THE MICROSCOPE

THE SEARCH FOR PARASITES IN FECAL SPECIMENS

The pre-analytical phase of a parasitological stool examination, for the microscopic identification of intestinal protozoa and microsporidia, is not very different from that for other parasites (e.g. helminths).  This procedure requires knowledge of:

-     Prior epidemiological data:  place of residence, origin, lifestyle, routine diet and behavior, water supply, elimination and treatment of human and animal wastes, cohabitation with animals, journeys abroad, Unde venis?

-     Prior clinical data. Some parasitoses present with clear signs and symptoms. It is useful to know about the patient's health and the presence of: diarrhea (type, evolution), anal itching, general itching, anorexia, weight loss, vomiting, respiratory disorders, fever (duration, course), hepatosplenomegaly, adenopathy, etc.

-     Life cycle of the parasite. The prepatent period of intestinal protozoa (amoebae and flagellates) can range generally from a few days to 2-3 weeks, whereas for helminths it can vary from a few weeks to several months. The prepatent period is time between host infection by the infective stage of the parasite and the earliest moment in which the parasite can be recovered from feces, blood or other body exudates. The incubation period, instead, is the time between host exposure to the infective stage of the parasite and the manifestation of clinical symptoms of infection.

Preparation of the patient

In the week before stool collection, patients should not take purgative oils or drugs containing charcoal or salts of barium, magnesium or bismuth.  In fact, the presence of these substances makes microscopic examination very difficult. Even antibiotics that inhibit the intestinal bacterial flora may reduce the number of protozoa, since protozoa feed off bacteria.

To make the microscopist's work easier, for three days before and on the same day of stool collection, patients should follow a diet which does not include food that leaves residues that can be confused with parasites (pseudoparasites) or that renders  microscopic preparation of the specimen difficult.  In particular, patients should avoid eating:

-     Legumes;

-     Fruit, especially hard-peel fruits;

-     Corn and cereals;

-     Leafy vegetables that may result in the formation of hard concretions;

-     Mushrooms;

-     Honey;

-     Infusions and phytotherapeutics.

It is also recommended to discontinue all drugs except those necessary to treat serious illnesses.  Any drugs taken should be reported. 

Specimen collection

Stool specimens are collected in clean, dry containers with a large opening and a closing stopper.  Do not collect feces from the toilet because contamination with urine inhibits the motility of trophozoites.  Moreover, the water may contain free-living amoebae, leading to diagnostic errors.

In epidemiological surveys, only one specimen is generally collected, but in other cases this is often not sufficient. Because elimination of parasites is not continuous and may have a “negative period”, it is recommended to collect at least three specimens on non-consecutive days within an interval of 10 days. It is also strongly recommended to not mix the three samples in the same container because, if parasites are present in only one sample, they will be diluted by the other negative samples, significantly decreasing the sensitivity of the examination.

If amoebiasis, giardiasis or microsporidiosis is suspected, it may be necessary to examine several specimens collected over a period of 7-10 days for maximum sensitivity.  Wet mount examination of diarrheal samples should be carried out, if  possible, at once or within 30 minutes of passage, to look for mobile forms of trophozoites. After that period of time, any trophozoites that were present will have become immobile and started to degenerate, and thus it is difficult or even impossible to identify them.

In amoebic colitis, ulcers may form in the rectosigmoid portion of the colon: in this case parasites are more numerous on the surface than inside the stool specimens.  Therefore, it is important to collect and examine, at once (see above), bloodied stools as well as any mucus present.

The recommended time limits for direct examination procedures are:

-     Watery and diarrheic stools:  examine within 30 minutes of passage;

-     Semiformed (soft) stools:  examine within 1 hour of passage;

-     Formed stools:  examine within 24 hours of passage.

If these recommended times are not observed, a portion of the specimen should be placed, at once, in an appropriate fixative and the remainder refrigerated (4° C) until direct examination;  specimens are not be stored in an incubator.

Specimen fixation

The use of fixatives, in which feces are homogenized immediately after passage (by the patient himself), preserves the parasites' exact morphology. Any delay in fixing a specimen leads to the loss of morphological details of the parasites; in the case of permanent staining, the result can be very poor. For any kind of fixative (formalin, SAF, PVA), the ratio that should always be used is three parts fixative to one part specimen.

