Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 17
ОглавлениеVery few clinical laboratories offer specific culture techniques for parasites. The methods for in vitro culture are often complex, while quality control is difficult and not really feasible for the routine diagnostic laboratory. In certain institutions, some techniques may be available, particularly when consultative services are provided (reference laboratory situation) and for research purposes. However, most laboratories do not offer these techniques.
Few parasites can be routinely cultured; however, methods are available for Entamoeba histolytica, Naegleria fowleri, Acanthamoeba spp., Trichomonas vaginalis, Dientamoeba fragilis, Toxoplasma gondii, Blastocystis spp., Trypanosoma cruzi, and the leishmanias. Often, when specimens are cultured for potential pathogens, nonpathogenic protozoa could also be recovered. These procedures are usually available only after consultation with the laboratory and on special request. For those who may be interested in trying these techniques, the several different media presented below are representative of those available. More extensive options can be found in the literature (1–13).
Cultures of parasites grown in association with an unknown microbiota are referred to as xenic cultures. A good example of this type of culture would be stool specimens cultured for E. histolytica. If the parasites are grown with a single known bacterium, the culture is referred to as monoxenic. An example of this type of culture would be clinical specimens (corneal biopsy specimens) cultured with Escherichia coli as a means of recovering species of Acanthamoeba and Naegleria. If parasites are grown as pure culture without any bacterial associate, the culture is referred to as axenic. An example of this type of culture would be the use of media for the isolation of Leishmania spp. or Trypanosoma cruzi. All three types of cultures are discussed in this chapter.
Pancreatic digests of casein are major ingredients of media used in the axenic cultivation of lumen-dwelling parasitic protozoa. Unfortunately, the digest used almost exclusively in the development of these media has not been available since the early 1980s. Many digest products have been tried in the interim with marginal results in supporting the growth of E. histolytica. Diamond et al. (14) have developed a casein-free medium, YI-S, consisting of a nutrient broth, vitamin mixture, and serum. This may serve as a replacement for TYI-S-33, widely used for the axenic culture of E. histolytica and other intestinal protozoa.
Specimens include stool material, mucus, or a combination of the two. The clinical specimen(s) should be no more than 24 h old; a maximum of 2 to 3 h is recommended.
Balamuth’s Aqueous Egg Yolk Infusion Medium for Amebae
Balamuth’s aqueous egg yolk infusion medium is used to detect the presence of amebae. The specific solutions required are phosphate buffer and whole-liver concentrate solution.
Balamuth’s Aqueous Egg Yolk Infusion Medium
Phosphate buffer
Mix the solution in the ratio of 3 parts tribasic (A) to 2 parts monobasic (B) to make 1 M phosphate buffer stock. Dilute the stock buffer to 0.067 M before use (add 492 ml of distilled H2O to 1 liter of 1 M phosphate buffer).
Whole-liver concentrate solution
Suspend the powder in cold water, and autoclave. Filter through a Büchner funnel to remove sediment, dispense in 10-ml quantities, and reautoclave.
Preparation of Complete Medium (15)
1. Using a blender, blend 12 fresh hard-boiled egg yolks with 375 ml of 0.8% sodium chloride.
2. Autoclave at 7 lb/in.2 pressure for 10 min, repressurize slowly, and stir.
3. Autoclave again for 45 min at 7 lb/in.2 pressure.
4. Allow to cool slightly, and add distilled water to replace evaporation loss. Transfer the material to a muslin bag, and express the liquid portion, saving all the fluid. Return the volume to 375 ml with 0.8% sodium chloride.
5. Autoclave for 20 min at 121°C; cool to 5°C. Do not agitate the fluid at this point or during filtration.
6. Decant the fluid carefully through gauze into a Büchner funnel with Whatman no. 3 filter paper. Filter papers can be replaced as necessary.
7. Measure the filtrate, add an equal volume of 0.067 M phosphate buffer, and autoclave for 20 min at 121°C. After cooling, add stock liver concentrate (1 part stock liver to 9 parts medium).
8. Material can then be decanted into sterile flasks, which are stored until the medium is dispensed.
Procedure
Tubes should contain 6 to 8 ml of fluid and should be incubated for 4 days at 37°C as a sterility check. Before inoculation, a loopful of sterile rice powder or starch (can be autoclaved in a screw-cap tube) is added to each tube. To each tube, add stool material, mucus, or a combination of the two (about the size of a small pea), break it up thoroughly in the medium, and incubate at 37°C. The cultures should be checked at 2, 3, and 4 days by examining 0.1 ml of sediment under the microscope (low-intensity light) for characteristic motility. Although the initial culture may appear to be negative, subcultures may reveal organisms (Fig. 8.1).
Figure 8.1 Protozoa from culture systems. (Upper left) Entamoeba histolytica/E. dispar trophozoite from liquid medium containing rice starch (note that there are no definitive erythrocytes within the cytoplasm, so that it is not possible to differentiate the true pathogen, E. histolytica, from the nonpathogen, E. dispar). (Upper right) Naegleria fowleri trophozoite from nonnutrient agar culture with bacterial overlay (note that this trophozoite has been stained). (Lower left) Acanthamoeba spp. trophozoite from nonnutrient agar culture with bacterial overlay (note the spiky acanthapodia). (Lower right) Acanthamoeba spp. cysts from nonnutrient agar culture with bacterial overlay (note the double hexagonal wall appearance. doi:10.1128/9781555819002.ch8.f1
According to Dolkart and Halpern (16), the addition of gastric mucin to the egg component is reported to improve the performance of Balamuth’s medium.
