Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 13
ОглавлениеAmong the diagnostic techniques used with stool specimens, the routine ova and parasite (O&P) examination is the best known. This technique has three components: the direct wet mount, the examination of material from a stool concentrate, and the permanent stained smear. This is an excellent procedure and is recommended for most intestinal parasites. However, several other diagnostic techniques are available for the recovery and identification of parasitic organisms. Most laboratories do not routinely offer all of these techniques, but many are relatively simple and inexpensive to perform (1–4). The clinician should be aware of the possibilities and the clinical relevance of information obtained from using such techniques. Occasionally, it is necessary to examine stool specimens for the presence of scolices and proglottids of cestodes and adult nematodes and trematodes to confirm the diagnosis and/or for species identification. A method for the recovery of these stages is also described in this chapter. Although not routinely performed, tests for fecal fat and reducing substances are also included.
Culture of Larval-Stage Nematodes
Nematode infections giving rise to larval stages that hatch in soil or in tissues may be diagnosed by using certain fecal culture methods to concentrate the larvae. Strongyloides stercoralis larvae are generally the most common larvae found in stool specimens. However, depending on the fecal transit time through the intestine and the patient’s condition, rhabditiform and, rarely, filariform larvae may be present. Also, if there is a delay in examination of the stool, embryonated ova as well as larvae of hookworm may be present. Culture of feces for larvae is useful to (i) reveal their presence when they are too scanty to be detected by concentration methods; (ii) distinguish whether the infection is due to S. stercoralis or hookworm on the basis of rhabditiform larval morphology by allowing hookworm egg hatching to occur, releasing first-stage larvae; and (iii) allow the development of larvae into the filariform stage for further differentiation.
The use of certain fecal culture methods (sometimes referred to as coproculture) is especially helpful for detection of light infections with hookworm, S. stercoralis, and Trichostrongylus spp. and for specific identification of parasites. The rearing of infective-stage nematode larvae also helps in the specific diagnosis of hookworm and trichostrongyle infections because the eggs of many of these species are identical and specific identifications are based on larval morphology. Additionally, such techniques are useful for obtaining a large number of infective-stage larvae for research purposes. Four culture techniques and one “enhanced-recovery” method are described in this chapter.
Harada-Mori Filter Paper Strip Culture
To detect light infections with hookworm, S. stercoralis, and Trichostrongylus spp., as well as to facilitate specific identification, the Harada-Mori filter paper strip culture technique is very useful (Fig. 4.1). This filter paper test tube culture technique was initially introduced by Harada and Mori in 1955 (5) and was later modified by others (6, 7).
Figure 4.1 Culture methods for the recovery of larval-stage nematodes: Harada-Mori tube method and petri dish culture method. Viable larvae are present if the specimen contains S. stercoralis or other nematodes. Wear gloves when handling the culture devices. (Illustration by Nobuko Kitamura.) doi:10.1128/9781555819002.ch4.f1
In a study looking at the prevalence of S. stercoralis in three areas of Brazil, the diagnostic efficacy of the agar plate culture method (discussed later in this chapter) was as high as 94.9% compared to only 28.5 and 26.5% by the Harada-Mori filter paper culture and fecal concentration methods, respectively, when fecal specimens were processed using all three methods (8). Among the 49 positive samples, about 60% were confirmed as positive by using the agar plate method alone. These results indicate that the agar plate approach is probably a much more sensitive diagnostic method than the other two and is recommended for the diagnosis of strongyloidiasis.
The technique requires filter paper to which fresh fecal material is added and a test tube into which the filter paper is inserted. Moisture is provided by adding water to the tube, which continuously soaks the filter paper by capillary action. Incubation under suitable conditions favors hatching of ova and/or development of larvae. Fecal specimens to be cultured should not be refrigerated, since some parasites (especially Necator americanus) are susceptible to cold and may fail to develop after refrigeration. Also, caution must be exercised in handling the filter paper strip itself, since infective Strongyloides larvae may migrate upward as well as downward on the paper strip. Always observe standard precautions and wear gloves when performing these procedures.
Quality Control for Harada-Mori Filter Paper Strip Culture
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitologic examination.
2. Examine known positive and negative samples of stools (from laboratory animals), if available, to gain some experience in using the procedure.
3. Review larval diagrams and descriptions for confirmation of larval identification.
4. The microscope should be calibrated, and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. If the microscope undergoes hard use or is moved around within the laboratory, it is strongly recommended that recalibration be performed every 12 months. However, if the microscope remains in the same location and receives normal use, such frequent recalibration may not be required. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
5. Record all quality control (QC) results.
Procedure for Harada-Mori Filter Paper Strip Culture
1. Smear 0.5 to 1 g of fresh feces in the center of a narrow strip of filter paper (3/8 by 5 in. [1 in. = 2.54 cm], slightly tapered at one end).
2. Add 3 to 4 ml of distilled water to a 15-ml conical centrifuge tube; identify the specimen on the tube.
3. Insert the filter paper strip into the tube so that the tapered end is near the bottom of the tube. The water level should be approximately 1/2 in. below the fecal spot. It is not necessary to cap the tube. However, a cork stopper or a cotton plug may be used.
4. Maintain the tube upright in a rack at 25 to 28°C. Add distilled water to maintain the original level (usually evaporation takes place over the first 2 days, and then the culture becomes stabilized).
5. Keep the tube for 10 days, and check it daily by withdrawing a small amount of fluid from the bottom of the tube. Prepare a smear on a glass slide, cover the slide with a coverslip, and examine the smear with the 10× objective.
6. Examine the larvae for motility and typical morphological features to reveal whether hookworm, Strongyloides, or Trichostrongylus larvae are present.
Results and Patient Reports from Harada-Mori Filter Paper Strip Culture
Larval nematodes of hookworm, S. stercoralis, or Trichostrongylus spp. may be recovered. If Strongyloides organisms are present, free-living stages and larvae may be found after several days in culture.
1. Report “No larvae detected” if no larvae could be detected at the end of the incubation.
2. Report larvae detected by fecal culture.
Example: Strongyloides stercoralis larvae detected by fecal culture
Procedure Notes for Harada-Mori Filter Paper Strip Culture
1. If the larvae are too active to observe under the microscope and morphologic details are difficult to see, the larvae can be heat killed within the tube or after removal to the slide; iodine can also be used to kill larvae.
2. Infective larvae may be found any time after the fourth day or even on the first day in a heavy infection. Since infective larvae may migrate upward as well as downward on the filter paper strip, caution must be exercised in handling the fluid and the paper strip itself to prevent infection. Handle the filter paper with forceps. Wear gloves when handling the cultures.
3. It is important to maintain the original water level to keep optimum humidity.
4. Fresh stool is required for this procedure; preserved fecal specimens or specimens obtained after a barium meal are not suitable.
Procedure Limitations for Harada-Mori Filter Paper Strip Culture
1. The Harada-Mori technique allows both parasitic and free-living forms of nematodes to develop. If specimens have been contaminated with soil or water containing these forms, it may be necessary to distinguish parasitic from free-living forms. This distinction is possible since parasitic forms are more resistant to slight acidity than are free-living forms. Proceed as follows (9, 10).
