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.
70% Ethanol plus Iodine
1. Prepare a stock solution by adding iodine crystals to 70% alcohol until a dark solution is obtained (1 to 2 g/100 ml).
2. To use, dilute the stock solution with 70% alcohol until a dark reddish brown strong-tea color is obtained. As long as the color is acceptable, working solution does not have to be replaced. Replacement time depends on the number of smears stained and the size of the container (1 week to several weeks). This dish is required ONLY if the fecal specimen has been preserved using a mercuric chloride-based fixative; it is not required for staining SAF-preserved fecal specimens.
90% Ethanol, Acidified
70% Isopropyl or Ethyl Alcohol
100% Ethyl Alcohol (Recommended)
or 95%/5% Commercial Absolute Alcohol (Second Choice)
This is ethyl alcohol that has been denatured with isopropanol and methanol, but is considered commercial absolute alcohol and does not require a license to purchase (unlike absolute ethanol that has not been denatured). However, this product does not dehydrate as well as actual absolute ethanol.
Xylene or Xylene Substitute
Quality Control for Trichrome Stain
1. Stool samples used for QC can be fixed stool specimens known to contain protozoa or preserved negative stools to which buffy coat cells (PMNs or macrophages) have been added. A QC smear prepared from a positive fecal sample or a fixative sample containing buffy coat cells should be used when new stain is prepared or at least once each week. Cultured protozoa can also be used.
2. A QC slide should be included with a run of stained slides at least monthly; more frequent QC is recommended for those who may be unfamiliar with the method (14).
3. If the xylene becomes cloudy or there is an accumulation of water in the bottom of the staining dish containing xylene, discard the old reagents, clean the dishes, dry thoroughly, and replace with fresh 100% ethanol and xylene or xylene substitute.
4. All staining dishes should be covered to prevent evaporation of reagents (screw-cap Coplin jars or glass lids).
5. Depending on the volume of slides stained, staining solutions will have to be changed on an as-needed basis.
6. When the smear is thoroughly fixed and the staining procedure is performed correctly, the cytoplasm of protozoan trophozoites is blue-green, sometimes with a tinge of purple. Cysts tend to be slightly more purple. Nuclei and inclusions (chromatoidal bars, RBCs, bacteria, and Charcot-Leyden crystals) are red, sometimes tinged with purple. The background material usually stains green, providing a nice color contrast with the protozoa. This contrast is more distinct than that obtained with the hematoxylin stain, which tends to stain everything in shades of gray-blue to black.
7. If appropriate, the microscope should be calibrated (within the last 12 months), and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
8. Known positive microscope slides and photographs (reference books) should be available at the workstation.
9. Record all QC results; the laboratory should also have an action plan for “out-of-control” results.
Procedure for Trichrome Stain with Mercury-Based Fixatives (Fig. 3.16)
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.
Figure 3.16 Trichrome staining. Option 1, for use with smears prepared from fixatives containing mercuric chloride. The iodine is used to remove the mercuric chloride, and the subsequent two alcohol rinse steps remove the iodine. Thus, prior to staining, both the mercuric chloride and iodine have been removed from the smear. Options 2 and 3, for use with smears prepared from fixatives containing no mercuric chloride (the user can select option 2 or 3; there is minimal to no difference). doi:10.1128/9781555819002.ch3.f16
1. Prepare the slide for staining as described above.
2. Remove the slide from liquid Schaudinn’s fixative, and place it in 70% ethanol for 5 min.
3. Place the slide in 70% ethanol plus iodine for 1 min for fresh specimens or 5 to 10 min for PVA air-dried smears. The exposure to iodine will remove the mercuric chloride from the smear prior to staining with the actual trichrome dyes (substitution of iodine for mercury).
4. Place the slide in 70% ethanol for 5 min.* This and the next step in 70% ethanol will remove the iodine from the smear.
5. Place it in a second container of 70% ethanol for 3 min.*
6. Place it in trichrome stain for 10 min. The fecal smear no longer contains either mercuric chloride or iodine and is now ready for staining.
7. Place it in 90% ethanol plus acetic acid for 1 to 3 s. Immediately drain the rack (see Procedure Notes), and proceed to the next step. Do not allow slides to remain in this solution. This is the destaining step.
8. Dip the slide several times in 100% ethanol. Use this step as a rinse.
9. Place it in two changes of 100% ethanol for 3 min each.* This is a dehydration step.
10. Place it in xylene or xylene substitute for 5 to 10 min.* This is a dehydration step.
11. Place it in a second container of xylene or xylene substitute for 5 to 10 min.* This step completes the dehydration process (removal of all water on the smear).
12. Mount the slide with a coverslip (no. 1 thickness), using mounting medium (e.g., Permount).
13. Allow the smear to dry overnight or after 1 h at 37°C.
14. Examine the smear microscopically with the 100× objective. Examine at least 200 to 300 oil immersion fields before reporting a negative result (Fig. 3.17).
