Diagnostic Medical Parasitology. Lynne Shore Garcia
(single and budding cells and pseudohyphae) and human cells (macrophages, PMNs, and RBCs) can be identified. The following quantitation chart can be used for examination of permanent stained smears with the oil immersion lens (100× objective; total magnification, ×1,000).
Figure 3.18 Intestinal protozoa (stained with Wheatley trichrome stain). Rows, from top to bottom (left and right), show Iodamoeba bütschlii trophozoite and I. bütschlii cyst; Entamoeba histolytica trophozoite (note the ingested RBCs) and Entamoeba histolytica/E. dispar cyst; Giardia lamblia (G. duodenalis, G. intestinalis) trophozoite and G. lamblia cyst; Entamoeba coli trophozoite and E. coli cyst; and Endolimax nana trophozoite and E. nana cyst. doi:10.1128/9781555819002.ch3.f18
Figure 3.19 Intestinal protozoa (preserved with TOTAL-FIX, stained with Wheatley trichrome stain). From top to bottom and left to right. Top row: Giardia lamblia (G. duodenalis, G. intestinalis), 2 trophozoites, 2 cysts. Second row: Giardia lamblia (G. duodenalis, G. intestinalis), 3 trophozoites, 1 cyst. Third row: Endolimax nana, 3 trophozoites, 1 cyst. Note the nuclear variation in the three trophozoites (very common). Fourth row: Entamoeba coli, 3 trophozoites and 1 cyst. Fifth row: Dientamoeba fragilis (3 trophozoites) and Entamoeba hartmanni (1 trophozoite). doi:10.1128/9781555819002.ch3.f19
| Quantitation of parasites, cells, yeasts, and artifacts | |
| Quantity | No. per 10 oil immersion fields (×1,000) |
| Few Moderate Many | ≤2 3–9 ≥10 |
1. Report the organism and stage (do not use abbreviations).
Examples: Entamoeba histolytica/E. dispar trophozoites
Giardia lamblia (G. duodenalis, G. intestinalis) trophozoites
2. Quantitate the number of Blastocystis spp. organisms seen (rare, few, moderate, many). Do not quantitate other protozoa.
Example: Moderate Blastocystis spp.
3. Note and quantitate the presence of human cells.
Example: Moderate WBCs, many RBCs, few macrophages, rare Charcot-Leyden crystals
4. Report and quantitate yeast cells.
Example: Moderate budding yeast cells and few pseudohyphae
Note Because yeast can continue to grow if the stool is not immediately preserved, some laboratories do not report yeast, since the report can be misleading. They elect to call the physician and discuss the findings. Another option is to add a report comment indicating that reports of yeast (budding and/or pseudohyphae) might be misleading due to a lag time between stool passage and specimen fixation.
5. Save positive slides for future reference. Label prior to storage (name, patient number, organisms present). Most laboratories discard their negative permanent stained smears after several weeks (rotate storage boxes and discard when full).
Procedure Notes for the Trichrome Staining Method
1. The single most important step in the preparation of a well-stained fecal smear is good fixation. If this has not been done, the protozoa may be distorted or shrunk, may not be stained, or may exhibit an overall pink or red color with poor internal morphology.
2. Slides should always be drained between solutions. Touch the end of the slide to a paper towel for 2 s to remove excess fluid before proceeding to the next step. This will maintain the staining solutions for a longer period. The slide can also be touched against the staining dish to drain off excess fluid before moving on to the next dish.
3. Incomplete removal of mercuric chloride (Schaudinn’s fixative with a mercuric chloride base [with or without PVA]) may cause the smear to contain highly refractive crystals or granules, which may prevent finding or identifying any organisms present. Since the 70% ethanol-iodine solution removes the mercury complex, it should be changed at least weekly to maintain the strong-tea color. A few minutes are usually sufficient to keep the slides in the iodine-alcohol; too long a time in this solution may also adversely affect the staining of the organisms.
4. When using non-mercury-based fixatives, the iodine-alcohol step (used for the removal of mercury) and the subsequent alcohol rinse (removal of the iodine) can be eliminated from the procedure. The smears for staining can be prerinsed with 70% alcohol and then placed in the trichrome stain, or they can be placed directly into the trichrome stain as the first step in the staining protocol (Fig. 3.16).
5. Smears that are predominantly green may be due to the inadequate removal of iodine by the 70% ethanol (steps 4 and 5). Lengthening the time of these steps or more frequent changing of the 70% ethanol will help.
6. To restore weakened trichrome stain, remove the cap and allow the ethanol to evaporate (ethanol carried over on the staining rack from a previous dish). After a few hours, fresh stock stain may be added to restore lost volume. Older, more concentrated stain produces more intense colors and may require slightly longer destaining times (an extra dip). Smears from fixatives containing PVA usually require a slightly longer staining time due to the presence of the plastic PVA powder.
7. Although the trichrome stain is used essentially as a “progressive” stain (that is, no destaining is necessary), best results are obtained by using the stain “regressively” (destaining the smears briefly in acidified alcohol after the initial overstaining). Good differentiation is obtained by destaining for a very short time (two dips only, approximately 2 to 3 s); prolonged destaining results in poor differentiation.
8. It is essential to rinse the smears free of acid to prevent continued destaining. Since 90% alcohol will continue to leach trichrome stain from the smears, it is recommended that after the acid-alcohol is used, the slides be quickly rinsed in 100% alcohol and then dehydrated through two additional changes of 100% alcohol.
9. In the final stages of dehydration (steps 9 to 11), the 100% ethanol and the xylenes (or xylene substitute) should be kept as free from water as possible. Coplin jars must have tight-fitting caps to prevent both evaporation of reagents and absorption of moisture. If the xylene becomes cloudy after addition of slides from the 100% ethanol, return the slides to fresh 100% ethanol and replace the xylene with fresh stock.
10. If the smears peel or flake off, the specimen might have been inadequately dried on the slide (in the case of specimens containing PVA), the smear may have been too thick, or the slide may have been greasy (fingerprints). However, slides generally do not have to be cleaned with alcohol prior to use.
11. On examination, if the stain appears unsatisfactory and it is not possible to obtain another slide to stain, the slide may be restained. Place the slide in xylene to remove the coverslip, and reverse the dehydration steps, adding 50% ethanol as the last step. Destain the slide in 10% acetic acid for several hours, and then wash it thoroughly first in water and then in 50 and 70% ethanol. Place the slide in the trichrome stain for 8 min, and complete the staining procedure (3).
Procedure Limitations for the Trichrome Staining Method
1. The permanent stained smear is not recommended for staining helminth eggs or larvae; they are often too dark (excess stain retention) or distorted. However, they are occasionally recognized and identified. The wet smear preparation from the concentrate is the recommended approach for identification of helminth eggs and larvae.
2. The smear should be examined with the oil immersion lens (100×) for the identification of protozoa, human cells, Charcot-Leyden crystals, yeast cells, and artifact material. Quantitation of these cells and other structures is normally done from the examination of the permanent stained smear, not the wet smear preparations (direct wet smear or concentration wet smear).
3. This high-magnification (oil