Add 0.3 ml of concentrated hydrochloric acid per 10 ml of water containing the larvae (adjust the volume accordingly to achieve a 1:30 dilution of acid). Free-living nematodes are killed, while parasitic species live for about 24 h.
2. Specimens that have been refrigerated or preserved are not suitable for culture. Larvae of certain species are susceptible to cold environments.
Filter Paper/Slant Culture Technique (Petri Dish)
An alternative technique for culturing Strongyloides larvae is a filter paper/slant culture on a microscope slide placed in a glass or plastic petri dish (Fig. 4.1), which was originally described by Little (11). As with previous techniques, sufficient moisture is provided by continuous soaking of the filter paper in water. Fresh stool material is placed on the filter paper, which is cut to fit the dimensions of a standard (1 by 3 in.) microscope slide. The filter paper is then placed on a slanted glass slide in a glass or plastic petri dish containing water. This technique allows direct examination of the culture system with a dissecting microscope to look for nematode larvae and free-living stages of S. stercoralis in the fecal mass or the surrounding water without having to sample the preparation. Always wear gloves when performing these procedures.
Quality Control for the Filter Paper/Slant Culture Technique (Petri Dish)
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitologic examination.
2. Examine known positive and negative samples of stools (from laboratory animals), if available, to make sure that the procedure is precise.
3. Review larval diagrams and descriptions for confirmation of larval identification.
4. The microscope should be calibrated, and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
5. Record all QC results.
Procedure for the Filter Paper/Slant Culture Technique (Petri Dish)
1. Cut a filter paper strip (1 by 3 in.), and smear a film of 1 to 2 g of fresh fecal material in the center of the strip.
2. Place the strip on a glass slide (1 by 3 in.). Place the slide inclined at one end of the petri dish by resting the slide on a piece of glass rod or glass tubing; identify the specimen on the dish (Fig. 4.1).
3. Add water to the petri dish so that at least the bottom one-fourth of the slide is immersed in water. The stool will be kept moist by capillary action. Cover the dish, and maintain it at 25 to 28°C. As needed, add water to maintain the original level.
4. Keep the dish for 10 days. Examine daily, either with the dissecting microscope or by withdrawing a small amount of fluid and placing it on a microscope slide. Cover with a coverslip, and examine microscopically with the 10× and 40× objectives.
5. Examine any larvae recovered for typical morphologic features.
Results and Patient Reports from the Filter Paper/Slant Culture Technique (Petri Dish)
Larval nematodes of hookworm, S. stercoralis, or Trichostrongylus spp. may be recovered. If Strongyloides organisms are present, free-living stages and larvae may be found after several days in culture.
1. Report “No larvae detected” if no larvae could be detected at the end of incubation.
2. Report larvae detected by fecal culture.
Example: Strongyloides stercoralis larvae detected by fecal culture
Procedure Notes for the Filter Paper/Slant Culture Technique (Petri Dish)
1. It is often difficult to observe details in rapidly moving larvae; a drop of iodine or formalin or slight heating can be used to kill the larvae.
2. Infective larvae may be found any time after the fourth day and occasionally after the first day in heavy infections. Since infective larvae may migrate anywhere on the filter paper strip, caution must be exercised in handling the fluid and the paper strip itself to prevent infection. Wear gloves when handling the cultures.
3. There may be infective larvae in the moisture that accumulates under the petri dish lid, so be careful not to allow the water to touch the skin when raising the lid.
4. It is important to maintain the original water level to keep optimum humidity.
5. Preserved fecal specimens or specimens obtained after a barium meal are not suitable; fresh stool specimens must be used.
Procedure Limitations for the Filter Paper/Slant Culture Technique (Petri Dish)
1. The filter paper/slant culture technique allows both parasitic and free-living forms of nematodes to develop. If specimens have been contaminated with soil or water containing these forms, it may be necessary to distinguish parasitic from free-living forms. This distinction is possible since parasitic forms are more resistant to slight acidity than are free-living forms. Proceed as follows (9, 10).
Add 0.3 ml of concentrated hydrochloric acid per 10 ml of water containing the larvae (adjust the volume accordingly to achieve a 1:30 dilution of acid). Free-living nematodes are killed, while parasitic species live for about 24 h.
2. Specimens that have been refrigerated or preserved are not suitable for culture. Larvae of certain species are susceptible to cold environments.
Another way to culture hookworm, Strongyloides, and trichostrongyle larvae is by using a granulated charcoal culture. The conditions of this culture provide an environment for larval development that mimics conditions in nature. It provides an efficient way to harvest large numbers of infective-stage larvae for use in experimental infections.
Quality Control for Charcoal Culture
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitologic examination.
2. Examine known positive and negative samples of stools (from laboratory animals), if available, to make sure that the procedure is precise.
3. Review larval diagrams and descriptions for confirmation of larval identification.
4. The microscope should be calibrated, and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
5. Record all QC results.
Procedure for Charcoal Culture
1. Mix 20 to 40 g of fresh fecal material in tap water until a thick suspension is obtained.
2. Add this suspension to a storage dish (4 by 3 in.) that is slightly more than half filled with no. 10 granulated hardwood charcoal. Mix thoroughly with a wooden tongue depressor until the fecal suspension is evenly distributed throughout the moistened charcoal. Water can be added to ensure that there is adequate moisture, but do not add so much water than it forms a layer on the bottom of the dish. The surface of the charcoal should glisten.
3. Cover the dish, and place it in the dark (in a drawer or cabinet).
4. Check the dish the next day to make sure that there is still sufficient moisture (i.e., that the charcoal still glistens); if water is needed, sprinkle it on the surface without further mixing.
5. Check the dish each day for moisture content. Caution must be used because moisture will accumulate on the underface of the lid, and it may contain infective-stage larvae.
6. Approximately 5 or 6 days after the culture has been prepared, hookworm and Strongyloides larvae will have reached the infective stage. This can occur earlier if the patient has a heavy infection with numerous larvae in the stool.
7. To harvest the larvae, prepare a round gauze pad of 10 to 12 layer thickness stapled at the edges and cut to fit the dish. Moisten the pad (not dripping wet), and apply it carefully with forceps so that it snugly covers the surface of the charcoal. Do not allow your hands to touch the charcoal—the larvae will be infective!
8. Expose the dish, with lid off, to a light source such as a gooseneck lamp. The lamp should be 6 to 8 in. from the surface of the charcoal. Make sure the lamp is not too close; the larvae can be killed by the heat.
9. After approximately 1 h, the pad can be carefully removed with forceps and inverted onto the surface of water in a pilsner glass filled with water. The gauze pad will remain at the top, and the larvae will now make their way through the pad, enter the water, and fall to the bottom of the glass, where they can be harvested with a pipette after another 30 to 60 min. With care, there will be no charcoal at the bottom of the glass, and the larvae will form a clean sediment.
Results and Patient Reports from Charcoal Culture
Larval nematodes of hookworm, S. stercoralis, or Trichostrongylus spp. may be recovered. If Strongyloides organisms are present, free-living stages and larvae may be found after several days in culture.
1. Report “No larvae detected” if no larvae could be detected at the end of incubation.
2. Report larvae detected by fecal culture.
Example: Strongyloides stercoralis larvae detected by fecal culture
Procedure Notes for Charcoal Culture
1. It is often difficult to observe details in rapidly moving larvae; a drop of iodine or formalin or slight heating can be used to kill the larvae.