Macroscopic and microscopic examinations of feces 

This section has the following structure:

-     Macroscopic examination

-     Microscopic examination

      Direct, with saline

      With stains for temporary (wet mount) preparations

     Lugol's iodine solution

     MIF solution

     Bailenger's stain

     Sargeaunt's stain

     Merbromin solution

     Nigrosin solution

      After concentration

     Ritchie's biphasic method

     Junod's method for the concentration of trophozoites

     Sheather's flotation-sedimentation method

      Permanent staining of fecal smears

     Trichrome

     Hematoxylin

     Giemsa, Field

     Modified Ziehl-Neelsen

     Kinyoun

     Trichrome and Giemsa stains for microsporidia

      Fluorescence microscopy (autofluorescence) for detecting coccidian oocysts

Macroscopic examination

This kind of examination is carried out on fixative-free feces.  For proper examination, see the preceding paragraphs about specimen collection.  If mucus, especially bloody mucus, is present, it should be collected to the permit microscopic examination for mobile forms of trophozoites.

Microscopic examination

Direct microscopic examination with saline

This examination is mainly used to evaluate the motility of trophozoites of amoebae and flagellates in diarrheic, watery or loose stools.  It also permits observing helminth eggs and larvae, protozoan cysts, and coccidian oocysts, although these forms are better detected after fecal concentration procedures.

Procedure:

-     Dilute a small quantity of feces (about 2 mg) with physiological saline (R2) on a slide and mix well;

-     With a coverslip held at an angle, touch the edge of the drop and lower the coverslip onto the slide, avoiding the formation of air bubbles.

The suspension has a correct density if it is possible to read a newspaper through it. Bloody mucus should not be diluted but delicately pressed under the coverslip. Carefully examine the whole preparation, particularly the mucous parts, since trophozoites can be trapped in some areas but entirely missing in others, especially in the liquid part or within fecal debris. This is also recommended for rectosigmoidoscopy specimens. 

Adjust the microscope for best interpupillary distance, eyepiece diopter setting and Köhler illumination, and focus on the sample first with the coarse adjustment knob and then with the fine adjustment.  Examine the whole preparation systematically, like a farmer ploughing a field.

The examination at low magnification (10x objective) should be carried out with the light-source voltage regulator at a minimum.  Any forms suspected of being protozoa should be examined at high magnification (40x objective), increasing the intensity of the light source.  The iris diaphragm must not be excessively closed, since this only appears to increase field depth and contrast: in fact, excessively closing of the iris diaphragm drastically reduces the resolution power of the objective.

The wet mount preparation should be examined immediately in order to assess the motility of the trophozoites.  This also avoids evaporation of the fecal suspension and the formation of air bubbles that make microscopic examination difficult. Alternatively, the preparation can be stored for a few hours, sealing the edges of the slide with nail polish to prevent dehydration. Given the small amount of feces examined, the result of the examination may be negative, especially when the parasite load is low.

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Microscopic examination with stains for temporary (wet mount) preparations

Staining with 1% Lugol's (iodine solution)

The sample is prepared as described earlier, although physiological saline is substituted by Lugol's iodine solution (R8). To avoid coagulation of the stool suspension by the Lugol's solution, mix immediately.

This stain gives the best results with fresh, not formalin-fixed feces. The nuclear structures (chromatin and the karyosome) and the fibrils (flagellar remnants) of the flagellates take on a yellowish-brown color and become clearly visible; iodophilic vacuoles or masses stain intensely brown.Food residues also stain, in particular partially digested starch becomes violet or black, while fully digested starch becomes pink. To obtain a deeper contrast, it is recommended to not use excessive iodine solution. As time goes by (<10 min), the preparation tends to dry out, rendering it unreadable. To avoid this, an iodine solution with glycerin (R9) can be used, which maintains the preparation clearly visible  for 2 days.

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Staining with MIF solution

Staining with merthiolate-iodine-formaldehdye (MIF) can be done directly on the slide or in a test tube.

Direct staining on the slide:

-     Dilute about 2 mg feces with saline in order to obtain a thicker preparation than for direct examination;

-     Add a drop of MIF solution (R7-a) and mix well with a coverslip.

Note that MIF staining of specimens fixed in formalin or concentrated with formalin-ether or formalin-ethyl acetate is sometimes of poor quality.

Staining in a test tube:

-     Prepare a test tube containing the MIF working solution (R7-b);

-     Add a pea-sized amount of fresh feces (~0.25 g) and mix well with a glass or plastic rod;

-     Let stand until a sediment forms.  Then, remove a drop of the top layer with a pipette and transfer it to a microscope slide;

-     Seal the test tube containing the specimen.  If it is necessary to examine the specimen again in the next few days, resuspend the sediment by inverting the tube, let it deposit for 15-20 minutes and prepare a new wet mount;

-     MIF solution enables simultaneous fixing and staining of trophozoites, protozoan cysts and helminthic eggs; by increasing the ratio of staining solution to feces, large amounts of specimen can be stored, for example for teaching purposes.

Microscopic examination and interpretation.

