Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 12
ОглавлениеIf the consistency of a stool specimen can be determined (formed, soft, or liquid), this information may give an indication of the organism stages that might be present. Trophozoites (potentially motile forms) of the intestinal protozoa are usually found in liquid specimens; both trophozoites and cysts might be found in a soft specimen; and the cyst forms are usually found in formed specimens. However, there are always exceptions to these general statements. Coccidian oocysts and microsporidian spores can be found in any type of fecal specimen; in the case of Cryptosporidium spp., the more liquid the stool, the more oocysts that are found in the specimen. Helminth eggs may be found in any type of specimen, although the chances of finding eggs in a liquid stool are reduced by the dilution factor. Tapeworm proglottids may be found on or beneath the stool on the bottom of the collection container. Adult pinworms and Ascaris lumbricoides are occasionally found on the surface or in the stool.
The presence of blood in or on the specimen may indicate several things and should always be reported. Dark stools may indicate bleeding high in the gastrointestinal tract, and fresh (bright red) blood most often is the result of bleeding at a lower level. In certain parasitic infections, blood and mucus may be present. Soft or liquid stool accompanied by blood is more suggestive of an amebic infection; these areas of blood and mucus should be carefully examined for the presence of trophic amebae. Occult blood in the stool may or may not be related to a parasitic infection and could result from a number of different conditions. Ingestion of various compounds may give a distinctive color to the stool (iron, black; barium, light tan to white).
Many laboratories prefer that stool specimens be submitted in some type of preservative. Rapid fixation of the specimen immediately after passage (by the patient) provides an advantage in terms of recovery and identification of intestinal protozoa. This advantage (preservation of organisms before distortion or disintegration) is thought to outweigh the limited motility information that might be gained by examining fresh specimens as direct wet mounts. Other laboratories still request a collection system that includes both a preserved specimen and the remainder of the fresh stool. Certainly cost is a factor, because several vials in the collection system cost more than a single vial containing preservative. Each laboratory will have to decide for itself, often basing the decision on the types of procedures ordered by the physicians who use the laboratory service, the test method selected (traditional methods, immunoassay detection kits, or molecular methods), and the lag time between specimen collection and submission to the laboratory.
With increased emphasis on continuous quality improvement, managed-care contracts, cost containment, and the clinical relevance of diagnostic test results generated, compliance with specimen acceptance or rejection criteria has become more important and a necessary part of overall quality performance. The generation of patient data begins with the quality of the specimen; anything that is done to compromise that quality should not be acceptable within the laboratory setting.
If the specimen has not been preserved immediately after passage, it is important to know the age of the specimen when it reaches the laboratory. Freshly passed specimens are necessary for the detection of trophic amebae, flagellates, and ciliates. Liquid specimens must be examined within 30 min of passage (not 30 min from the time the specimen reaches the laboratory or is clocked in by the computer). Soft specimens should be examined within 1 h of passage. Immediate examination of a formed specimen is not as critical; however, if the stool cannot be examined on the day of collection, portions of the specimen should be preserved. In a routine laboratory setting, these time frames are often neither practical nor possible. Thus, the routine use of stool preservatives for diagnostic parasitology is highly recommended and has been widely accepted.
Microscopic Examination (Ova and Parasite Examination)
The microscopic examination of the stool specimen, normally called the ova and parasite examination, consists of three separate techniques: the direct wet smear, the concentration, and the permanent stained smear. Each of these methods is designed for a particular purpose and forms an integral part of the total examination (1–22, 49). With increased emphasis on proper specimen collection and cost containment, the approach to the ova and parasite examination has changed somewhat during the last few years. Many laboratories are requesting that all fecal specimens be collected in preservatives prior to delivery to the laboratory to decrease the lag time between specimen passage and fixation, thus providing better organism morphology and subsequent identification.
Because preserved organisms do not exhibit motility, the direct wet smear is no longer considered a mandatory part of the routine ova and parasite examination. However, if fresh fecal specimens are delivered to the laboratory, the direct wet smear should be performed, particularly on liquid or very soft stools.
In addition to normal specimen debris, the microscopic examination of fecal material may reveal the following:
1. Trophozoites and cysts of intestinal protozoa
2. Oocysts of coccidia and spores of microsporidia
3. Helminth eggs and larvae
4. Red blood cells (RBCs), which may indicate ulceration or other hemorrhagic problems
5. White blood cells (WBCs), specifically polymorphonuclear leukocytes (PMNs), which may indicate inflammation
6. Eosinophils, which usually indicate the presence of an immune response (which may or may not be related to a parasitic infection)
7. Macrophages, which may be present in bacterial or parasitic infections
8. Charcot-Leyden crystals, which may be found when disintegrating eosinophils are present (and may or may not be related to a parasitic infection)
9. Fungi (Candida spp.) and other yeasts and yeastlike fungi
10. Plant cells, pollen grains, or fungal spores, which may mimic some helminth eggs, protozoan cysts, coccidian oocysts, or microsporidial spores
11. Plant fibers or root or animal hairs, which may mimic helminth larvae
Normal mixing in the intestinal tract usually ensures an even distribution of organisms. However, depending on the level of infection, examination of the fecal material as a direct smear may or may not reveal organisms. The direct wet smear is prepared by mixing a small amount of stool (about 2 mg) with a drop of 0.85% NaCl; this mixture provides a uniform suspension under a 22- by 22-mm coverslip. Some workers prefer a 1.5- by 3-in. (1 in. = 2.54 cm) slide for the wet preparations rather than the standard 1- by 3-in. slide, which is routinely used for the permanent stained smear. A 2-mg sample of stool forms a low cone on the end of a wooden applicator stick. If more material is used for the direct mount, the suspension is usually too thick for an accurate examination; any sample of less than 2 mg results in the examination of too thin a suspension, thus decreasing the chances of finding organisms. If present, blood or mucus should always be examined as a direct mount. The entire 22- by 22-mm coverslip should be systematically examined with the low-power objective (10×) and low light intensity (Fig. 3.1); any suspicious objects may then be examined with the high dry objective (40×). At least one-third to one-half of the coverslip should be examined under high dry power (total magnification, ×400), even if nothing suspicious has been seen. Use of an oil immersion objective (100×) on mounts of this kind is not routinely recommended unless the coverslip is sealed to the slide (a no. 1 thickness coverslip is recommended for oil immersion). For a temporary seal, a cotton-tipped applicator stick dipped in equal parts of heated paraffin and petroleum jelly should be used. Nail polish can also be used to seal the coverslip. Many workers think that the use of the oil immersion objective on this type of preparation is impractical, especially since morphological detail is more readily seen by oil immersion examination of the permanent stained smear. This is particularly true in a busy clinical laboratory situation.
Figure 3.1 Method of scanning direct wet film preparation with a 10× objective. Note that the entire coverslip preparation should be examined before indicating the examination is negative. (Illustration by Nobuko Kitamura.) Note: All methods contained in the figures can be found in reference 48. doi:10.1128/9781555819002.ch3.f1
The direct wet mount is used primarily to detect motile protozoan trophozoites. These organisms are very pale and transparent, two characteristics that require the use of low light intensity. Protozoan organisms in a saline preparation usually appear as refractile objects. If suspicious objects are seen on high dry power, at least 15 s should be allowed to detect motility of slowly moving protozoa. Application of heat by placing a hot penny on the edge of a slide may enhance the motility of trophic protozoa. Tapping on the coverslip can also stimulate the fluid to move; objects will roll over, thus providing a better view of the parasite or artifact. Helminth eggs and/or larvae, protozoan cysts, and coccidian oocysts may also be seen on the wet film, although these forms are more likely to be detected after fecal concentration procedures (Fig. 3.2).
Figure 3.2 Direct wet smear with saline. (Top row) Giardia lamblia (G. duodenalis, G. intestinalis) trophozoite (left), 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 maintenance, yearly calibration is recommended. The calibration factors for all objectives should be posted on the microscope or close by for easy access.
6. All QC results should be appropriately recorded; the laboratory should also have an action plan for “out-of-control” results.
Procedure for Direct Wet Smear
1. Place 1 drop of 0.85% NaCl on the left side of the slide and 1 drop of iodine (working solution) on the right side of the slide. If preferred, two slides can be used instead of one. One drop of Nair’s methylene blue can also be placed on a separate slide, although this technique is much less common.
2. Take a small amount of fecal specimen (the amount picked up on the end of an applicator stick when introduced into the specimen), and thoroughly emulsify the stool in the saline and iodine preparations (use separate sticks for each).
3. Place a 22-mm coverslip (no. 1) on each suspension.
4. Systematically scan both suspensions with the 10× objective. The entire coverslip area should be examined under low power (total magnification, ×100).
5. If something suspicious is seen, the 40× objective can be used for more detailed study. At least one-third to one-half of the coverslip should be examined under high dry power (total magnification, ×400), even if nothing suspicious has been seen.
6. Another approach is to prepare and examine the saline mount and then add iodine at the side of the coverslip. The iodine will diffuse into the stool-saline mixture, providing some stain for a second examination. Remember, the iodine will kill any organisms present; thus, no motility will be seen after the iodine is added to the preparation. The use of iodine is not mandated, but is based on personal preference.
Results and Patient Reports from Direct Wet Smear
Protozoan trophozoites and/or cysts and helminth eggs and larvae may be seen and identified. In a heavy infection with Cryptosporidium spp., Cyclospora cayetanensis, or Cystoisospora (Isospora) belli, oocysts may be seen in a direct smear; however, some type of modified acid-fast stain or fecal immunoassay is normally used to detect Cryptosporidium spp., particularly when few oocysts are present. Cyclospora oocysts are often confirmed using autofluorescence or the modified acid-fast stain. Spores of the microsporidia are too small, and the shape resembles other debris within the stool; therefore, they are not readily visible in a direct smear.
1. Motile trophozoites and protozoan cysts may or may not be identified to the species level (depending on the clarity of the morphology) and should be confirmed using the permanent stained smear.
Examples: Giardia lamblia (G. duodenalis, G. intestinalis) trophozoites
Entamoeba coli cysts
2. Helminth eggs and/or larvae may be identified.
Examples: Ascaris lumbricoides eggs
Strongyloides stercoralis larvae
3. Cystoisospora (Isospora) belli oocysts may be identified; however, Cyclospora and Cryptosporidium oocysts are generally too small to be recognized or identified without subsequent immunoassays or modified acid-fast staining.
Example: Cystoisospora (Isospora) belli oocysts
4. Artifacts and/or other structures may also be seen and reported as follows.
Note These crystals and cells are quantitated; however, the quantity is usually assessed when the permanent stained smear is examined under oil immersion.
Examples: Moderate Charcot-Leyden crystals
Few RBCs
Moderate PMNs
Procedure Notes for Direct Wet Smear
1. In preserved specimens, the formalin, sodium acetate-acetic acid-formalin (SAF), or Universal Fixative replaces the saline and can be used as a direct smear; however, no organism motility will be visible (organisms are killed by the fixatives). Consequently, the direct wet smear is usually not performed when the specimen (already preserved) arrives in the laboratory. The technical time is better spent performing the concentration and permanent stained smear. This approach is recommended for specimens submitted to the laboratory in preservative (2).
2. As mentioned above, some workers prefer to make the saline and iodine mounts on separate slides and on 2- by 3-in. slides. Often, there is less chance of getting fluids on the microscope stage if separate slides (less total fluid on the slide and under the coverslip) or larger slides are used. Selection of slide size and the use of iodine depend on the personal preference of laboratory personnel.
3. The microscope light should be reduced for low-power observations, since most organisms are overlooked with bright light due to limited contrast of the internal morphology. This is particularly true when the preparation is being examined without the use of iodine. Illumination should be regulated so that some of the cellular elements in the feces show refraction (lower the condenser). Most protozoan cysts and some coccidian oocysts are refractile under these light conditions.
Procedure Limitations for Direct Wet Smear
1. As mentioned above, because motility is lost when specimens are placed in preservatives, many laboratories are no longer performing the direct wet smear (the primary purpose is to see motility) but are proceeding directly to the concentration and permanent stained smear procedures as a better, more cost-effective use of personnel time, as well as a more clinically relevant approach.
2. Most of the time, results obtained from wet smear examinations should be confirmed by permanent stained smears. Some protozoa are very small and difficult to identify to the species level using just the direct wet smear technique. Confirmation is particularly important for Entamoeba histolytica/E. dispar versus Entamoeba coli. Findings from the direct wet smear examination can be reported as “preliminary, based on the direct wet mount examination only,” and the final report can be submitted after the concentration and permanent stain procedures are completed. However, if the laboratory turnaround time is less than 24 h, there is no need to send out a preliminary report; the final report can be submitted once the complete ova and parasite examination has been performed.
Concentration (Sedimentation and Flotation)
Fecal concentration has become a routine procedure as a part of the complete ova and parasite examination for parasites; it allows the detection of small numbers of organisms that may be missed by using only a direct wet smear (11, 21). There are two types of concentration procedures, sedimentation and flotation, both of which are designed to separate protozoan organisms and helminth eggs and larvae from fecal debris by centrifugation and/or differences in specific gravity (Fig. 3.5) (2, 4).
Figure 3.5 Fecal concentration procedures: various layers seen in tubes after centrifugation. (A) Formalin-ether (or ethyl acetate). The sediment should be well mixed, and a drop of sediment should be examined using the 10× low-power objective and the 40× high dry power objective. (B) Zinc sulfate (the surface film should be within 2 to 3 mm of the tube rim). Material from both the surface film and the sediment must be examined before the specimen is indicated as negative. The amount of sediment should not be excessive in either the sedimentation or flotation procedure. Heavy or operculated helminth eggs do not float. (Illustration by Sharon Belkin.) doi:10.1128/9781555819002.ch3.f5
Sedimentation methods (by centrifugation) lead to the recovery of all protozoa, oocysts, eggs, and larvae present; however, the concentration sediment that will be examined contains more debris. Although some workers recommend using both flotation and sedimentation procedures for every stool specimen submitted for examination, this approach is impractical for most laboratories. If one technique is selected for routine use, the sedimentation procedure is recommended as being the easiest to perform and the least subject to technical error (Fig. 3.6).
Figure 3.6 Sedimentation concentration. (Left) Unfertilized Ascaris lumbricoides egg. (Right) Hymenolepis diminuta egg. doi:10.1128/9781555819002.ch3.f6
A flotation procedure permits the separation of protozoan cysts, coccidian oocysts, and certain helminth eggs and larvae through the use of a liquid with a high specific gravity. The parasitic elements are recovered in the surface film, and the debris remains in the bottom of the tube. This technique yields a cleaner preparation than does the sedimentation procedure; however, some helminth eggs (operculated eggs and/or very dense eggs such as unfertilized Ascaris eggs) do not concentrate well in the flotation method (Fig. 3.7). The specific gravity may be increased, although this may produce more distortion in the eggs and protozoa. Laboratories that use only flotation procedures may fail to recover all of the parasites present; to ensure detection of all organisms in the sample, both the surface film and the sediment should be carefully examined. Directions for any flotation technique must be followed exactly to produce reliable results.
