Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 15
ОглавлениеSince Pneumocystis jirovecii (previously P. carinii) has been reclassified with the fungi, this organism is no longer included in this book or discussed in terms of organism, disease, pathogenesis, and diagnosis. However, other organisms such as the microsporidia can be stained using silver stains, so these methods and references have been kept in this chapter and will periodically refer to P. jirovecii.
Although it is not one of the more common specimens, expectorated sputum may be submitted for examination for parasites. Organisms in sputum that may be detected and may cause pneumonia, pneumonitis, or Loeffler’s syndrome include the migrating larval stages of Ascaris lumbricoides, Strongyloides stercoralis, and hookworm; the eggs of Paragonimus spp.; Echinococcus granulosus hooklets; and P. jirovecii (now classified with the fungi), Entamoeba histolytica, Entamoeba gingivalis, Trichomonas tenax, Cryptosporidium spp., and possibly the microsporidia (now classified with the fungi) (1). In a Paragonimus infection, the sputum may be viscous and tinged with brownish flecks, which are clusters of eggs (“iron filings”), and may be streaked with blood (Fig. 6.1).
Figure 6.1 Paragonimus spp. eggs. (Left) Operculated eggs in a sputum specimen. (Right) Eggs photographed at a higher magnification. doi:10.1128/9781555819002.ch6.f1
Collection and Examination of the Specimen
Sputum is usually examined as a wet mount (saline or iodine), using low and high dry power (×100 and ×400). The specimen is not concentrated before preparation of the wet mount. If the sputum is thick, an equal amount of 3% sodium hydroxide (or undiluted chlorine bleach) can be added; the specimen is thoroughly mixed and then centrifuged. NaOH should not be used if Entamoeba spp. or T. tenax is being sought; the protozoa will be destroyed. After centrifugation, the supernatant fluid is discarded and the sediment can be examined as a wet mount with saline or iodine. If examination has to be delayed for any reason, the sputum should be fixed in 5 or 10% formalin to preserve helminth eggs or larvae or in one of the fecal fixatives to be stained later for protozoa. Usually, sputum is not a recommended specimen for the diagnosis of P. jirovecii; however, if such a specimen is accepted by the laboratory, other stains such as silver methenamine, Giemsa, or immunoassay reagents are used. If Cryptosporidium is suspected (rare), acid-fast or immunoassay techniques normally used for stool specimens can be used (see chapters 3 and 15) (2–5). Trichrome stains of material may aid in differentiating E. histolytica from E. gingivalis, and Giemsa stain may better define larvae and juvenile worms.
A sputum specimen should be collected properly so the laboratory receives a “deep sputum” sample for examination rather than a specimen that is primarily saliva from the mouth rather than from the lower respiratory passages. If the sputum is not induced, the patient can be instructed as follows.
1. Expectorated sputum specimens are collected after the patient is instructed in the appropriate measures to ensure high-quality specimens, including prior mouth washing with hydrogen peroxide and exclusion of saliva from specimens. Try to obtain the specimen early in the morning, when the chances of obtaining a deep sputum specimen are increased.
2. Transport the specimen to the laboratory in clean, closed containers as quickly as possible. Select any blood-tinged, viscous areas for sampling.
3. If the specimen is uniformly mucoid:
A. Wear gloves when handling specimens.
B. Remove a 1.0-ml portion to a 15-ml conical tube.
C. Add 1.0 ml of mucolytic agent such as Sputalysin (Sputalysin Stat-Pack dithiothreitol solution [Behring Diagnostics, Inc.]) which has been prepared as specified by the manufacturer. This reagent can be stored unopened at room temperature until the expiration date. Date and store working solution at 4°C (include the expiration date). Discard working solution after 48 h. Prepare working solution by removing 1.0 ml from the 10-ml bottle and diluting with 9.0 ml of sterile water.
D. Incubate at room temperature for 15 min.
E. Add 2.0 ml of phosphate buffer (pH 6.8; M/15 [0.067 M]).
F. Centrifuge the material at 500 × g for 5 min.
G. Decant supernatant fluid, and use sediment to prepare wet mounts and smears.
H. Quality control measures should include the following.
a. Ensure that the saline and mucolytic agent are free of contamination (particulate matter) as determined by a clear appearance. If cloudy, discard and make new working solution.
b. Control the trichrome stain for each new set of reagents with a specimen containing blood to ensure that white cells stain with purple nuclei and blue-green cytoplasm. If cells do not stain appropriately, change reagents.
c. With each new lot of Giemsa stain or new buffer, check the stain with a specimen containing blood to ensure that red cells stain grayish, white cell nuclei stain red-purple, and cytoplasm stains bluish. If cells do not stain appropriately, check the stock stain and buffer to find cause.
d. Yeast-containing material may be used as a positive control for silver staining.
e. Both P. jirovecii-positive and P. jirovecii-negative and yeast-positive material should be used for immune-specific staining (check with suppliers).
f. Stains cannot be evaluated if controls do not stain appropriately. A stain is not within acceptable results when:
(i) P. jirovecii does not stain or stains black without delineation of “parentheses.”
(ii) Other fungi and actinomycetes do not stain.
g. Store stock stain solutions in area away from light; discard any reagent (prior to the expiration date) that is cloudy or appears contaminated.
h. Evaluate the fluorescence microscope for correct light wavelength, using commercially available quality control slides.
i. Record all quality control results.
Procedure for Examination of Sputum
1. Wear gloves when performing these procedures.
2. For expectorated sputum (no addition of mucolytic agent):
A. Using a Pasteur pipette, place 1 or 2 drops (50 µl) on one side of a 2- by 3-in. (1 in. = 2.54 cm) glass slide, and cover with a 22- by 22-mm (no. 1) coverslip.
B. Place a second drop on the slide, add 1 drop of saline, and cover with a coverslip.
3. Expectorated sputum (treated with a mucolytic agent) can be resuspended in 100 µl of saline; place 1 drop on a 2- by 3-in. slide, and cover with a coverslip.
4. Save the untreated specimen and remaining treated specimen for permanent smear preparation if stains are required.
5. Examine the entire 22- by 22-mm coverslip wet preparation under low light with the 10× objective to detect eggs, larvae, oocysts, or amebic trophozoites.
6. If results are inconclusive, prepare smears for permanent staining.
A. Place 1 drop of sediment in the center of each of three 1- by 3-in. glass slides, add 2 to 3 drops of fecal fixative to one of the slides, and mix/spread the material with the tip of the pipette.
B. Allow all three smears to air dry a minimum of 30 min (they can be incubated for drying at 37°C).
C. Trichrome stain the slide fixed in fecal fixative.
D. Fix the other two air-dried smears in methanol; stain one with Giemsa and the other with a modified acid-fast stain. The modified trichrome stain may have to be used if infection with microsporidia is suspected (6, 7).
E. Put immersion oil on stained smears; examine the Giemsa-stained smear with the 10× objective and the trichrome-stained smear with the 50× oil objective if available; otherwise, use the 100× oil objective. Read at least 300 oil immersion fields (total magnification, ×1,000) before reporting the specimen as negative.
F. The methenamine silver stain can be used; however, expectorated sputum is generally not recommended as an acceptable specimen for the recovery and identification of P. jirovecii.
7. Report any organisms found as follows.
A. Give genus and species, if necessary, after confirmation with permanent stain.
B. “No parasites found” in expectorated sputum is considered a normal/negative report; therefore, call the physician if any organisms are found.
