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 immersion; total magnification, ×1,000) examination is recommended for protozoa, particularly for confirming species identification.
4. With low magnification (10× objective), one might see eggs or larvae; however, this is not recommended as a routine approach.
5. In addition to helminth eggs and larvae, C. belli oocysts are best seen in wet preparations (concentration wet smears prepared from formalin-preserved, not PVA-preserved, material).
6. Cryptosporidium and Cyclospora oocysts are generally not recognized on a trichrome-stained smear (modified acid-fast stains or the immunoassay reagent kits are recommended). Microsporidial spores do not stain sufficiently for recognition by the regular trichrome method; modified trichrome stains are required.
Trichrome Stain (Modified for Use with SAF-Preserved Fecal Specimens) (Dr. Norbert Ryan, VIDRL Modification)
It is generally recognized that stained fecal films are the single most productive means of stool examination for intestinal protozoa. The permanent stained smear facilitates detection and identification of cysts and trophozoites and affords a permanent record of the protozoa encountered. Small protozoa missed by direct smear and concentration techniques are often seen on the stained smear. Many reference texts mention that the combination of trichrome and SAF is unsuitable, because the green background stain of trichrome dominates, perhaps interacting with the albumin used to bind material to the slide. In this method the trichrome stain has been modified to reduce background staining. The modification is based on the theory that phosphotungstic acid functions both as a mordant and as a “colorless” stain of medium size. By increasing the content of this agent the intensity of background staining is reduced, without compromising the staining of structures with small pore size such as protozoa and tissue cells.
Note All other sections related to the trichrome stain presented above are relevant for the VIDRL modification of the trichrome stain for SAF-preserved fecal specimens presented here.
Trichrome Stain (VIDRL Modification for SAF-fixed smears)
1. Prepare the stain by adding 1.0 ml of acetic acid to the dry components. Allow the mixture to stand (ripen) for 15 to 30 min at room temperature.
2. Add 100 ml of distilled water. Properly prepared stain is purple.
3. Store in a glass or plastic bottle at room temperature. The shelf life is 24 months.
Procedure for Trichrome Stain (VIDRL Modification)
Note In all staining procedures for fecal and gastrointestinal tract specimens, the term “xylene” is used in the generic sense. Xylene substitutes are recommended for the safety of all personnel performing these procedures.
1. Place the slide in 70% ethanol for 5 min.*
2. Place it in a second container of 70% ethanol for 3 min.*
3. Place it in trichrome stain (VIDRL modification) for 10 min. The fecal smear no longer contains either mercuric chloride or iodine and is now ready for staining.
4. Place it in 90% ethanol plus 0.5% acetic acid for 1 to 3 s. Immediately drain the slide (see Procedure Notes), and proceed to the next step. Do not allow slides to remain in this solution. This is the destaining step.
5. Place the slide in 95% ethanol for 1 to 3 s. Use this step as a rinse.
6. Place it in two changes of 100% ethanol for 3 min each.* This is a dehydration step.
7. Place it in xylene or xylene substitute for 10 min.* This is a dehydration step.
8. Mount the slide with a coverslip (no. 1 thickness), using mounting medium (e.g., Permount).
9. Allow the smear to dry overnight or after 1 h at 37°C.
10.