Figure 3.7 Flotation concentration. (Upper) Fasciolopsis buski egg (left), Diphyllobothrium latum egg (right). Note that both of these eggs in the top row are operculated and WILL NOT float in the zinc sulfate flotation concentration method; the opercula pop open, and the eggs fill with fluid and sink to the bottom of the tube. (Lower) Hookworm egg (left), Trichuris trichiura egg (right). These eggs concentrate using the flotation method and can be seen in the surface film. However, remember that both the surface film and the sediment must be examined by this method before reporting the final ova and parasite examination results. doi:10.1128/9781555819002.ch3.f7
Formalin-Ethyl Acetate Sedimentation Concentration
By centrifugation, the formalin-ethyl acetate sedimentation concentration procedure leads to the recovery of all protozoa, eggs, larvae, coccidia, and microsporidia present; however, the preparation contains more debris than is found in the flotation procedure. Ethyl acetate is used as an extractor of debris and fat from the feces and leaves the parasites at the bottom of the suspension in the sediment. The formalin-ethyl acetate sedimentation concentration procedure is recommended as being the easiest to perform, allowing recovery of the broadest range of organisms, and being the least subject to technical error.
The specimen must be fresh or formalinized stool (5 or 10% buffered or nonbuffered formalin or SAF or the Universal Fixative). Many of the single-vial preservative systems are also acceptable; however, the formulas are proprietary (e.g., UNIFIX; Medical Chemical Corp., Torrance, CA). Specimens preserved in fixatives containing polyvinyl alcohol (PVA) can also be used. However, PVA preservative formulations are rarely used for concentration methods in most laboratories but are recommended for the preparation of permanent stained smears.
5 or 10% Formalin
Formaldehyde (USP)
..................100 ml (for 10%) or
....................50 ml (for 5%)
Saline solution 900 ml (for 10%) or
0.85% NaCl 950 ml (for 5%)
Note Formaldehyde is normally purchased as a 37 to 40% HCHO solution; however, for dilution, it should be considered to be 100%.
Dilute 100 ml of formaldehyde with 900 ml of 0.85% NaCl solution. (Distilled water may be used instead of saline solution.)
Quality Control for Sedimentation Concentration
1. Check the liquid reagents each time they are used; the formalin and saline should appear clear, without any visible contamination.
2. The microscope should be calibrated (within the last 12 months), and the objectives and oculars used for the calibration procedure should be in place on the microscope when objects are measured. The calibration factors for all objectives should be posted on the microscope or close by for easy access. Some researchers feel that a microscope does not require calibration every 12 months; however, if the microscope is moved periodically, can be easily bumped, or does not receive adequate maintenance, it should be rechecked yearly for calibration accuracy.
3. Known positive specimens should be concentrated and organism recovery should be verified at least quarterly and particularly after the centrifuge has been recalibrated. Human WBCs (buffy coat cells) mixed with negative stool can be used as a QC specimen. These human cells, when mixed with negative stool, can mimic protozoan parasites. The human cells concentrate just like human parasites, such as protozoa and helminth eggs and larvae.
4. All QC results should be appropriately recorded; the laboratory should also have an action plan for “out-of-control” results.
Procedure for Sedimentation Concentration
1. Transfer 1/2 teaspoon (about 4 g) of fresh stool into 10 ml of 5 or 10% formalin in a shell vial, unwaxed paper cup, or round-bottom tube (the container may be modified to suit individual laboratory preferences). Mix the stool and formalin thoroughly, and let the mixture stand for a minimum of 30 min for fixation. If the specimen is already in 5 or 10% formalin (or SAF or other non-PVA single-vial preservatives), stir the stool-preservative mixture.
2. Depending on the amount and viscosity of the specimen, strain a sufficient quantity through wet gauze (no more than two layers of gauze or one layer if the new “pressed” gauze [e.g., Johnson & Johnson nonsterile three-ply gauze, product 7636] is used) into a conical 15-ml centrifuge tube to give the desired amount of sediment (0.5 to 1 ml) for step 3 below. Usually, 8 ml of the stool-formalin mixture prepared in step 1 is sufficient. If the specimen is received in a vial of preservative (5 or 10% formalin, SAF, or other single-vial preservatives), approximately 3 to 4 ml of the preservative-stool mixture is sufficient for testing. If the vial contains very little specimen, then the entire amount may be used in the procedure. If the specimen contains a lot of mucus, do not strain through gauze but immediately fix in 5 or 10% formalin for 30 min and centrifuge for 10 min at 500 × g. Proceed directly to step 10.
3. Add 0.85% NaCl or 5 or 10% formalin (some workers prefer to use formalin for all rinses) almost to the top of the tube, and centrifuge for 10 min at 500 × g. The amount of sediment obtained should be approximately 0.5 to 1 ml.
4. Decant and discard the supernatant fluid, and resuspend the sediment in saline or formalin; add saline or formalin almost to the top of the tube, and centrifuge again for 10 min at 500 × g. This second wash may be eliminated if the supernatant fluid after the first wash is light tan or clear. Some prefer to limit the washing to one step (regardless of the clarity or color of the supernatant fluid after centrifugation) to eliminate additional manipulation of the specimen prior to centrifugation. The more the specimen is manipulated and/or rinsed, the more likely it is that some organisms will be lost and accidentally discarded prior to examination of the sediment.
5. Decant and discard the supernatant fluid, and resuspend the sediment on the bottom of the tube in 5 or 10% formalin. Fill the tube half full only. If the amount of sediment left in the bottom of the tube is very small or the original specimen contained a lot of mucus, do not add ethyl acetate in step 6; merely add the formalin, spin, decant, and examine the remaining sediment.
6. Add 4 to 5 ml of ethyl acetate. Stopper the tube, and shake it vigorously for at least 30 s. Hold the tube so that the stopper is directed away from your face.
7. After a 15- to 30-s wait, carefully remove the stopper.
8. Centrifuge for 10 min at 500 × g. Four layers should result: a small amount of sediment (containing the parasites) in the bottom of the tube; a layer of formalin; a plug of fecal debris on top of the formalin layer; and a layer of ethyl acetate at the top (Fig. 3.5).
9. Free the plug of debris by ringing the plug with an applicator stick; decant and discard all of the supernatant fluid. After proper decanting, a drop or two of fluid remaining on the side of the tube may run down into the sediment. Mix this fluid with the sediment.
10. If the sediment is still somewhat solid, add 1 or 2 drops of saline or formalin to the sediment, mix, add a small amount of material to a slide, add a coverslip (22 by 22 mm, no. 1), and examine.
11. Systematically scan with the 10× objective. The entire coverslip area should be examined under low power (total magnification, ×100).
12. If something suspicious is seen, the 40× objective can be used for more detailed study. At least one-third to one-half of the coverslip should be examined under high dry power (total magnification, ×400), even if nothing suspicious has been seen. As in the direct wet smear, iodine can be added to enhance morphological detail, and the coverslip can be tapped to see objects move and turn over. The use of iodine is optional.
Results and Patient Reports from Sedimentation Concentration
Protozoan trophozoites and/or cysts and helminth eggs and larvae may be seen and identified. Protozoan trophozoites are less likely to be seen. In a heavy infection with Cryptosporidium spp. or C. cayetanensis, oocysts may be seen in the concentrate sediment; oocysts of C. belli can also be seen. Spores of the microsporidia are too small, and the shape resembles that of other debris within the stool; therefore, they are not readily visible in the concentration sediment. However, special stains can be performed on the sediment for the identification of coccidian oocysts and microsporidian spores.
1. Protozoan cysts may or may not be identified to the species level (depending on the clarity of the morphology).
Examples: Entamoeba coli cysts
Giardia lamblia (G. duodenalis, G. intestinalis) cysts
2. Helminth eggs and/or larvae may be identified.
Examples: Ascaris lumbricoides eggs
Hookworm larvae
3. C. belli oocysts may be identified; however, Cyclospora and Cryptosporidium oocysts are generally too small to be recognized and can be identified using appropriate immunoassays or modified acid-fast staining.
Example: Cystoisospora (Isospora) belli oocysts
4. Artifacts and/or other structures may also be seen and reported as follows.
Note These crystals and cells are quantitated; however, the quantity is usually assessed when the permanent stained smear is examined under oil immersion).
Examples: Moderate Charcot-Leyden crystals
Few RBCs
Moderate PMNs
Procedure Notes for Sedimentation Concentration
1. The gauze should never be more than one (pressed gauze) or two (woven gauze) layers thick; more gauze may trap mucus (containing Cryptosporidium oocysts and/or microsporidial spores).
2. Tap water may be substituted for 0.85% NaCl throughout this procedure, although the addition of water to fresh stool causes Blastocystis spp. cyst (central body) forms to rupture and is not recommended. In addition to the original 5 or 10% formalin fixation, some workers prefer to use 5 or 10% formalin for all rinses throughout the procedure.
3. Ethyl acetate is widely recommended as a substitute for ether (16). It can be used in the same way in the procedure and is much safer. Hemo-De can also be used and is thought to be safer than ethyl acetate (23).
A. After the plug of debris is rimmed and excess fluid is decanted, the sides of the tube can be swabbed with a cotton-tipped applicator stick while the tube is still upside down to remove excess ethyl acetate. This is particularly important if you are working with plastic centrifuge tubes or the plastic commercial concentrators. If the sediment is too dry after the tube has been swabbed, add several drops of saline before preparing the wet smear for examination.
B. If there is excess ethyl acetate in the smear of the sediment prepared for examination, bubbles will be present, which will obscure the material of interest.
4. If specimens are received in SAF, begin the procedure at step 2.
5. If specimens are received in fixative containing PVA, the first two steps of the procedure should be modified as follows.
A. Immediately after stirring the stool-PVA mixture with applicator sticks, pour approximately half of the mixture into a tube (container optional) and add 0.85% NaCl (or 5 or 10% formalin) almost to the top of the tube.
B. Filter the stool-PVA-saline (or formalin) mixture through wet gauze into a 15-ml centrifuge tube. Follow the standard procedure from here to completion, beginning with step 3.
6. Too much or too little sediment will result in an ineffective concentration sediment examination. See the section on Commercial Fecal Concentration Devices later in this chapter.
7. The centrifuge should reach the recommended speed before the centrifugation time is monitored. However, since most laboratories have their centrifuges on automatic timers, the centrifugation time in this protocol takes into account the fact that some time will be spent coming up to speed prior to full-speed centrifugation. If the centrifugation time at the proper speed (10 min at 500 × g) is reduced, some of the organisms (Cryptosporidium and Cyclospora oocysts or microsporidian spores) may not be recovered in the sediment.
Procedure Limitations for Sedimentation Concentration
1. Results obtained with wet smears (direct wet smears or concentration sediment wet smears) should usually be confirmed by permanent stained smears. Some protozoa are very small and difficult to identify as to species with just the direct wet smears. Also, special stains are sometimes necessary for organism identification.
2. Confirmation is particularly important for E. histolytica/E. dispar versus E. coli.
3. Certain organisms [G. lamblia (G. duodenalis, G. intestinalis), hookworm eggs, and occasionally Trichuris eggs] may not concentrate as well from PVA-preserved specimens as they do from those preserved in formalin. However, if enough Giardia organisms are present to concentrate from formalin, PVA should contain enough for detection on the permanent stained smear. In clinically important infections, the number of helminth eggs present would ensure detection regardless of the type of preservative used. Also, the morphology of Strongyloides stercoralis larvae is not as clear from specimens in PVA as from specimens fixed in formalin.
4. For unknown reasons, C. belli oocysts are routinely missed in the concentrate sediment when concentrated from PVA-preserved specimens. The oocysts would be found if the same specimen were preserved in formalin rather than PVA.
5. In past publications, recommended centrifugation times have not taken into account potential problems with the recovery of Cryptosporidium oocysts. There is anecdotal evidence strongly indicating that Cryptosporidium oocysts may be missed unless the centrifugation speed is 500 × g for a minimum of 10 min.
6. Adequate centrifugation time and speed have become very important for recovery of microsporidial spores. In some of the earlier publications, use of uncentrifuged material was recommended. However, we have found that centrifugation for 10 min at 500 × g definitely increases the number of microsporidial spores available for staining and subsequent examination.
Iodine-Trichrome Stain for Sediment
A combination of Lugol’s iodine solution and trichrome stain can be used to stain fecal sediment from the concentration procedure (24). Coloring the eggs and cysts yellow-brown (iodine) and the debris green (trichrome) provides contrast which facilitates the detection of parasites. The use of such an approach usually depends on personal preferences and the results of parallel trials of the current method and new methods being considered. This iodine-trichrome wet examination can be used as an adjunct procedure but does not take the place of the unstained wet examination of the sediment.
Quality Control for Iodine-Trichrome Stain for Sediment
1. Check the working iodine solution each time it is used or periodically (once a week). The iodine and trichrome stain 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 in a saline mount. Human WBCs (buffy coat cells) mixed with negative stool can be used as a QC specimen. The human cells stain with the same color as that seen in the protozoa. The background debris stains green from the components of the trichrome stain.
4. If appropriate due to extensive use and/or lack of routine maintenance, 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 being measured. The calibration factors for all objectives should be posted on the microscope or close by for easy access.
5. All QC results should be appropriately recorded; the laboratory should also have an action plan for “out-of-control” results.
Procedure for Iodine-Trichrome Stain for Sediment
1. Place 4 drops of Lugol’s iodine solution into a test tube.
2. Place 4 drops of fecal concentrate into the test tube. Mix well.
3. Place 2 drops of the Lugol’s iodine solution-fecal concentrate mixture from step 2 on a glass slide.
4. Add 1 drop of trichrome stain. Mix with a wooden applicator stick, and cover with a coverslip (22 by 22 mm, no. 1).
5. Microscopically examine the entire preparation under low power (×100) and at least one-third to one-half of the area under high dry power (×400).
Results and Patient Reports from Iodine-Trichrome Stain for Sediment
Protozoan trophozoites and/or cysts and some helminth eggs and larvae may be seen and identified. Lugol’s iodine stains A. lumbricoides and Taenia eggs quite dark; these eggs are difficult to recognize and may be mistaken for debris, hence the need for the unstained sediment examination. In a heavy infection with Cryptosporidium spp., oocysts may be seen in a direct smear; however, some type of modified acid-fast stain or fecal immunoassay is normally used to detect these organisms, particularly when few oocysts are present. Oocysts of C. belli can also be seen in a direct smear. Cyclospora oocysts may not be recognized because they resemble debris. Spores of the microsporidia are too small, and the shape resembles that of other debris within the stool; therefore, they are not readily visible in a direct smear.
1. Protozoan cysts may or may not be identified to the species level (depending on the clarity of the morphology).
Example: Iodamoeba bütschlii cysts
2. Helminth eggs and/or larvae may be identified.
Example: Hymenolepis nana eggs
3. C. belli oocysts may be identified; however, Cyclospora and Cryptosporidium oocysts are generally too small to be recognized or identified. Subsequent immunoassays or modified acid-fast staining is recommended.
Example: Cystoisospora (Isospora) belli oocysts
4. Artifacts and/or other structures may also be seen and reported as follows.
Note These cells are quantitated; however, the quantity is usually assessed when the permanent stained smear is examined under oil immersion.