The addition of rice flour (prerinsed with distilled water) to Balamuth’s medium is reported to support abundant growth of Balantidium coli.
Boeck and Drbohlav’s Locke-Egg-Serum (LES) Medium for Amebae
Boeck and Drbohlav’s LES medium is another culture medium used to diagnose the presence of amebae.
Boeck and Drbohlav’s LES Medium
Locke’s solution
Autoclave before storage.
Preparation of Complete Medium
1. Wash four eggs, wipe the shells with 70% alcohol, and break the eggs into a sterile flask containing glass beads.
2. Add 50 ml of Locke’s solution, and shake until homogenous.
3. Dispense the medium so that a slant of 1 to 1.5 in. (1 in. = 2.54 cm) is produced in the bottom of the tube. (Tube size is not critical.)
4. Plug the tubes, and place them in a slant position in an inspissator at 70°C until the slant solidifies. Inspissator conditions may be achieved in the autoclave by leaving the door ajar (nonpressurized system).
5. Autoclave the tubes at 121°C for 20 min. Discard any damaged slants.
6. Prepare a mixture of 8 parts sterile Locke’s solution to 1 part sterile inactivated human serum. Sterilize the mixture by filtration, and incubate at 37°C for 24 to 48 h as a sterility check before use. Cover the slants to a depth of <1 cm with the sterile solution, and inoculate in the same manner as for Balamuth’s medium. LES medium should have a loopful of sterile rice powder added before inoculation.
TY1-S-33 Medium for Entamoeba histolytica (17)
TYI Broth
Note Biosate may be replaced with 20 g of Trypticase (BBL) and 10 g of yeast extract (BBL). Some lots of Biosate, Trypticase, yeast extract, or serum may inhibit growth.
1. Adjust pH to 6.8 with 1 N NaOH, and filter through Whatman no. 1 paper. Autoclave at 121°C for 15 min.
2. Cool, and add:
3. Aseptically dispense 13 ml of complete medium into 16- by 125-mm screw-cap tubes. To prepare Special 107 Vitamin Mix, aseptically combine the following:
Total volume of complete Special 107 Vitamin Mix is 120 ml.
Note The shelf life of solutions 1, 2, and 3 is 22 months.
TYI-S-33 (Keister’s Modification) for Giardia lamblia
For TYI-S-33 (Keister’s modification [2]), prepare TYI broth exactly as for E. histolytica with the following changes.
(i) Increase the amount of L-cysteine hydrochloride to 2.0 g/liter.
(ii) Add 500 mg of dehydrated bovine bile per liter.
(iii) Adjust the pH to 7.0 to 7.1.
(iv) Sterilize by filtration through a 0.22-µm-pore-size filter. Do not autoclave! The complete medium is made by addition of bovine serum to 10%.
TYSGM-9 Medium for Entamoeba histolytica
Nutrient Broth
The nutrient broth may be stored for several months at –2°C.
5% Tween 80 Solution
1. Vigorously stir, with a magnetic stirrer, 95 ml of glass-distilled water in a bottle.
2. Add 5 g of Tween 80 (very thick solution; must be weighed), and keep stirring for a few minutes.
3. Filter sterilize through a 0.22-µm-pore-size membrane.
4. Aseptically dispense into a number of sterile screw-cap test tubes, 10 ml per tube.
5. Label as 5% Tween 80 solution, with the preparation date and an expiration date of no longer than 1 month. Store at 4°C.
Phosphate-Buffered Saline (PBS no. 8), pH 7.2
1. Dissolve the salts in the distilled water with a magnetic stirrer.
2. Autoclave for 15 min at 121°C.
3. When cool, label as PBS no. 8 with the preparation date and an expiration date of 3 months.
Rice Starch
For best results use rice starch obtained from BDH, Ltd. (Merck, Ltd.) or Gailard Schlesinger, Inc.
1. Dispense 500 mg of rice starch into each of several 16- by 125-mm screw-cap tubes; do not tighten the caps.
2. Place the tubes horizontally in a dry-heat sterilizer or an oven. Make sure that the rice starch is uniformly distributed loosely over the undersurface of the tubes.
3. Heat the tubes for 2.5 h at 150°C.
4. When cool, tighten caps and label as rice starch with the date of preparation and an expiration date of 3 months.
Rice Starch Suspension
1. Add 9.5 ml of sterile PBS no. 8 to each tube of rice starch.
2. Shake vigorously or use a vortex machine to uniformly suspend the rice starch at the time of use.
Stock Antibiotic Solution
1. Using a 6-ml syringe and 20-gauge needle, add 5 ml of sterile distilled water to a vial of penicillin G sodium (106 U).
2. Using a 6-ml syringe and 20-gauge needle, add 5 ml of sterile distilled water to a vial of streptomycin sulfate (106 µg/ml).
3. Shake gently, and let stand for 30 min to dissolve the antibiotics completely in the distilled water.
4. Mix the two antibiotics in a graduated flask or cylinder, and bring the volume to 125 ml with distilled water. The stock concentration of antibiotics is 8,000 U of penicillin/ml and 8,000 µg of streptomycin/ml.