2. Infective larvae may be found any time after the fourth day and occasionally after the first day in heavy infections. Since infective larvae may be present, use caution when handling the fluid, gauze pad, and charcoal to prevent infection. Wear gloves when handling the cultures.
3. It is important to maintain the moisture on the charcoal to keep optimum humidity (the charcoal should glisten).
4. Preserved fecal specimens or specimens obtained after a barium meal are not suitable; fresh stool specimens must be obtained.
Procedure Limitations for Charcoal Culture
1. This technique allows both parasitic and free-living forms of nematodes to develop. If specimens have been contaminated with soil or water containing these forms, it may be necessary to distinguish parasitic from free-living forms. This distinction is possible since parasitic forms are more resistant to slight acidity than are free-living forms. Proceed as follows (9, 10).
Add 0.3 ml of concentrated hydrochloric acid per 10 ml of water containing the larvae (adjust the volume accordingly to achieve a 1:30 dilution of acid). Free-living nematodes are killed, while parasitic species live for about 24 h.
2. Specimens that have been refrigerated or preserved are not suitable for culture. Larvae of certain species are susceptible to cold environments.
Another method of examining a stool specimen suspected of containing small numbers of Strongyloides larvae is the use of a modified Baermann apparatus (Fig. 4.2). The Baermann technique, in which a funnel apparatus is used, relies on the principle that active larvae will migrate from a fresh fecal specimen that has been placed on a wire mesh with several layers of gauze which are in contact with tap water (1, 12). Larvae migrate through the gauze into the water and settle to the bottom of the funnel, where they can be collected and examined. The main difference between this method and the Harada-Mori and petri dish methods is the greater amount of fresh stool used, possibly providing a better chance of larval recovery in a light infection. Besides being used for patient fecal specimens, this technique can be used to examine soil specimens for the presence of larvae.
Figure 4.2 (Upper) Baermann apparatus. (Illustration by Nobuko Kitamura.) (Lower) Baermann apparatus set up in the laboratory. doi:10.1128/9781555819002.ch4.f2
Quality Control for the Baermann Technique
1. Follow routine procedures for optimal collection and handling of fresh specimens for parasitologic examination.
2. Examine known positive and negative samples of stools (from laboratory animals), if available, to make sure that the procedure is precise.
3. Review larval diagrams for confirmation of larval identification.
4. The microscope should be calibrated, and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
5. Record all QC results.
Procedure for the Baermann Technique
1. If possible, use a fresh fecal specimen that has been obtained after administration of a mild saline cathartic, not a stool softener. Soft stool is recommended; however, any fresh fecal specimen is acceptable.
2. Set up a clamp supporting a 6-in. glass funnel. Attach rubber tubing and a pinch clamp to the bottom of the funnel. Place a collection beaker underneath (Fig. 4.2).
3. Place a wire gauze or nylon filter over the top of the funnel, followed by a pad consisting of two layers of gauze.
4. Close the pinch clamp at the bottom of the tubing, and fill the funnel with tap water until it just soaks the gauze padding.
5. Spread a large amount of fecal material on the gauze padding so that it is covered with water. If the fecal material is very firm, first emulsify it in water.
6. Allow the apparatus to stand for 2 h or longer; then draw off 10 ml of fluid into the beaker by releasing the pinch clamp, centrifuge the fluid for 2 min at 500 × g, and examine the sediment under the microscope (magnification, ×100 and ×400) for the presence of motile larvae. Make sure that the end of the tubing is well inside the beaker before slowly releasing the pinch clamp. Infective larvae may be present; wear gloves when performing this procedure.
Results and Patient Reports from the Baermann Technique
Larval nematodes (hookworm, S. stercoralis, or Trichostrongylus spp.) may be recovered. Both infective and noninfective Strongyloides larvae may be recovered, particularly in a heavy infection.
1. Report “No larvae detected” if no larvae could be detected at the end of incubation.
2. Report larvae detected by fecal culture.
Example: Strongyloides stercoralis larvae detected by fecal culture
Procedure Notes for the Baermann Technique
1. It may be difficult to observe morphologic details in rapidly moving larvae; a drop of iodine or formalin or slight heating can be used to kill the larvae.
2. Infective larvae may be found any time after the fourth day and occasionally after the first day in heavy infections. Caution must be exercised in handling the fluid, gauze pad, and beaker to prevent infection. Wear gloves when using this technique.
3. Remember to make sure that the pinch clamp is tight until you want to release some of the water.
4. Preserved fecal specimens or specimens obtained after a barium meal are not suitable for processing by this method; fresh stool specimens must be obtained.
Procedure Limitations for the Baermann Technique
1. The Baermann technique allows both parasitic and free-living forms of nematodes to develop. If specimens have been contaminated with soil or water containing these forms, it may be necessary to distinguish parasitic from free-living forms. This distinction is possible since parasitic forms are more resistant to slight acidity than are free-living forms. Proceed as follows (9, 10).
Add 0.3 ml of concentrated hydrochloric acid per 10 ml of water containing the larvae (adjust the volume accordingly to achieve a 1:30 dilution of acid). Free-living nematodes are killed, while parasitic species live for about 24 h.
2. Specimens that have been refrigerated or preserved are not suitable for culture. Larvae of certain species are susceptible to cold environments.
3. Gloves should be worn when this procedure is performed.
4. Release the pinch clamp slowly to prevent splashing; have the end of the tubing close to the bottom of the beaker for the same reason.
Modification of the Baermann Method
A simple modification of the Baermann method for diagnosis of strongyloidiasis has been developed (13). For this modification, the funnel used in the original version is replaced by a test tube with a rubber stopper, perforated to allow insertion of a plastic pipette tip (Fig. 4.3). The tube containing the fecal suspension is inverted over another tube containing 6 ml of saline solution and incubated at 37°C for at least 2 h. The saline solution from the second tube is centrifuged, and the pellet is observed microscopically as a wet mount. Larvae of S. stercoralis can be found in the pellet. Although the method is almost identical to the original Baermann method, the amount of stool used in the modified method is smaller.
Figure 4.3 Baermann apparatus modification. (Drawing by Sharon Belkin; adapted from reference 13.) doi:10.1128/9781555819002.ch4.f3
Agar Plate Culture for Strongyloides stercoralis
Agar plate cultures are also recommended for the recovery of S. stercoralis larvae and tend to be more sensitive than some of the other diagnostic methods (14–22). It is important to remember that more than half of S. stercoralis-infected individuals tend to have low-level infections (23). The agar plate method continues to be documented as a more sensitive method than the usual direct smear or fecal concentration methods (16–19). Daily search for furrows on agar plates for up to six consecutive days results in increased sensitivity for diagnosis of both S. stercoralis and hookworm infections. Also, a careful search for S. stercoralis should be performed for all patients with comparable clinical findings before a diagnosis of idiopathic eosinophilic colitis is made, because the consequent steroid treatment may have a fatal outcome by inducing widespread dissemination of the parasite (24). Human T-cell leukemia virus type 1 (HTLV-1) infection is endemic in a number of Latin American countries. HTLV-1-associated myelopathy/tropical spastic paraparesis and adult T-cell leukemia-lymphoma are emerging diseases in the region. S. stercoralis hyperinfection syndrome and therapeutic failure in apparently healthy patients with nondisseminated strongyloidiasis may be markers of HTLV-1 infection (25).