Example: Moderate PMNs
Procedure Notes for Iodine-Trichrome Stain for Sediment
1. As mentioned above, some workers prefer to make the wet mounts on larger slides. Often, there is less chance of getting fluids on the microscope stage if larger slides are used.
2. This stain is darker than the traditional iodine stain. The microscope light should be increased over that used for the unstained wet smear examination.
Procedure Limitations for Iodine-Trichrome Stain for Sediment
1. Because this is a darker stain than routine iodine stains, it is important to also examine a saline wet smear. This is particularly important because A. lumbricoides and Taenia eggs stain too dark with the iodine and may not be recognized as helminth eggs.
2. Results obtained with wet smears should usually be confirmed by permanent stained smears. Some protozoa are very small and difficult to identify to the species level with just the direct wet smears. Confirmation is particularly important for E. histolytica/E. dispar versus E. coli. These findings can be reported as “preliminary report, based on direct wet smear examination only,” and the final report can be submitted after the concentration and permanent stain procedures are completed. However, if the examination turnaround time is approximately 24 h or less, there is no need for a preliminary report; the final report can be submitted after completion of the concentration and permanent stained smear examinations.
Zinc Sulfate Flotation Concentration
The flotation procedure permits the separation of protozoan cysts and eggs of certain helminths from excess debris through the use of a liquid (zinc sulfate) with a high specific gravity. The parasitic elements are recovered in the surface film, and the debris and some heavy parasitic elements remain in the bottom of the tube. This technique yields a cleaner preparation than does the sedimentation procedure; however, some helminth eggs (operculated and/or very dense eggs, such as unfertilized Ascaris eggs) do not concentrate well in the flotation method; a sedimentation technique is recommended to detect these infections.
When the zinc sulfate solution is prepared, the specific gravity should be 1.18 for fresh stool specimens; it must be checked with a hydrometer. This procedure may be used on formalin-preserved specimens if the specific gravity of the zinc sulfate is increased to 1.20; however, this usually causes more distortion in the organisms present and is not recommended for routine clinical use. To ensure detection of all possible organisms, both the surface film and the sediment must be examined. For most laboratories, this is not a practical approach.
The specimen must be fresh or formalinized stool (5 or 10% buffered or nonbuffered formalin, SAF, or other non-PVA single-vial preservatives). PVA-preserved specimens can also be used; however, this approach is not commonly used or recommended.
Zinc Sulfate (33% Aqueous Solution)
1. Using a magnetic stirrer, dissolve the zinc sulfate in distilled water in an appropriate flask or beaker.
2. Adjust the specific gravity to 1.20 by the addition of more zinc sulfate or distilled water. Use a specific gravity of 1.18 with fresh stool (nonformalinized).
3. Store in a glass-stoppered bottle with an expiration date of 24 months.
Quality Control for Flotation Concentration
1. Check the reagents each time they are used. The formalin, saline, and zinc sulfate should appear clear, without any visible contamination.
2. 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 or close by for easy access. As mentioned above, some workers feel that recalibration of the microscope is not necessary each year; however, this would depend on the use and maintenance of that particular piece of equipment.
3. Known positive specimens should be concentrated and organism recovery should be verified at least quarterly, particularly after the centrifuge has been recalibrated. Human WBCs (buffy coat cells) mixed with negative stool can be used as a QC specimen. These human cells, when mixed with negative stool, mimic human parasites. The human cells concentrate just like human parasites, such as protozoa and helminth eggs and larvae.
4. All QC results should be appropriately recorded; the laboratory should also have an action plan for “out-of-control” results.
Procedure for Flotation Concentration
1. Transfer 1/2 teaspoon (about 4 g) of fresh stool into 10 ml of 5 or 10% formalin in a shell vial, unwaxed paper cup, or round-bottom tube (the container may be modified to suit individual laboratory preferences). Mix the stool and formalin thoroughly. Let the mixture stand for a minimum of 30 min for fixation. If the specimen is already in 5 or 10% formalin (or SAF or Universal Fixative), stir the stool-formalin (or SAF) mixture.
2. Depending on the size and density of the specimen, strain a sufficient quantity through wet gauze (no more than two layers of gauze or one layer if the new “pressed” gauze [e.g., Johnson & Johnson nonsterile three-ply gauze, product 7636] is used) into a conical 15-ml centrifuge tube to give the desired amount of sediment (0.5 to 1 ml) in step 3 below. Usually, 8 ml of the stool-formalin mixture prepared in step 1 is sufficient. If the specimen is received in vials of preservative (5 or 10% formalin, SAF, or other single-vial preservatives), approximately 3 to 4 ml of the mixture is sufficient unless the specimen has very little stool in the vial. If the specimen contains a lot of mucus, do not strain through gauze but immediately fix in 5 or 10% formalin or other fixative (does not contain PVA) for 30 min and centrifuge for 10 min at 500 × g. Proceed directly to step 5.
3. Add 0.85% NaCl almost to the top of the tube, and centrifuge for 10 min at 500 × g. Approximately 0.5 to 1 ml of sediment should be obtained. Too much or too little sediment results in an ineffective concentration examination. See the section on Commercial Fecal Concentration Devices later in this chapter.
4. Decant and discard the supernatant fluid, resuspend the sediment in 0.85% NaCl almost to the top of the tube, and centrifuge for 10 min at 500 × g. This second wash may be eliminated if the supernatant fluid after the first wash is light tan or clear. Some prefer to limit the washing to one step (regardless of the color and clarity of the supernatant fluid) to eliminate additional manipulation of the specimen prior to centrifugation. The more the specimen is manipulated and/or rinsed, the more likely it is for parasitic elements to be lost.
5. Decant and discard the supernatant fluid, and resuspend the sediment on the bottom of the tube in 1 to 2 ml of zinc sulfate. Fill the tube within 2 to 3 mm of the rim with additional zinc sulfate.
6. Centrifuge for 2 min at 500 × g. Allow the centrifuge to come to a stop without interference or vibration. Two layers should result: a small amount of sediment in the bottom of the tube, and a layer of zinc sulfate (Fig. 3.5). The protozoan cysts and some helminth eggs are found in the surface film; some operculated and/or heavy eggs are found in the sediment.
7. Without removing the tube from the centrifuge, remove 1 or 2 drops of the surface film with a Pasteur pipette or a freshly flamed (and allowed to cool) wire loop and place them on a slide. Do not use the loop as a “dipper”; simply touch the surface (bend the loop portion of the wire 90° so that the loop is parallel with the surface of the fluid). Make sure the pipette tip or wire loop is not below the surface film (Fig. 3.8).
8. Add a coverslip (22 by 22 mm, no. 1) to the preparation. Iodine may be added to the preparation (optional).
9. Systematically scan with the 10× objective. The entire coverslip area should be examined under low power (total magnification, ×100).
10. If something suspicious is seen, the 40× objective can be used for more detailed study. At least one-third to one-half of the coverslip should be examined with high dry power (total magnification, ×400), even if nothing suspicious has been seen. As in the direct wet smear, iodine can be added to enhance morphological detail, and the coverslip can be gently tapped to observe objects moving and turning over.
Figure 3.8 Method used to remove the surface film in the zinc sulfate flotation concentration procedure. (A) A wire loop is gently placed on (not under) the surface film. (B) The loop is then placed on a glass slide. (Illustration by Nobuko Kitamura.) doi:10.1128/9781555819002.ch3.f8
Results and Patient Reports from Flotation Concentration
Protozoan trophozoites and/or cysts and some helminth eggs and larvae may be seen and identified. Heavy helminth eggs and operculated eggs do not float in zinc sulfate; they are seen in the sediment within the tube. The high specific gravity of the zinc sulfate causes the opercula to pop open; the eggs fill with fluid and sink to the bottom. Protozoan trophozoites are less likely to be seen. In a heavy infection with Cryptosporidium spp. or C. cayetanensis, oocysts may be seen in the concentrate sediment; oocysts of C. belli can also be seen. Spores of the microsporidia are too small, and the shape resembles other debris within the stool; therefore, they are not readily visible in the concentration sediment.
1. Protozoan cysts may or may not be identified to the species level (depending on the clarity of the morphology).
Example: Giardia lamblia (G. duodenum, G. intestinalis) cysts
2. Helminth eggs and/or larvae may be identified.
Example: Hookworm eggs
3. C. belli oocysts may be identified; however, Cyclospora and Cryptosporidium oocysts are generally too small to be recognized or identified. Subsequent immunoassays or modified acid-fast staining is recommended.
Example: Cystoisospora (Isospora) belli oocysts
4. Artifacts and/or other structures may also be seen and reported as follows.
Note These cells are quantitated; however, the quantity is usually assessed when the permanent stained smear is examined.
Examples: Few macrophages
Moderate PMNs
Procedure Notes for Flotation Concentration
1. The gauze should never be more than one or two layers thick; more gauze may trap mucus (containing Cryptosporidium oocysts, Cyclospora oocysts, and/or microsporidial spores). A round-bottom tube is recommended rather than a centrifuge tube.
2. Tap water may be substituted for 0.85% NaCl throughout this procedure, although the addition of water to fresh stool causes Blastocystis spp. (Blastocystis hominis) cyst (central body) forms to rupture and is not recommended. In addition to the original 5 or 10% formalin fixation, some workers prefer to use 5 or 10% formalin for all rinses throughout the procedure.
3. If fresh stool is used (nonformalin preservatives), the zinc sulfate should be prepared with a specific gravity of 1.18. If formalinized specimens are to be concentrated, the zinc sulfate should have a specific gravity of 1.20.
4. If specimens are received in SAF or other single-vial preservatives, begin the procedure at step 2.
5. If fresh specimens are received, the standard procedure requires the stool to be rinsed in distilled water prior to the addition of zinc sulfate in step 4. However, the addition of fresh stool to distilled water will destroy any Blastocystis spp. (Blastocystis hominis) cysts present and is not a recommended approach.
6. Some workers prefer to remove the tubes from the centrifuge prior to sampling the surface film. This is acceptable; however, there is more chance that the surface film will be disturbed prior to sampling.
7. Some workers prefer to add a small amount of zinc sulfate to the tube so that the fluid forms a slightly convex meniscus. A coverslip is then placed on top of the tube so that the undersurface touches the meniscus. It is left undisturbed for 5 min. The coverslip is then carefully removed and placed on a slide for examination. This approach tends to be somewhat messy, particularly if too much zinc sulfate has been added.
8. When using the hydrometer (solution at room temperature), mix the solution well. Float the hydrometer in the solution, giving it a slight twist to ensure that it is completely free from the sides of the container. Read the bottom meniscus and correct the figure for temperature, if necessary. Most hydrometers are calibrated at 20°C. A difference of 3°C between the solution temperature (room temperature) and the hydrometer calibration temperature requires a correction of 0.001, to be added if above and subtracted if below 20°C.
Procedure Limitations for Flotation Concentration
1. Results obtained with wet smears (direct wet smears or concentrated specimen wet smears) should usually be confirmed by permanent stained smears. Some protozoa are very small and difficult to identify to the species level with just the direct wet smears. Also, special stains are sometimes necessary for organism identification.
2. Confirmation is particularly important for E. histolytica/E. dispar versus E. coli.
3. Protozoan cysts and thin-shelled helminth eggs are subject to collapse and distortion when left for more than a few minutes in contact with the high-specific-gravity zinc sulfate. The surface film should be removed for examination within 5 min of the time the centrifuge comes to a stop. The longer the organisms are in contact with the zinc sulfate, the more distortion will be seen on microscopic examination of the surface film.
4. Since most laboratories have their centrifuges on automatic timers, the centrifugation time in this protocol takes into account the fact that some time will be spent coming up to speed prior to full-speed centrifugation.
5. If zinc sulfate is the only concentration method used, both the surface film and the sediment must be examined to ensure detection of all possible organisms.
Commercial Fecal Concentration Devices
There are a number of commercially available fecal concentration devices which may help a laboratory to standardize the concentration technique. Standardization is particularly important when personnel rotate throughout the laboratory and may not be familiar with parasitology techniques. These devices help ensure consistency, thus leading to improved parasite recovery and subsequent identification. Some of the systems are enclosed and provide a clean, odor-free approach to stool processing, features that may be important to nonmicrobiology personnel processing such specimens. Both 15- and 50-ml systems are available. It is important to remember that a maximum of 0.5 to 1.0 ml of sediment is needed in the bottom of the tube. Often, when the 50-ml systems are used, there is too much sediment in the bottom of the tube. This problem can be solved by adding less of the fecal specimen to the concentration system prior to centrifugation. Since the sediment is normally mixed thoroughly and 1 drop is taken to a coverslip for examination, good mixing may not occur if too much sediment is used. There also appears to be layering in the bottom of the tubes; again, adding less material to the concentrator at the beginning should help eliminate this problem (Fig. 3.9 through 3.12).
Figure 3.9 (Upper) FPC JUMBO large concentration tubes and connector system (Evergreen Scientific). (Lower) Small concentration tubes and FPC HYBRID connector system (Evergreen Scientific). doi:10.1128/9781555819002.ch3.f9
Figure 3.10 Stool collection vial and funnel used in fecal concentration (Hardy Diagnostics). doi:10.1128/9781555819002.ch3.f10
Figure 3.11 (Top) PARA-SED concentration system with both small and large tubes (Medical Chemical Corp.). (Middle) SED-CONNECT system with collection vials and various reagents (Medical Chemical Corp.). (Bottom) Filter attachment system (Medical Chemical Corp.). Note the screen, which is a substitute for the gauze in the traditional gauze/filter concentration method. See also MICRO-SED. doi:10.1128/9781555819002.ch3.f11
Figure 3.12 (Upper left) MACRO-CON concentration system (Meridian Bioscience). (Upper right) SPINCON concentration system (Meridian Bioscience). (Lower) Funnel used in fecal concentration (Meridian Bioscience). doi:10.1128/9781555819002.ch3.f12
Automated Workstation for the Microscopic Analysis of Fecal Concentrates
The FE-5 (Apacor Ltd USA, Brooklyn, NY) is a countertop workstation that automates the microscopic analysis of fecal concentrates (Fig. 3.13). The system automates the aspiration, resuspension, staining or diluting (based on user preference), transfer, presentation, and disposal of fecal concentrates. When the sample button is pressed, within 5 s two samples of fecal concentrate are automatically and simultaneously aspirated from the concentrate tube and transported to the glass dual-flow-cells of the Optical Slide Assembly (Fig. 3.14). Based on user preference, the FE-5 also simultaneously stains or dilutes one of the two samples to be examined. After the microscopic examination of the fecal suspension within the glass viewing chambers, the flow chambers can be purged and cleaned and are ready for the next specimen. The dual-flow-cell Optical Slide Assembly is designed to fit within the stage clips of any standard upright microscope. The Optical Slide Assembly accommodates bright-field, phase-contrast, polarized-light, and other common forms of microscopy. The system can be moved from one microscope to another or can be set up as a semipermanent station for fecal concentrate microscopy. Removal to another microscope just involves removing the Optical Slide Assembly from the microscope stage.