5. Filter sterilize the antibiotic solution through a 0.22-µm-pore-size membrane filter, dispense the filtrate into a number of sterile screw-cap vials or sterile cryovials (1 ml per vial), and label as stock antibiotic solution with the preparation date and an expiration date of 6 months. Store at 120°C in a cryovial box.
Buffered Methylene Blue Solution
Solution A, 0.2 M acetic acid
Add the acetic acid to the water, mix, and store in a glass-stoppered bottle. Label with the date of preparation and an expiration date of 1 year.
Solution B, 0.2 M sodium acetate
Dissolve the sodium acetate in 400 ml of distilled water in a volumetric flask, bring the volume to the 1,000-ml mark, mix well, and store in a glass-stoppered bottle. Label with the date of preparation and an expiration date of 1 year.
Acetate buffer, pH 3.6
Mix solutions A and B in a volumetric flask, and bring the volume to 100.0 ml with distilled water. The pH should be 3.6. Store in a glass-stoppered bottle. Label with the date of preparation and an expiration date of 1 year.
Methylene blue stain
Dissolve the dye in the buffer, and store in a glass-stoppered bottle. Label with the date of preparation and an expiration date of 1 year.
Complete Medium (TYSGM-9 Medium)
1. Place 200 mg of gastric mucin (U.S. Biochemical Corp., catalog no. 16025) in a 125-ml screw-cap bottle or Erlenmeyer flask.
2. Add 97 ml of nutrient broth; using a magnetic stirrer, stir vigorously for at least 1 h or until the medium becomes clear.
3. Autoclave for 15 min at 121°C; cool to room temperature.
4. Add aseptically, in a biological safety cabinet, 5.0 ml of heat-inactivated bovine serum.
5. Add 0.1 ml of the 5% Tween 80 solution.
6. Dispense aseptically, in a biological safety cabinet, into a number of sterile 16- by 125-mm screw-cap tubes, 8 ml per tube.
7. Add 0.25 ml of rice starch solution after vigorously shaking the tube.
8. Store the tubes at 4°C for not more than 1 month.
9. The final pH of the medium should be 7.2.
Robinson’s Culture
Robinson’s medium is a complex medium that has nevertheless found widespread use for the isolation of enteric amebae. To prepare Robinson’s medium, prepare the six following stock solutions (2).
Solution 1 (0.5% Erythromycin) (Filter Sterilized)
Prepare 0.5% erythromycin in distilled water and filter sterilize. Refrigerate.
Solution 2 (20% Bacto Peptone) (Autoclave)
Prepare 20% Bacto Peptone in distilled water. Autoclave and refrigerate.
Solution 3 (10× Phthalate Solution, Stock) (Autoclave)
Bring to 1 liter at pH 6.3. Autoclave for 15 min at 121°C under a pressure of 15 lb/in.2. Store at room temperature. Dilute 1:10 with sterile water before use.
A stock solution of phthalate-Bacto Peptone can be made by adding 1.25 ml of 20% Bacto Peptone per 100 ml of 1× phthalate solution. Store refrigerated.
Solution 4 (10× R Medium Stock) (Autoclave)
Bring to 500 ml. Dilute stock 1:10, adjusting pH to 7.0. Autoclave for 15 min at 121°C under a pressure of 15 lb/in.2 in 20-ml amounts.
Solution 5 (BR Medium)
To prepare BR medium, inoculate 1× R medium with a standard Escherichia coli strain such as O111. Incubate at 37°C for 48 h, and store at room temperature (good for several months).
Solution 6 (BRS Medium)
To prepare BRS medium, add an equal volume of heat-inactivated bovine serum to BR medium and incubate at 37°C for 24 h. Store at room temperature (good for several months).
1. To prepare agar slants, many people use screw-cap glass bijou bottles (total volume, 7 ml), but standard culture tubes also work well.
2. Autoclave a solution of 1.5% Noble agar in 0.7% NaCl-distilled water for 15 min at 121°C under a pressure of 15 lb/in.2.
3. Dispense in 5-ml (tube) or 3-ml (bottle) amounts, reautoclave, and slant until cool and set. For slants in tubes, use an angle that produces a 12- to 15-mm (ca. 0.5-in.) butt.
4. When cool, tighten lids and store at room temperature or refrigerated.
5. To one tube or bottle, add the following: 3 ml of 1× phthalate-Bacto Peptone, 1 ml of BRS medium, and 50 µl of erythromycin. This must be done on the same day as the inoculation. Note that although erythromycin is added to Robinson’s medium at every subculture, this does not lead to a monoxenic culture as occasionally stated. Additional antibiotic treatment would be needed for this to be the case.
Xenic Culture
When initiating xenic cultures, the stool samples should be inoculated into at least two tubes, one with and the other without antibiotics. In some cases, some component of the natural bacterial flora may be helpful or even necessary for the amebae to become established.
1. Warm several tubes of TYSGM-9 medium in the incubator (35°C for 1 to 2 h).
2. Add 0.1 ml of stock antibiotics to each tube of medium. The final concentration of antibiotics is 100 U of penicillin per ml and 100 µg of streptomycin per ml.