Stool is placed onto agar plates, and the plates are sealed to prevent accidental infections and held for 2 days at room temperature. As the larvae crawl over the agar, they carry bacteria with them, thus creating visible tracks over the agar (Fig. 4.4). The plates are examined under the microscope for confirmation of the presence of larvae, the surface of the agar is then washed with 10% formalin, and final confirmation of larval identification is made via wet examination of the sediment from the formalin washings (Fig. 4.5).
Figure 4.4 Agar plate culture method for Strongyloides stercoralis. (Upper) Agar plate showing bacterial growth after being distributed over the plate by the movement of the larval worms. (Lower) More random pattern of bacterial growth from the inoculation of bacteria over the agar from the movement of the larval worms (image courtesy of Audrey N. Schuetz, Weill Cornell Medical College). doi:10.1128/9781555819002.ch4.f4
Figure 4.5 Agar culture method for Strongyloides stercoralis. (1) Agar plates are prepared. (2) Agar is dried for 4 to 5 days on the bench top. (3) Plates are stored in plastic bags. (4) Fresh stool is submitted to the laboratory. (5) Approximately 2 g of stool is placed onto an agar plate. (6) The plate is sealed with tape. (7) The culture plate is incubated at 26 to 33°C for 2 days. (8) The plate is examined microscopically for the presence of tracks (bacteria carried over agar by migrating larvae). (9) 10% formalin is placed onto agar through a hole made in the plastic with hot forceps. (10) Material from the agar plate is centrifuged. (11) The material is examined as a wet preparation for rhabditiform or filariform larvae (high dry power; magnification, ×400). (Illustration by Sharon Belkin.) doi:10.1128/9781555819002.ch4.f5
Occasionally, finding nematode larvae in sputum or bronchoalveolar lavage fluid specimens may be very suggestive of a potential infection with S. stercoralis. In Fig. 4.6, larvae can be seen stained with Giemsa stain or a Gram stain. Once larvae are seen in respiratory specimens, fecal specimens can be collected for agar plate cultures to confirm strongyloidiasis.
Figure 4.6 Strongyloides stercoralis larvae. (Upper) Giemsa-stained larvae in a specimen of bronchoalveolar lavage fluid (courtesy of Marc Long). (Middle) Gram-stained larva in thoracic fluid. (Lower) Gram-stained larva in a respiratory specimen (sputum). doi:10.1128/9781555819002.ch4.f6
Agar
1.5% agar
0.5% meat extract
1.0% peptone
0.5% NaCl
Note Positive tracking on agar plates has been seen on a number of different types of agar. However, the most appropriate agar formula is that listed above.
Quality Control for Agar Plate Culture for Strongyloides stercoralis
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitologic examination.
2. Examine agar plates to ensure that there is no cracking and the agar pour is sufficient to prevent drying. Also, make sure that there is no excess water on the surface of the plates.
3. Review larval diagrams and descriptions for confirmation of larval identification.
4. The microscope should be calibrated, and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
5. Record all QC results (condition of agar plates).
Procedure for Agar Plate Culture for Strongyloides stercoralis
1. Place approximately 2 g of fresh stool in the center of the agar plate (area approximately 1 in. in diameter).
2. Replace the lid, and seal the plate with cellulose tape.
3. Maintain the agar plate (right side up) at room temperature for 2 days.
4. After 2 days, examine the sealed plates through the plastic lid under the microscope for microscopic colonies that develop as random tracks on the agar and evidence of larvae at the ends of the tracks away from the stool.
Note It has been documented that daily search for tracks on agar plates for up to six consecutive days results in increased sensitivity for diagnosis of both S. stercoralis and hookworm infections (17). When trying to rule out strongyloidiasis in immunocompromised patients or in those who may receive immunosuppressive drugs, it is recommended that two plates be set up, one that can be examined after 2 days and one that can be examined after the full 6 days.
5. With the ends of hot forceps, make a hole in the top of the plastic petri dish.
6. Gently add 10 ml of 10% formalin through the hole onto the agar surface, and swirl to cover the surface and rinse the agar plate. Allow to stand for 30 min.
7. Remove the tape and lid of the agar plate. Pour the 10% formalin through a funnel into a centrifuge tube. Do not try and pour the formalin off directly into the centrifuge tube—the tube opening is too small, and formalin will be spilled onto the counter.
8. Centrifuge the formalin rinse fluid 5 min at 500 × g.
9. Prepare a wet smear preparation from the sediment and examine with a 10× objective (low power) for presence of larvae. If larvae are found, confirm the identification with a 40× objective (high dry power).
Results and Patient Reports from Agar Plate Culture for Strongyloides stercoralis
Larval nematodes of hookworm, S. stercoralis, or Trichostrongylus spp. may be recovered. If Strongyloides organisms are present, free-living stages and larvae may be found after several days on the agar plates.
1. Report “No larvae detected” if no larvae could be detected at the end of incubation and rinse procedure.
2. Report larvae detected by agar plate culture.
Example: Strongyloides stercoralis larvae detected by agar plate culture.
Procedure Notes for Agar Plate Culture for Strongyloides stercoralis
1. If the larvae are too difficult to observe under the microscope and morphologic details are difficult to see, the larvae can be formalin killed within the plate and examined in the formalin-concentrated sediment.
2. Infective larvae may be found any time after the first or second day or even on the first day in a heavy infection. Since infective larvae may be present on the agar, caution must be exercised in handling the plates once the cellulose tape is removed. Wear gloves when handling the cultures.
3. It is important to maintain the plates upright at room temperature. Do not incubate or refrigerate them at any time; this also applies to the fresh stool specimen.
4. Fresh stool is required for this procedure; preserved fecal specimens or specimens obtained after a barium meal are not suitable.
Procedure Limitations for Agar Plate Culture for Strongyloides stercoralis
1. The agar plate culture technique is successful if any larvae present are viable. If the fresh stool specimen is too old, larvae may not survive and a negative result will be reported.
2. Specimens that have been refrigerated or preserved are not suitable for culture. Larvae of certain species are susceptible to cold environments.
Estimation of Worm Burdens and Kato-Katz Thick Film
The only human parasites for which it is reasonably possible to correlate egg production with adult worm burdens are Ascaris lumbricoides, Trichuris trichiura, and the hookworms (Necator americanus and Ancylostoma duodenale). The specific instances in which information on approximate worm burdens is useful are when one is determining the intensity of infection, deciding on possible chemotherapy, and evaluating the efficacy of the drugs administered. With current therapy, the need for monitoring therapy through egg counts is no longer as relevant. However, several methods that can be used if necessary are discussed below. Remember that egg counts are estimates; you will obtain count variations regardless of how carefully you follow the procedure. If two or more fecal specimens are being compared, it is best to have the same individual perform the technique on both samples and to do multiple counts.
Direct-Smear Method of Beaver
The direct-smear method of Beaver is the easiest to use and is reasonably accurate when performed by an experienced technologist. In the original method, Beaver (26) used a calibrated photoelectric cell to prepare a direct smear of exactly 2 mg. For routine purposes, this is impractical, and the procedure has subsequently been modified (27) such that a direct smear of 2 mg (enough fresh fecal material to form a low cone on the end of a wooden applicator stick) of stool is prepared. Egg counts on the direct smear are reported as eggs per smear, and the appropriate calculations can be made to determine the number of eggs per gram of stool.