Figure 3.13 Countertop workstation that automates the microscopic analysis of fecal concentrates (Apacor Ltd. USA, Brooklyn, NY). doi:10.1128/9781555819002.ch3.f13
Figure 3.14 Dual-flow-cell Optical Slide Assembly (Apacor Ltd USA, Brooklyn, NY) that fits into the stage clips of any standard upright microscope. doi:10.1128/9781555819002.ch3.f14
The detection and correct identification of many intestinal protozoa frequently depend on the examination of the permanent stained smear with the oil immersion lens (100× objective). These slides not only provide the microscopist with a permanent record of protozoan organisms identified but also may be used for consultations with specialists when unusual morphologic characteristics are found. Considering the morphologic variations that are possible, organisms that are very difficult to identify and do not fit the pattern for any one species may be found. A routine work flow diagram including the permanent stained smear is shown in Fig. 3.15.
Figure 3.15 Work flow diagram for fecal specimens. The total examination includes the direct mount, concentrate, and permanent stained smear (fresh specimen) or the concentrate and permanent stained smear (preserved specimens). doi:10.1128/9781555819002.ch3.f15
Although an experienced microscopist can occasionally identify certain organisms on a wet preparation, most identifications should be considered tentative until confirmed by the permanent stained slide. The smaller protozoan organisms are frequently seen on the stained smear when they are easily missed with only the direct smear and concentration methods. For these reasons, the permanent stain is recommended for every stool sample submitted for a routine parasite examination. It is also important to remember that the fecal immunoassays for specific organisms have proven to be more sensitive than the routine or specialized stains for Giardia lamblia (G. duodenalis, G. intestinalis) or Cryptosporidium spp. (25, 26).
There are a number of staining techniques available; selection of a particular method may depend on the degree of difficulty of the procedure and the amount of time necessary to complete the stain. The older classical method is the long Heidenhain iron hematoxylin method; however, for routine diagnostic work, most laboratories select one of the shorter procedures, such as the trichrome method or one of the modified methods involving iron hematoxylin. Other procedures are available (4, 6–8, 13, 17, 20, 24, 27, 30, 49), and some of them are presented here.
Most problems encountered in the staining of protozoan trophozoites and cysts in fecal smears occur because the specimen is too old, the smears are too dense, the smears are allowed to dry before fixation, or fixation is inadequate. There is variability in fixation in that immature cysts fix more easily than mature cysts, and E. coli cysts require a longer fixation time than do those of other species. It is critical that adequate mixing occur between the fecal specimen and preservative (27–29, 31, 32).
Preparation of Material for Staining
Fresh Material
1. When the specimen arrives, prepare two slides with applicator sticks or brushes and immediately (without drying) place them in Schaudinn’s fixative [(This approach is rarely used because the formulation contains mercury). Preserved stool collected in fecal fixatives (with or without PVA) can be smeared on a slide and allowed to air dry prior to staining.] Allow the slides to fix for a minimum of 30 min; overnight fixation is acceptable. The amount of fecal material smeared on the slide should be thin enough that newsprint can be read through the smear. Smears preserved in liquid Schaudinn’s fixative should be placed in 70% alcohol to remove the excess fixative prior to placement in iodine-alcohol (used for mercury-based fixatives).
2. If the fresh specimen is liquid, place 3 or 4 drops of fixative (Schaudinn’s fixative to which has been added a plastic powder [PVA], which serves as an adhesive to “glue” the fecal material onto the slide) on the slide, mix several drops of fecal material with the PVA fixative, spread the mixture, and allow it to dry for several hours in a 37°C incubator or overnight at room temperature.
3. Proceed with the trichrome staining procedure by placing the slides in iodine-alcohol.
Preserved Material Containing PVA
1. Stool specimens that are preserved in fixative (mercuric chloride, copper sulfate, or zinc sulfate bases) containing PVA should be allowed to fix for at least 30 min. Thoroughly mix the contents of the vial or bottle (fixative/stool/PVA specimen) with two applicator sticks.
2. Pour some of the well-mixed fixative/stool/PVA mixture onto a paper towel, and allow it to stand for 3 min to absorb out the excess PVA. Do not eliminate this step.
3. With an applicator stick (or brush), apply some of the stool material from the paper towel to two slides and allow them to dry for several hours in a 37°C incubator or overnight at room temperature.
Note The fixative/stool/PVA mixture should be spread to the edges of the glass slide; this will cause the film to adhere to the slide during staining. It is also important to thoroughly dry the slides in order to prevent the material from washing off during staining.
4. The dry slides may then be placed into iodine-alcohol. There is no need to give them a 70% alcohol rinse before placing them in the iodine-alcohol, because the PVA smears are already dry (unlike the wet smears coming out of Schaudinn’s fixative). The iodine-alcohol is used to remove the mercuric chloride from the slides before they are stained. When reviewing the trichrome staining procedure, you will note that this step is not required if the stool specimen is preserved in a copper sulfate or zinc sulfate-based preservative.
NOTE Polyvinyl alcohol (PVA) is a plastic powder that is often added to fecal fixatives to help “glue” the stool material onto the glass slide. PVA is inert and has no fixing properties. Its main purpose is to cause the fecal material to adhere to the glass once the material is dry. HOWEVER, the Universal Fixatives do not require PVA or albumin to serve as adhesives.
California State Department of Health Modification (1) for PVA-Preserved Material
1. Thoroughly mix the fixative/stool/PVA mixture, and strain it through damp (with tap water) gauze into 15-ml centrifuge tubes.
2. After centrifugation (2 min at 500 × g), decant and discard the fluid and swab the sides of the tubes to remove any excess liquid. Although stains for Cryptosporidium spp. are not recommended from preserved material containing PVA, the centrifuge speed indicated here is probably not sufficient to recover the oocysts or microsporidian spores for special staining, fecal immunoassays using fluorescence, or other diagnostic procedures; use 500 × g for 10 min.
3. Use the remaining sediment to prepare coverslip smears (not 3- by 1-in. slides), which are immediately (without drying) stained, beginning with the iodine-alcohol step of the trichrome staining procedure (for removal of mercuric chloride).
4. The staining times can be reduced to 2 to 4 min, and dehydration steps can be reduced to 1 min each.
Note This procedure will work quite well for most stool specimens preserved in fixative containing PVA. However, for very liquid specimens or those containing large amounts of mucus, drying the smears before staining will be advantageous.
SAF-Preserved Material (31)
1. Thoroughly mix the SAF-stool mixture, and strain it through gauze into a 15-ml centrifuge tube.
2. After centrifugation (1 min at 500 × g), decant and discard the supernatant fluid. Although stains for coccidian oocysts and microsporidian spores are not recommended from PVA-preserved material, the centrifuge speed indicated here is probably not sufficient to recover the oocysts or spores. Centrifugation time and speed are recommended as 10 min at 500 × g. The final amount of sediment should be about 0.5 to 1.0 ml. If necessary, adjust by repeating step 1 or by resuspending the sediment in saline (0.85% NaCl) and removing part of the suspension.
3. Prepare a smear from the sediment for later staining by placing 1 drop of Mayer’s albumin on the slide and adding 1 drop of SAF-preserved fecal sediment. Allow the smear to air dry at room temperature for 30 min prior to staining.
4. After being dried, the smear can be placed directly into 70% alcohol (step 4) of the staining procedure (the iodine-alcohol step can be eliminated).
Merthiolate-Iodine-Formalin (MIF)-Preserved Material
1. Prepare a Mayer’s albumin-coated slide and allow it to air dry at room temperature for 30 min.
2. From the MIF vial (material allowed to settle in the vial for at least 1 h undisturbed), remove a portion of sediment and place this material onto the albumin-coated slide. Allow the slide to remain flat for 5 min; if there is still fluid left on the slide after this time, stand it up and allow the excess fluid to run off onto a paper towel.
3. After all excess fluid is removed, the slides are ready for staining in polychrome IV stain.
Universal Fixative (TOTAL-FIX)-Preserved Material
The TOTAL-FIX stool collection kit is a single-vial system that provides a standardized method for untrained personnel to properly collect and preserve stool specimens for the detection of helminth larvae and eggs, protozoan trophozoites and cysts, coccidian oocysts, and microsporidian spores. Concentrations, permanent stains, most fecal immunoassays, special stains, and some molecular testing can be performed from a TOTAL-FIX-preserved specimen. TOTAL-FIX is a mercury-, formalin-, and PVA-free fixative that preserves parasite morphology and helps with disposal and monitoring problems encountered by laboratories. The product contains zinc sulfate, acetic acid, and alcohols in water; however, the formula is proprietary and is patented (see Algorithm 3.1).
Algorithm 3.1 Method for the preparation of TOTAL-FIX-preserved fecal specimens (Universal Fixative) for the concentration, permanent stained smear, fecal immunoassays, and special stains for the coccidia and microsporidia. (a) This fixative contains NO mercury, NO formalin, and NO polyvinyl alcohol (PVA); use one-third stool and two-thirds fixative and MIX WELL. Allow 30 min prior to processing. (b) The EIA and rapid cartridge detect antigen; sample the clear fluid at the top of the vial containing the stool/fixative mixture. If the vial is shaken, make sure it stands on the counter for 5 to 10 min to allow the particulate material to settle out; work with the clear fluid only. (c) TOTAL-FIX has been tested with a number of molecular methods; however, there is no confirmation that the specimen will work with every possible available method. (d) Rinse fluids (water, saline, formalin) will PREVENT permanent stained smears from staining correctly; rinse fluids can ONLY be used for the final concentration step (after original sediment has been removed for permanent stained slide preparations—STEP 2). (e) Since the direct fluorescent antibody immunoassay (DFA) detects the actual Giardia cysts (rare trophozoites—much less intense fluorescence) and Cryptosporidium oocysts, this test sensitivity is enhanced if centrifuged sediment is used for testing; the same advantage is seen using modified acid-fast stains for Cryptosporidium (use sediment). Remember, do not use RINSED sediment, but sediment from the original centrifugation. (f) Examine entire 22-by-22-mm coverslip at low power (10× objective); examine one-third to one-half coverslip at high dry power (40× objective).
Trichrome Stain
The trichrome technique of Wheatley (22) for fecal specimens is a modification of Gomori’s original staining procedure for tissue (7). It is a rapid, simple procedure which produces uniformly well-stained smears of the intestinal protozoa, human cells, yeast cells, and artifact material in about 45 min or less.
The specimen usually consists of fresh stool smeared on a microscope slide, which is immediately fixed in liquid Schaudinn’s fixative, preserved stool (with or without PVA), the single-vial fixatives, or the Universal Fixative smeared on a slide and allowed to air dry.
Trichrome Stain
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.
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).
Figure 3.17 Guess the identity of these intestinal protozoa! See p. 76 for the answers. doi:10.1128/9781555819002.ch3.f17
*Slides may be held for up to 24 h in these solutions without harming the quality of the smear or stainability of organisms.
Procedure for Trichrome Stain with Non-Mercury-Based Fixatives (Iodine-Alcohol Step and Alcohol Rinse Not Required) (Fig. 3.16)
1. Prepare the slide for staining as described above.
2. Place the slide in 70% ethanol for 5 min.*
3. Place it in the trichrome stain for 10 min. Some people prefer to place the dry smear directly into the stain and eliminate step 2 from the protocol.
4. 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.
5. Dip the slide several times in 100% ethanol. Use this step as a rinse.
6. Place it in two changes of 100% ethanol for 3 min each.*
7. Place it in xylene or xylene substitute for 5 to 10 min.*
8. Place it in a second container of xylene or xylene substitute for 5 to 10 min.*
9. Mount with a coverslip (no. 1 thickness), using mounting medium (e.g., Permount).
10. Allow the smear to dry overnight or after 1 h at 37°C.
11. Examine the smear microscopically with the 100× objective. Examine at least 200 to 300 oil immersion fields before reporting a negative result.
*Slides may be held for up to 24 h in these solutions without harming the quality of the smear or stainability of organisms.
Results and Patient Reports from the Trichrome Staining Method
Protozoan trophozoites and cysts are readily seen, although helminth eggs and larvae may not be easily identified because of excess stain retention (wet smears from the concentration procedure[s] are recommended for detection of these organisms) (Fig. 3.18 and 3.19). Yeasts (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 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. 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).
*Slides may be held for up to 24 h in these solutions without harming the quality of the smear or stainability of organisms.
Note An alternative method to using mounting medium is as follows (2).
A. Remove the slide from the last container of xylene, place it on a paper towel (flat position), and allow it to air dry. Remember that some of the xylene substitutes may take a bit longer to dry than xylene itself does.
B. Approximately 5 to 10 min before you want to examine the slide, place a drop of immersion oil on the dry fecal film. Allow the oil to sink into the film for a minimum of 10 to 15 min. If the smear appears to be very refractile on examination, you have not waited long enough for the oil to sink into the film or you need to add a bit more oil to the film.
C. Once you are ready to examine the slide, place a no. 1 (22- by 22-mm) coverslip onto the oiled smear, add another drop of immersion oil onto the top of the coverslip (as you would normally do for any slide with a coverslip), and examine with the oil immersion lens (100× objective).
D. Do not eliminate adding the coverslip; the dry fecal material on the slide often becomes very brittle after dehydration. Without the addition of the protective coverslip, you might scratch the surface of the oil immersion lens. Coverslips are much cheaper than oil immersion objectives!
Iron Hematoxylin Stain
The iron hematoxylin stain is one of a number of stains that allow one to make a permanent stained slide for detecting and quantitating parasitic organisms. Iron hematoxylin was the stain used for most of the original morphologic descriptions of intestinal protozoa found in humans (3) (Fig. 3.20). On oil immersion power (×1,000), one can examine the diagnostic features used to identify the protozoan parasites. Although there are many modifications of iron hematoxylin techniques, only two methods are outlined below: the Spencer-Monroe (19) and Tompkins-Miller (20) procedures. Both methods can be used with either fresh specimens or those preserved with SAF, preservatives containing PVA, single-vial systems, or the Universal Fixatives.
Figure 3.20 Intestinal protozoa stained with iron hematoxylin stain. (Top row) Dientamoeba fragilis trophozoite (left), Giardia lamblia (G. duodenalis, G. intestinalis) cysts (right); (middle row) Iodamoeba bütschlii cyst (note the large glycogen vacuole) (left), Entamoeba histolytica trophozoite (note ingested RBCs) (right); (bottom row) Entamoeba coli cyst (note more than five nuclei) (left), Entamoeba histolytica/E. dispar cyst (right). doi:10.1128/9781555819002.ch3.f20
The specimen usually consists of fresh stool smeared on a microscope slide, which is immediately fixed in Schaudinn’s fixative; stool in fixative containing PVA smeared on a slide and allowed to air dry; SAF-preserved stool smeared on an albumin-coated slide and allowed to air dry; single-vial-preserved stool smeared on an albumin-coated slide and allowed to air dry; or stool smeared on a slide and allowed to air dry (Universal Fixative, no adhesive is required).
Iron Hematoxylin Stain (Spencer-Monroe Method) (19)
Solution 1
Place solution in a stoppered clear flask or bottle, and allow it to ripen in a lighted room for at least 1 week at room temperature.
Solution 2
Ferrous ammonium sulfate
Ferric ammonium sulfate
Working Solution
Mix equal volumes of solutions 1 and 2. The working solution should be made fresh every week.
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, new working solution does not have to be made. The replacement time will depend on the number of smears stained and the size of the container (1 week to several weeks).