3. After vortexing or vigorously shaking the tube, use a Pasteur pipette to add 3 drops of the starch suspension to each tube of the medium.
4. Place a pea-sized portion of fresh stool sample into the bottom of the tube, and break up the stool gently with the pipette.
5. Tightly cap the tubes, and incubate at a 45 to 50° angle at 35°C for 48 h.
6. Examine the tubes with an inverted microscope and the 10× objective for the presence of amebae. Amebae, if present, are usually seen attached to the underside of the tubes, interspersed with the fecal material and rice starch. Sometimes it is necessary to gently invert the tubes in order to disperse the stool material and rice starch to uncover the amebae. If you do not have an inverted microscope, proceed to step 13.
7. If amebae are not seen, stand the tubes upright for about 30 min at 35°C.
8. With a Pasteur pipette, remove from the bottom of each tube the entire sediment and inoculate the sediment into fresh tubes containing rice starch and antibiotics.
9. Incubate as above for another 48 h.
10. Examine the tubes as before and discard the tubes if amebae are still not seen. Report patient results as negative.
Note The patient results should not be reported unless the quality control organisms and cultures are growing, thus indicating the culture system is performing according to expected results.
11. If amebae are present in small numbers, chill the tube in ice-water for 5 min and centrifuge the tube for 5 min at 250 × g. Aspirate and discard the supernatant, and inoculate the sediment into a fresh tube as before.
12. If amebae are present in large numbers, let the tube stand upright for 30 min and remove about 0.2 ml of sediment from the bottom. Inoculate the sediment into fresh tubes as before.
13. If you do not have an inverted microscope, stand the tubes upright for about 30 min at 35°C. With a sterile pipette, remove about 0.5 ml of sediment from the bottom of the tube and place a couple of drops onto each of two slides. Add 2 drops of methylene blue solution to one of the slides. Cover both with coverslips, and examine the slides under the microscope for amebae. Amebae may appear rounded or with pseudopodial extrusions. The nuclei may be clearly seen in the methylene blue preparation. Proceed to steps 7 through 12.
Axenic Culture
Axenic culture is used for research when strains of organisms are necessary for work requiring a culture system free from bacterial contaminants. If the axenic culture tubes become contaminated, 1,000 U of penicillin per ml and 1,000 µg of streptomycin or 50 µg of gentamicin per ml can be added to each tube. If, however, the contaminant happens to be Pseudomonas spp., it is probably better to discard the tube and use an uncontaminated tube for subculture purposes (3, 15).
1. Remove tubes containing TYI-S-33 medium from 4°C, and incubate at 35°C for 1 to 2 h.
2. With an inverted microscope, examine stock culture tubes of E. histolytica for any signs of bacterial contamination (no longer acceptable for use). Select one or several tubes showing good growth of amebae. Since the tubes are incubated in a slanted position, usually at an angle of 5 to 10°, a thick button of amebae will be seen at the bottom of the tube. Gently invert the tubes once or twice to disperse the amebae uniformly, and examine the tubes again. A majority of the amebae should be attached to the tube walls and show pseudopodial motility. If you do not have an inverted microscope, examine organisms from the bottom of the tube (as a wet smear). If you can see pseudopodial motility, proceed to step 3.
3. Immerse the tubes in a bucket of ice-cold water for about 5 to 10 min to dislodge the amebae from the tube walls. Invert the tubes several times to distribute the amebae.
4. Remove, with a Pasteur pipette, about 1.0 to 1.5 ml of culture medium; inoculate 0.5 to 1 ml into a fresh tube. Inoculate the rest of the fluid into nutrient agar, brain heart infusion, and thioglycolate broth for routine monitoring of bacterial contamination. Inoculate several tubes this way, and incubate the cultures slanted at 5 to 10° at 35°C as before.
5. If amebic growth is not good but some amebae are attached to the tube walls, remove about 10 ml of medium from the bottom with a serologic pipette and add 10 ml of fresh medium.
6. If amebic growth is not good and only few amebae are present along with a lot of debris, centrifuge the tube at 250 × g for 10 min, aspirate the supernatant fluid, and transfer the sediment to a fresh tube and incubate as before.
Quality Control for Intestinal Protozoan Cultures
The following control strains should be available when using these cultures for clinical specimens (3): ATCC 30925 (Entamoeba histolytica HU-1:CDC), ATCC 30015 (Entamoeba histolytica HK-9), and ATCC 30042 (Entamoeba histolytica-like, Laredo strain, culture at 25°C) (17).
1. Check all reagents and media (Balamuth’s aqueous egg yolk infusion medium, Boeck and Drbohlav’s LES medium, PBS solution no. 8, rice starch suspension, Tween 80 solution, and TYSGM-9 and TYI-S-33 media) each time they are used or periodically (once a week). The media and all solutions should be free of any signs of precipitation and bacterial and/or fungal contamination.
2. Maintain stock cultures of E. histolytica at 35°C (ATCC 30925 [strain HU-1:CDC] and ATCC 30015 [strain HK-9]). Maintain E. histolytica-like Laredo strain (ATCC 30042) at 25°C.
A. Transfer stock culture (ATCC 30925) every other day with TYSGM-9 medium.
B. Transfer stock culture (ATCC 30015) once every 3 days with TYI-S-33 medium.
C. Transfer stock culture (ATCC 30042) once a month with TYSGM-9 medium.
D. E. histolytica trophozoites measure 10 to 60 µm and demonstrate directional motility by extruding hyaline, finger-like pseudopodia from the cytoplasm. Cysts are not usually found in cultures.
