DIAGNOSTIC METHODS

Direct Wet Examinations

What is the purpose of the direct wet examination?

This procedure is designed to allow the viewer to detect motile protozoa; this procedure should not be performed on preserved specimens and should be reserved for fresh stool specimens that are very soft or liquid.

How should the direct wet preparation be examined?

The entire coverslip preparation (22 x 22mm) should be examined under low magnification (X100); approximately 1/3 of the coverslip preparation should be examined under high dry magnification (X400).  It is not practical to examine this preparation using oil immersion magnification (X1000).  Saline and/or iodine mounts can be examined.

What do you expect to see during a wet preparation examination?

Helminth eggs and/or larvae can be seen, as well as some protozoan cysts, WBCs, some yeast, and fecal debris.  Many of the intestinal protozoa will need to be confirmed using the oil immersion magnification (X1000) for the permanent stained smear.

Concentrations

What is the recommended time and speed for centrifugation for the concentration method?  Why is this important?

The current recommendation is for every centrifugation step in the concentration method (sedimentation) to be performed for 10 min at 500 Xg.  If this recommendation is not followed, then small coccidian oocysts and microsporidial spores may not be recovered in the concentrate sediment.  The number of organisms is greatly increased over taking the sample from the unspun specimen.

What is the purpose of the concentration procedure?

The purpose of the concentration is to concentrate the parasites present, either through sedimentation or flotation.  The concentration is specifically designed to allow recovery of protozoan cysts, coccidian oocysts, microsporidian spores and helminth eggs and larvae.

Why is the flotation concentration used less frequently than the sedimentation concentration?

There are several reasons.  First, not all parasites will float; therefore, you need to examine both the surface film and the sediment before indicating the concentration examination is negative.  Second, the organisms must not be left in contact with the high specific gravity zinc sulfate for too long or protozoa will tend to become distorted, so the timing of the examination is more critical.  Also, the specific gravity of the fluid will need to be checked periodically.

What specific gravity zinc sulfate should be used for the flotation concentration procedure?

If the concentration is being performed on fresh stool, the specific gravity of the solution should be 1.18.  However, if the concentration is being performed on stool preserved in a formalin-based fixative, the specific gravity of the zinc sulfate should be 1.20.

How should the concentration wet preparation be examined?

Formalin-ethyl acetate sedimentation concentration is the most commonly used.  Zinc sulfate flotation will not detect operculated or heavy eggs; both the surface film and sediment will need to be examined before reporting a negative result.  Smears prepared from concentrated stool are normally examined as for the direct wet mount using the low power objective (10X) and the high dry power objective (40X); use of the oil immersion objective (100X) is not recommended (organism morphology not that clear).  The addition of too much iodine may obscure helminth eggs (will mimic debris).

What semi-automated methods are available to read the concentration sediments?

The traditional sampling approach using pipettes and the preparation of wet smears using glass slides and coverslips can be replaced with a semi-automated sampling and viewing system from DiaSys Corporation.  The specimen is drawn through tubing from the mixed concentrated stool sediment into two viewing chambers that fit onto the microscope stage.  The quality of the glass is excellent and organism morphology can be easily seen within the viewing chambers. Selection of such a system often depends on the laboratory work load.

Are there any tips for specimen processing for detection of the microsporidia?

The earlier studies on microsporidia were mainly performed at CDC.  When early comparisons were performed on methods, the authors used a slower centrifugation compared with using unspun material.  Using these methods, they felt using unspun material was better for recovery of microsporidial spores.  However, at UCLA when we looked carefully at this approach compared with the 500 Xg for 10 min (standard centrifugation time and speed), we found considerably more spores in the sediment.  If the stool contains a lot of mucus or is runny, just add formalin and centrifuge.  Don't bother with ethyl acetate since it may pull much of the material you want to examine up into the mucus layer.  If the stool is not particularly watery or doesn't contain a lot of mucus, you can treat it just like a regular concentration (but remembering to make every spin at 500 Xg for 10 min).  Also, the more you manipulate a stool, the more likely you will love some organisms (applies to all parasites in stool).  So, you may want to eliminate the wash steps and work with the first sediment you obtain from the first spin.

Remember to make the smears pretty thin - this will help you see the spores, but also remember not to decolorize too much.  Filtration is not a problem using any of the commercial concentration systems.  If you are using gauze, make sure you use woven gauze and use only two layers (do not use pressed gauze, which is too thick).

Comment on the use of formalin within the microbiology laboratory.

The formalin regulations were originally developed for industry (plywood, etc.) where great amounts of formalin are used in the manufacturing process.  The amount of formalin we are exposed to in the laboratory is very minimal; we've never heard of any microbiology laboratory (including a full-service parasitology service) even coming close to the limits.