Prepare by combining.
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 Iron Hematoxylin Stain
1. Stool samples used for quality control can be fixed stool specimens known to contain protozoa or preserved negative stools containing PVA to which buffy coat cells (PMNs or macrophages) have been added. A QC smear prepared from a positive sample or a 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.
3. If the xylene or xylene substitute becomes cloudy or there is an accumulation of water in the bottom of the staining dish, discard the old reagents, clean the dishes, dry them 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 should to be changed on an as-needed basis.
6. When the smear is thoroughly fixed and the stain is performed correctly, the cytoplasm of protozoan trophozoites will be blue-gray, sometimes with a tinge of black. Cysts tend to be slightly darker. Nuclei and inclusions (chromatoidal bars, RBCs, bacteria, and Charcot-Leyden crystals) are dark gray-blue, sometimes almost black. The background material usually stains pale gray or blue, providing some color intensity contrast with the protozoa. This contrast is less distinct than that obtained with the trichrome stain, which tends to stain everything with multiple colors (pink, red, purple, green, blue).
7. The microscope should be calibrated (within the last 12 months) (recommended but not always required, depending on the use and care of the microscope), 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 Iron Hematoxylin Stain with Mercury-Based Fixatives
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. Prepare the slide for staining as previously described (for SAF smears or smears prepared from other non-mercury single-vial preservatives, proceed to step 4).
2. Place the slide in 70% ethanol for 5 min.
3. Place the slide in the iodine-70% ethanol (70% alcohol to which is added enough D’Antoni’s iodine to obtain a strong-tea color) solution for 2 to 5 min. The iodine is designed to remove the mercury from the smear.
4. Place it in 70% ethanol for 5 min (this rinse step removes the iodine). Begin the procedure for SAF-fixed slides at this point.*
5. Wash the slide in running tap water (constant stream of water into the container) for 10 min.
6. Place the slide in iron hematoxylin working solution for 4 to 5 min.
7. Wash the slide in running tap water (constant stream of water into the container) for 10 min.
8. Place the slide in 70% ethanol for 5 min.*
9. Place the slide in 95% ethanol for 5 min.*
10. Place the slide in two changes of 100% ethanol for 5 min each.*
11. Place the slide in two changes of xylene for 5 min each.*
12. Add Permount to the stained area of the slide, and cover with a coverslip.
Note An alternative method to using mounting medium is given on p. 53.
13. Examine the smear microscopically with the 100× objective. Examine at least 200 to 300 oil immersion fields before reporting a negative result.
*Slides may be held for up to 24 h in these solutions without harming the quality of the smear or stainability of organisms.
Procedure for Iron Hematoxylin Stain with Non-Mercury-Based Fixatives
1. Prepare the slide for staining as described above.
2. Place it in 70% ethanol for 5 min.*
3. Wash the slide in running tap water (constant stream of water into the container) for 10 min.
4. Place the slide in iron hematoxylin working solution for 4 to 5 min.
5. Wash the slide in running tap water (constant stream of water into the container) for 10 min.
6. Place the slide in 70% ethanol for 5 min.*
7. Place the slide in 95% ethanol for 5 min.*
8. Place the slide in two changes of 100% ethanol for 5 min each.*
9. Place the slide in two changes of xylene for 5 min each.*
10. Add Permount to the stained area of the slide and cover it with a coverslip.
Note An alternative method to using mounting medium is given on p. 53.
11. Examine the smear microscopically with the 100× objective. Examine at least 200 to 300 oil immersion fields before reporting a negative result.
*Slides may be held for up to 24 h in these solutions without harming the quality of the smear or stainability of organisms.
Results and Patient Reports from the Iron Hematoxylin Staining Method
Protozoan trophozoites and cysts are readily visible, although helminth eggs and larvae may not be easily identified because of excess stain retention (wet smears from the concentration procedure[s] are recommended for detection of these organisms). Yeasts (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).
| 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 coli trophozoites
Dientamoeba fragilis trophozoites
2. Quantitate the number of Blastocystis spp. organisms seen (rare, few, moderate, many). Do not quantitate other protozoa.
Example: Many Blastocystis spp.
3. Note and quantitate the presence of human cells.
Example: Moderate WBCs, few macrophages, few RBCs, rare Charcot-Leyden crystals
4. Report and quantitate yeast cells.
Example: Many 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 hold their negative smears for several weeks and then discard them; slide boxes that are rotated can be used to batch, store, and discard negative smears.
Procedure Notes for the Iron Hematoxylin 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 gray or blue-gray 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 slides can also be drained against the edge of each container before being moved to the next container.
3. Incomplete removal of mercuric chloride (Schaudinn’s fixative and PVA fixative prepared with a mercuric chloride base) 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 (used for the removal of iodine) can be eliminated from the procedure. The smears for staining can be prerinsed with 70% alcohol and then placed in the water step prior to the hematoxylin stain as the first step in the staining protocol.
5. For staining large numbers of slides, the working hematoxylin solution may be diluted and affect the quality of the stain. If dilution occurs, discard the working solution and prepare a fresh working solution.
6. The shelf life of the stock hematoxylin solutions may be extended by keeping the solutions in the refrigerator at 4°C. Because of crystal formation in the working solutions, it may be necessary to filter them before preparing a new working solution.
7. In the final stages of dehydration (steps 9 to 11), the 100% ethanol and the xylenes 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.
8. If the smears peel or flake off, the specimen might have been inadequately dried on the slide (in the case of fixed 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.
9. 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 iron hematoxylin stain for 8 min, and complete the staining procedure (2, 3).
Procedure Limitations for the Iron Hematoxylin Staining Method
1. The permanent stained smear is not recommended for staining helminth eggs or larvae; these structures 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, 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 preserved stool containing PVA).
6. Cryptosporidium and Cyclospora oocysts are generally not recognized on an iron hematoxylin-stained smear (modified acid-fast stains or the fecal immunoassay for Cryptosporidium spp. is recommended).
Iron Hematoxylin Stain (Tompkins-Miller Method) (20)
A longer iron hematoxylin method was described by Tompkins and Miller (20). Since differentiation of overstained slides is critical in most iron hematoxylin staining procedures, Tompkins and Miller have described a method that employs phosphotungstic acid to destain the protozoa and that gives excellent results, even in unskilled hands.
1. Prepare the slide for staining as described above (for SAF or non-mercury-based smears, proceed to step 4).
2. Place the slide in 70% ethanol for 5 min.
3. Place the slide in the iodine−70% ethanol (70% alcohol to which is added enough D’Antoni’s iodine to obtain a strong-tea color) solution for 2 to 5 min.
4. Place it in 50% ethanol for 5 min. Begin the procedure for SAF- or non-mercury-fixed slides at this point.*
5. Wash the slide in running tap water (constant stream of water into the container) for 3 min.
6. Place the slide in 4% ferric ammonium sulfate mordant for 5 min.
7. Wash the slide in running tap water (constant stream of water into the container) for 1 min.
8. Place the slide in 0.5% aqueous hematoxylin for 2 min.
9. Wash the slide in tap water for 1 min.
10. Place the slide in 2% phosphotungstic acid for 2 to 5 min.
11. Wash the slide in running tap water for 10 min.
12. Place the slide in 70% ethanol (plus a few drops of saturated aqueous lithium carbonate) for 3 min.
13. Place the slide in 95% ethanol for 5 min.*
14. Place the slide in two changes of 100% ethanol for 5 min each.*
15. Place the slide in two changes of xylene for 5 min each.*
16. Add Permount to the stained area of the slide, and cover it with a coverslip.
Note An alternative method to using mounting medium is as follows.
A. Remove the slide from the last container of xylene, place it on a paper towel (flat position), and allow it to air dry. Remember that some of the xylene substitutes may take a bit longer to dry.
B. Approximately 5 to 10 min before you want to examine the slide, place a drop of immersion oil on the dry fecal film. Allow the oil to sink into the film for a minimum of 10 to 15 min. If the smear appears to be very refractile on examination, you have not waited long enough for the oil to sink into the film or you need to add a bit more oil onto the film.
C. Once you are ready to examine the slide, place a no. 1 (22- by 22-mm) coverslip onto the oiled smear, add another drop of immersion oil to the top of the coverslip (as you would normally do for any slide with a coverslip), and examine with the oil immersion lens (100× objective).
D. Do not eliminate adding the coverslip; the dry fecal material on the slide often becomes very brittle after dehydration. Without the addition of the protective coverslip, you might scratch the surface of your oil immersion lens. Coverslips are much cheaper than oil immersion objectives!
17. Examine the smear microscopically with the 100× objective. Examine at least 200 to 300 oil immersion fields before reporting a negative result.
*Slides may be held for up to 24 h in these solutions without harming the quality of the smear or stainability of organisms.
Modified Iron Hematoxylin Stain (Incorporating the Carbol Fuchsin Step)
The following combination staining method for SAF-preserved fecal specimens was developed to allow the microscopist to screen for acid-fast organisms in addition to other intestinal parasites (Fig. 3.21). For laboratories using iron hematoxylin stains in combination with SAF-fixed material and modified acid-fast stains for Cryptosporidium, Cyclospora, and Cystoisospora, this modification represents another staining option (49). This combination stain provides a saving in both time and personnel use. However, some laboratories prefer to use these two stains as single methods; the overall result may be better than the combination approach. The selection of one approach or the other would be up to the laboratory and would be based on personal preferences.
Figure 3.21 Iron hematoxylin stain incorporating the carbol fuchsin step. Note the modified acid-fast Cryptosporidium oocysts (4–6 µm) and the Giardia lamblia cyst. doi:10.1128/9781555819002.ch3.f21
Any fecal specimen submitted in SAF, other non-mercury single-vial-system preservatives, or the Universal Fixative can be used. Fresh fecal specimens after fixation in SAF for 30 min can also be used. This combination stain approach is not recommended for specimens preserved in Schaudinn’s fixative using a mercuric chloride base (with or without PVA).
Mayer’s Albumin
Add an equal quantity of glycerin to a fresh egg white. Mix gently and thoroughly. Store at 4°C, and indicate an expiration date of 3 months. Mayer’s albumin from commercial suppliers can normally be stored at 25°C for 1 year.
Stock Solution of Hematoxylin Stain
1. Mix well until dissolved.
2. Store in a clear-glass bottle in a light area. Allow to ripen for 14 days before use.
3. Store at room temperature with an expiration date of 1 year.
Mordant
Ferrous ammonium sulfate
Ferric ammonium sulfate
Working Solution of Hematoxylin Stain
1. Mix equal quantities of stock solution of stain and mordant.
2. Allow the mixture to cool thoroughly before use (prepare at least 2 h prior to use). The working solution should be made fresh every week.
Picric Acid
Mix equal quantities of distilled water and an aqueous saturated solution of picric acid to make a 50% saturated solution.
Add enough ammonia to bring the pH to approximately 8.0.
Carbol Fuchsin
1. To make basic fuchsin (solution A), dissolve 0.3 g of basic fuchsin in 10 ml of 95% ethanol.
2. To make phenol (solution B), dissolve 5 g of phenol crystals in 100 ml of distilled water. (Gentle heat may be needed.)
3. Mix solution A with solution B.
4. Store at room temperature. The solution is stable for 1 year.
Procedure for Modified Iron Hematoxylin Stain (Carbol Fuchsin Step)
1. Prepare slide.
A. Place 1 drop of Mayer’s albumin on a labeled slide.
B. Thoroughly mix the sediment from the fecal concentration with an applicator stick.
C. Add approximately 1 drop of the fecal concentrate to the albumin, and spread the mixture over the slide.
2. Allow the slide to air dry at room temperature (the smear appears opaque when dry).
3. Place the slide in 70% alcohol for 5 min.
4. Wash the slide in a container of tap water (not under running water) for 2 min.
5. Place the slide in Kinyoun’s stain for 5 min.
6. Wash the slide in running tap water (constant stream of water into container) for 1 min.
7. Place the slide in acid-alcohol decolorizer for 4 min.*
8. Wash the slide in running tap water (constant stream of water into container) for 1 min.
9. Place the slide in iron hematoxylin working solution for 8 min.
10. Wash the slide in distilled water (in container) for 1 min.
11. Place the slide in picric acid solution for 3 to 5 min.
12. Wash the slide in running tap water (constant stream of water into container) for 10 min.
13. Place the slide in 70% alcohol plus ammonia for 3 min.
14. Place the slide in 95% alcohol for 5 min.
15. Place the slide in 100% alcohol for 5 min.
16. Place the slide in two changes of xylene for 5 min.
*This step can also be performed as follows.
A. Place the slide in acid-alcohol decolorizer for 2 min.
B. Wash the slide in running tap water (constant stream of water into container) for 1 min.
C. Place the slide in acid-alcohol decolorizer for 2 min.
D. Wash the slide in running tap water (constant stream of water into container) for 1 min.
E. Continue the staining sequence with step 9 (iron hematoxylin working solution).
Procedure Notes for Modified Iron Hematoxylin Stain (Carbol Fuchsin Step)
1. The first 70% alcohol step acts with the Mayer’s albumin to “glue” the specimen to the glass slide. The specimen may wash off if insufficient albumin is used or if the slides are too thick and are not completely dry before being stained.
2. The working hematoxylin stain should be checked each day of use by adding a drop of stain to alkaline tap water. If a blue color does not develop, prepare a fresh working stain solution.
3. The picric acid differentiates the hematoxylin stain by removing more stain from fecal debris than from the protozoa and removing more stain from the organism cytoplasm than from the nucleus. When properly stained, the background should be various shades of gray-blue and the protozoa, with medium blue cytoplasm and dark blue-black nuclei, should be easily seen.
Polychrome IV Stain
Polychrome IV stain can be used in place of trichrome for staining fecal smears by the MIF, PVA, or SAF fixative method. Both the stain and staining directions are available commercially (Devetec, Inc., P.O. Box 10275, Bradenton, FL 34282). Another source for the stain is Scientific Device Laboratory, Inc., P.O. Box 88, Glenview, IL 60025. Polychrome IV stain has been used primarily to stain permanent smears prepared from MIF-preserved fecal specimens.
Chlorazol Black E Stain
Chlorazol black E staining, developed by Kohn (9), is a method in which both fixation and staining occur in a single solution. This approach is used for fresh specimens, but it is not recommended for preserved material containing mercuric chloride (6) because it does not include an iodine-alcohol step, which is used to remove the mercuric chloride compound found in Schaudinn’s fixative (with and without PVA) prepared with mercuric chloride. The optimal staining time must be determined for each batch of fixative-stain. The length of time for which the fixative-stain can be used depends on the number of slides run through the solution within a 30-day period. If the slides appear visibly red, the solution must be changed. This stain is not widely used.
Modified Kinyoun’s Acid-Fast Stain (Cold Method)
Cryptosporidium spp. and C. belli have been recognized as causes of severe diarrhea in immunocompromised hosts but can also cause diarrhea in immunocompetent hosts (33). Oocysts in clinical specimens may be difficult to detect without special staining (34). Modified acid-fast stains are recommended to demonstrate these organisms. Unlike the Ziehl-Neelsen modified acid-fast stain, the modified Kinyoun’s stain does not require the heating of reagents for staining (35). With additional reports of diarrheal outbreaks due to Cyclospora spp., it is also important to remember that these organisms are modified acid-fast variable and can be identified by this staining approach. Although the microsporidial spores are also acid fast, their size (1 to 2 µm) makes identification very difficult without special modified trichrome stains or the use of immunoassay reagents (36–41).