E. Trophozoites are uninucleate and characterized by finely granular, uniform, evenly distributed peripheral chromatin. The nucleolus is small and usually centrally located but may be eccentric.
3. Depending on its use, the microscope(s) may need to be calibrated (within the last 12 months), and the original optics used for the calibration should be in place on the microscope(s). The calibration factors for all objectives should be posted on the microscope for easy access.
Note If the tubes containing fecal material are positive for amebae after 48 h of incubation, confirm the identification with the permanent stained smear. If the tubes do not show any amebae, subculture the contents of the tubes as described above and incubate for an additional 48 h. If the tubes are still negative for amebae, report the specimen as negative and discard the tubes. Even when the culture system is within quality control guidelines, a negative culture is still not definitive in ruling out the presence of E. histolytica. Xenic cultures of E. histolytica serve only as a supplemental procedure and never replace the primary diagnosis by microscopic examination of concentration sediments and permanent stained smears. Axenic culture is used to maintain quality control strains and for research purposes.
Specimens would include cerebrospinal fluid (CSF), biopsy tissue, and autopsy tissue of the brain; for Acanthamoeba spp., corneal scrapings or biopsy material, contact lenses and contact lens paraphernalia such as lens cases and solutions, skin abscess material, ear discharge, or feces can also be used. All clinical specimens should be processed within 24 h; 2 to 3 h is recommended. However, eye-related specimens can be shipped by mail with apparently few problems (18–21). The procedure for growing Naegleria and Acanthamoeba from clinical specimens involves the use of a nonnutrient agar spread with E. coli or some other nonmucoid bacteria. Amebas begin feeding on bacteria and soon grow to cover the agar surface in 1 to 2 days at 37°C. The presence of the protozoa can be confirmed by examining the agar surface using an inverted microscope or with a conventional microscope by inverting the plate on the stage and focusing through the agar with a 10× objective (Fig. 8.1).
Diagnostic methods include direct microscopy of wet mounts of CSF or stained smears of CSF sediment, light or electron microscopy of tissues, in vitro cultivation of Acanthamoeba, and histologic assessment of frozen or paraffin-embedded sections of brain or cutaneous lesion biopsy material. Immunocytochemistry, chemifluorescent-dye staining, PCR, and analysis of DNA sequence variation also have been used for laboratory diagnosis. Several approaches to specimen handling and diagnostic methods can be seen in Table 8.1.
Environmental Issues
There continue to be ongoing discussions and publications regarding the association of free-living amebae and their intracellular bacterial flora, particularly within the context of environmental transmission of infection, contamination of equipment, and overall environmental concerns. The recognition that Acanthamoeba spp. can sequester a variety of bacteria with known potential for causing human disease suggests that these amebae serve as reservoirs for bacterial pathogens. The increase in the reported incidence of Acanthamoeba infections may be due to greater recognition of the disease potential of these amebae. Also, a number of factors may account for an increased incidence of infection, such as a large number of HIV-infected individuals and more patients undergoing chemotherapy or immunosuppressive therapy for organ transplantation (22).
Acanthamoeba Medium
For the isolation of Naegleria or Acanthamoeba spp. from tissues or soil samples, the following procedure is recommended.
Acanthamoeba Medium
Page’s saline (10×)
1. Autoclave at 121°C for 15 min.
2. Store refrigerated in a glass bottle for up to 6 months.
Nonnutrient agar
1. Dissolve agar in Page’s saline and distilled water with gentle heating; stir or swirl.
2. Aliquot 20 ml into screw-cap tubes (20 by 150 mm).
3. Autoclave at 15 lb/in.2 for 15 min; label deeps with 12-month expiration date, and store in the refrigerator.
4. Melt agar deeps, and pour into petri dishes as needed. Plates may be stored in the refrigerator for up to 3 months.
Monoxenic culture
1. Remove the nonnutrient agar plates from the refrigerator, and place them in a 37°C incubator for 30 min.
2. Add 0.5 ml of ameba saline to a slant bacterial culture of E. coli or Enterobacter aerogenes. Gently scrape the surface of the slant (do not break the agar surface). Suspend the bacteria uniformly by gently pipetting with a Pasteur pipette, and add 2 or 3 drops of this suspension to the middle of the warmed agar plate. Spread the bacteria on the surface of the agar with a bacteriological loop.
3. Inoculate the specimen on the center of the agar plate as described below.
A. For CSF samples, centrifuge the CSF at 250 × g for 10 min. With a sterile serologic pipette, carefully transfer all but 0.5 ml of the supernatant to a sterile tube and store at 4°C (for possible future use). Mix the sediment in the rest of the fluid, and, with a Pasteur pipette, place 2 or 3 drops in the center of the nonnutrient agar plate that has been precoated with bacteria. After the fluid has been absorbed, seal the plates with a 5- to 6-in. length of 1-in.-wide Parafilm strip. Incubate the plate inverted at 37°C in room air. Using a wax pencil or laboratory marker, you may want to make a circle on the underside of the plate to indicate exactly where the specimen was inoculated onto the agar.