Note Care should be taken not to confuse E. gingivalis, which may be found in the mouth and saliva, with E. histolytica, which could result in an incorrect suspicion of pulmonary abscess. E. gingivalis usually contains ingested polymorphonuclear leukocytes, while E. histolytica may contain ingested red blood cells but not polymorphonuclear leukocytes (Fig. 6.2). T. tenax would also be found in saliva from the mouth and thus would be an incidental finding and normally not an indication of pulmonary problems (Fig. 6.3).
Figure 6.2 (Upper) Entamoeba gingivalis containing ingested polymorphonuclear leukocytes (PMNs) within large vacuoles. (Lower) Entamoeba histolytica containing ingested red blood cells (RBCs). Note: the nuclear characteristics are very similar; however, the vacuole sizes are quite different (large vacuoles contain ingested PMNs, small vacuoles contain ingested RBCs). doi:10.1128/9781555819002.ch6.f2
Figure 6.3 (Left) Trichomonas tenax from the mouth (stained with Giemsa stain). (Right) Trichomonas vaginalis from a genital specimen (stained with Giemsa stain). Note the large nucleus in T. tenax and the fact that this flagellate is somewhat smaller than T. vaginalis. doi:10.1128/9781555819002.ch6.f3
Concentrated stained preparations of induced sputum specimens are commonly used to detect P. jirovecii and differentiate trophozoite and cyst forms from other possible causes of pneumonia, particularly in AIDS patients (8). Organisms must be differentiated from other fungi such as Candida spp. and Histoplasma capsulatum. Although induced sputum specimens have been used successfully in the diagnosis of P. jirovecii in some institutions, they have not been useful in others. This difference may be due in part to careful adherence to specimen rejection criteria. If the clinical evaluation of a patient suggests P. jirovecii pneumonia and the induced sputum specimen is negative, a bronchoalveolar lavage fluid specimen should be evaluated with the stains presented in this protocol or newer molecular methods (9).
Collection and Examination of the Specimen
Induced sputum specimens are collected by pulmonary or respiratory therapy staff after patients have used appropriate cleansing procedures to reduce oral contamination. Nebulizing procedures are generally determined by the respiratory therapy staff collecting the specimens. The induction protocol is critical for the success of the procedure, and it is mandatory for well-trained individuals to be involved in the recovery of organisms. Patients with Pneumocystis pneumonia usually have dry, nonproductive coughs. Organisms are rarely detected in expectorated sputum, which is not accepted by many laboratories as a clinically relevant specimen. In cooperation with the pulmonary staff, the laboratory processing the specimens must establish a protocol for this diagnostic procedure. Induced sputum specimens are most useful for detection of P. jirovecii in HIV-infected individuals, because others have fewer, less readily detected organisms. Stains that can be used for organism detection include the rapid methenamine silver stain, the rapid Giemsa stain (Diff-Quik or Giemsa Plus), the Giemsa stain, and immune-specific staining (10).
Preparation of the Specimen Prior to Staining
1. Wear gloves when handling specimens.
2. Specimens which contain mucus should be treated with a mucolytic agent by adding the agent in a 1:1 ratio with the specimen, usually 2 or 3 ml, and incubating the specimen at room temperature for 15 min. Large-volume specimens usually are watery. These specimens (up to 20 to 25 ml) should be concentrated by centrifugation prior to the addition of a mucolytic agent.
3. Centrifuge the specimen at 500 × g for 5 min in capped centrifuge tubes and closed carriers.
4. Decant supernatant fluids into a disinfectant solution (1:10 dilution of bleach).
5. If the sediment contains a significant amount of blood, treat a portion (one-half to one-third) with a red-cell lytic agent such as saponin or Lyse in one-half to one-third of the volume of the sediment, leave at room temperature for 5 min, and recentrifuge.
6. Decant supernatant fluid from treated specimens.
7. Use Pasteur pipettes to resuspend remaining sediment.
8. Using 1- by 3-in. glass slides, place drops of sediment in the center of each slide. For specimens treated to lyse red cells, prepare two smears, one from material before lytic treatment and one from the treated specimen.
9. Spread the drops with the pipette so that they are thin and even.
10. Air dry slides, and dip them in methanol prior to Giemsa or silver staining. Fix slides for immune-specific staining as specified in the package insert directions.
Reagents Used in Staining Procedures
1. Mucolytic agent: Sputalysin Stat-Pack dithiothreitol solution (Behring Diagnostics, Inc.). Red cell lytic agent: saponin (Aldrich, Milwaukee, WI; ICN Biochemicals, Costa Mesa, CA), Lyse (Curtin Matheson Scientific, Inc.), or Hematall LA-Hgb reagent (Fisher Scientific).
2. Rapid Giemsa stain: Diff-Quik (Baxter Scientific Products) or Wright’s Dip Stat (Medical Chemical). Follow the manufacturer’s instructions or use Giemsa stain.
3. Giemsa stain: azure B alcoholic stock (Harleco, Philadelphia, PA) diluted 1:20 with phosphate buffer containing 0.01% Triton X-100.
4. Buffers prepared from stock buffers for the Giemsa stain:
A. Alkaline buffer NaHPO4 M/15 solution is 9.5 g of Na2HPO4 dissolved in 1 liter of distilled water.
B. Acid buffer NaHPO4 M/15 solution is 9.2 g of N Na2HPO4 · H2O dissolved in 1 liter of distilled water.
These buffers can be kept for 12 months. To prepare buffered water for the stain, add 39 ml of acid buffer and 61 ml of alkaline buffer to 900 ml of water.
Reagents for Rapid Methenamine Silver Stain (Microwave) (6)
Chromic Acid (10% Solution)
Solution is stable for up to 1 year.
Methenamine (3% Solution)
Solution is stable for up to 6 months.
Silver Nitrate (5% Solution)
Solution is stable for 1 to 6 months at 4°C.
Sodium Borate (Borax) (5% Solution)
Solution is stable for up to 1 year.
Sodium Bisulfite (1% Solution)
Solution is stable for up to 1 year.
Gold Chloride (1% Solution)
Solution is stable for up to 1 year.
Sodium Thiosulfate (5% Solution)
Solution is stable for up to 1 year.
Stock Light Green
Solution is stable for up to 1 year.
Working Light Green
Solution is stable for up to 1 month.
Ethanol, 100% (Absolute) and 95%
Methanol, 100% (Absolute)
Xylene or Xylene Substitute
All reagents can be stored at 25°C except for the silver nitrate, which must be refrigerated. Although stable for up to 6 months, silver nitrate may have to be discarded and replaced after storage for 1 month.
Procedure for Rapid Methenamine Silver Stain (Microwave) (11)
1. Place specimen smear slides and control slides on stain rack.
2. Add 10% chromic acid to cover smears, and let stand for 10 min.
3. During this time, prepare working methenamine-silver nitrate by placing the following in order in a plastic Coplin jar: 20 ml of 3% methenamine, 1 ml of 5% silver nitrate, 1.5 ml of 5% sodium borate, and 17 ml of distilled deionized water.
4. Wash slides with distilled deionized water.
5. Cover slides with 1% sodium bisulfite for 1 min.
6. Wash slides with water, place them in the plastic Coplin jar containing methenamine-silver nitrate, and cover with cap.
7. Place in microwave oven at 50% power for approximately 35 s, rotate 90°, and heat for another 35 s; leave slides in hot liquid for 2 min. Solution should turn brown (fresh reagent may appear almost clear if the stock methenamine is freshly prepared; with time and use, the working solution will turn more brown).
