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 (usually within 10 to 14 days).
Nair’s Buffered Methylene Blue Stain for Trophozoites (Direct Smear)
Although not commonly used, Nair’s buffered methylene blue stain is effective in showing nuclear detail in the trophozoite stages when used at a low pH; a pH range of 3.6 to 4.8 allows more active penetration of dye into the organism (15). After 5 to 10 min, the cytoplasm is stained a pale blue, with the nuclei being a darker blue; the slide should be examined within 30 min. Methylene blue (0.06% in an acetate buffer at pH 3.6) usually gives satisfactory results.
Acetate Buffer Solution Stock Solution A (0.2 M)
Acetate Buffer Solution Stock Solution B (0.2 M)
Mix the quantity of stock solutions A and B shown in the following table and dilute with distilled water to a total of 100 ml.
Quality Control for Direct Smear
1. Check the working iodine solution each time it is used or periodically (once a week). The iodine and Nair’s methylene blue solutions should be free of any signs of bacterial or fungal contamination.
2. The iodine should be the color of strong tea (discard if it is too light).
3. Protozoan cysts stained with iodine should contain yellow-gold cytoplasm, brown glycogen material, and paler refractile nuclei. The chromatoidal bodies may not be as clearly visible as they are in a saline mount. Human WBCs (buffy coat cells) mixed with negative stool can be used as a quality control (QC) specimen. These human cells, when mixed with negative stool, mimic protozoan parasites. The human cells stain with the same color as that seen in the protozoa.
4. Protozoan trophozoite cytoplasm should stain pale blue and the nuclei should stain a darker blue with the methylene blue stain. Human WBCs mixed with negative stool should stain the same colors as seen with the protozoa.
5. The microscope should be calibrated (within the last 12 months), and the original optics used for the calibration should be in place on the microscope when objects are measured. Some microbiologists feel that calibration is not required on a yearly basis; however, if the microscope receives heavy use, is in a position where it can be bumped, or does not receive routine