B. For tissue samples, triturate a small piece of the tissue (brain, lung, skin abscess, corneal biopsy, or similar specimens) in a small quantity (ca. 0.5 ml) of ameba saline. Process as above. Corneal smear, ear discharge material, etc., may be placed directly on the agar surface. Incubate central nervous system tissues at 37°C (room air) and tissues from other sites at 30°C.
C. Water samples of 10 to 100 ml may be processed to isolate these amebae. First, filter the water sample through three layers of sterile gauze or cheesecloth to remove leaves, dirt, etc. Next, either (i) filter the sample through a sterile 5.0-µm cellulose acetate membrane (47 mm in diameter), invert the membrane over a nonnutrient agar plate precoated with bacteria, seal, and incubate the plates as above; or (ii) centrifuge the water sample for 10 min at 250 × g. Aspirate the supernatant, suspend the sediment in about 0.5 ml of ameba saline, and deposit this suspension in the center of the nonnutrient agar plate precoated with bacteria. Seal and incubate the plate at 37°C as before.
D. For soil samples, mix about 1 g of the soil sample with enough ameba saline (ca. 0.5 to 1 ml) to make a thick slurry. Inoculate this slurry in the center of the nonnutrient agar plate precoated with bacteria, and incubate as above.
E. For contact lens solutions, small volumes (ca. 1 to 2 ml) may be inoculated directly onto the nonnutrient agar plates precoated with bacteria. Larger volumes (2 to 50 ml) should be centrifuged as in step 3, and the sediment should be inoculated onto the center of the nonnutrient agar plate and incubated as above.
4. Using the low-power (10×) objective, examine the plates microscopically for amebae (cysts or trophozoites) every day for 10 days. Thin linear tracks (areas where amebae have ingested bacteria) might also be seen. If amebae are seen, circle that area with a wax pencil, carefully remove the Parafilm seal in a biological safety cabinet, open the lid of the petri dish, and carefully cut out the marked area from the agar with a spatula that has been heated to red hot and cooled before use to prevent contamination. Transfer the piece face down onto the surface of a fresh agar plate coated with bacteria, seal the plate with Parafilm, and incubate as before. Naegleria and Acanthamoeba spp. can be cultured by this method and, with periodic transfers, can be maintained in the laboratory indefinitely. In lieu of subcultures, the organisms can also be frozen for long-term storage. Under the microscope, the amebae resemble small uneven spots; observation for several seconds may reveal organism motility. After 4 to 5 days of incubation, the amebae begin to encyst and both trophozoites and cysts are visible. Unfortunately, Balamuthia spp. cannot be grown by using this system; they can be grown by using tissue culture methods on monkey kidney or lung fibroblast cell lines.
5. The enflagellation experiment is carried out as follows.
A. Examine the plates every day for signs of amebae. If present, amebae will feed on bacteria, multiply, and cover the entire surface of the plate within a few days. Once the food supply is exhausted, the amebae will differentiate into cysts.
B. Use a wax pencil to mark the area containing a large number of amebic trophozoites.
C. Using a bacteriological loop, scrape the surface of the agar at the marked area and transfer several loopfuls of the scraping to a sterile tube containing about 2 ml of sterile distilled water. Alternately, flood the surface of the agar plate with about 10 ml of sterile distilled water, gently scrape the agar surface with a loop, transfer the liquid to a sterile tube, and incubate at 37°C.
E. Periodically examine the tube with an inverted microscope for the presence of flagellates.
a. N. fowleri, the causal agent of primary amebic meningoencephalitis, undergoes transformation to a pear-shaped flagellate, usually with two flagella but occasionally with three or four flagella; the flagellate stage is a temporary nonfeeding stage and usually reverts to the trophozoite stage (Fig. 8.2). N. fowleri trophozoites are typically ameba-like and move in a sinuous way. They are characterized by a nucleus with a centrally located, large nucleolus. The trophozoites are also characterized by the presence of a contractile vacuole that appears once every 45 to 50 s and discharges its contents. The contractile vacuole looks like a hole or a dark depression inside the trophozoite and can be easily seen when examining the plate under the 10× or 40× objective. When the food supply is exhausted, N. fowleri trophozoites differentiate into spherical, smooth-walled cysts.
b. In contrast, Acanthamoeba spp., which cause keratitis and granulomatous amebic encephalitis, do not transform into the flagellate stage. Acanthamoeba trophozoites are characterized by the presence of fine, thorn-like processes that are constantly extended and retracted. The trophozoites produce double-walled cysts characterized by a wrinkled outer wall (ectocyst) and a polygonal, stellate, oval, or even round inner wall (endocyst). The trophozoites are also characterized by the presence of the contractile vacuole, which disappears and reappears at regular intervals (45 to 50 s).
c. The cysts of both Acanthamoeba and Naegleria spp. are uninucleate.
d. Immunofluorescence and immunoperoxidase tests with monoclonal or polyclonal antibodies (available at the Centers for Disease Control and Prevention) are helpful in differentiating Acanthamoeba and Balamuthia spp. in fixed tissue.