8. If a water bath is used, heat it to 80°C. Place plastic Coplin jar containing stain reagents in bath for 6 min prior to placing slides in jar, add slides, and leave jar in bath for an additional 5 min. Solution should turn brown (fresh reagent may appear almost clear if the stock methenamine is freshly prepared; with time and use, the working solution will turn more brown).
9. Remove slides; wash in distilled deionized water.
10. Dip slides up and down in 1% gold chloride, wash in distilled deionized water, and place on rack again.
11. Cover slides with 5% sodium thiosulfate for 1 min.
12. Wash in distilled deionized water.
13. Cover with 0.2% fast green in acetic acid, counterstain for 1 min; use working solution.
14. Wash with distilled deionized water, and stand slides on end to drain and dry.
15. Examine with oil immersion or mount slides.
16. Fill Coplin jar used for staining with bleach (full strength), and let stand for at least 1 h.
Reagents for Rapid Methenamine Silver Stain (No Microwave) (12)
In another rapid methenamine silver-staining procedure, a 5% (rather than 10%) chromic acid solution is used, the gold chloride is present at 0.2% (rather than 1%), the sodium thiosulfate is present at 2% (rather than 5%), and the microwave oven is not used (12). Another difference from the microwave method involves fixation of the smears prior to staining. Heat fix all smears at 70°C for 10 min on a heating block for specimens that could contain Mycobacterium tuberculosis. Place the air-dried smears in absolute methyl alcohol for 5 min. Fix a control smear (microsporidia, fungus, or P. jirovecii) at the same time. It is mandatory that control smears be used every time patient specimens are stained.
Chromic Acid (5% Solution)
Solution is stable for up to 1 year.
Methenamine (3% Solution)
Solution is stable for up to 1 year.
Sodium Bisulfite (1% Solution)
Solution is stable for up to 1 year.
Sodium Thiosulfate (Hypo) (2% Solution)
Solution is stable for up to 1 year.
Stock Light Green
Solution is stable for up to 1 year.
Silver Nitrate (5% Solution)
Solution is stable for 1 to 6 months at 4°C.
Sodium Borate (Borax) (5% Solution)
Solution is stable for up to 1 year.
Gold Chloride (0.2% Solution)
This solution is stable for 1 year; however, when toning begins to fail and the organisms come out too black, it should be changed.
Note The 1% gold chloride solution is made from ampoules and is diluted as specified in the accompanying directions (dilute with tap water).
Stock Methenamine-Silver Nitrate
A white precipitate forms but immediately dissolves on shaking. Clear solutions remain stable for months at refrigerator temperature (4°C).
Working Light Green
Solution is stable for up to 1 month.
Working Methenamine-Silver Nitrate
Caution The working methenamine-silver nitrate solution must be prepared fresh each time the stain is run. Do not try to reuse, even for a stain run immediately following one previously performed.
Procedure for Rapid Methenamine Silver Stain (No Microwave) (12)
1. While slides are fixing, fill one Coplin jar (with a lid) with 5% chromic acid. In a second, screw-cap Coplin jar, prepare the working methenamine-silver nitrate solution.
2. Place fixed slides into chromic acid, and put both Coplin jars into the 80°C water bath. Warm the jars and their contents in a stream of hot water for approximately 30 s before placing them in the water bath, to prevent the jars from cracking. Place smears into the heated 5% chromic acid, and incubate them in the water bath for 2 min (oxidation step).
3. Wash slides briefly in running tap water.
4. Place slides into 1% sodium bisulfite for 30 s.
5. Rinse slides in three changes of distilled water.
6. Place slides into the jar of working methenamine-silver nitrate solution in the 80°C water bath for 4.5 min (reduction step).
7. Rinse slides with three changes of distilled water. Wipe backs of slides with a paper towel to remove excess methenamine-silver nitrate.
8. Tone in 0.2% gold chloride for 30 s.
9. Rinse in three changes of distilled water.
10. Place slides into 2% sodium thiosulfate for 30 s (removes reduced silver).
11. Rinse in three changes of distilled water.
12. Counterstain in working light green solution for 30 s.
13. Rinse in three changes of distilled water.
14. Dehydrate and clear for 30-s intervals in two changes each of 95% ethyl alcohol, 100% ethyl alcohol, and xylene or xylene substitute, respectively.
15. Mount slides in Permount.
The fungi generally stain gray to black; P. jirovecii exhibits a delicately stained wall, usually brownish or grayish, which is somewhat transparent. Structures described as “parentheses” are usually seen and are stained dark gray or black (Fig. 6.4). If the organisms are too dark, it is probably time to change the solutions. It is also important to prepare the smears so that the material is thin and evenly spread on the glass slide. Glycerin, red blood cells, and mucin stain rose taupe to dark gray. The inner parts of mycelia and hyphae stain “old rose.” The background usually appears pale green. Microsporidial spores stain dark gray to black; the horizontal or diagonal “stripe” or polar filament is not visible in every spore (Fig. 6.4). Various background colors are visible, depending on the stain used, thickness of the specimen, and tissue source of the specimen.
Figure 6.4 (Upper) Pneumocystis jirovecii. Note that the “parentheses” are visible within the cell wall (methenamine silver stain). (Middle) Microsporidial spores in a stool specimen stained with silver stain (methenamine silver stain). Note the spore with a polar tubule (horizontal line through the spore) within the black box; there are other examples of spores containing the polar tubule throughout the image. (Bottom) Microsporidian spores within an ocular biopsy specimen (methenamine silver stain). In this image the spores are clustered together and polar tubules are not visible. doi:10.1128/9781555819002.ch6.f4
Note The examination of specimens for the diagnosis of microsporidia is not often considered to be a STAT procedure and may be handled in the anatomic pathology division. The stained smears can be difficult to read and interpret, hence the need for available experienced personnel. However, special modified trichrome stains for the microsporidia are often performed in the microbiology laboratory; the procedure is not difficult, but examination of the stained smears is very labor intensive.
Rapid Giemsa Stain (Diff-Quik or Giemsa Plus)
1. Keep stain solutions in dropper bottles. Place 1 or 2 drops of red stain (solution 1) on specimen smear and control slide (normal blood film), hold for 10 s, and drain.
2. Add drops of blue (solution 2), hold for 10 s, drain, and rinse very briefly with deionized water.
3. Stand slides on end to drain and air dry.
4. Slides must be examined with oil or mounted with mounting medium.
Giemsa Stain
Make Giemsa working solution fresh each day. Discard and make new solution after 10 slides have been stained in a Coplin jar. Additional directions can be found in chapter 7.
1. Place 2 ml of Giemsa (azure B) stain in a Coplin jar. Remove stain from stock stain in bottle with a clean, dry pipette.
2. Add 40 ml of phosphate buffer (pH 7.0 to 7.2) containing 0.01% Triton X-100.
3. Place fixed specimen smears and control smears in Giemsa stain for 30 min.
4. Remove slides, dip in phosphate buffer, stand slides on end, and allow to drain and air dry.
5. Examine with oil or mount slides.
Giemsa Staining for Microsporidia
1. Internal morphology may be difficult to interpret (spores with polar tubules).
2. Giemsa stain may be more relevant for specimens other than stool (urine, sputum, eye).
3. Perform modified trichrome (Ryan blue or Weber green) stains on stool.
Potential Problems with Stain Interpretation
In the procedures in which the spores are stained, spores and fungi may appear very much alike. If rare organisms are seen, it may be almost impossible to differentiate microsporidia from various fungi, particularly if no budding organisms are seen. Although the microsporidia are now classified with the fungi, they will continue to be covered in both sections of this book.
