prepared D’Antoni’s iodine; most commercial suppliers can provide this iodine solution. Do NOT USE Gram’s iodine for the parasitology procedures. doi:10.1128/9781555819002.ch3.f4
Lugol’s Iodine
1. Follow the directions listed above for D’Antoni’s iodine, including the expiration date of 1 year.
2. Dilute a portion 1:5 with distilled water for routine use (working solution).
3. Place this working solution into a brown dropper bottle. The working solution should have a strong-tea color and should be discarded when the color lightens