Figure 8.2 Naegleria fowleri flagellate stage. (Upper) When placed in distilled water (enflagellation test), N. fowleri, the causal agent of primary amebic meningoencephalitis, undergoes transformation to a pear-shaped flagellate, usually with two flagella but occasionally with three or four flagella; the flagellate stage is a temporary nonfeeding stage and usually reverts to the trophozoite stage. (Lower) Naegleria flagellate stage (microtubules are highlighted in green, basal bodies in red, and DNA is stained blue). (Courtesy of Lillian Fritz-Laylin; from http://genome.jgi-psf.org/Naegr1/Naegr1.home.html.) doi:10.1128/9781555819002.ch8.f2
Quality Control for Pathogenic Free-Living Amebae
The following control strains should be available when using these cultures for clinical specimens: ATCC 30010 (Acanthamoeba castellanii) and ATCC 30215 (N. fowleri). ATCC strains of E. coli or E. aerogenes are not necessary. Any routine clinical isolate or stock organism is acceptable.
1. Check all reagents and media (ameba saline, distilled water, and nonnutrient agar plates) each time they are used or periodically (once a week).
A. The media and all solutions should be free of any visible signs of precipitation and bacterial and/or fungal contamination.
B. Examine the nonnutrient agar plates under the 4× objective of an inverted or binocular microscope, and make sure that no fungal contamination has occurred.
2. Maintain stock cultures of A. castellanii and N. fowleri at 25°C.
A. Transfer stock cultures monthly, using nonnutrient agar plates prepared with Page’s ameba saline and the bacterial overlay of E. coli.
B. N. fowleri measures 10 to 35 µm and demonstrates an eruptive locomotion by producing smooth hemispherical bulges. The cyst produces smooth (7- to 15-µm) walls. The flagellate stage does not have a cytostome.
C. A. castellanii is 15 to 45 µm and produces fine, tapering, hyaline projections called acanthapodia. It has no flagellate stage but produces a double-walled cyst with a wrinkled outer wall (10 to 25 µm).
D. Trophozoites of Naegleria and Acanthamoeba spp. are uninucleate, characterized by a large, dense central nucleolus.
E. For staining, a slide prepared from a stock strain of amebae is run in parallel with the patient slide. Staining results are acceptable when the control amebae stain well.
F. Plate both stock cultures onto fresh media, and incubate at 37°C parallel with the patient culture. Culture results are acceptable when growth appears by day 7.
G. Run N. fowleri in parallel with the patient culture being observed for enflagellation. The test results are acceptable when free-swimming, pear-shaped flagellates with two flagella are observed in 2 to 24 h on the control slide.
3. Depending on use, the microscope(s) may have to be calibrated (within the last 12 months), and the original optics used for the calibration should be in place on the microscope(s). The calibration factors for all objectives should be posted on the microscope for easy access.
Note For patient specimens, if a plate is positive for amebae and the amebae transform into flagellates, the specimen should be reported as positive for N. fowleri. If the amebae do not transform into flagellates even after overnight incubation, if the trophozoites possess the characteristic acanthapodia, if they show a large centrally placed nucleolus in the nucleus on trichrome stain, and if the trophozoites differentiate into the characteristic double-walled cysts, the specimen should be reported as positive for Acanthamoeba sp. For contact lens solution, if the plates are positive for amebae and the amebae do not transform into flagellates but differentiate into cysts with a wrinkled outer ectocyst and an inner stellate, polygonal, oval, or round endocyst, the specimen should be reported as positive for Acanthamoeba sp. Naegleria spp. have not been isolated from contact lens solutions; however, small amebae (e.g., hartmannellid or vahlkampfiid amebae, which produce smooth-walled cysts), probably contaminants, have occasionally been isolated from these solutions. Plates inoculated with water samples are usually positive for many genera and species of small free-living amebae (freshwater is their normal habitat). Therefore, the sample should be reported as positive for small free-living amebae. The physician should be notified immediately if patient specimens are positive for Acanthamoeba or Naegleria spp.
Most patient specimens, especially CSF, should be examined microscopically as soon as they arrive in the laboratory. However, if the specimen is very clear with no visible sediment, it should be cultured without being examined microscopically.
1. After centrifugation, remove a small drop of the CSF sediment, place it on a microscope slide, cover it with a no. 1 coverslip, seal the edges of the coverslip with Vaspar (optional), and examine it immediately with the 10× or 40× objective (phase-contrast/differential interference contrast optics are preferred). If bright-field microscopy is used, reduce the illumination by adjusting the iris diaphragm.
A. The N. fowleri trophozoite is highly motile and can be identified by its sinuous movement. A warmed penny may be applied to the bottom surface of the slide to activate the movement of the trophozoite. Occasionally, a flagellate is seen traversing the field.
B. Acanthamoeba trophozoites are rarely seen in the CSF. If present, they may be recognized by their characteristic acanthapodia, which are constantly extended and retracted. Both amebae, especially Acanthamoeba spp., may be recognized by the contractile vacuole.
2. Contact lens care solutions (opened) can be processed and examined like CSF. Testing of unopened commercial lens care solutions should be handled by the FDA.
3. If very small amounts of tissue are received, they should be reserved for culture. If the specimen received is visibly contaminated with bacteria and/or fungi, the tube containing the specimen should be placed in an ice-water bath for 3 min prior to centrifugation. This causes any trophozoites attached to the walls of the tube to come off and drop into the fluid prior to centrifugation.