Although a second type of stain can be used (Giemsa, which stains the spores), the organisms may be very difficult to differentiate from cellular debris. Consequently, few laboratories rely on the Giemsa stain alone, especially when staining specimens for Pneumocystis or the microsporidia (Fig. 6.5).
Figure 6.5 Pneumocystis jirovecii trophozoites within the cyst wall (stained with Giemsa stain). Note that the cyst wall is not visible when Giemsa stain is used. Other trophozoites can also be seen in the background. doi:10.1128/9781555819002.ch6.f5
Other methods such as fluorescent-antibody and immunoperoxidase techniques have been used for P. jirovecii (13–17). The development of tissue culture procedures for P. jirovecii may lead to additional diagnostic procedures for this infection. Immunoassay reagents are also available (14–18) and are being widely used in many diagnostic laboratories.
When noninduced sputum and the direct fluorescent-antibody technique were used, the sensitivity was 55%; this is within the range reported in the literature for the diagnosis of Pneumocystis pneumonia from induced sputum (19). Although detection of P. jirovecii has increased with the use of PCR, particularly in sputum specimens, some workers recommend that sputum samples of HIV-infected patients be tested by both PCR and immunofluorescence. The results of PCR in this patient group could be misleading without careful clinical evaluation (20). However, the level of sensitivity seen using PCR and other molecular tests indicates that these procedures should be considered for patients in whom Pneumocystis pneumonia is suggestive clinically and the specimen is negative by the immunofluorescence test (21, 22).
Quality Control Measures
Quality control measures for all stains should include the following, and control slides must be incorporated into all stain procedures.
1. Examine a control slide for each stain prior to examination of specimen stains. The stain intensity of controls will be a guide to the stain appearance of organisms in specimens.
2. Examine stained specimen smears by systematically moving from field to field until the majority of the smear has been covered (total area for silver stain).
3. In Giemsa stains, Pneumocystis trophozoite clumps of various sizes may be detected. In large clumps, it may be difficult to differentiate individual organisms. Look at organisms at the edges of clumps, and look for small, more dispersed clumps.
4. In silver- or other cyst wall-stained smears, look for the various cyst forms, including those that show dark centers, cup-shaped cysts, and cysts with foldlike lines (they look like “punched-in” ping-pong balls). If dark-staining organisms appear more oval, look carefully for budding forms, which indicate that the organisms are yeasts. It is important to review thinner areas of the smear for microsporidial spores.
A. P. jirovecii cysts: 70% should have delicately stained walls, usually brown or gray. They appear somewhat transparent, with structures described as “parentheses” staining black; these curved structures are usually thick (much thicker than the cyst wall) rather than thin like a line drawing.
B. Other fungi and actinomycetes: gray to black. Microsporidial spores stain dark gray to black; some of the spores may contain the polar tubule (horizontal or diagonal stripe), but they are more difficult to see in the silver stain than in the modified trichrome stains.
C. Glycogen, mucin, and red blood cells: rose taupe to gray.
D. Background: pale green (from the light green counterstain).
5. Detection of P. jirovecii in specimen smears from one type of stain always suggests a careful examination of the other stain, hopefully to confirm the identification.
6. Retain all positive stained specimen slides and control slides for reference.
Reporting Smear Results
1. Report P. jirovecii as follows (high dry power; total magnification, ×400).
A. “No Pneumocystis jirovecii seen.”
B. “Pneumocystis jirovecii seen” (no quantitation should be included).
2. Report other fungi that may be present as follows (low power; total magnification, ×100).
A. “No mycotic elements seen.”
B. “Budding yeasts” or “Budding yeasts and pseudohyphae resembling Candida species” are quantitated as few, moderate, or many.
C. Hyphae are reported with no quantitation:
a. “Septate hyphae seen.”
b. “Nonseptate hyphae seen.”
3. Report actinomycetes (oil immersion; total magnification, ×1,000) as follows.
A. “No filamentous branching bacteria seen.”
B. “Filamentous branching bacteria seen” (no quantitation).
4. Report microsporidia (oil immersion; total magnification, ×1,000) as follows.
A. “No microsporidial spores seen.”
B. “Microsporidial spores seen; identification to genus not possible.” However, if the specimen is stool or other gastrointestinal tract specimens, the organisms are probably Enterocytozoon bieneusi or Encephalitozoon intestinalis; a note containing this information can be attached to the report.
Notes on Staining Procedures
1. Use at least two stains for detection and identification of P. jirovecii. With traditional histochemical stains, a trophozoite stain such as Giemsa and a cyst wall stain such as methenamine-silver nitrate are recommended.
2. There are many cyst wall stains in addition to the one described, and there are other modifications of the silver stain (12, 23).
3. Stain effectiveness varies. Other counterstains may be used; a counterstain with Giemsa is useful if referral slides will be submitted for examination.
4. When selecting a cyst wall stain, consider stain quality, reagent stability, and potential testing frequency (including STAT requests). Toluidine blue O stains vary in dye lots and in the stability of sulfation reagents (24). In the modified procedure, the sulfation reagent made of glacial acetic acid and concentrated sulfuric acid presents disposal problems.
5. The rapid silver stain described is a modification of the procedure described by Brinn (11). Heating the chromic acid may cause nonspecific staining, making the background too dark. Also, buildup of silver on the stain container may interfere with staining. Bleach should be added to the stain jar after staining, and jars should be scrubbed periodically with a brush. Additional tips for getting good stain results include the following.
A. Before use, inspect glassware to ensure that it does not have residual silver deposits.
B. Use fresh reagents for each run (5% chromic acid, 1% sodium bisulfite, methenamine-silver nitrate working solution, and 2% sodium thiosulfate).
C. Distilled deionized water must be used throughout the procedure, including sliderinses (microwave procedure).
D. Methenamine-silver nitrate working solution must be clear; if it becomes opaque at any time, the reduction step may take longer or may not occur at all. Check the distilled deionized water source, and prepare fresh methenamine-silver nitrate working solution.
6. If mounted slides appear opaque or cloudy, the dehydration and clearing with xylene (or xylene substitute) were not adequate. Soak slides in xylene to remove the coverslips. Using fresh ethanol and xylene, repeat the dehydration steps.
7. Degenerating polymorphonuclear leukocytes may resemble P. jirovecii.
8. Monoclonal antibodies specific for human strain P. jirovecii are now available. The commercial systems vary; some are indirect stains, and some are direct stains. Reports with all systems have been variable. Many laboratories are now using these immunospecific stains.
9. Select an organism stain and a cyst wall stain or immunospecific stain; use of a pair of stains will help avoid both false-negative and false-positive reporting.
10. In addition to organism detection, cytocentrifuge preparations of sputum can be used to determine cell populations for further patient evaluation.
The examination of aspirated material for the diagnosis of parasitic infections may be extremely valuable, particularly when routine testing methods have failed to demonstrate the organisms. These types of specimen should be transported to the laboratory immediately after collection. Aspirates include liquid specimens collected from a variety of sites where organisms might be found. Aspirates most commonly processed in the parasitology laboratory include fine-needle aspirates and duodenal aspirates. Fluid specimens collected by bronchoscopy include bronchoalveolar lavage fluid and bronchial washings.
Procedural details for sigmoidoscopic aspirates and scrapings for the recovery of E. histolytica are presented in chapter 5. Techniques for preparation of duodenal aspirate material are also presented in that chapter.