Concentrated (500 × g for 10 min) sediment of fresh, formalin-preserved, or other single-vial fixative-preserved stool may be used. Other types of clinical specimens such as duodenal fluid, bile, and pulmonary specimens (induced sputum, bronchial washings, or biopsy specimens) may also be stained (Fig. 3.22).
Figure 3.22 Work flow diagram for fecal specimens (coccidia). The most important part of the procedure would be the concentrate (Cytoisospora sp.) and the permanent stained smear, using one of the modified acid-fast techniques (Cryptosporidium and Cyclospora). doi:10.1128/9781555819002.ch3.f22
50% Ethanol
1. Add 50 ml of absolute ethanol to 50 ml of distilled water.
2. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
Kinyoun’s Carbol Fuchsin
1. Dissolve 4 g of basic fuchsin in 20 ml of 95% ethanol (solution A).
2. Dissolve 8 g of phenol crystals in 100 ml of distilled water (solution B).
3. Mix solutions A and B.
4. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
1% Sulfuric Acid
1. Add 1 ml of concentrated sulfuric acid to 99 ml of distilled water.
2. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
Loeffler Alkaline Methylene Blue
1. Dissolve 0.3 g of methylene blue in 30 ml of 95% ethanol.
2. Add 100 ml of dilute (0.01%) potassium hydroxide.
3. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
Quality Control for Kinyoun’s Acid-Fast Stain
1. A control slide of Cryptosporidium from a 10% formalin-preserved specimen is included with each staining batch run. If the Cryptosporidium slide stains well, any Cystoisospora or Cyclospora oocysts present will also take up the stain, although Cyclospora oocysts tend to be acid-fast variable (Fig. 3.23).
2. Cryptosporidium spp. stain pink-red. Oocysts measure 4 to 6 µm, and four sporozoites may be visible within some oocysts. The background should stain uniformly blue.
3. The specimen is also checked for adherence to the slide (macroscopically).
4. 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). If the microscopes receive adequate maintenance and are not moved frequently, yearly recalibration may not be necessary.
5. Known positive microscope slides and photographs (reference books) should be available at the workstation.
6. Record all QC results; the laboratory should also have an action plan for “out-of-control” results.
Figure 3.23 Quality control slides for performing modified acid-fast stains (Cryptosporidium and Cyclospora) (Medical Chemical Corp.). doi:10.1128/9781555819002.ch3.f23
Procedure for Kinyoun’s Acid-Fast Stain
1. Smear 1 to 2 drops of concentrated specimen sediment on the slide, and allow it to air dry. Do not make the smears too thick (you should be able to see through the wet material before it dries). Prepare two smears.
2. Fix with absolute methanol for 1 min.
3. Flood the slide with Kinyoun’s carbol fuchsin and stain for 5 min.
4. Rinse the slide briefly (3 to 5 s) with 50% ethanol.
5. Rinse the slide thoroughly with water.
6. Decolorize with 1% sulfuric acid for 2 min or until no more color runs from the slide.
7. Rinse the slide with water. Drain.
8. Counterstain with methylene blue for 1 min.
9. Rinse the slide with water. Air dry.
10. Examine the slide with the low or high dry objective. To see the internal morphology, use an oil objective (100×).
Results and Patient Reports from Kinyoun’s Acid-Fast Staining Method
The oocysts of Cryptosporidium, Cyclospora, and Cystoisospora spp. stain pink to red to deep purple. Some of the four sporozoites may be visible in the Cryptosporidium oocysts. Some of the Cystoisospora immature oocysts (entire oocyst) will stain, while those that are mature usually appear with the two sporocysts within the oocyst wall stained pink to purple and a clear area between the stained sporocysts and the oocyst wall. The background stains blue. If Cyclospora oocysts are present (uncommon unless in an outbreak situation), they tend to be approximately 8 to 10 µm, they resemble Cryptosporidium spp. but are larger, and they have no definite internal morphology; the acid-fast staining tends to be more variable than that seen with Cryptosporidium or Cystoisospora spp. (Fig. 3.24 and 3.25). Some of the Cyclospora oocysts tend to look like “wrinkled cellophane,” with tremendous variation in color intensity ranging from clear to pink to red to deep purple to almost black. The stain intensity also depends on the thickness of the smear, the percentage of acid in the decolorizer (1% sulfuric acid recommended), and the length of time the smear is in contact with the decolorizer. If the patient has a heavy infection with microsporidia (immunocompromised patient), small (1- to 2-µm) spores may be seen but may not be recognized as anything other than bacteria or small yeast cells.
Figure 3.24 Cyclospora cayetanensis oocysts (8–10 µm). (Top) Autofluorescent oocysts; (bottom) oocysts stained with modified acid-fast stain. Note the range of color intensity; some oocysts resemble “wrinkled cellophane.” doi:10.1128/9781555819002.ch3.f24
Figure 3.25 (Left) Cyclospora (8 to 10 µm), Cryptosporidium (4 to 6 µm), and artifact (∼2 µm) stained with modified acid-fast stain. Note that one of the Cyclospora oocysts did not stain (modified acid-fast variable) and has the “wrinkled-cellophane” appearance, while the other oocyst displays the typical purple/red color; the Cryptosporidium oocyst and the artifact did stain modified acid-fast positive. These artifacts are frequently seen; it is very important to measure objects seen in the modified acid-fast smears to confirm that they are actual parasites and not artifacts. (Right) Cyclospora stained with a hot safranin stain (higher magnification than the image on the left). (Courtesy of CDC.) doi:10.1128/9781555819002.ch3.f25
There is usually a range of color intensity in the organisms present; not every oocyst appears deep pink to purple. The greatest staining variation will be seen with Cyclospora oocysts; do not decolorize too long.
1. Report the organism and stage (oocyst). Do not use abbreviations.
Examples: Cryptosporidium spp. oocysts
Cystoisospora belli oocysts
2. Call the physician when these organisms are identified.
3. Save positive slides for future reference. Label prior to storage (name, patient number, organisms present).
Procedure Notes for Kinyoun’s Acid-Fast Staining Method
1. Routine stool examination stains (trichrome and iron hematoxylin) are not recommended; however, sedimentation concentration (500 × g for 10 min) is recommended for the recovery and identification of the coccidia after the concentration sediment has been stained with one of the modified acid-fast stains. The routine concentration (formalin-ethyl acetate) can be used to recover Cystoisospora oocysts (wet sediment examination and/or modified acid-fast stains), but routine permanent stains (trichrome and iron hematoxylin) are not reliable for this purpose.
2. Preserved specimens containing PVA are not acceptable for staining with the modified acid-fast stain. However, specimens preserved in SAF, other non-mercury single-vial-system fixatives, or those preserved in Universal Fixative are perfectly acceptable.
3. Avoid the use of wet-gauze filtration (an old, standardized method of filtering stool prior to centrifugation) with too many layers of gauze that may trap organisms and not allow them to flow into the fluid to be concentrated. It is recommended that no more than two layers of gauze be used; another option is to use the commercially available concentrators that use plastic or metal screens instead of gauze.
4. Other organisms that stain positive include acid-fast bacteria, Nocardia spp., and the microsporidia (which are very difficult to find and identify even when they appear to be acid fast).
5. It is very important that smears not be too thick. Thicker smears may not adequately destain.
6. Concentration of the specimen is essential to demonstrate organisms. The number of organisms seen in the specimen may vary from many to very few; therefore, the organisms are concentrated in the sediment, which is used to prepare the smears for subsequent staining.
7. Some specimens require treatment with 10% KOH because of their mucoid consistency. Add 10 drops of 10% KOH to the sediment, and vortex until homogeneous. Rinse with 10% formalin, and centrifuge (500 × g for 10 min). Without decanting the supernatant, take 1 drop of the sediment and smear it thinly on a slide.
8. Commercial concentrators and reagents are available. (See Appendix 1.)
9. Concentrations of sulfuric acid of 1.0 to 3.0% are normally used. Higher concentrations remove too much stain. The use of acid-alcohol (routinely used in the Ziehl-Neelsen acid-fast staining method for the mycobacteria) decolorizes all organisms; therefore, one must use the modified decolorizer (1 to 3% H2SO4) for good results. In general, 1% acid is recommended; this approach will provide excellent staining results for all the coccidia.
10. There is some debate whether organisms lose their ability to take up the acid-fast stain after long-term storage in 10% formalin. Some laboratories have reported this diminished staining.
11. Specimens should be centrifuged in capped tubes, and gloves should be worn during all phases of specimen processing.
Procedure Limitations for Kinyoun’s Acid-Fast Staining Method
1. Light infections with Cryptosporidium spp. may be missed (small number of oocysts). The fecal immunoassay methods are more sensitive.
2. Multiple specimens must be examined, since the numbers of oocysts present in the stool vary from day to day. A series of three specimens submitted on alternate days is recommended.
3. Cyclospora may be suspected if the organisms appear to be Cryptosporidium but are about twice the size (about 10 µm) (Fig. 3.25). The microsporidial spores are extremely small (1 to 2 µm) and will probably not be recognized unless they are very numerous and appear to have a somewhat different morphology from the bacteria in the preparation.
Modified Ziehl-Neelsen Acid-Fast Stain (Hot Method)
Cryptosporidium and Cystoisospora have been recognized as causes of severe diarrhea in immunocompromised hosts but can also cause diarrhea in immunocompetent hosts. Oocysts in clinical specimens may be difficult to detect without special staining. Modified acid-fast stains are recommended to demonstrate these organisms. Application of heat to the carbol fuchsin assists in the staining, and the use of a milder decolorizer allows the organisms to retain their pink-red color (37). With continued reports of diarrheal outbreaks due to Cyclospora, it is also important to remember that these organisms are acid fast and can be identified by using this staining approach (42, 43). Although the microsporidial spores are also acid fast, their size (1 to 2 µm) makes identification very difficult without special stains or the use of monoclonal antibody reagents.
Concentrated sediment of fresh, formalin, other non-mercury single-vial fixative-preserved stool, or those preserved in Universal Fixative may be used. Other types of clinical specimens such as duodenal fluid, bile, and pulmonary specimens (induced sputum, bronchial washings, or biopsy specimens) may also be stained.
Carbol Fuchsin
1. To make basic fuchsin (solution A), dissolve 0.3 g of basic fuchsin in 10 ml of 95% ethanol.
2. To make phenol (solution B), dissolve 5 g of phenol crystals in 100 ml of distilled water. (Gentle heat may be needed.)
3. Mix solution A with solution B.
4. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
1–3% Sulfuric Acid
1. Add 1 to 3 ml of concentrated sulfuric acid to 99 or 97 ml of distilled water.
2. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
Methylene Blue
1. Dissolve 0.3 g of methylene blue chloride in 100 ml of distilled water.
2. Store at room temperature. The solution is stable for 1 year. Note the expiration date on the label.
Quality Control for the Modified Ziehl-Neelsen Acid-Fast Staining Method
QC guidelines are the same as those for the Kinyoun’s acid-fast stain and are given on p. 53.
Procedure for the Modified Ziehl-Neelsen Staining Method
1. Smear 1 to 2 drops of specimen on the slide, and allow it to air dry. Do not make the smears too thick (you should be able to see through the wet material before it dries). Prepare two smears.
2. Dry on a heating block (70°C) for 5 min.
3. Place the slide on a staining rack, and flood it with carbol fuchsin.
4. With an alcohol lamp or Bunsen burner, gently heat the slide to steaming by passing a flame under the slide. Discontinue heating once the stain begins to steam. Do not boil.
5. Allow the specimen to stain for 5 min. If the slide dries, add more stain without additional heating.
6. Rinse thoroughly with water. Drain.
7. Decolorize with 1–3% sulfuric acid for 30 s. (Thicker slides may require a longer destain.)
8. Rinse the slide with water. Drain.
9. Flood the slide with methylene blue for 1 min.
10. Rinse the slide with water, drain, and air dry.
11. Examine with low or high dry objective. To see internal morphology, use the oil objective (100×).
Results and Patient Reports from the Modified Ziehl-Neelsen Acid-Fast Staining Method
The oocysts of Cryptosporidium and Cystoisospora spp. stain pink to red to deep purple. Some of the four sporozoites may be visible in the Cryptosporidium oocysts. Some of the Cystoisospora immature oocysts (entire oocyst) stain, while those that are mature usually appear with the two sporocysts within the oocyst wall stained a pink to purple color and a clear area between the stained sporocysts and the oocyst wall (Fig. 3.26). The background stains blue. If Cyclospora oocysts are present (uncommon), they tend to be approximately 8 to 10 µm, they resemble Cryptosporidium oocysts but are larger, and they have no definite internal morphology; the acid-fast staining tends to be more variable than that seen with Cryptosporidium or Cystoisospora spp. If the patient has a heavy infection with microsporidia (immunocompromised patient), small (1- to 2-µm) spores may be seen but may not be recognized as anything other than bacteria or small yeast cells.
Figure 3.26 (Upper) Immature Cystoisospora belli oocyst stained with modified acid-fast stain. (Lower) C. belli immature oocyst (left) (note that the entire oocyst retains the stain) and C. belli mature oocyst containing two sporocysts (right). doi:10.1128/9781555819002.ch3.f26
There is usually a range of color intensity in the organisms present; not every oocyst appears deep pink to purple. The greatest staining variation would be seen with Cyclospora organisms.
1. Report the organism and stage (oocyst). Do not use abbreviations.
Examples: Cryptosporidium spp. oocysts
Cystoisospora belli oocysts
Cyclospora cayetanensis oocysts
2. Call the physician when these organisms are identified.
3. Save positive slides for future reference. Label prior to storage (name, patient number, organisms present).
Procedure Notes for the Modified Ziehl-Neelsen Acid-Fast Staining Method
1. Routine stool examination stains (trichrome and iron hematoxylin) are not recommended; however, sedimentation concentration (500 × g for 10 min) is acceptable for the recovery and identification of the coccidia, particularly after the concentration sediment has been stained with one of the modified acid-fast stains. The routine concentration (formalin-ethyl acetate) can be used to recover Cystoisospora oocysts (wet sediment examination and/or modified acid-fast stains), but routine permanent stains (trichrome and iron hematoxylin) are not reliable for this purpose.
2. Preserved specimens containing PVA are not acceptable for staining with the modified acid-fast stain. However, specimens preserved in SAF, other single-vial preservatives, or the Universal Fixative are perfectly acceptable.
3. Avoid the use of wet-gauze filtration (an old, standardized method of filtering stool prior to centrifugation) with too many layers of gauze that may trap organisms and not allow them to flow into the fluid to be concentrated. It is recommended that no more than two layers of gauze be used. Another option is to use the commercially available concentration systems in which metal or plastic screens are used for filtration.
4. Other organisms that stain positive include acid-fast bacteria, Nocardia spp., and the microsporidia (which are very difficult to find and identify even when they appear to be acid fast).