Examine the entire preparation with the 10x and 40x objectives. Vegetative forms of amoebae and flagellates take on a yellow or pale brown color, cysts are seen as refractile and colorless areas on a red-pink background, and the nuclear membrane becomes brownish-black. As time passes, the action of the iodine gradually decreases; cysts become pink and the brown staining of iodophilic vacuoles or masses tends to disappear.

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Staining with Bailenger's stain

-     On a slide, mix a drop of fecal suspension (prepared in saline, formalin or SAF) or a drop of sediment obtained by concentration with a small quantity (5 μl!) of Bailenger's stain (R10);

-     Mix well with a coverslip.

Microscopic examination and interpretation.

Examine the preparation with the 10x and 40x objectives. Staining is immediate for trophozoites while cysts, especially mature ones, stain only gradually. The small amount of staining solution does not dilute the preparation and enables slow, gradual staining. Cytoplasm and chromatoid bodies become red-violet, while nuclear structures and fibrils (flagellar remnants) of flagellates become deep violet or black. The nuclei of Dientamoeba fragilis trophozoites do not stain at all.

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Staining with Sargeaunt's stain

This is a stain for chromatoid bodies of amoebic cysts. It should be used only on fecal sediment obtained by the Ritchie's concentration method using ethyl ether (not ethyl acetate).

-     Mix together a drop of fecal sediment and a drop of Sargeaunt's stain (R11) on a slide;

-     Cover with a coverslip.

Microscopic examination and interpretation.

Examine the preparation with the 40x objective. Chromatoid bodies of amoebic cysts will stain dark green while nuclei and cytoplasm will stain light green.

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Staining with merbromin solution

This is a stain for Cryptosporidium oocysts.

-     On a slide, mix a drop of fecal sediment obtained by concentration and a drop of merbromin solution (R12) ;

-     Using a rod, form a thin, uniform smear on about half the slide;

-     Let stand until almost completely dry;

-     Add one or two drops of immersion oil and cover with a coverslip.

Microscopic examination and interpretation.

Examine the preparation with the 40x and 100x objectives. Oocysts appear colorless on a red-pink background. This preparation can also done using fresh feces or feces fixed in formalin or SAF.

Staining with nigrosin solution (R13)

This is a stain for Cryptosporidium oocysts. The staining procedure is the same as that for merbromin solution.  Oocysts appear colorless on a black background.

Microscopic examination after concentration

If the specimen contains few parasites, direct microscopic examination may give a false-negative result.  Therefore, a concentration method should be used. Although many concentration methods have been described in the literature, Ritchie’s biphasic method and Sheather's flotation-sedimentation method (for Cryptosporidium oocysts) are the most commonly used.

Ritchie’s biphasic concentration method (modified)

This procedure is used by the Epidemiology Service and Laboratory for Tropical Diseases of the Sacro Cuore – Don Calabria Hospital in Negrar, Verona (Italy).  It is used on fresh, formalin-fixed or SAF-fixed feces.

Fresh feces specimens.

Collect a sample of stool about the size of a walnut from various parts (outside and inside of feces).

Gradually add a 10% or 5% formalin solution, or SAF;  mix well so the specimen becomes semi-liquid. Let the specimen fix for at least 30 minutes.

Specimens fixed in formalin or SAF.

Carefully stir the specimen with a plastic or a glass rod and check the consistence of the fecal suspension. If it is too dense, add more fixative and stir again until the specimen is semi-liquid.

Elimination of coarse residue

1-   Filter the suspension through a single layer of surgical gauze, previously moistened with saline and placed in a funnel above a 15 ml conical centrifuge tube of glass or ether-resistant plastic. The quantity of filtered suspension should be about 12 ml.

2-   Centrifuge for 3 min at 500 x g.

3-   Discard the supernatant fluid and check the quantity of sediment obtained: it should be about 2 ml; if not, filter some more fecal suspension and centrifuge again.

Alternatively to filtration through gauze, leave the test tube suspension to sediment spontaneously for less than 1 minute and pour off the supernatant into another test tube; proceed with point 2.

Elimination of water-soluble substances

4-   After obtaining the right amount of sediment, add saline and resuspend;   centrifuge again as described in point 2. Repeat until the supernatant is clean.

Elimination of fatty substances

5-   After eliminating the supernatant of the last centrifugation, resuspend the sediment with 10 ml saline, add 3 ml ethyl ether, cap the tube with a plastic bung and shake vigorously from top to bottom for at least 30 seconds. Remove the bung carefully to prevent spraying (due to the pressure of ether) and centrifuge immediately for 3 min at 500 x g.

6-   After centrifugation, four layers will be visible in the test tube: from the top, the ether with yellow or yellowish brown color, an interface of fecal debris, a layer of saline-formalin, and then the fecal pellet.