Specimens Obtained from Aspiration Procedures
Fine-needle aspirates are often collected by cytopathology staff, who process the specimens, or they may be collected and sent to the laboratory directly for slide preparation and/or culture. Suggested stains are Giemsa and methenamine silver for P. jirovecii, Giemsa for Toxoplasma gondii (25), trichrome for amebae, modified acid-fast stains for Cryptosporidium (2), and modified trichrome stains for the microsporidia (see chapter 3).
Aspirates of cysts and abscesses to be evaluated for amebae may require concentration by centrifugation, digestion, microscopic examination for motile organisms in direct preparations, and cultures and microscopic evaluation of stained preparations.
Duodenal aspirates to be evaluated for S. stercoralis, Giardia lamblia (G. duodenalis, G. intestinalis). Cryptosporidium spp., Cyclospora cayetanensis, or the microsporidia require concentration by centrifugation prior to microscopic examination for motile organisms and permanent stains (26–29).
Cyst aspirates to be evaluated for hydatid “sand” (daughter cysts, small protoscolices, hooklets) can be examined as wet direct mounts after centrifugation. Several stains can be used to visualize the hooklets; one of the best is the Ryan blue modified trichrome stain for the microsporidia (see chapter 3). The stained smears can be examined using routine microscopy with transmitted light (30).
Bone marrow aspirates to be evaluated for Leishmania amastigotes, Trypanosoma cruzi amastigotes, or Plasmodium spp. require staining with Giemsa or another blood stain (31, 32).
Fluid specimens collected by bronchoscopy may be lavages or washings, with bronchoalveolar lavages preferred. Specimens usually are concentrated by centrifugation prior to microscopic examination of stained preparations. Organisms included here which may be detected are P. jirovecii, Toxoplasma gondii, and Cryptosporidium spp.
Pneumocystosis
Although formerly classified with the sporozoa, P. jirovecii has been reclassified with the fungi. It is an important cause of pulmonary infections, particularly in patients who are immunosuppressed as a result of therapy or from congenital or acquired immunologic deficiencies (33, 34). Clinically, the infection may involve both lungs diffusely, may be localized, or may be disseminated. For immunosuppressed patients, in whom the disease may progress very quickly, correct specimen collection and rapid diagnostic techniques are very important. The organisms can be demonstrated in stained impression smears of lung material obtained by open or brush biopsy. P. jirovecii can also be seen in stained smears of tracheobronchial aspirates, although examination of lung tissue is more likely to reveal the organisms. Sputum specimens are generally considered unacceptable for the recovery of P. jirovecii; however, in patients with severe, progressive disease, such specimens may be acceptable. To avoid the possibility of false-negative results, acceptance of sputum specimens should be carefully monitored and reviewed. Also, even for patients with fulminant disease, multiple specimens may have to be submitted to recover and identify the organisms. With a number of the stains that are available, the cyst walls stain but the cyst contents do not (Fig. 6.4); however, only the methenamine silver stain is discussed here (15).
Amebiasis
Examination of aspirates from lung or liver abscesses may reveal trophozoites of E. histolytica (Fig. 6.6); demonstration of the organisms is often very difficult. In many cases, serologic confirmation is recommended (35). Liver aspirate material should be taken from the margin of the abscess rather than the necrotic center (Fig. 6.7). The organisms are often trapped in the viscous pus or debris and may not exhibit typical motility. The Amoebiasis Research Unit, Durban, South Africa, has recommended using proteolytic enzymes to free the organisms from the aspirate material.
Figure 6.6 Entamoeba histolytica containing ingested red blood cells within the cytoplasm; morphologically this organism is E. histolytica, and this would be the species designation if organisms were isolated from liver abscess material (trichrome stain). doi:10.1128/9781555819002.ch6.f6
Figure 6.7 (Upper) Liver abscess caused by Entamoeba histolytica. Amebae would be found at the advancing margin of the lesion; the last portion of the aspirated material might reveal the organisms. (Illustration by Sharon Belkin.) (Lower) Amebic abscess; note the “flask”-shaped ulcer. doi:10.1128/9781555819002.ch6.f7
The digestion technique is performed as follows.
1. A minimum of two separate portions of exudate should be removed (more than two are recommended). The first portion of the aspirate, usually yellowish white, rarely contains organisms. The last portion of the aspirated abscess material is reddish and is more likely to contain amebae. The best material to examine is that obtained from the actual wall of the abscess.
2. Add 10 U of streptodornase per ml of thick pus; incubate this mixture for 30 min at 37°C, and shake repeatedly.
3. Centrifuge the mixture at 500 × g for 5 min. The sediment may be examined microscopically as wet mounts or used to inoculate culture media. Some of the aspirate can be mixed directly with a fecal fixative on a slide, allowed to air dry, stained, and examined as a permanent stained smear.
Hydatid Disease
Aspiration of cyst material for the diagnosis of hydatid disease is a dangerous procedure and is normally performed only when open surgical techniques are used for cyst removal. Aspirated fluid usually contains hydatid sand (intact and degenerating protoscolices, hooklets, and calcareous corpuscles) (Fig. 6.8). Some older cysts contain material that resembles curded cottage cheese, and the hooklets may be very difficult to see. Some of this material can be diluted with saline or 10% KOH; usually, protoscolices or daughter cysts will have disintegrated. However, the diagnosis can be made by seeing the hooklets, which can be stained using the Ryan blue modified trichrome stain (see chapter 3) (Fig. 6.9).
Figure 6.8 Hydatid disease (Echinococcus granulosus). (Upper left) Hydatid cyst with protoscolices budding off from the germinal layer. (Upper right) Immature protoscolices, with the dark area being the hooklets. (Lower left) Higher magnification of the protoscolices taken from the hydatid cyst fluid. (Lower right) Hooklets from disintegrating protoscolices. Reprinted from reference 3. doi:10.1128/9781555819002.ch6.f8
Figure 6.9 Hydatid disease (Echinococcus granulosus). (Upper) Hydatid protoscolex revealed by Ryan modified trichrome stain. (Lower) Hydatid hooklets revealed by Ryan blue modified trichrome stain (reprinted from reference 30). doi:10.1128/9781555819002.ch6.f9
Examination of hydatid cyst material is carried out as follows.
1. If the cyst material is fluid, centrifuge at 500 × g for 3 min.
2. Carefully remove some of the sediment and prepare a wet mount.
3. Examine the material under low (100×) and high dry (430×) power (remember to use low light intensity, since some of the material may be very transparent).
4. If the cyst material is more viscous or solid, the material can be mixed with saline or 10% KOH and then centrifuged at 500× g for 3 min.
5. Some of the very viscous material can be placed on a glass slide (undiluted). Another glass slide can be placed on top of the first (material will now lie between the two slides). Rub the glass slides back and forth over each other and listen for a grating sound (like grains of sand being scratched). If this occurs, you may be hearing evidence of the presence of calcified hooklets. Place the material from the smears (may be diluted with saline) under the microscope to try to confirm the presence of hooklets (which may be extremely difficult to see with only low power).
6. A dried smear of the cyst aspirate can be stained using the Ryan blue modified trichrome stain (see chapter 3). The hooklets will stain reddish-purple in color; remember they are quite small, measuring 20 to 40 µm long.
Note Remember, the absence of protoscolices or hooklets does not rule out the possibility of hydatid disease, since some cysts are sterile and contain no protoscolices and/or daughter cysts. Review of the cyst wall from pathology (tissue sections) should be able to confirm the diagnosis.
Lymph Nodes, Spleen, Liver, Bone Marrow, Spinal Fluid, Eyes, and Nasopharynx
African Trypanosomiasis, Leishmaniasis, Chagas’ Disease, Primary Amebic Meningoencephalitis, Granulomatous Amebic Encephalitis, Amebic Keratitis, and Microsporidial Keratitis
Material from lymph nodes, spleen, liver, bone marrow, spinal fluid, eye specimens, or nasopharynx may be examined for the presence of parasites and should be processed as follows.