5. It is very important that smears not be too thick. Thicker smears may not adequately destain.
6. Concentration of the specimen is essential to demonstrate organisms. The number of organisms seen in the specimen may vary from numerous to very few. Therefore, the sensitivity of the method is enhanced if the stains are performed on concentrated sediment.
7. Some specimens require treatment with 10% KOH because of their mucoid consistency. Add 10 drops of 10% KOH to the sediment, and vortex until homogeneous. Rinse with 10% formalin, and centrifuge (500 × g for 10 min). Without decanting the supernatant, take 1 drop of the sediment and smear it thinly on a slide.
8. Commercial concentrators and reagents are available.
9. Do not boil the stain. Gently heat until steam rises from the slide. Do not allow the stain to dry on the slide.
10. Various concentrations of sulfuric acid (0.25 to 10%) may be used; the destaining time varies according to the concentration used. Generally, a 1 or 3% solution is used. The use of acid-alcohol (routinely used in the Ziehl-Neelsen acid-fast staining method for the mycobacteria) decolorizes all organisms; therefore, the modified decolorizer must be used for good results.
11. There is some debate whether organisms lose their ability to take up the acid-fast stain after long-term storage in 10% formalin. Some laboratories have reported this diminished staining.
12. Specimens should be centrifuged in capped tubes, and gloves should be worn during all phases of specimen processing.
Procedure Limitations for the Modified Ziehl-Neelsen Acid-Fast Staining Method
1. Light infections with Cryptosporidium or Cyclospora may be missed (small number of oocysts). When available, fecal immunoassay methods for Cryptosporidium are more sensitive.
2. Multiple specimens must be examined, since the numbers of oocysts present in the stool vary from day to day. A series of three specimens submitted on alternate days is recommended.
3. The identification of both Cyclospora organisms and microsporidia may be difficult. Cyclospora may be suspected if the organisms appear to be Cryptosporidium but are about twice the size (about 8 to 10 µm). The microsporidial spores are extremely small (1 to 2 µm) and will probably not be recognized unless they are very numerous and appear to have a somewhat different morphology from the bacteria in the preparation.
4. Often, artifact material may be seen in these stained smears (Fig. 3.25). The artifacts may resemble the oocysts of Cryptosporidium or Cyclospora; therefore, it is very important that any “parasites” seen in the stained smears be measured for confirmation.
There are three other stains that can be used for the coccidia, although they may not be as common as the Kinyoun or hot acid-fast method. They are the carbol fuchsin negative stain, the safranin stain, and the auramine O stain.
Carbol Fuchsin Negative Stain for Cryptosporidium (from W. L. Current)
1. Mix thoroughly an equal volume (3 to 10 µl) of fresh or formalin-fixed stool and Kinyoun’s carbol fuchsin on a slide.
2. Spread out as a thin film, and allow to air dry at room temperature.
3. Add immersion oil directly to the stained smear, and then cover with a coverslip.
4. Observe under bright-field microscopy (×400). Everything but the oocysts stains darkly. The oocysts are bright and refractile because they contain water and everything else is oil soluble.
Rapid Safranin Method for Cryptosporidium (36)
1. Smear fresh or formalin-fixed feces on a slide, and allow the film to air dry at room temperature.
2. Fix briefly with one pass through the Bunsen burner flame.
3. Fix for 3 to 5 min with 3% HCl in methanol.
4. Wash with tap water (brief rinse).
5. Stain with 1% aqueous safranin for 1 min (heat until steam appears) (the authors indicate by personal communication that boiling may be beneficial).
6. Rinse in tap water.
7. Counterstain with 1% methylene blue for 30 s (0.1% aqueous crystal violet was almost as good, but malachite green was unsatisfactory).
Rapid Safranin Method for Cyclospora, Using a Microwave Oven (40, 44)
Another rapid safranin method uniformly stains Cyclospora oocysts a brilliant reddish orange. However, the fecal smears must be heated in a microwave oven before being stained. This stain is fast, reliable, and easy to perform (40) (Fig. 3.25).
1. Using a 10-µl aliquot of concentrated stool, prepare the smear by spreading the material thinly across the slide.
2. Allow the smear to dry on a 60°C slide warmer.
3. Cool the slide to room temperature before staining.
4. Place the slide in a Coplin jar containing acidic alcohol (3% [vol/vol] HCl in methanol), and let it stand for 5 min.
5. Wash off excess acidic alcohol with cold tap water.
6. Place the slide(s) into the Coplin jar containing the safranin solution in acidified water (pH 6.5), and microwave at full power (650 W) for 1 min. (Place the staining jar in another container to catch the overflow of stain because of boiling.)
7. Wash off excess stain with cold tap water.
8. Place the slide(s) for 1 min in a Coplin jar containing 1% methylene blue.
9. Rinse gently with cold tap water.
10. Air dry.
11. Add a coverslip to the slide by using Cytoseal 60 or other mounting medium; the immersion oil mounting method can also be used.
12. Examine the smear with low-power or high dry power objectives. To see additional morphology, use the oil immersion objective (100×).
Auramine O Stain for Coccidia (from Thomas Hänscheid)
Coccidia are acid-fast organisms and also stain well with auramine O (phenolized auramine O). The size and typical appearance of Cryptosporidium, Cyclospora, and Cystoisospora oocysts enable auramine O-stained slides to be examined at low power using the 10× objective. The entire sample area can usually be examined in less than 30 s. The low cost of the reagents, the simple staining protocol, and the rapid microscopic examination also make this staining method suitable for screening unconcentrated fecal specimens.
Concentrated sediment from fresh or non-PVA-preserved stool may be used. Other stool samples may also be used, such as unconcentrated stool submitted for culture in a bacteriology transport medium. However, to increase the sensitivity of the test, small numbers of oocysts are more easily detected using concentrated stools.
Auramine O Stain
Solution 1
Solution 2
Combine solutions 1 and 2. Store in a dark bottle at room temperature for up to 3 months.
Destaining Agent (0.5% Acid-Alcohol)
Store at room temperature for up to 3 months.
Counterstain (0.5% Potassium Permanganate)
Quality Control for the Auramine O Stain for Coccidia
QC guidelines are the same as those for the Kinyoun’s acid-fast stain and are given on p. 53.
Procedure for the Auramine O Stain for Coccidia
1. Using a 10- to 20-µl aliquot of concentrated stool, prepare the smear by spreading the material across the slide.
2. Heat fix the slides either on a 65 to 75°C heat block for at least 2 h or with the flame of a Bunsen burner. However, do not overheat. Another fixation option would be to fix the slide in absolute methanol for 1 min, air dry, and then proceed with staining.
3. Cool the slide to room temperature before staining.
4. Flood the slide with the phenolized auramine O solution.
5. Allow the smear to stain for ca. 15 min. Do not heat.
6. Rinse the slide in water. Drain excess water from the slide.
7. Flood the slide with the destaining solution (0.5% acid-alcohol).
8. Allow to decolorize for 2 min.
9. Flood with counterstain (potassium permanganate) solution.
10. Stain for 2 min. The timing of this step is critical.
11. Rinse the slide in water. Drain excess water from the slide.
12. Allow the smear to air dry. Do not blot.
13. Examine the smear with a fluorescence microscope with a 10× objective and fluorescein isothiocyanate optical filters (auramine O: excitation max., ∼435 nm in water; emission max., ∼510 nm). Screen the whole sample area for the presence of fluorescent oocysts. Suspicious objects can be reexamined using a 20× or 100× objective.
14. Smears can be restained by any of the carbol fuchsin (modified acid-fast) staining procedures to allow examination with light microscopy.
Results and Patient Reports from the Auramine O Staining Method
Cryptosporidium and Cyclospora oocysts fluoresce brightly and have a regular round appearance (“starry sky” appearance with the 10× objective). In contrast to the large majority of fluorescent artifacts, the oocysts do not stain homogeneously. Thus, the fluorescence is heterogeneously distributed in the interior of the oocyst (Fig. 3.27). Cystoisospora oocysts fluoresce brightly with three patterns: (i) a more or less brightly but heterogeneously stained interior of the whole oocyst, (ii) one brightly staining sporocyst, or (iii) two brightly staining sporocysts within the oocyst wall.
Figure 3.27 Cryptosporidium spp. oocysts stained with auramine O fluorescent stain (using different filters). doi:10.1128/9781555819002.ch3.f27
1. Report the organism and stage (oocyst). Do not use abbreviations.
Examples: Cryptosporidium spp. oocysts
Cystoisospora belli oocysts
Cyclospora cayetanensis oocysts
2. Call the physician when these organisms are identified.
3. Save positive slides for future reference. Label prior to storage (name, patient number, organisms present). These slides can be kept at room temperature in the dark, and the fluorescence remains stable for up to 3 to 4 weeks.
Procedure Notes for the Auramine O Staining Method
1. It is mandatory that positive control smears be stained and examined each time patient specimens are stained and examined.
2. For best results, examine the auramine O solution for deposits and remove them by filtration or centrifugation. This problem can also be avoided by preparing smaller volumes more frequently.
3. Slides should be observed as soon as possible after staining. However, they can be kept at room temperature in the dark, and fluorescence remains stable for up to 3 to 4 weeks.
Procedure Limitations for the Auramine O Staining Method
1. Light infections might be missed, particularly if unconcentrated stool is used; it is always recommended that concentrated stool sediment be used for staining.
2. Use of the 40× high dry objective often causes a blurred image (fluorescent “halo” around the image, hazy contours), which appears to be the effect of interfering fluorescence from the auramine O stain located outside the plane of focus (Fig. 3.27). The 100× oil immersion objective gives better-quality images. Immersion oils used for light microscopy may be autofluorescent, and special low-fluorescence immersion oil should be used.
3. If the fluorescence is not clear or definitive, a suspicious slide can be restained with a modified acid-fast stain and reexamined using light microscopy and the 100× oil immersion objective.
4. If protected from sunlight, auramine O slides can be kept on the bench at room temperature for up to 2 to 3 weeks, with only minor loss of fluorescence (photo bleaching).
Modified Trichrome Stain for the Microsporidia (Weber—Green) (41)
The diagnosis of intestinal microsporidiosis (Enterocytozoon bieneusi, Encephalitozoon intestinalis) has depended on the use of invasive procedures and subsequent examination of biopsy specimens, often by electron microscopy. However, the need for a practical method for the routine clinical laboratory has stimulated some work in the development of additional methods. Slides prepared from fresh, formalin-fixed, or stools preserved in the Universal Fixative can be stained by a chromotrope-based technique and can be examined under light microscopy. This staining method is based on the fact that stain penetration of the microsporidial spore is very difficult; therefore, the dye content in the chromotrope 2R is greater than that routinely used to prepare Wheatley’s modification of Gomori’s trichrome method, and the staining time is much longer (90 min) (Fig. 3.28 and 3.29) (39, 41). At least several of these stains are available commercially from a number of suppliers.
Figure 3.28 (Left) Microsporidian spores in a nasopharyngeal specimen stained with a Ryan modified trichrome stain; (right) microsporidian spores in stool stained with a Weber modified trichrome stain. The spores range from about 1.5 to 2.0 µm in diameter. Note the horizontal lines, indicating the presence of a polar tubule. doi:10.1128/9781555819002.ch3.f28
Figure 3.29 (Left) Microsporidian spores in a urine sediment, stained with calcofluor white stain; (right) Encephalitozoon intestinalis spores stained with an organism-specific fluorescent antibody reagent. Note that some of the spores are intracellular. A urine specimen tends to be “cleaner” than stool; therefore the spores may be easier to see and identify in urine or any specimen that contains less artifact material than stool. doi:10.1128/9781555819002.ch3.f29
The specimen can be fresh stool or stool that has been preserved in 5 or 10% formalin, SAF, or the newer single-vial-system fixatives (Universal Fixative—no PVA). Actually, any specimen other than tissue thought to contain microsporidia could be stained by this method.
Trichrome Stain (Modified for Microsporidia) (Weber—Green) (41)
*(10 times the normal trichrome stain formula)
1. Prepare the stain by adding 3.0 ml of acetic acid to the dry ingredients. Allow the mixture to stand (ripen) for 30 min at room temperature.
2. Add 100 ml of distilled water. Properly prepared stain is dark purple.
3. Store in a glass or plastic bottle at room temperature. The shelf life is at least 24 months.
Acid-Alcohol
Prepare by combining the two solutions.
Quality Control for the Modified Trichrome Staining Method (Weber—Green)
1. Unfortunately, the only way to perform acceptable QC procedures for the modified trichrome staining method is to use actual microsporidial spores as the control organisms. Obtaining these positive controls may be somewhat difficult. It is particularly important to use the actual organisms, because the spores are difficult to stain and they are very small (1 to 2.5 µm).
2. A QC slide should be included with each run of stained slides, particularly if the staining setup is used infrequently.
3. All staining dishes should be covered to prevent evaporation of reagents (screw-cap Coplin jars or glass lids).
4. Depending on the volume of slides stained, staining solutions must be changed on an as-needed basis.
5. When the smear is thoroughly fixed and the stain is performed correctly, the spores are ovoid and refractile, with the spore wall being bright pinkish red. Occasionally, the polar tube can be seen either as a stripe or as a diagonal line across the spore. The majority of the bacteria and other debris tend to stain green. However, some bacteria and debris still stain red.
6. The specimen is also checked for adherence to the slide (macroscopically).
7. 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). Although recalibration every 12 months may not be necessary, this will vary from laboratory to laboratory, depending on equipment care and use.
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 Modified Trichrome Staining Method (Weber—Green)
1. Using a 10-µl aliquot of unconcentrated (concentrated recommended), preserved liquid stool (5 or 10% formalin or SAF or one of the non-PVA single-vial preservatives), prepare the smear by spreading the material over an area 45 by 25 mm.
2. Allow the smear to air dry.
3. Place the smear in absolute methanol for 5 min.
4. Allow the smear to air dry.
5. Place in trichrome stain for 90 min.
6. Rinse in acid-alcohol for no more than 10 s.
7. Dip slides several times in 95% alcohol. Use this step as a rinse.
8. Place in 95% alcohol for 5 min.
9. Place in 100% alcohol for 10 min.
10. Place in xylene substitute for 10 min.
11. Mount with a coverslip (no. 1 thickness), using mounting medium.
12. Examine smears under oil immersion (×1,000), and read at least 300 fields; the examination time will probably be at least 10 min per slide.
Modified Trichrome Stain for the Microsporidia (Ryan—Blue) (39)
A number of variations to the modified trichrome stain (Weber—green) have been tried in an attempt to improve the contrast between the color of the spores and the background staining. Optimal staining was achieved by modifying the composition of the trichrome solution. This stain is also available commercially from a number of suppliers.
The specimen can be fresh stool or stool that has been preserved in 5 or 10% formalin, SAF, or the newer single-vial-system fixatives. Actually, any specimen other than tissue thought to contain microsporidia could be stained by this method.
Trichrome Stain (Modified for Microsporidia) (Ryan—Blue)
*(10 times the normal trichrome stain formula)
1. Prepare the stain by adding 3.0 ml of acetic acid to the dry ingredients. Allow the mixture to stand (ripen) for 30 min at room temperature.
2. Add 100 ml of distilled water, and adjust the pH to 2.5 with 1.0 M HCl. Properly prepared stain is dark purple. The staining solution should be protected from light.