Dilution Egg Count
The Stoll count (28) is probably the most widely used dilution egg-counting procedure for the purpose of estimating worm burdens. However, because of cost containment and clinical relevance (therapy is often initiated with no egg count data), most laboratories do not offer this procedure.
Stool displacement flasks for use in this procedure are available commercially. These flasks have a long neck with etched lines at 56 and 60 ml to facilitate proper filling with sodium hydroxide and fecal material. If commercial Stoll flasks are unavailable, any flask that can hold the sodium hydroxide solution, a weighed amount of 4 g of stool, and a few small glass beads can be used for a container.
1. In a calibrated Stoll flask, add 0.1 N sodium hydroxide to the 56-ml mark.
2. Add fresh fecal material to the flask so that the level of fluid rises to the 60-ml mark. This amount of feces is equivalent to 4 g of feces.
3. Add a few glass beads, and shake vigorously to make a uniform suspension. If the specimen is hard, the mixture may be placed in a refrigerator overnight before shaking to aid in mixing.
4. With a calibrated pipette, quickly remove 0.15 ml of suspension and transfer it to a slide.
5. Do not use a coverslip; place the slide on a mechanical stage, and count all of the eggs.
6. Multiply the egg count by 100 to obtain the number of eggs per gram of stool.
7. The estimate (eggs per gram) obtained will vary according to the consistency of the stool. The following correction factors should be used to convert the estimate to a formed-stool basis:
Egg counts on liquid specimens are generally unreliable; the most accurate counts are obtained with use of formed or semiformed specimens.
Modified Stoll Dilution Method
1. Fill a 15-ml centrifuge tube to the 14-ml mark with 0.1 N sodium hydroxide.
2. Add stool to bring the liquid contents up to the 15-ml mark.
3. Mix thoroughly with a wooden applicator stick.
4. If the stool is hard, allow the mixture to stand for several hours.
5. Shake to thoroughly mix, and quickly withdraw exactly 0.15 ml from the middle of the suspension.
6. Transfer the material to a slide, cover (it may be easier to count without a coverslip, since occasionally the eggs flow to the outside of the coverslip), and count the eggs in the entire preparation.
7. Multiply the egg count per preparation by 100 to give an uncorrected count of eggs per milliliter. Corrections may be made as in the Stoll dilution method for original stool consistency.
Kato-Katz Thick Smear
Although this method of counting helminth eggs has been modified many times since it was originally published, the protocol seen below provides an acceptable approach (29). Currently, the Kato-Katz is the recommended method of WHO, with the McMaster method being considered an alternative for the quantification of soil-transmitted helminths eggs in human stool (primarily Ascaris, Trichuris, hookworm, Schistosoma mansoni) (35). Although this thick film method has been adapted to estimate worm burden calculations, it is rarely used in routine clinical laboratories. Also, it is unsuitable for the diagnosis of protozoa or helminth larvae and is difficult to perform on hard and/or liquid stools.
1. Coverslips made of wettable cellophane, medium thickness, cut into 22 × 30 mm rectangles (No. 124 PD, E.I. DuPont de Nemours, Inc., Film Department, Wilmington, DE).
2. Glycerine-malachite green solution: 10 ml pure glycerine and 1 ml 3% aqueous malachite green in 100 ml distilled water. Allow the cellophane coverslips to soak in the glycerine mixture for at least 24 h. Although the malachite green is recommended, it is not essential for the procedure.
3. Some procedures require a metal or synthetic fiber filter mesh; however, this method is more complex. Just remember that if the fecal specimen is formed, hard, and somewhat dry, you will have to add a few drops of water so the final stool/water emulsion becomes mushy. To obtain 10-, 20-, or 50-mg samples, metal templates containing holes calibrated to deliver these fecal amounts can be used.
Procedure
1. Using an applicator stick, transfer approximately 50 mg of fresh stool to a standard 1 × 3-in. microscope slide. (A 4-mm cube of feces weighs about 65 mg.)
2. Cover the stool with the cellophane coverslip.
3. Turn the preparation upside down on a paper towel (flat, absorbent surface), and press it down until the fecal film covers approximately 20 to 25 mm in diameter. If the amount of feces is too much, the excess usually flows from under the cellophane coverslip and is absorbed by the paper towel.
4. Allow the preparation to stand for 1 h at room temperature to allow clearing of the fecal material but not the eggs. Do not overclear; thin-shelled eggs (hookworm) will tend to collapse and may disappear. Examine the slide at the end of the clearing period (usually not to exceed 1 h). It has been recommended that the clearing time be extended to 24 h for S. mansoni eggs; however, hookworm eggs will probably have collapsed and disappeared after this amount of time.
5. Examine the entire fecal film using the low power objective (10×).
Correlation with Treatment
The following numbers have been compiled to indicate the correlation between egg counts and the need for treatment. The two helminths listed below are generally the only ones for which the egg count will determine whether the patient is treated for the initial infection. As mentioned above, with the current therapeutic agents available, this information has become less clinically relevant, at least in many parts of the world. However, it has been recognized that egg counts do not always correlate with or accurately predict the worm burden of the host.
1. For T. trichiura, about 30,000 eggs/g indicates the presence of several hundred worms, which may cause definite symptoms.
2. For hookworm, about 2,500 to 5,000 eggs/g usually indicates a clinically significant infection.
The effectiveness of therapy for any helminth infection may be evaluated by doing repeated egg counts after treatment. Low egg counts for T. trichiura and hookworm are generally reflected by a lack of clinical signs and symptoms in individuals harboring these parasites. However, the presence of even one Ascaris worm is potentially dangerous because of the active migrating habits of this parasite, which may result in serious clinical manifestations.
When schistosome eggs (Fig. 4.7) are recovered from either urine or stool, they should be carefully examined to determine viability. The presence of living miracidia within the eggs indicates an active infection that may require therapy. The viability of the miracidia can be determined in two ways: (i) the cilia of the flame cells (primitive excretory cells) may be seen on a wet smear by using high dry power and are usually actively moving, and (ii) the miracidia may be released from the eggs by a hatching procedure (Fig. 4.8) (1, 28, 30, 31). The eggs usually hatch within several hours when placed in 10 volumes of dechlorinated or spring water (hatching may begin soon after contact with the water). The eggs that are recovered in the urine (24-h specimen collected with no preservatives) are easily obtained from the sediment and can be examined under the microscope to determine viability. McMullen and Beaver (31) recommended the use of a sidearm flask, but an Erlenmeyer flask is an acceptable substitute.
Figure 4.7 Schistosoma mansoni egg showing flame cells. (Illustration by Nobuko Kitamura; modified from E. C. Faust et al., Craig and Faust's Clinical Parasitology, 8th ed., Lea & Febiger, Philadelphia, PA, 1970.) doi:10.1128/9781555819002.ch4.f7
Figure 4.8 (Upper) Sidearm hatching flask used to recover miracidia from viable schistosome eggs (illustration by Nobuko Kitamura). (Lower) Miracidium larva released from egg during hatching procedure (Armed Forces Institute of Pathology photograph). doi:10.1128/9781555819002.ch4.f8
Quality Control for Hatching of Schistosome Eggs
1. Make sure that the water used is chlorine free; chlorine kills the miracidia. You can use bottled water or leave tap water in an open pan overnight before use (eliminates chlorine in tap water).