7-   Keeping the test tube vertical, delicately detach the plug of debris from the wall with a glass or plastic rod; then rapidly turn the test tube upside down, eliminating the top three layers. Bring the test tube back to vertical position immediately and resuspend the pellet with a few drops of formalin (not too much, in order not to overly dilute the sediment in which the parasites are concentrated).

Microscopic examination: see the previous section “Direct microscopic examination”.

Ethyl ether can be replaced with ethyl acetate, which is less flammable and therefore safer.  Ethyl acetate is also more effective than ethyl ether in the concentration of Giardia sp. cysts (as well as eggs from Taenia sp. and Hymenolepis nana), which tend to be trapped in the layer of fecal debris. However, the use of ethyl acetate to extract fatty substances and mucus provides less transparent preparations that are harder to read. The limit of Ritchie’s method is represented by the poor efficiency in concentrating vegetative forms of intestinal protozoa.

Trophozoite concentration method for SAF-fixed stool samples

The original method by L. Junod (1972) comprises:

-     Fixation of trophozoites and differentiation of nuclear structures in SAF solution (R4);

-     Flotation of trophozoites in a potassium iodomercurate solution with density 1.20.

-     Precipitation of trophozoites in a low-density solution.

Without requiring any type of staining, this method provides excellent results both for the concentration of trophozoites and for the differentiation of nuclei of all species of amoeba. However, potassium iodomercurate is highly toxic if inhaled, touched or ingested, and it is also dangerous for the environment. An alternative method that does not concentrate trophozoites but gives clean samples is as follows:

-     Carefully stir the specimen fixed in SAF and allow it to sediment spontaneously in a conical centrifuge tube for less than 1 minute;

-     Pour off the supernatant into another test tube and centrifuge at 500 x g for 2 minutes;

-     Discard the supernatant, add saline and centrifuge at 500 x g for 2 minutes; repeat this step;

-     Discard the supernatant and add a little SAF solution;

-     Resuspend the pellet.

Prepare a wet mount for direct examination, without or with an appropriate stain (e.g. Lugol's iodine solution or Bailenger's stain).

Calculation of the relative centrifugal force (g), based on rotations per minute (RPM), is described in the section “Reagents” (R35) . For standard bench-top centrifuges, 500 x g corresponds to about 2000 RPM.

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Sheather's flotation-sedimentation concentration method for Cryptosporidium oocysts

1-   In a conical centrifuge tube containing 7-8 ml sucrose solution (R3), place 0.5-1 g solid feces (or 1-2 ml diarrheal feces or 1-2 ml formalin- or SAF-fixed stool).

2-   Stir well and fill with sucrose solution to about 1-2 mm from the top of the centrifuge tube.

3-   Centrifuge at 500 x g for 5 minutes.  Stop the centrifuge without causing vibrations. For this method, a swinging bucket centrifuge is recommended.

4-   Without removing the tube from the centrifuge, touch the surface of the liquid with a metal or plastic loop and transfer the drop to a slide.  Repeat several times.

5-   Cover the drops with a coverslip.

6-   Examine under the microscope, preferably in dark field or phase contrast. Experienced microscopists can identify Cryptosporidium oocysts even with a bright field microscope (100x objective).


Permanent staining of fecal smears

The samples (e.g. feces, duodenal aspirates, rectosigmoidoscopy specimens and other materials) that are meant for permanent staining must be fixed immediately after passage to maintain the morphological characteristics of the intestinal protozoa and obtain a correct staining.

Schaudinn’s fixed specimen

Sample preparation:

-     Smear the specimen on a slide using a wood, plastic or glass applicator rod, making thicker (rolled) and thinner (smeared) areas.

-     Without allowing the smear to dry, place the slide immediately in Schaudinn's working solution (R5) for at least 1 hour (overnight is better).

It is recommended to fix the remaining material in SAF or PVA solutions.

For trichrome or iron-hematoxylin staining of specimens fixed in Schaudinn's solution, proceed from point 1 below.

Trichrome stain (Wheatley's modification of the Gomori method)

Preparation of smears from PVA-fixed specimens

Fix the specimen (after homogenization) in PVA solution for at least 30 min. Stir well, then remove 1-2 ml of the suspension with a pipette, pour it onto a sheet of paper towel and allow the excess PVA to be absorbed for 3 min (do not skip this step). With an applicator rod (plastic, wood or glass), take the fecal material from the paper towel and put it on a slide.  Then, with the same rod, form thicker (rolled) and thinner (smeared) areas on the slide. Leave to dry for at least 2 h in a 37 °C incubator or overnight at room temperature. These slides can be stored for at least one month at room temperature.

Staining procedure.

For occasional use (or when few slides are stained at one time), staining can be carried out using a Coplin jar.