"The Occupational Safety and Health Administration (OSHA) amended the original regulations for occupational exposure to formaldehyde in May of 1992 (1:Fed Regist 1992 May 27;57(102):22290-328).  The final amendments lower the permissible exposure level for formaldehyde from 1 ppm (part per million) as an 8-hour time-weighted average (TWA) to an 8-hour time-weighted average of 0.75 ppm.  The amendments also add medical removal protection provisions to supplement the existing medical surveillance requirements for those employees suffering significant eye, nose or throat irritation and for those suffering from dermal irritation or sensitization from occupational exposure to formaldehyde.  In addition, changes have been made to the standard’s hazard communication and employee training requirements.  These amendments establish specific hazard labeling for all forms of formaldehyde, including mixtures and solutions composed of 0.1% or greater of formaldehyde in excess of 0.1 ppm.  Additional hazard labeling, including a warning that formaldehyde presents a potential cancer hazard, is required where formaldehyde levels, under reasonably foreseeable conditions of use, may potentially exceed 0.5 ppm.  The final amendments also provide for annual training of all employees exposed to formaldehyde at levels of 0.1 ppm or higher."

Those laboratories that have been monitored have not come close to either measurement.  Once a laboratory has been measured and the results are on file, this information does not need to be generated again.  No badges are required.  Even without a fume hood (many labs do not use a fume hood), performing the routine formalin-ethyl acetate concentration does not seem to be a problem.  A number of people who have indicated they want to remove formalin from the laboratory probably don't really understand the history of the regulation or the actual issues.  The only possible problem seen in the clinical laboratory/pathology setting might be a routine anatomical pathology laboratory where very large amounts of formalin were used, and with sloppy use.  However, within the microbiology laboratory (even large laboratories), it does not seem to be a problem.

Permanent stains

What is the purpose of the permanent stained smear?

The purpose of the permanent stained smear is to provide contrasting colors for both the background debris and parasites present; it is designed to allow examination and recognition of detailed organism morphology under oil immersion examination (100X objective for a total magnification of X1000).  This examination is primarily designed to allow recovery and identification of the intestinal protozoa.

How long should the permanent stained smear be examined?

Rather than responding with a specific number of minutes, the recommendation is to examine at least 300 oil immersion (X1000 total magnification) fields; additional fields may be required if suspect organisms have been seen in the wet preparations from the concentrated specimen.

What recent changes have influenced the overall quality of the permanent stained smear?

The use of mercury substitutes in PVA generally leads to diminished overall morphology quality of the intestinal protozoa.  However, some of the mercury substitutes provide morphologic quality that is close to mercury and allows identification of most of the intestinal protozoa.  Differences in detection and identification are usually comparable unless there are very few organisms present; some organisms may be missed using mercury substitutes.

What is the purpose of the iodine dish in Wheatley’s trichrome stain protocol?

Mercury is removed from the smear when placed in the iodine dish; there is a chemical substitution of iodine for mercury.  The iodine is removed during the next two alcohol rinses; at the point the slide is placed in trichrome stain, neither mercury nor iodine is left on the smear.

Why don’t you need to use the iodine dish when staining fecal smears prepared from specimens preserved in the newer single vial systems (zinc sulfate based PVA)?

The zinc sulfate-based PVA is water soluble, so the dry smears can be placed directly into the trichrome dish without having to go through the iodine and subsequent rinse steps.  The zinc sulfate will be removed by the water in the trichrome stain.

Why might you have to use the iodine dish and subsequent rinses in your staining set up when staining slides from the proficiency testing agencies (AAB, various states, etc.)?

Smears for proficiency testing are prepared from fecal specimens that have been preserved in mercury-based fixatives, so the iodine dish and subsequent rinse steps are required to remove mercury and iodine prior to staining with either trichrome or iron hematoxylin stains.  For several years, CAP proficiency testing specimens have been preserved in non-mercury fixatives; thus the iodine dish is not required.

What role does the acetic acid play in the trichrome stain?

Both the trichrome and iron hematoxylin stains are considered regressive stains; the fecal smears are overstained and then destained.  The acetic acid in the 90% alcohol rinse step in the trichrome stain removes some of the stain and provides better contrast.  However, in some cases differences in the quality of staining between stained protozoa that have been subjected to the 90% alcohol rinse with and without the acetic acid may be difficult to detect.

What causes the xylene (or xylene substitute) dehydration solutions to turn cloudy when a slide from the previous alcohol dish is moved forward into the xylene dish?

If there is too much water carryover from the last alcohol dish, the xylene solution may turn cloudy.  When this occurs, replace the 100% alcohol dishes, back up the slide into 70% alcohol (you can also use a series of steps - 95%, then 70%), allow it to stand for 15 min and then move the slide forward through the 100% alcohol steps and xylene steps.

Why is absolute ethanol (100%) recommended as the best approach?

Although many laboratories use the commercially available 95/5% denatured alcohol mix, the dehydration of stained fecal smears will not be as good as that obtained with 100% ethanol.