1. A portion of the fluid material can be examined under low (×100) and high dry (×400) power as a wet mount (diluted with saline) for the presence of motile organisms. Spinal fluid should not be diluted before examination.
2. Impression smears from tissues should be prepared and stained with Giemsa or another blood stain. The material is pressed between two slides, and a smear is formed when the slides are pulled apart (one across the other).
3. The smears are allowed to air dry and then processed like a thin blood film (fixed in absolute methanol and stained with Giemsa stain or one of the other blood stains).
4. Appropriate culture media should be inoculated with any remaining material (see chapter 8).
5. If microsporidia are suspected, modified trichrome stains (Ryan blue, Weber green) can be used; calcofluor white and immunoassay methods (currently under development) are also excellent options (36–43).
Primary amebic meningoencephalitis is rare, but the examination of spinal fluid may reveal the amebae, usually Naegleria fowleri (Fig. 6.10). Granulomatous amebic encephalitis is caused by Acanthamoeba spp., Balamuthia mandrillaris, or Sappinia diploidea. Uncentrifuged sedimented spinal fluid should be placed under a coverslip on a slide and observed for motile amebae; smears can also be stained with Wright’s, Giemsa, or one of the other blood stains (including the rapid stains). Spinal fluid, exudate, or tissue fragments can be examined by light microscopy or phase-contrast microscopy. Care must be taken not to confuse leukocytes with actual organisms and vice versa. The appearance of the spinal fluid may vary from cloudy to purulent (with or without red blood cells), with a cell count from a few hundred to over 20,000 white blood cells (primarily neutrophils) per ml. Failure to find bacteria in this type of spinal fluid should alert one to the possibility of primary amebic meningoencephalitis. These organisms can be isolated from tissues or soil with special media (see chapter 8).
Figure 6.10 Naegleria fowleri. Note the large karyosome and lobate pseudopodia. Spinal fluid should be examined on a slide, not in a counting chamber, where protozoan trophozoites could mimic white blood cells. (Armed Forces Institute of Pathology photograph.) doi:10.1128/9781555819002.ch6.f10 (Armed Forces Institute of Pathology photograph.)
Note When spinal fluid is placed in a counting chamber, any organisms that settle to the bottom of the chamber will tend to round up and look very much like white blood cells. For this reason, it is better to examine the spinal fluid on a slide directly under a coverslip, not in a counting chamber.
A rapid method for the diagnosis of Acanthamoeba or microsporidial keratitis involves the use of calcofluor white, which is a chemofluorescent dye with an affinity for the polysaccharide polymers of amebic cysts and microsporidial spores (Fig. 6.11 and 6.12). The following method has proven to be very successful for examination of corneal scrapings or biopsy material (42).
Figure 6.11 Acanthamoeba cyst. Note the fluorescence of the cyst wall. Trophozoites do not fluoresce when calcofluor white is used. doi:10.1128/9781555819002.ch6.f11
Figure 6.12 Microsporidial spores. (Upper left) Spores in a corneal scraping specimen; (upper right) spores in a fecal specimen (both images stained with silver stain). (Lower left) Microsporidial spores in a urine sediment; (lower right) spores from nasopharyngeal aspirate (both images stained using calcofluor white). doi:10.1128/9781555819002.ch6.f12
1. Place corneal scrapings on slides, and air dry them.
2. Fix the slides in absolute methyl alcohol for 3 to 5 min.
3. Add several drops of solution (0.1% calcofluor white and 0.1% Evans blue dissolved in distilled water); leave on for 5 min.
4. Turn the slide on its side, and allow excess stain to run off onto paper towels.
5. Add a coverslip, and examine under the fluorescence microscope for pale blue chemofluorescence of the amebic cysts (will not stain trophozoite). The microsporidial spores usually stain brighter; however, fluorescence can vary between 1+ and 3+.
Note Select UV irradiation with an exciter filter which transmits the 365-nm group of intense mercury spectral emission lines (Zeiss UGI or G365). View through a barrier filter which removes UV while emitting visible blue light and longer wavelengths (Zeiss no. 41 or LP420).
Leishmaniasis
Material containing intracellular Leishmania organisms must be aspirated from below the ulcer bed through the uninvolved skin, not from the surface of the ulcer (Fig. 6.13). It is very important that the surface of the ulcer be thoroughly cleaned before specimens are taken; any contamination of the material with bacteria or fungi may prevent recovery of organisms from culture. Cutaneous ulcers can be seen in Fig. 6.14.
Figure 6.13 Proper way to aspirate material from below the ulcer bed (Leishmania spp.); sterile saline (0.5 to 1.0 ml) can be injected under the ulcer prior to aspiration. (Illustration by Sharon Belkin.) doi:10.1128/9781555819002.ch6.f13
Figure 6.14 Leishmania cutaneous lesions. doi:10.1128/9781555819002.ch6.f14
Biopsy specimens are recommended for the diagnosis of tissue parasites. The following procedures may be used for this purpose in addition to standard histologic preparations: impression smears and teased and squash preparations of biopsy tissue from the skin, muscle, corneas, intestines, liver, lungs, and brain. Tissue to be examined by permanent sections or electron microscopy should be fixed as specified by the laboratories that will process the tissue. In certain cases, examination of a biopsy specimen may be the only means of confirming a suspected parasitic infection. Specimens that are going to be examined as fresh material rather than as tissue sections should be kept moist in saline and submitted to the laboratory immediately.
Importance of Biopsy Specimens
Success in detection of parasites in tissue depends in part on specimen collection and on the presence of sufficient material to perform the recommended diagnostic procedures. Biopsy specimens are usually quite small and may not be representative of the diseased tissue. Examination of multiple tissue samples often improves diagnostic results. To optimize the yield from any tissue specimen, examine all areas and use as many procedures as possible. Tissues are obtained by invasive procedures, many of which are very expensive and lengthy; consequently, these specimens deserve the most comprehensive procedures possible. It is also important to remember that if the tissue specimen is not sufficient for all diagnostic procedures requested, the procedures should be prioritized after consultation with the physician.
Submission of Specimens
Tissue submitted on a sterile sponge dampened with saline in a sterile container may be used for cultures of protozoa after mounts for direct examination or impression smears for staining have been prepared. If cultures for parasites will be included, sterile slides should be used for smear and mount preparation.
1. Use sterile slides for impression smears; slides can be either autoclaved or prepared by an alternative method in which they are soaked in 95% ethyl alcohol and flamed prior to use.
2. Use sterile (autoclaved or flamed) forceps for handling tissue.
3. Place tissue in sterile petri dish to examine macroscopically and select a sample for microscopic evaluation. Provided that it is kept sterile, minced tissue can be used.
A. If biopsy tissue is several millimeters to a centimeter in size, select the specimen from an abnormal area (granulomatous lung or ulcerated area of intestinal tissue). Generally, the tissue fragments are too small to permit such a judgment to be made.
B. If several small pieces of tissue that look alike are submitted, use one piece. However, if they look different, use one of each type for microscopic examination. Again, sometimes this assessment to make.
4. Prepare impression smears.
A. Blot the tissue sample on sterile toweling. If the sample size is sufficient, cut the tissue with a scalpel (very sharp cut) and use the cut surface to touch the slide.
B. Press tissue against the sterile slide, lift, and press again. Turn the sample over, and press the other end against the slide to make two more impressions. Keep impressions close together to speed the screening process with the microscope.