2. Check the saline solution (used to prepare the stool concentration) for the presence of any free-living organisms (flagellates or ciliates). This is normally not a problem.
3. Since it is neither realistic nor practical for the majority of laboratories to perform parallel positive-control procedures, review drawings and size measurements of schistosome eggs and/or miracidia.
4. The microscope should be calibrated, and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope for easy access (multiplication factors can be pasted on the body of the microscope).
5. Record all QC results.
Procedure for Hatching Schistosome Eggs
1. Thoroughly homogenize a fresh stool specimen (40 to 50 g) in 50 to 100 ml of 0.85% NaCl.
2. Strain through two layers of gauze placed on a funnel. Collect material in a small beaker or 50-ml centrifuge tube.
3. Allow the suspension to settle for 1 h. Pour off and discard the supernatant fluid, and repeat this rinse process at least twice.
4. Decant the saline solution, resuspend the sediment in a small quantity of chlorine-free (spring) water (10 to 20 ml), and pour the suspension into a 500-ml sidearm flask or an Erlenmeyer flask.
5. Add chlorine-free water to the flask so that the fluid level rises to 2 to 3 cm in the side arm or to the top 2 cm of the Erlenmeyer flask. Cover the flask with aluminum foil or black paper, leaving the side arm of the flask exposed to light; if an Erlenmeyer flask is used, cover to 1 cm below the level of fluid in the neck of the flask.
6. Allow the flask to stand at room temperature for several hours or overnight in subdued light.
7. Place a bright light at the side of the flask opposite the surface of exposed water. Do not place the light against the glass, to avoid generation of excess heat. As the eggs hatch, the liberated miracidia will swim to the upper layers and collect in the side arm (or neck region of an Erlenmeyer flask). Make sure that the exposed part of the flask or side arm of the hatching flask does not get too warm.
8. Examine the illuminated area with a magnifying lens (hand lens) to look for minute white organisms swimming rapidly in a straight line (placing dark cardboard or black paper behind the flask will facilitate observation of the white miracidia against a black background).
Results and Patient Reports from Hatching Schistosome Eggs
Living miracidia may be seen; however, failure to see these larvae does not rule out schistosomiasis.
1. Report as “Miracidia of schistosomes detected, indicating the presence of viable eggs.”
2. Report as “No miracidia of schistosomes detected; presence of eggs is not ruled out by this procedure.”
Procedure Notes for Hatching Schistosome Eggs
1. Both urine and stool specimens must be collected without preservatives and should not be refrigerated before being processed. Regardless of the species suspected, BOTH URINE AND STOOL SHOULD BE EXAMINED FOR EVERY PATIENT WITH POTENTIAL SCHISTOSOMIASIS.
2. Hatching does not occur until the saline is removed and nonchlorinated water is added. If a stool concentration is performed, use saline throughout the procedure to prevent premature hatching.
3. Make sure that the light is not too close to the side arm or top layer of water in the Erlenmeyer flask. Excess heat kills the miracidia.
Procedure Limitations for Hatching Schistosome Eggs
1. The absence of live miracidia does not rule out the presence of schistosome eggs. Nonviable eggs or eggs that failed to hatch are not detected by this method. Microscopic examination of direct or concentrated specimens should be used to demonstrate the presence or absence of eggs.
2. Egg viability can be determined by placing some stool or urine sediment (same material as used for the hatching flask) on a microscope slide. Low-power magnification (×100) can be used to locate the eggs. Individual eggs can be examined under high dry magnification (×400); the detection of moving cilia on the flame cells (primitive excretory system) confirms egg viability.
3. Free-living ciliates may be present in soil-contaminated water. Therefore, it may be necessary to perform the following steps to differentiate those forms from the parasitic miracidia (26).
A. Transfer a few drops of the suspension containing the organisms to a 3 x 2-in. slide, and examine under the microscope.
B. Add a drop of weak iodine solution (weak-tea color) or dilute methylene blue (pale blue).
C. Parasitic miracidia will stop moving, while free-living forms continue to move.
Since the medication used for treatment of tapeworms is usually very effective, a search for tapeworm scolices is rarely requested and no longer clinically relevant. However, stool specimens may have to be examined for the presence of scolices and gravid proglottids of cestodes for proper species identification. This procedure requires mixing a small amount of feces with water and straining the mixture through a series of wire screens (graduated from coarse to fine mesh) to look for scolices and proglottids. Remember to use standard precautions and wear gloves when performing this procedure. The appearance of scolices after therapy is an indication of successful treatment. If the scolex has not been passed, it may still be attached to the mucosa; the parasite is capable of producing more segments from the neck region of the scolex, and the infection continues. If this occurs, the patient can be retreated when proglottids begin to reappear in the stool.
After treatment for tapeworm removal, the patient should be instructed to take a saline cathartic and to collect all stool material passed for the next 24 h. The stool material should be immediately placed in 10% formalin, thoroughly broken up, and mixed with the preservative (1-gal [3.8-liter] plastic jars, half full of 10% formalin, are recommended).
For additional information, see chapter 8 (Fixation and Special Preparation of Fecal Parasite Specimens and Arthropods).
Quality Control for the Tapeworm Scolex Search
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitologic examination.
2. Review diagrams and sizes of proglottids and scolices of tapeworms.
Procedure for the Tapeworm Scolex Search
1. Mix a 24-h stool specimen (fresh or preserved in 10% formalin) with water, and thoroughly break up the specimen to make a watery suspension.
2. Strain the suspension (or the purged stool) through a double layer of screen wire or a sieve (a coarse-mesh screen placed over a fine-mesh screen).
3. Wash off the sediment remaining from each portion by running a slow flow of tap water over it.
4. Examine the cleansed debris with a hand lens to look for scolices and proglottids (the Taenia scolex is 0.5 to 1 cm long and 1 to 2 mm wide).
5. Repeat steps 3 and 4 for each portion of the suspension strained.
6. Collect the strained sediment in a glass petri dish, and place the dish over a black surface to increase the contrast of organisms against the background.
7. Observe with a magnifying hand lens, and pick pieces of worms with an applicator stick.
8. Rinse gravid proglottids and/or scolices with tap water, blot them dry on paper towels, and place them between two microscope slides separated at the edges by thin pieces of cardboard.
9. Fasten the preparation by placing rubber bands at each end of the slides so that the segments become somewhat flattened. One can also perform the India ink injection of the proglottid (see next protocol below).
10. Observe under the low power of a dissecting microscope for the number of uterine branches and genital pores in the segments and the presence or absence of a rostellum of hooks on the scolex. Quite often, the scolex is broken off from the rest of the strobila (chain of proglottids) and is ∼1 cm long.
Results and Patient Report from the Tapeworm Scolex Search
Tapeworm proglottids may be recovered (either singly or several attached together), and a scolex may or may not be seen.