1-     Place slides in iodine tincture for 5 min. (R14)

2-     Place slides in 70% ethanol for 2 min.

3-     Place slides in 50% ethanol for 2 min.

4-     Place slides in undiluted stock trichrome stain for 15 min. (R15)

5-     Drain slides after staining by blotting on paper towel and dip them for a few seconds in destaining solution (R16) (only 2 immersions).

6-     Place in 95% ethanol, shaking for 30 s.

7-     Place in absolute ethanol for 5 min.

8-     Place in absolute ethanol for 5 min.

9-     Place in xylene for 5-10 min.

Mount in resinous mounting medium or using the alternative method (see below).

Preparation of smears from SAF-fixed specimens

1-   Fix the thoroughly homogenized fecal specimen in SAF solution (R4) for at least 30 min.

2-   Mix well, take 2-3 ml of suspension and allow to sediment spontaneously in a centrifuge tube for less than 1 minute, then pour off the supernatant into another tube.

3-   Fill the tube with saline and centrifuge for 2 min at 500 x g. Discard the supernatant and repeat washing at least 4-5 times.

4-   After the last centrifugation, discard all the saline.

5-   Carefully homogenize the washed sediment with a rod, take a drop of sediment and mix it with a drop of Mayer's albumin (R17) previously placed on a slide.

6-   With the same rod, smear the material on the slide and form thicker (rolled) and thinner (smeared) areas. Leave to dry completely at room temperature; generally a few hours are enough, depending on the amount of saline present in the sediment, on the quantity of Mayer's albumin, and on the room temperature and humidity. Do not stain the slides if they are still shiny or wet, because the material can detach from the slide during staining. Slides can be stored for at least two weeks at room temperature away from dust.

7-   Stain slides with trichrome as described for PVA-fixed specimens, above, starting from point 2.

Microscopic examination and interpretation

Even if all the procedures described above are carried out properly, unexpected problems may generate disappointing results and make the preparation difficult to read. Therefore, for all permanent staining procedures, it is important to include a control slide with known protozoa (preferably a specimen containing Dientamoeba fragilis, for its particularly delicate nuclear structure). However, control slides stored for a long time may dehydrate (the moisture of the specimen is retained by the glycerin within Mayer's albumin). If this occurs, the result is a pale staining of the control slide. If specimens with known protozoa are not available, include a negative control slide of PVA- or SAF-fixed stool without protozoa, but to which a buffy coat of white blood cells has been added.

Examine the smear at low power (40x objective) identifying the areas of uniform thickness in order to assess the overall staining. Then, proceed to oil immersion observation (100x objective) and examine at least 300 microscopic fields before concluding that a specimen is negative.

Results:

-     In well-stained smears, the chromatic features of the organisms stand out against the background of the preparation, making them more visible than with hematoxylin staining;

-     The cytoplasm of the trophozoites is stained blue-green or green, although in some cases it takes on purple shades;

-     Entamoeba coli cysts show a brighter red-purple color than cysts of Entamoeba histolytica/dispar/moshovskii and other amoebae;

-     Cysts are often surrounded by a clear halo because they shrink during the fixing process: to determine the original size of the cyst, always measure the diameter of the surrounding halo;

-     Nuclear chromatin, the karyosome and inclusion bodies (chromatoid bodies, red blood cells and bacteria) stain red or red-purple, as do the fibrils (flagellar remnants) of flagellates;

-     Charcot-Leyden crystals stain red;

-     Yeast and Blastocystis stain green, the latter with red nuclei on a thin rim of cytoplasm;

-     Vacuoles or glycogen masses do not stain.

Trichrome stain provides best results using specimens fixed with Schaudinn's solution or PVA; the use of SAF fixative gives poor results.

Notes on trichrome staining

-     Slides can remain for 24 hour in the solutions indicated in points 2, 8 and 9 without affecting the quality of the staining.

-     The removal of mercuric chloride from specimens fixed with Schaudinn's solution or PVA is achieved using an iodine tincture. This solution should be made fresh at least weekly or whenever its color becomes clear.  If this is not done, a highly refractive precipitate of crystals or granules will form and make smear examination difficult or impossible.

-     Incomplete removal of iodine causes a predominant green staining of the smear. Change frequently the 70% ethanol solution at point 2 or lengthen the time of this step.

-     The transition from point 3 to point 4 of the staining process may cause ethanol to enter the trichrome stain, diluting it. To avoid this, drain smears on a paper towel before placing them in the trichrome stain.

-     To check the quality of the staining solution, tilt the tank slightly to see if the moist rim is red;if not, regenerate the solution by leaving the tank uncovered overnight to evaporate off any ethanol present; then refill to the original level with new stock trichrome solution.