3. Store in a glass or plastic bottle at room temperature. The shelf life is at least 24 months.
Acid-Alcohol
Prepare by combining the two solutions.
Quality Control for the Modified Trichrome Staining Method (Ryan—Blue)
1. Unfortunately, the only way to perform acceptable QC procedures for this method is to use actual microsporidial spores as the control organisms. Obtaining these positive controls may be somewhat difficult. It is particularly important to use the actual organisms, because the spores are difficult to stain and are very small (1 to 2.5 µm).
2. A QC slide should be included with each run of stained slides, particularly if the staining setup is used infrequently.
3. All staining dishes should be covered to prevent evaporation of reagents (screw-cap Coplin jars or glass lids).
4. Depending on the volume of slides stained, staining solutions must be changed on an as-needed basis.
5. When the smear is thoroughly fixed and the stain is performed correctly, the spores are ovoid and refractile, with the spore wall being bright pinkish red. Occasionally, the polar tube can be seen either as a stripe or as a diagonal line across the spore. The majority of the bacteria and other debris tend to stain blue. However, some bacteria and debris still stain red.
6. The specimen is also checked for adherence to the slide (macroscopically).
7. 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). Although recalibration every 12 months may not be necessary, this will vary from laboratory to laboratory, depending on equipment care and use.
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 the Modified Trichrome Staining Method (Ryan—Blue)
1. Using a 10-µl aliquot of concentrated (10 min at 500 × g), preserved liquid stool (5 or 10% formalin, SAF, one of the non-PVA single-vial preservatives, zinc based, or Universal Fixative), prepare the smear by spreading the material over an area 45 by 25 mm.
2. Allow the smear to air dry.
3. Place the smear in absolute methanol for 5 or 10 min.
4. Allow the smear to air dry.
5. Place in trichrome stain for 90 min.
6. Rinse in acid-alcohol for no more than 10 s.
7. Dip the slides several times in 95% alcohol. Use this step as a rinse (no more than 10 s).
8. Place in 95% alcohol for 5 min.
9. Place in 95% alcohol for 5 min.
10. Place in 100% alcohol for 10 min.
11. Place in xylene substitute for 10 min.
12. Mount with a coverslip (no. 1 thickness), using mounting medium.
13. Examine smears under oil immersion (×1,000), and read at least 300 fields; the examination time will probably be at least 10 min per slide.
Results and Patient Reports from Modified Trichrome Staining Methods (Weber or Ryan)
The microsporidial spore wall should stain pinkish to red, with the interior of the spore being clear or perhaps showing a horizontal or diagonal stripe which represents the polar tube. The background appears green or blue, depending on the method. Bacteria, some yeast cells, and some debris stain pink to red; the shapes and sizes of the various components may be helpful in differentiating the spores from other structures. The results of this staining procedure should be reported only if the positive control smears are acceptable. The production of immunoassay reagents should provide a more specific and sensitive approach to the identification of the microsporidia in fecal specimens.
1. Report the organism.
Examples: Microsporidial spores present. The following information can be added to the report to assist the physician in treating and following the patient:
The organisms are most probably Enterocytozoon bieneusi or Encephalitozoon intestinalis (if from fecal specimen or urine).
The organisms are most probably Encephalitozoon intestinalis (identification to species highly likely) (generally the organism involved in disseminated cases from the gastrointestinal tract to other body sites).
Procedure Notes for Modified Trichrome Staining Methods (Weber or Ryan)
1. It is mandatory that positive control smears be stained and examined each time patient specimens are stained and examined.
2. Because of the difficulty in getting the stain to penetrate the spore wall, prepare thin smears and do not reduce the staining time in trichrome. Also, make sure the slides are not left too long in the decolorizing agent (acid-alcohol). If the control organisms are too light, leave them in the trichrome longer and shorten the time to two dips in the acid-alcohol solution. Also, remember that the 95% alcohol rinse after the acid-alcohol should be performed quickly to prevent additional destaining from the acid-alcohol reagent.
3. When you purchase the chromotrope 2R, obtain the highest dye content available. Two sources are Harleco, Gibbstown, NJ, and Sigma Chemical Co., St. Louis, MO (dye content among the highest [85%]). Fast green and aniline blue can be obtained from Allied Chemical and Dye, New York, NY. See also appendix 5.
4. In the final stages of dehydration, the 100% ethanol and the xylenes (or xylene substitutes) 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 or xylene substitute becomes cloudy after addition of slides from 100% alcohol, return the slides to fresh 100% alcohol and also replace the xylene or xylene substitute with fresh stock.
Procedure Limitations for Modified Trichrome Staining Methods (Weber or Ryan)
1. Although this staining method stains the microsporidia, the range of stain intensity and the small size of the spores cause some difficulty in identifying these organisms. Since this procedure results in many other organisms or objects staining in stool specimens, differentiation of the microsporidia from surrounding material is still very difficult. There also tends to be some slight size variation among the spores.
2. If the patient has severe watery diarrhea, there is less artifact material in the stool to confuse with the microsporidial spores; however, if the stool is semiformed or formed, the amount of artifact material is much greater and the spores are much harder to detect and identify. Also, the number of spores varies according to the stool consistency (generally, the more liquid the stool, the more spores will be present). However, remember that there is also a dilution factor if the stool is very liquid.
3. The workers who developed some of these procedures think that concentration procedures result in an actual loss of microsporidial spores; therefore, there is a recommendation to use unconcentrated, formalinized stool. However, there are no data indicating which centrifugation speeds, etc., were used in the study.
4. In the UCLA Clinical Microbiology Laboratory, we have generated data (unpublished) to indicate that centrifugation at 500 × g for 10 min dramatically increases the number of microsporidial spores available for staining (from the concentrate sediment). This is the same method we use for centrifugation of all stool specimens, regardless of the suspected organism.
5. Avoid the use of wet-gauze filtration (an old, standardized method of filtering stool prior to centrifugation) with too many layers of gauze that may trap organisms and not allow them to flow into the fluid to be concentrated. It is recommended that no more than two layers of gauze be used. Another option is to use the commercially available concentration systems in which metal or plastic screens are used for filtration.
Modified Trichrome Stain for the Microsporidia (Kokoskin—Hot Method) (38)
Changes in temperature from room temperature to 50°C and in the staining time from 90 to 10 min have been recommended as improvements for the modified trichrome staining methods. The procedure is as follows.
1. Using a 10-µl aliquot of concentrated, preserved stool (5 or 10% formalin, SAF, or Universal Fixative), prepare the smear by spreading the material over an area 45 by 25 mm.
2. Allow the smear to air dry.
3. Place the smear in absolute methanol for 5 min.
4. Allow the smear to air dry.
5. Place in trichrome stain for 10 min at 50°C.
6. Rinse in acid-alcohol for no more than 10 s.
7. Dip the slide several times in 95% alcohol. Use this step as a rinse (no more than 10 s).
8. Place in 95% alcohol for 5 min.
9. Place in 100% alcohol for 10 min.
10. Place in xylene substitute for 10 min.
11. Mount with a coverslip (no. 1 thickness), using mounting medium.
12. Examine the smear under oil immersion (×1,000) and read at least 300 fields; the examination time will probably be at least 10 min per slide.
Acid-Fast Trichrome Stain for Cryptosporidium and the Microsporidia
The detection of Cryptosporidium spp. and the microsporidia from stool specimens has depended on two separate stains. However, a method is now available that stains both organisms, an important improvement since dual infections have been demonstrated in AIDS patients (44). This acid-fast trichrome stain yields results comparable to those obtained by the Kinyoun and modified trichrome methods and considerably reduces the time necessary for microscopic examination (30, 45, 46) (Fig. 3.30). Also, it appears that modified trichrome stains and staining with fluorochromes are equally useful in the diagnosis of microsporidiosis; however, a combination of the two methods may be more sensitive in cases where the number of spores is very small (41).
Figure 3.30 Cystoisospora (Isospora) belli oocyst and microsporidian spores in an acid-fast trichrome stain. Note the size differential. doi:10.1128/9781555819002.ch3.f30
The specimen can be fresh stool or stool that has been preserved in 5 or 10% formalin, SAF, or some of the newer single-vial-system fixatives. Actually, any specimen other than tissue thought to contain microsporidia could be stained by this method.
Trichrome Stain (Modified for Microsporidia) (44)
*(10 times the normal trichrome stain formula)
1. Prepare the stain by adding 3.0 ml of acetic acid to the dry ingredients. Allow the mixture to stand (ripen) for 30 min at room temperature.
2. Add 100 ml of distilled water, and adjust the pH to 2.5 with 2.0 N HCl. Properly prepared stain is dark purple. The staining solution should be protected from light.
3. Store in a glass or plastic bottle at room temperature. The shelf life is at least 24 months.
Carbol Fuchsin Solution
Phenol solution
Saturated alcoholic fuchsin solution
Add the mixture of phenol and water to 25.0 ml of the saturated alcoholic fuchsin solution.
Acid-Alcohol
Prepare by combining the two solutions.
Quality Control for the Acid-Fast Trichrome Staining Method
1. Unfortunately, the only way to perform acceptable QC procedures for this method is to use actual microsporidian spores as the control organisms. Obtaining these positive controls may be somewhat difficult. It is particularly important to use the actual organisms because the spores are difficult to stain and are very small (1 to 2.5 µm).
2. A QC slide should be included with each run of stained slides, particularly if the staining setup is used infrequently.
3. All staining dishes should be covered to prevent evaporation of reagents (screw-cap Coplin jars or glass lids).
4. Depending on the volume of slides stained, staining solutions must be changed on an as-needed basis.
5. When the smear is thoroughly fixed and the stain is performed correctly, the spores are ovoid and refractile, with the spore wall being bright pinkish red. Occasionally, the polar tube can be seen either as a stripe or as a diagonal line across the spore. The majority of the bacteria and other debris tend to stain blue; however, some bacteria and debris stain red.
6. The specimen is also checked for adherence to the slide (macroscopically).
7. The microscope should be calibrated (within the last 12 months if it has received heavy use), 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). Although recalibration every 12 months may not be necessary, this varies from laboratory to laboratory, depending on equipment care and use.
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 the Acid-Fast Trichrome Staining Method
1. Using a 10-µl aliquot of concentrated (10 min at 500 × g), preserved liquid stool (5 or 10% formalin, SAF, one of the single-vial preservatives, zinc based or Universal Fixative), prepare the smear by spreading the material over an area 45 by 25 mm (47, 48).
2. Allow the smear to air dry.
3. Place the smear in absolute methanol for 5 or 10 min.
4. Allow the smear to air dry.
5. Place in carbol fuchsin solution for 10 min (no heat required).
6. Rinse briefly with tap water.
7. Decolorize with 0.5% acid-alcohol.
8. Briefly rinse with tap water.
9. Place in trichrome stain for 30 min at 37°C.
10. Rinse in acid-alcohol for no more than 10 s.
11. Dip the slide several times in 95% alcohol. Use this step as a rinse (no more than 10 s).
12. Place in 95% alcohol for 30 s.
13. Allow the slide to air dry.
14. Examine the smear under oil immersion (×1,000) and read at least 300 fields; the examination time will probably be at least 10 min per slide.
Results and Patient Reports from the Acid-Fast Trichrome Staining Method
The microsporidial spore wall should stain pink, with the interior of the spore being clear or perhaps showing a horizontal or diagonal stripe that represents the polar tube; a vacuole may also be visible in some spores. The Cryptosporidium oocysts stain bright pink or violet. The results of this staining procedure should be reported only if the positive control smears are acceptable. The production of immunoassay reagents should provide a more specific and sensitive approach to the identification of the microsporidia in fecal specimens.
1. Report the organism.
Examples: Microsporidial spores present. The following information can also be added to the report to assist the physician in treating and following the patient:
The organisms are probably Enterocytozoon bieneusi or Encephalitozoon intestinalis (if from fecal specimens or urine).
The organisms are probably Encephalitozoon intestinalis (identification to species highly likely) (generally the organism involved in disseminated cases from the gastrointestinal tract to kidneys; organisms are recovered in urine).
Procedure Notes for the Acid-Fast Trichrome Staining Method
1. It is mandatory that positive control smears be stained and examined each time patient specimens are stained and examined.
2. Because of the difficulty in achieving stain penetration through the spore wall, prepare thin smears and do not reduce the staining time in trichrome. Also, make sure the slides are not left too long in the decolorizing agent (acid-alcohol). If the control organisms are too light, leave them in the trichrome longer and shorten the time to two dips in the acid-alcohol solution. Also, remember that the 95% alcohol rinse after the acid-alcohol step should be performed quickly to prevent additional destaining from the acid-alcohol reagent.
3. When you purchase the chromotrope 2R, obtain the highest dye content available. Two sources are Harleco, Gibbstown, NJ, and Sigma Chemical Co., St. Louis, MO. (the dye content is among the highest [85%]). Fast green and aniline blue can be obtained from Allied Chemical and Dye, New York, NY. See also appendix 5.
4. In the final stages of dehydration, the 95% ethanol 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.
Procedure Limitations for the Acid-Fast Trichrome Staining Method
1. Although this staining method stains the microsporidia, the range of stain intensity and the small size of the spores may still cause some difficulty in identifying these organisms. Since this procedure results in many other organisms or objects staining in stool specimens, differentiation of the microsporidia from surrounding material is still very difficult. There also tends to be some slight size variation among the spores.
2. If the patient has severe watery diarrhea, there is less artifact material in the stool to confuse with the microsporidial spores; however, if the stool is semiformed or formed, the amount of artifact material is much greater and so the spores are much harder to detect and identify. Also, the number of spores may vary according to the stool consistency (generally, the more liquid the stool, the more spores are present).
3. The workers who developed some of these procedures think that concentration procedures result in an actual loss of microsporidial spores; therefore there is a recommendation to use unconcentrated, formalinized stool. However, there are no data indicating which centrifugation speeds were used in the study.
4. In the UCLA Clinical Microbiology Laboratory, we have generated data (unpublished) to indicate that centrifugation at 500 × g for 10 min dramatically increases the number of microsporidial spores available for staining (from the concentrate sediment). We use the same protocol for centrifugation of all stool specimens, regardless of the suspected organism. The acid-fast trichrome procedure presented here also recommended the use of centrifuged fecal specimens.
5. Avoid the use of wet-gauze filtration (an old, standardized method of filtering stool prior to centrifugation) with too many layers of gauze that may trap organisms and not allow them to flow into the fluid to be concentrated. It is recommended that no more than two layers of gauze be used. Another option is to use the commercially available concentration systems that use metal or plastic screens for filtration.
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The correct answers for Fig 3.17 are as follows: top row left and right, Giardia lamblia (G. duodenalis, G. intestinalis) trophozoite, Chilomastix mesnili cyst; next row left and right, Endolimax nana or Iodamoeba bütschlii trophozoite (both look essentially the same in the trophozoite form), Entamoeba coli cyst; next row left and right, Dientamoeba fragilis trophozoite, Blastocystis spp.; next row left and right, Entamoeba histolytica trophozoite, Blastocystis spp.; bottom row left and right, Endolimax nana cyst, Entamoeba histolytica/E. dispar trophozoite.