1. Report as “A search for adult worms reveals the presence/absence of . . . (finding).”
Example: Taenia saginata scolex present
Procedure Notes for the Tapeworm Scolex Search
1. Remember that Taenia solium eggs are infective (cysticercosis), as are the eggs of Hymenolepis nana.
2. Wear gloves when performing this procedure.
3. Specimens preserved in 10% formalin are recommended.
4. If the patient has received niclosamide or praziquantel, a purged specimen is required, which should be immediately preserved in 10% formalin.
5. Wood’s lamp may be used to reveal the scolices if the patient has been given quinacrine dyes; the worms will have absorbed the dye, and they fluoresce at a wavelength of 360 nm. Also, after the use of quinacrine, tapeworm proglottids appear yellow.
Procedure Limitations for the Tapeworm Scolex Search
1. Niclosamide or praziquantel therapy leads to dissolution of the tapeworm. Therefore, the scolex and other proglottids may be difficult to recover unless the patient receives a saline purge soon after taking the medication.
2. It is often difficult to identify proglottids without staining. Identification may be achieved by staining with India ink.
India Ink Injection Procedure for Tapeworm Proglottids
Identification of adult worms usually involves examination of a tapeworm proglottid. A Taenia proglottid must be gravid, containing the fully developed uterine branches. Using a syringe (1 ml or less) and a 25-gauge needle, India ink is injected into the central uterine stem of the proglottid, filling the uterine branches with ink, or into the uterine pore. The proglottid can then be rinsed in water or saline, blotted dry on paper towels, pressed between two slides, and examined. After the identification has been made, the proglottid can be left between the two slides (place a rubber band around the slides), dehydrated through several changes of ethyl alcohol (50, 70, 90, and 100%), cleared in two changes of xylene, and mounted with Permount for a permanent record. After the xylene step, the proglottid will be stuck to one of the two slides; do not try to remove it (it is very brittle and will crack) but merely add the Permount to the proglottid and then add the coverglass (Fig. 4.9).
Figure 4.9 Taenia gravid proglottids after India ink injection of the uterine branches. (Left) Taenia fresh gravid proglottid prior to India ink injection. (Middle) Taenia saginata gravid proglottid. (Right) Taenia solium gravid proglottid. doi:10.1128/9781555819002.ch4.f9
It is important to remember that even when submitted in fixative, the tapeworm eggs within the branched uterine structure may not be killed and may remain infective. Since this procedure is usually limited to tapeworms in the genus Taenia, if eggs of Taenia solium (pork tapeworm) are present, they present a potential danger in terms of possible egg ingestion/inhalation/swallowing leading to cysticercosis. This becomes a possibility when performing the actual ink injection. If the needle accidentally pokes through the tapeworm proglottid, an aerosol can be created that may contain infectious eggs. ONE SHOULD WEAR A MASK, GLOVES, AND LABORATORY COAT AT ALL TIMES WHEN PERFORMING THIS PROCEDURE.
Procedure for the Tapeworm India Ink Injection (MASK, LABORATORY COAT, AND GLOVES MUST BE WORN AT ALL TIMES WHEN PERFORMING THIS PROCEDURE)
1. Using a 1-ml syringe and a 25-gauge needle, fill the syringe with approximately 0.5 ml of India ink.
2. Rinse the proglottid with tap water, and blot it dry on paper towels.
3. To prepare for the injection, hold the proglottid between your thumb and first finger or on the surface of the counter (harder to do).
5. Very carefully, penetrate either the central uterine stem at the end where the proglottid has become unattached from the chain or penetrate the side uterine pore with the end of the needle.
6. GENTLY inject some India ink; it should penetrate the uterine branches very quickly. IF THE INJECTION DOES NOT IMMEDIATELY OCCUR, DO NOT FORCE THE INJECTION.
7. You can try the other entry location, again GENTLY injecting the ink.
8. After the injection, rinse the gravid proglottid with tap water, blot dry on paper towels, and place the proglottid between two microscope slides separated at the edges by thin pieces of cardboard.
9. Fasten the preparation by placing rubber bands at each end of the slides so that the segments become somewhat flattened.
10. Observe the number of uterine branches on ONE SIDE ONLY where they come off the main uterine stem; generally Taenia saginata has 12 or more branches, while Taenia solium has approximately 8 or fewer branches.
11. After the identification has been made, the proglottid can be left between the two slides (place a rubber band around the slides), dehydrated through several changes of ethyl alcohol (50, 70, 90, and 100%), cleared in two changes of xylene, and mounted with Permount for a permanent record.
12. After the xylene step, the proglottid will be stuck to one of the two slides; do not try to remove it (it is very brittle and will crack) but merely add the Permount to the proglottid and then add the coverglass.
Results and Patient Report from the India Ink Injection Procedure
Tapeworm proglottids may be recovered (either singly or several attached together).
1. Report as:
Example: Taenia saginata gravid proglottid present
Procedure Notes for the India Ink Injection Procedure
1. Remember that T. solium eggs are infective (cysticercosis). The identification to the species level may not be possible until after the ink injection is completed.
2. Wear mask, laboratory coat, and gloves when performing this procedure.
3. Fresh, unpreserved proglottids are recommended; preserved proglottids can be used, but they are harder to inject.
4. Make sure the proglottid is blotted dry prior to the injection; otherwise, it may be difficult to hold onto the proglottid—it can slip through your fingers.
5. DO NOT FORCE THE INK INJECTION. If the ink does not easily flow into the branches, it may be because the proglottid is not gravid, but only mature. Also, the needle may not be in the right place.
6. IF THE INK INJECTION IS FORCED, it is more likely you may create an aerosol, thus increasing the danger for exposure to viable T. solium eggs.
7. Remember that after the xylene step, the proglottid will be stuck to one of the two slides; do not try to remove it (it is very brittle and will crack) but merely add the Permount to the proglottid and then add the coverglass.
Procedure Limitations for the India Ink Injection Procedure
1. If the proglottid is fresh, it may have begun to disintegrate, thus making the ink injection more difficult.
2. If the proglottid is mature AND NOT YET GRAVID, the branches may not inject properly.
3. If the ink injection is not successful, one can try some of the mounting methods found in chapter 9.
Qualitative Test for Fecal Fat
The microscopic examination of stool with the addition of Sudan III is a very simple, quick, and widely used technique to screen the specimen for fat. Fresh, unpreserved fecal material is required. If there is a time delay prior to testing, the specimen should be refrigerated. Specimens collected more than 48 h earlier or specimens that are dried out should be discarded, and fresh specimens should be collected. Although this technique is quite old and the original paper may be difficult to obtain, it was published in 1961 (32).
Sudan Black IV Stain, also known as scarlet red, was introduced by Michaelis in 1901 as a fat stain. It is a dimethyl derivative of Sudan III, which makes it a deeper and more intense stain, but it has similar physical properties and is fat soluble. This stain has been widely used as a screening method because it is easy to use and correlates well with quantitative methods.
Sudan Black IV Stain is used as a qualitative method to detect the presence of fecal fat. Normally the stool does not contain more than 20 g of fat daily. In patients with steatorrhea, fat malabsorption occurs, and greater quantities of fat are detected in the stool. This procedure, when performed carefully and consistently, is a simple method of detecting this condition in the patient (33). Other dyes such as Sudan II, Sudan III, Oil Red O, and Sudan Black B can be used as well.