-     Although trichrome stain is progressive, the best results are obtained with regressive staining, i.e. using a destaining solution. Excessive destaining (it is easy to over-differentiate - point 5) causes poor differentiation of the parasites, even when the specimens stain well. For example, the granular nuclear chromatin appearance of Dientamoeba fragilis will be difficult to detect and may generate confusion with other amoeba species (e.g. Entamoeba hartmanni). Contrarily, in over-stained specimens (little differentiation), the nuclear chromatin may appear as a compact blob, not formed of granules, thereby causing confusion with Endolimax nana or Iodamoeba buetschlii. 

-     In the final dehydration phase, the ethanol solution should be absolute (100%), i.e. completely free of water.  If, after the passage from 100% ethanol to xylene, the latter becomes opaque, dip the slide once more in new 100% ethanol and then in new xylene.

-     If the result of the staining is unsatisfactory and if it is not possible to get a new specimen, restain the slides as follows:

o  Place smears mounted with resinous mounting medium into xylene for at least 12-24 hours, until the coverslips detach from the slides;

o  Dip slides in absolute ethanol (1 min), 95% ethanol (1 min), 70% ethanol (1 min) and finally in 50% ethanol (1 min);

o  Destain smears in 10% acetic acid for several hours;

o  Wash by soaking in tap water for 3 min (change at least 4-5 times);

o  Dip in 70% ethanol (1 min) and 50% ethanol (1 min);

o  Now repeat trichrome staining from point 4.

If you wish to examine the smear with the oil immersion objective, immediately after staining without permanent mounting – which requires a waiting time of at least 24-36 hours for drying, an alternative mounting method can be used and permanent mounting can be carried out later.

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Alternative mounting method

-     Remove slides from the xylene and leave to air dry in horizontal position.

-     Place some drops of immersion oil on the smear, allow the oil to spread evenly and leave for about 10 min;

-     Place a coverslip, add one more drop of immersion oil, and examine at high power (100x objective).

Never examine smears without coverslip. After dehydration, the stained fecal smears are very fragile. In addition, without using a protective coverslip, the lens of the high power objective (100x) can be irreparably damaged. In any case, the transparency and brightness of the preparation mounted with this alternative method are lower than that achieved with permanent mounting.

Iron-hematoxylin stain

Hematoxylin is a natural stain extracted from Hematoxylon campechianum wood.  It consists of colorless crystals lacking staining ability, which is then acquired after oxidation and transformation into hematein.

Many hematoxylin staining methods are available.  Regressive methods are time consuming and require experience, especially in the destaining step; the preparation must be checked several times under the microscope when wet, in order to follow the destaining and differentiation processes. These methods, however, are able to reveal fine cytological details with excellent results. On the other hand, progressive methods are rapid, do not involve destaining, and provide nuclear staining which is sufficient for identifying amoeba species. 

Preparation of smears

Specimens fixed in Schaudinn's solution (see trichrome stain)

Specimens fixed in SAF (see trichrome stain)

Specimens fixed in PVA (see trichrome stain)

Staining with iron-hematoxylin gives excellent results with specimens fixed with SAF or Schaudinn's solution.

Iron-hematoxylin stain (Heidenhain's regressive method)

For occasional use (or when few slides are stained at one time), staining can be carried out using a Coplin jar.

For specimens fixed in Schaudinn's solution, proceed from point 1.

For specimens fixed in PVA, proceed from point 1.

For specimens fixed in SAF, proceed from point 2.

Procedure:

1-     Dip slides in iodine tincture for 5 min (R14).

2-     Place slides in 70% ethanol for 5 min.

3-     Place slides in 50% ethanol for 2 min.

4-     Wash well with distilled water.

5-     Place slides in hematoxylin working solution (R20) for 10 min .

6-     Place slides under running tap water (best if tepid) for 10 min.

7-     Differentiate slides (one by one) in destaining solution (R21) or (R22) for 30 s.

8-     Wash with tap water and check the wet slide under the microscope (40x objective): if the result is unsatisfactory, replace the slide in destaining solution and then in tap water, until the desired result is achieved. If differentiation in destaining solution is excessive (slide too destained or poor differentiation of parasites), restain with hematoxylin starting from point 5. 

9-     Place slides under running tap water (best if tepid) for 10 min.

10-   Place slides in 95% ethanol for 5 min.

11-   Place slides in 100% ethanol for 5 min.

12-   Place slides in 100% ethanol for 3 min.

13-   Place slides in xylene for 5-10 min.

Mount slides with resinous mounting medium or use the alternative mounting method (see above).

During staining, smears should never dry.