C. Air dry and fix smears in absolute methanol for 1 min for subsequent blood stain, methenamine-silver nitrate, modified acid-fast, and modified trichrome staining. If the amount of tissue is sufficient, multiple smears should be prepared for each stain (Fig. 6.15).
Figure 6.15 (Upper) Leishmania donovani amastigotes in a cell; the one on the right is stained with Giemsa stain. (Lower left) Leishmania amastigote; note the nucleus and bar (primitive flagella). (Lower right) Cryptosporidium spp. oocysts stained with modified acid-fast stain. Note that the sporozoites are visible within some of the oocysts. doi:10.1128/9781555819002.ch6.f15
D. Place wet slide in Schaudinn’s fixative for subsequent trichrome staining.
E. Fix the slide as specified by the manufacturer for immunospecific staining.
5. Teased preparations are made as follows.
A. Place sample in the bottom of a plastic petri dish, and cover with 2 to 4 drops of saline.
B. Gently tease tissue apart with needles, or hold the tissue with forceps while pulling apart with a scalpel or needles.
C. Put a cover on dish, and leave at room temperature for 30 min.
6. Squash preparations are made as follows.
A. Using a scalpel, cut selected tissue portions into very fine pieces.
B. Place a piece of tissue on a 1- by 3-in. slide, add 1 drop of saline, cover with a second 1- by 3-in. slide, and hold together with membrane clips (from a surgical supply company). If these are not available, paper clips can be used but are not as efficient. You can also press the slides together with your fingers, but always wear gloves and be careful not to contaminate the microscope stage with tissue fluids.
7. Skin scrapings should be submitted in a small vial.
8. Prepare cultures to demonstrate the following organisms (see chapter 8): E. histolytica, Acanthamoeba and Naegleria spp., and Leishmania spp.
9. Mouse passage for Toxoplasma gondii:
A. Grind tissue in 0.85% NaCl into a fine suspension.
B. Inject 0.2 to 0.4 ml of suspension intraperitoneally into three to five mice of any laboratory strain weighing ∼20 g.
C. Maintain mice in isolation.
D. Every day, check for signs of central nervous system dysfunction; if symptoms are detected, perform an autopsy on the animal.
Examination of Specimens
1. Examination of impression smears is summarized in Table 6.1.
2. Examine skin snips in teased preparations for detection of microfilariae of Onchocerca volvulus and Mansonella streptocerca as follows.
A. Tease the small bit of tissue apart in a few drops of saline to release the microfilariae.
B. Remove drops of saline to a 1- by 3-in. glass slide, cover with a no. 1 coverslip, and examine under low light for microfilariae.
C. For a permanent preparation, place 100% methanol under the coverslip to fix filariae, partially dry, remove the coverslip, and stain with Giemsa or one of the other blood stains (including the rapid stains).
3. Examine squash preparations for detection of Trichinella spp. in muscle microscopically at low power (×100) and under low light.
4. Examine scrapings of skin for Sarcoptes scabiei (scabies) microscopically at low power (×100) and under low light. Confirmation at high dry power (×400) may be necessary (Fig. 6.16). See chapter 35 on arthropods for specific/detailed skin-scraping protocol for scabies.
Figure 6.16 Sarcoptes scabiei mites from skin scrapings. (Upper) Note the two mites to the left of center. (Lower) Note the two eggs, the small nymph, and the adult mite (photographed at a higher magnification). If the light is too strong when examining the scrapings in saline, the mites will probably not be seen. doi:10.1128/9781555819002.ch6.f16
5. Inoculate cultures with ground tissue suspensions (to release organisms from the cells).
A. Place a small tissue sample in a sterile tissue grinder (Ten Broeck or Dounce) in 0.5 ml of sterile saline, and grind until tissue is broken up but not totally liquefied.
B. Add several drops of ground tissue to culture medium as follows.
a. NNN medium (see chapter 8): add drops of tissue to the bottom of the slant, where they will “pool” with condensed moisture. Incubate at room temperature (isolation of Leishmania spp.).
b. TYSGM-9 medium (see chapter 8): add drops of tissue to the liquid medium, add 3 drops of the starch suspension, and incubate at a 45 to 50° angle at 35°C for 48 h (isolation of E. histolytica).
c. Nonnutrient agar plate seeded with bacteria (see chapter 8): add drops to center of seeded agar plate, and incubate at 35 to 37°C (isolation of Acanthamoeba or Naegleria spp.).
6. Examine cultures (wear gloves at all times).
A. NNN agar for promastigotes of Leishmania spp. (see chapter 8): using a Pasteur pipette, remove a drop of fluid from interface of agar slant and culture tube, place on glass slide, cover with coverslip, and examine microscopically (×400) under low light for motile promastigotes (Fig. 6.17).
Figure 6.17 Leishmania culture sediment. Note the promastigotes from the culture sediment. The typical “rosette” formation in the right image is frequently seen. doi:10.1128/9781555819002.ch6.f17
B. TYSGM-9 medium for amebae (see chapter 8): using a sterile Pasteur pipette, remove 1 or 2 drops of material from interphase of agar and overlay, place on glass slide, cover with coverslip, and examine microscopically (×100) under low light for trophozoite motility.
C. Nonnutrient agar plate with lawn of Escherichia coli or Enterobacter sp. for detection of free-living amebae (see chapter 8): examine microscopically at low power (×100) for changes in bacterial lawn, particularly patches and tracks, indicating that the protozoan trophozoites have ingested bacteria as they move over the agar.
D. Mouse passage for detection of Toxoplasma gondii:
a. Wear canvas gloves to handle mice; sacrifice the animal(s).
b. Pin mouse to board, spray with 70% ethyl alcohol, and open peritoneal cavity with sterile scissors.
c. Using a sterile Pasteur pipette, remove fluid from the peritoneal cavity, and place in small tube or prepare smears.
d. Using one of the blood stains, prepare stained smears of peritoneal exudate, and examine microscopically at a magnification of ×1,000 for T. gondii tachyzoites.
e. Place mice and all contaminated disposable materials in bag to be autoclaved and destroyed. Place nondisposable materials (scissors) in bag to be autoclaved prior to washing.
7. Correlate all examination results (wet mount, stains, and cultures) to determine presence of organisms.
Results
The majority of the protozoa are found on the permanent stained smears (impression smears, touch or squash preparations, or teased preparations). When culture is used, permanent stained smears of the culture medium or sediment may also reveal some of the protozoa. Although infrequently used, material from animals (at autopsy) can be examined as both wet and permanent stained preparations for confirmation of protozoa. Filarial infections may be confirmed by the recovery and identification of microfilariae in skin scrapings and/or biopsy specimens.
If identifications are certain, report the organisms detected; if a presumptive identification is made, confirmation by another laboratory is suggested.
Onchocerca volvulus and Mansonella streptocerca
The use of skin snips is the method of choice for the diagnosis of human filarial infections with O. volvulus and M. streptocerca. Microfilariae of both species occur chiefly in the skin, although O. volvulus microfilariae are on rare occasions found in the blood and occasionally in the urine. For best results, the skin snip specimens should be thick enough to include the outer part of the dermal papillae. Snips may be taken in various ways. A small slice may be cut (using a razor blade) from a skin fold held between thumb and forefinger, or a slice may be taken from a small “cone” of skin pulled up by a needle. The skin snip should be so thin that significant bleeding does not occur, just a slight oozing of fluid. Corneal-scleral punches (either Holth or Walser type) have been found to be successful in taking skin snips of uniform size and depth and an average weight of 0.8 mg (range, 0.4 to 1.2 mg); this procedure is easy to perform and is painless. It has been demonstrated that in African onchocerciasis, it is preferable to take skin snips from the buttock region (above the iliac crest); in Central American onchocerciasis, the preferred skin snip sites are from the shoulders (over the scapula).