Sudan Stain
Quality Control for the Qualitative Test for Fecal Fat
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitology.
2. Run a QC sample with each batch of patient tests as in procedure below.
3. As a positive control, mayonnaise is used; red-stained fat globules should be observed microscopically.
4. As a negative control, water is used; no fat globules are observed.
5. Record all QC results. If the QC results are unacceptable, the test must be repeated and documented on the corrective action sheet.
Procedure for the Qualitative Test for Fecal Fat (Sudan III)
1. Mix a small amount of stool with an equal amount of 95% ethanol in a test tube. Mix well.
2. Add 2 drops of Sudan III stain to the stool-ethanol mixture. Mix well, and let stand for a few minutes.
3. Using a pipette, remove a drop from the test tube and place it on a slide. Cover with a coverslip.
4. Using the microscope, examine the slide for globules of fat stained red (neutral fat) and needle-like crystals (fatty acid).
5. Add several drops of glacial acetic acid to the test tube. Remove a drop from the test tube and place it on a slide. Cover with a coverslip.
A. Gently heat the slide over a flame. (Do not boil.)
B. Observe again on the microscope for globules of fat stained red (Fig. 4.10).
Figure 4.10 Fat droplets seen in the fecal fat test. The color can range from red to orange (pale to more intense), and the droplets can vary in size. doi:10.1128/9781555819002.ch4.f10
Results and Patient Reports for the Qualitative Test for Fecal Fat (Sudan III)
Red-stained fat globules or fatty acid crystals are seen microscopically. Report the specimen as “fat not increased” or “fat increased” depending on the number of globules seen.
1. If fewer than 100 globules/high-power field (hpf) are seen before or after heating, report as “fat not increased.”
2. If more than 100 globules/hpf of fat are seen before or after heating, report as “fat increased.”
3. If fatty acid crystals have been seen before heating and globules appear after heating, report as “fatty acids increased.”
Procedure for the Qualitative Test for Fecal Fat (Sudan Black IV)
1. Place a small aliquot of stool suspension on a clean glass slide.
2. Mix 2 drops of 95% ethanol with the suspension on the slide.
3. Add 2 drops of Sudan Black IV Stain to the suspension on the slide, and mix well.
4. Cover the suspension with a coverslip, and examine microscopically for the presence of large orange or red droplets.
Results and Patient Reports for the Qualitative Test for Fecal Fat (Sudan Black IV)
1. Fatty acids are present as lightly staining flakes or needle-like crystals that do not stain.
2. Soaps appear as nonstaining formless flakes, coarse crystals, or rounded masses.
3. Neutral fats appear as large orange or red droplets. If 60 or more stained droplets (neutral fats) are seen per 40× (high-power) field, it is a presumptive finding that the patient has steatorrhea.
Procedure Limitations for the Qualitative Test for Fecal Fat
1. The formation of large needle-like crystals as the preparation cools after heating does not necessarily mean that the original globules were fatty acid. Sudan III forms very short needle-like crystals in bunches as it dries.
2. Very few, if any, neutral fat globules are seen in a normal stool specimen. The presence of large amounts of neutral fat should raise suspicion that the patient has ingested mineral oil or castor oil, thus causing a false-positive result.
3. Do not count the fat that is present in vegetable cells.
Quantitation of Reducing Substances (Clinitest)
Clinitest is a reagent tablet test based on the classic Benedict’s copper sulfate reduction reaction, combining ingredients with an integral heat-generating system. Clinitest provides clinically useful information about carbohydrate metabolism by determining the amount of reducing substance in urine or stool. Reducing substances convert the cupric sulfate (CuSO4) to cuprous (Cu2O) oxide and cause a change in solution color ranging from blue through green to orange (Fig. 4.11). Unpreserved stool is required; the specimen should be placed in the refrigerator if there is a delay in testing. Specimens collected more than 48 h earlier or specimens that are dried out should be discarded, as fresh specimens should be collected. Although this is a relatively old method, there are references in the literature (34).
Figure 4.11 Color range for stool Clinitest reactions: blue (negative) to orange (positive). doi:10.1128/9781555819002.ch4.f11
Clinitest Tablets
Clinitest reagent tablets (store tablets at room temperature in a plastic bag)
Comparative color chart that comes with the tablets
Deionized water
Chek-Stix positive control
Quality Control for Quantitation of Reducing Substances (Clinitest)
1. Follow routine procedures for optimal collection and handling of fresh fecal specimens for parasitology.
2. Run a QC sample with each batch of patient tests as in procedure below.
3. As a positive control, Chek-Stix is used; the development of a green, yellow, or orange color with a yellow or red precipitate is considered a positive result.
4. As a negative control, 0.5 ml of deionized water is used; blue color is considered a negative result.
5. Record all QC results. If the QC results are unacceptable, the test must be repeated and documented on the corrective action sheet.
Procedure for Quantitation of Reducing Substances (Clinitest)
1. All testing on clinical specimens should be performed in a biological safety cabinet by personnel wearing gloves and a laboratory coat.
2. Add 1 volume of stool to 2 volumes of deionized water, and mix thoroughly.
3. Using a disposable transfer pipette, transfer 15 drops of this suspension into a clean test tube.
4. Drop one Clinitest tablet reagent into the test tube.
5. Observe the reaction. Do not shake the tube while the chemical reaction is occurring.
6. Wait 15 s after the reaction stops, then gently shake contents to mix.
7. Compare the color of the liquid to the color chart in the package insert of the Clinitest tablet reagent (Fig. 4.11).
8. Discard supplies in appropriate biohazard containers.
Results and Patient Reports for Quantitation of Reducing Substances (Clinitest)
1. Negative: clear to cloudy blue color
2. Positive: compare the liquid color to the color chart that comes with the tablets, and grade the degree of color development to the color chart (trace, 1+, 2+, 3+, or 4+). These results equate to the grams per deciliter of the reducing substance present per sample. The colors range from blue through green through yellow/orange to orange (negative to 4+).
3. Positive: report as Trace (0.25 g/dl), 1+ (0.5 g/dl), 2+ (0.75 g/dl), 3+ (1.0 g/dl), or 4+ (equal to or greater than 2 g/dl)
Example: 1+ (0.5 g/dl)
4. Negative
Example: Negative
Procedure Limitations for Quantitation of Reducing Substances (Clinitest)
1. Clinitest is not specific for glucose and reacts with any reducing substance in stool, including lactose, fructose, galactose, and pentoses.
2. Interfering substances may affect the results. These include salicylates, penicillin, large quantities of ascorbic acid, nalidixic acid, and cephalosporins.
3. Failure to observe the reaction at all times may lead to erroneously low results if reducing substances are present in large amounts. If more than 2% sugar is present, a rapid color change may occur during boiling, causing the color to pass rapidly through bright orange to a dark brown or greenish brown.
References
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4. Isenberg HD (ed). 1995. Essential Procedures for Clinical Microbiology. American Society for Microbiology, Washington, DC.
5. Harada U, Mori O. 1955. A new method for culturing hookworm. Yonago Acta Med 1:177–179.
6. Hsieh HC. 1962. A test-tube filter-paper method for the diagnosis of Ancylostoma duodenale, Necator americanus, and Strongyloides stercoralis. WHO Tech Rep Ser 255:27–30.
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