Microscopic examination and interpretation

-     Parasites stain blue to black, according to how old the hematoxylin working solution is;

-     Trophozoitic cytoplasm stains dark blue, gray or black;

-     Cysts stain dark blue, gray or black, and very often are surrounded by a clear halo which should be considered when measuring cyst size; 

-     Peripheral nuclear chromatin, the karyosome and inclusion bodies (chromatoid bodies, red blood cells and bacteria) stain dark blue, gray or black, as do the fibrils (flagellar remnants) of flagellates;

-     Charcot-Leyden crystals stain black;

-     Yeast and Blastocystis stain dark blue, gray or black, the latter with the nuclei darker on the thin rim of cytoplasm;

-     Glycogen does not stain, but a clear area should be seen.

Examine at least 300 microscopic fields (100x objective) before considering the specimen negative.

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Mayer's hematoxylin stain (rapid progressive method)

Albeit not providing the same quality results as the regressive method, the rapid progressive method can be a valid alternative for beginners.

Sample preparation (see above).

Staining procedure

1-     Place slides in 70% ethanol for 5 min and then in 50% ethanol for 1 min.

2-     Wash slides in tap water for 5 min.

3-     Quickly wash slides twice in distilled water.

4-     Place slides in Mayer's hematoxylin (R23) for 40 s. This time must be checked for every new stock solution.

5-     Wash slides with running tap water for 5 min.

6-     Place slides for 30 s in a solution of water and ammonia (R24).

7-     Place slides in 95% ethanol for 1 min; then in new 95% ethanol for 30 s.

8-     Dehydrate slides in 100% ethanol for 5 min.

9-     Place slides in xylene for 5-10 min.

10-     Mount slides using a resinous mounting medium or the alternative mounting method.

During staining, slides should never dry.

With this method, all parasites stain blue.

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Notes on trichrome and hematoxylin staining

These staining methods were developed in order to make vegetative forms of intestinal protozoa (amoebae, flagellates) easy to recognize and identify.  In specimens containing cysts, the results can be poor due to the presence of collapsed, shrunken, deformed or over-stained forms. This is not the result of bad staining, but it is due to the fact that cysts are poorly fixed in Schaudinn's, PVA and SAF solutions.

During fixation, vegetative forms may be in any one of the different phases of the cell cycle, including the rest period (interphase) and the various phases of nuclear division.  Therefore, it is possible to observe forms with 2 nuclei or an apparently deformed nucleus undergoing division (anaphase).

See “Trophozoite division (mitosis)”

Giemsa and Field stains

These stains are used only on fresh, unfixed specimens (e.g. feces, duodenal aspirates) to identify trophozoites of Dientamoeba fragilis or of flagellates, with excellent results. Other intestinal protozoa are difficult or impossible to identify.

Giemsa stain

Sample preparation

1-     Spread the specimen on a slide to obtain a thin smear; if the feces have a normal consistence, dilute with saline (not with water).

2-     Allow the slide to air dry.

3-     Fix in 100% methanol for 1 minute.

4-     Allow the slide to air dry.

Staining

1-     Place the slides in 10% Giemsa solution (R33) for 30 minutes.

2-     Wash slides well with tap water.

3-     Allows the slides to air dry.

Microscopic examination

Put some drops of immersion oil onto the stained slide, allow the oil to spread evenly and leave for about 10 min.  Cover with a coverslip and add one more drop of immersion oil.  Then, examine at high power (100x objective).  Do not examine specimens without a coverslip because the lens of the objective can be damaged. 

Interpretation

-     Nuclei stain red-purple or violet;

-     Cytoplasm stains blue or blue-gray.

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Field stain

Sample preparation (see Giemsa stain)

Staining

1-     Flood slide with 1 ml Field stain B (diluted 1:4 with distilled water).

2-     Immediately add 1 ml Field stain A and, with the same pipette, mix the two stains together.

3-     Stain for 1 minute.

4-     Wash slide well with tap water and allow to air dry.

Microscopic examination and interpretation (see Giemsa stain)

Ziehl-Neelsen stain, modified (hot method)

This staining method highlights the acid resistance of intestinal coccidia.

Procedure

1-     The fecal sample is spread on the slide to produce a thin smear.  For this procedure, use feces fixed in formalin or SAF or feces that have previously been concentrated (in the last case, to recover Cryptosporidium oocysts, all centrifugation phases must be lengthened by at least 10 min).

2-     Allow the slide to air dry.

3-     Fix in 100% methanol for 1 minute.

4-     Allow the slide to air dry.

5-     Flood slide with carbol-fuchsin working solution (R25) for 5 minutes, gently heating on a Bunsen flame or ethanol lamp.  Do not boil or dry the staining solution; if it dries, add more staining solution to the slide.

6-     Decolorize with acid solution (R26) until the stain (red-pink) no longer drips from the slide.