Skin snips are placed immediately in a drop of normal saline or distilled water and covered so that they will not dry; teasing the specimen with dissecting needles is not necessary but may facilitate release of the microfilariae. Microfilariae tend to emerge more rapidly in saline; however, in either fluid, the microfilariae usually emerge within 30 min to 1 h and can be examined under low-intensity light with the 10× objective of the microscope. To see definitive morphologic details of the microfilariae, allow the snip preparation to dry, fix it in absolute methyl alcohol, and stain it with Giemsa or one of the other blood stains (Fig. 6.18). For the differential diagnosis of O. volvulus and M. streptocerca, see chapter 23.
Figure 6.18 Microfilariae (Giemsa stain). (Upper) In this image, the microfilariae from a skin snip saline preparation are visible after staining. (Lower) Microfilariae from blood; note that the characteristic nuclei are more easily seen in this preparation. doi:10.1128/9781555819002.ch6.f18
Cutaneous Amebiasis and Cutaneous Leishmaniasis
Skin biopsy specimens for the diagnosis of cutaneous amebiasis and cutaneous leishmaniasis should be processed for tissue sectioning and subsequently stained by the hematoxylin-eosin technique.
Itch Mites (Sarcoptes scabiei)
Several genera infect the skin of mammals, with S. scabiei being found in humans. S. scabiei is microscopic and lives in cutaneous burrows, where the fertilized female deposits eggs (Fig. 6.16). Scabies is transmitted by close contact with infested individuals, including touching, shaking hands, sexual contact, or day care centers with children or the elderly. The usual skin sites that are susceptible to infection are the interdigital spaces, backs of the hands, elbows, axillae, groin, breasts, umbilicus, penis, shoulder blades, small of the back, and buttocks. The most dramatic symptom is intense itching. Scratching commonly causes weeping, bleeding, and sometimes secondary infection. Specific and detailed diagnostic methods for the recovery of mites, eggs, or scybala (fecal pellets) can be found in the chapter on arthropods (chapter 35).
Trypanosomiasis, Leishmaniasis, Chagas’ Disease, and Toxoplasmosis
Material obtained from lymph nodes should be processed for tissue sectioning and as impression smears that should be processed as thin blood films and stained with Giemsa or other blood stains. Appropriate culture media can also be inoculated, again making sure that the specimen has been collected under sterile conditions.
Trichinosis
The presumptive diagnosis of trichinosis is often based on a combination of facts: history of ingestion of raw or rare pork, walrus meat, horse meat, or bear meat; diarrhea followed by edema and muscle pain; and the presence of eosinophilia. Usually, by the time the patient is symptomatic, the suspected food is no longer available for examination. The diagnosis may be confirmed by finding larval Trichinella spp. in a muscle biopsy specimen. The encapsulated larvae (which may be few) are easily seen in fresh muscle if small pieces are pressed between two slides and examined under the microscope (Fig. 6.19). Larvae are usually most abundant in the diaphragm, masseter muscle, or tongue and may be recovered from these muscles at necropsy. Routine histologic sections can also be prepared.
Figure 6.19 Trichinella spp. larvae in a squash preparation of muscle biopsy tissue; note the encysted larvae. doi:10.1128/9781555819002.ch6.f19
A portion of the specimen can be digested in artificial digestive fluid at 37°C, with the larvae recovered by centrifugation; however, very young larvae might also be digested.
Artificial Digestive Fluid
1. Prepare the tissue by grinding (with a tissue grinder or commercial meat grinder or blender). The tissue should be medium grind; do not overgrind, particularly in a blender.
2. Add the tissue to the digestive fluid in the ratio of 1 part tissue to 20 parts fluid in an Erlenmeyer flask.
3. Place the flask in an incubator at 37°C, on a shaker table or magnetic stirrer if available; leave for 12 to 24 h.
4. Add warm water (37°C) to the digestive fluid mixture. (Add enough water to triple the original volume.)
5. Pour the diluted mixture into a Baermann funnel, and add water up to the screen.
6. Allow the mixture to stand for 1 to 2 h; the larvae will settle out in the lower part of the funnel.
7. Remove a few drops of fluid, and examine the material under the microscope (10× objective) for the presence of larvae.
8. If no larvae are present, centrifuge 50 ml and examine the sediment.
Cestode Larval Stages
Human infection with any of the larval cestodes may present diagnostic problems, and frequently the larvae are referred for identification after surgical removal. In addition to Echinococcus granulosus (hydatid disease) and the larval stage of Taenia solium (cysticercosis), other larval cestodes occasionally cause human disease. The larval stage of tapeworms of the genus Multiceps, a parasite of dogs and wild canids, is called a coenurus and may cause human coenurosis. The coenurus resembles a cysticercus but is larger and has multiple scolices developing from the germinal membrane surrounding the fluid-filled bladder. These larvae occur in extraintestinal locations, including the eye, central nervous system, and muscle (see chapter 25).
Human sparganosis is caused by the larval stages of tapeworms of the genus Spirometra, which are parasites of various canine and feline hosts; these tapeworms are closely related to the genus Diphyllobothrium. Sparganum larvae are elongated, ribbonlike larvae without a bladder and with a slightly expanded anterior end lacking suckers. They are usually found in superficial tissues or nodules, although they may cause ocular sparganosis, a more serious disease (see chapter 25).
The diagnosis of larval cestodes is frequently facilitated by the recognition of prominent calcareous corpuscles occurring in the tapeworm tissue; specific identification usually depends on referral to specialists.
Schistosomiasis
Often when a patient has an old, chronic infection or a light infection with Schistosoma mansoni or S. japonicum, the eggs may not be found in the stool and an examination of the rectal mucosa may reveal the presence of eggs. The fresh tissue should be compressed between two microscope slides and examined under the low power of the microscope (low-intensity light). Critical examination of these eggs should be made to determine whether living miracidia are still found within the egg. Treatment may depend on the viability of the eggs; for this reason, the condition of the eggs should be reported to the physician.
Mucosa from the bladder wall may reveal eggs of Schistosoma haematobium when they are not being recovered in the urine. As with rectal biopsy specimens, the eggs in the bladder wall should be checked for viability (fresh, unfixed tissue) (Fig. 6.20).
Figure 6.20 Schistosoma haematobium eggs seen in a squash preparation of a bladder biopsy specimen. Note the pointed terminal spine. (Armed Forces Institute of Pathology photograph.) doi:10.1128/9781555819002.ch6.f20
Viability Testing of Schistosome Eggs
1. With careful observation of the egg, using the 40× objective (low-intensity light), the cilia of the flame cells of the miracidium within the shell can be seen to move in a rapid flickering motion.
2. The eggs within the tissue can be removed (carefully tease the tissue apart in saline solution) and subjected to a hatching procedure. If the miracidia are released from the egg and swim to the top of the hatching flask, this movement is also proof of viability. (The procedure can be found in chapter 4.)
Digestion Procedure in Diagnosis of Schistosomiasis
Small pieces of tissue may be digested in 4% potassium hydroxide (4% sodium hydroxide may be used) for 2 to 3 h at 60 to 80°C. The material may then be concentrated by sedimentation or centrifugation and examined under the microscope for eggs (low power with low-intensity light).
References
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