7-     Wash slide well with tap water.

8-     Flood slide with a methylene blue solution (R27) for 30 s.

9-     Wash well with tap water, and allow the slide to air dry.

Microscopic examination and interpretation

Examine slides at low (40x objective) and high (100x objective) power. Oocysts stain pink to red or deep purple.

Cryptosporidium: inside the roundish or oval oocysts, black punctiform or comma-like formations (sporozoites) are seen; the bright red color of oocysts is readily recognizable against a blue background.

Isospora belli: in mature oocysts, two red sporocysts are visible, surrounded by a clear halo separating them from the oocyst wall.

Cyclospora: on average only one oocyst out of four becomes stained, taking on a red to pale pink color; the other oocysts can be seen as roundish and colorless “ghosts”. To stain all the oocysts, a safranin method can be used which, however, requires a microwave oven.

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Kinyoun stain (cold method)

This stain highlights the acid resistance of intestinal coccidia.

Procedure

1-     Spread stool, formalin- or SAF-fixed specimens, or concentrated samples to obtain a thin smear. (If it is necessary to recover Cryptosporidium oocysts, all centrifugation phases of the concentration method must be lengthened by at least 10 min).

2-     Allow the slide to air dry.

3-     Fix in 100% methanol for 1 minute.

4-     Allow the slide to air dry.

5-     Flood the slide with Kinyoun carbol-fuchsin solution (R28) for 5 min.

6-     Decolorize with acid solution ((R29) until the stain (red-pink) no longer drips from the slide.

7-     Wash slide well with tap water.

8-     Flood slide with a methylene blue solution (R30) for 30 s.

9-     Wash the slide well with tap water and allow to air dry.

Microscopic examination and interpretation

Examine slides under the microscope at low (40x objective) and high (100x objective) power. Oocysts stain pink to red or deep purple, but less bright than with the modified Ziehl-Neelsen hot method. 

Weber's trichrome stain for microsporidia

Preparation

-     Prepare a thin smear with 10 μl of feces fixed with 10% formalin or SAF. Specimens that had been concentrated with Ritchie's method cannot be used;

-     Fix in 100% methanol for 10 minutes and allow the slide to air dry.

Staining procedure

1-     Place the slide for 90 minutes in modified trichrome solution (R31);

2-     Destain in acid alcohol solution for 5-10 s (R32);

3-     Briefly rinse in 95% ethanol for 2-4 s (2 immersions);

4-     Dehydrate twice in absolute ethanol for 5 minutes;

5-     Place slide in xylene for 5 minutes (twice);

6-     Mount with resinous mounting medium or the alternative mounting method.

Microscopic examination and interpretation

Carefully examine at least 300 microscopic fields with the oil immersion lens (100x objective). Microsporidia spores are refractile, oval, with reddish-pink walls, but they are difficult to identify due to their small size. Sometimes the content of the spore does not stain at all, while in other cases a reddish-pink diagonal or equatorial band can be seen. The background of the preparation, comprising bacteria and fecal debris, stains weakly green. 

To avoid erroneous interpretations, use of a positive control is mandatory.

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Giemsa stain for microsporidia

Sample preparation (see Weber's trichrome staining for microsporidia)

Staining procedure

1-     Dip fixed slides in Giemsa solution (R33) for 35 minutes;

2-     Wash with running tap water for 30 s;

3-     Allow slides to air dry.

Microscopic examination and interpretation

Carefully examine at least 300 microscopic fields with the oil immersion lens (100x objective). When present, microsporidia spores appear either isolated or clustered in groups. The nucleus stains red, the cytoplasm pale blue; a clear intracytoplasmic vacuole is visible.  To avoid erroneous interpretations, use of a positive control is recommended.

Fluorescence microscopy procedure for coccidian oocysts

Coccidian (Cyclospora, Isospora, Sarcocystis, Toxoplasma, but not Cryptosporidium) oocysts exhibit blue autofluorescence when excited with light at wavelengths less than 400 nm. However, microscopes with quartz-iodine bulbs do not emit UV radiation at these wavelengths, so oocystic autofluorescence cannot be detected. Microscopes equipped with mercury or xenon vapor lamps do emit UV radiation under 400 nm, but oocystic autofluorescence can be detected only with the use of a 340-380 nm excitation filter.    

Sample preparation 

Fresh specimens, those fixed with 10% formalin or SAF, as well as concentrated stool can be examined.

Procedure

1.     Place few drops of sample onto a slide and cover with a coverslip;

2.     Examine the slide with fluorescence microscopy using a 10x or 40x objective as well as the 340-380 nm excitation filter.

Interpretation

Oocysts retain their normal morphology.  The inside appears light blue while the wall appears brightly blue fluorescent.

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