4. Material from the surface of a positive agar plate can be removed, fixed, and stained with trichrome for microscopic examination at a higher magnification (×1,000). Results obtained with wet mounts should always be confirmed by performing permanent stained smears (trichrome, iron hematoxylin) for nuclear characteristics to differentiate the amebae from host cells. Organisms may not be recovered if appropriate centrifugation speeds and times are not used.
Liquid Culture Media for Pathogenic Free-Living Amebae
Although Acanthamoeba spp. can be grown in serum-free liquid medium, N. fowleri requires the addition of fetal calf serum or brain extract to grow in a liquid medium. Peptone-yeast extract-glucose medium for Acanthamoeba spp. and Nelson’s medium for N. fowleri are described below.
Peptone-Yeast Extract-Glucose (PYG) Medium for Acanthamoeba spp.
Final pH is 6.5 ± 0.2.
1. Dissolve all ingredients except CaCl2 in about 900 ml of distilled water in a bottle or flask with a magnetic stirrer.
2. Add CaCl2 while stirring.
3. Bring the volume to 1,000 ml with distilled water.
4. Dispense into 16- by 125-mm screw-cap tubes, 5 ml per tube.
5. Autoclave at 121°C for 15 min.
6. When tubes have cooled, label the tubes as Acanthamoeba medium with the preparation date and an expiration date of 3 months.
7. Store at 4°C.
Nelson’s Medium for N. fowleri
1. Add ameba saline to distilled water to make 1× ameba saline.
2. Dissolve the ingredients in ameba saline with a magnetic stirrer.
3. Dispense into 16- by 125-mm screw-cap tubes, 10 ml per tube.
4. Autoclave for 15 min at 15 lb/in.2.
5. Cool, and label as Nelson’s medium with the preparation date and an expiration date of 3 months.
6. Store at 4°C.
7. Add 0.2 ml of heat-inactivated fetal calf serum to each tube before inoculating with the amebae.
Cell Culture
Acanthamoeba, Balamuthia, Naegleria spp., and Sappinia spp. can be inoculated onto a number of different mammalian cell lines. The organisms grow very well and demonstrate cytopathic effects, similar to those seen in routine viral cultures. However, most laboratories do not offer these methods on a routine basis.
Standard Commercial Media for Growth and Isolation of Acanthamoeba spp.
Several studies have reported on the use of standard commercial media used for growth and isolation of Acanthamoeba spp. Buffered charcoal-yeast extract agar (BCYE) is an excellent commercially available culture medium for the recovery of Acanthamoeba spp. Good trophozoite recovery was obtained using BBL Trypticase soy agar (TSA) with 5% rabbit blood, TSA with 5% horse blood, and Remel TSA with 5% sheep blood. BBL TSA with 5% horse blood or 5% rabbit blood yielded good recovery of cysts. Nonnutrient agar with either live or dead bacteria yielded good recovery of trophozoites; however, live bacteria may be required for good cyst recovery.
Blastocystis spp. (Blastocystis hominis)
Blastocystis is a single-celled enteric protozoan that has a worldwide distribution. Blastocystis belongs to the phylum Stramenopila, is found in humans and animals, and is the most common parasite isolated from human stool samples throughout the world. Rates of infection vary from <5% in developed countries to >50% in developing countries. Specimens include stool material, mucus, or a combination of the two. The clinical specimen(s) should be no more than 24 h old; a maximum of 2 to 3 h is recommended. Xenic or monoxenic laboratory cultures of Blastocystis isolates, which are cultures of Blastocystis cells grown in association with nonstandardized or single known species of microorganisms, respectively, can be maintained in TYGM-9 medium or Boeck and Drbohlav’s inspissated egg medium (see below) (12).
Tryptone/Yeast Extract/Glucose/Methionine) TYGM-9 Xenic Medium
See TYSGM-9 Medium for Entamoeba histolytica earlier in this chapter.
Diphasic LE Medium (Blastocystis spp.)
LE medium is the NIH modification of Boeck and Drbohlav’s medium.
Locke’s Solution
Autoclave for 15 min at 121°C under a pressure of 15 lb/in.2. Cool to room temperature, and remove any precipitate by filtration (Whatman no. 1 paper). Reautoclave to sterilize.
Note The overlay can also be prepared using 90% PBS, 9% sterile horse serum, 1% or 20% [wt/vol] bacteriologic peptone, 1 mg of rice starch, and 500 µl of penicillin-streptomycin solution (12).
Preparation of Complete Medium
1. To prepare the egg slant, surface sterilize fresh hen eggs by flaming in 70% ethanol and break into a graduated cylinder.
2. Add 12.5 ml of Locke’s solution per 45 ml of egg.
3. Emulsify in a Waring-type blender, and filter through gauze into a flask.
4. Place under vacuum to draw out all air bubbles.
5. Add 5-ml amounts of the emulsified egg to standard culture tubes (16 by 125 mm), and autoclave at 100°C for 10 min with the tubes at an angle that produces a 12- to 15-mm (ca. 0.5-in.) butt. The resulting egg slants should be free of bubbles.
6. Cool to room temperature, overlay slants with 6 ml of Locke’s solution, and autoclave for 15 min at 121°C under a pressure of 15 lb/in.2.
7. Let the slants cool to room temperature, tighten the caps, and refrigerate the slants for up to 6 months.
The culture should be examined every 2 days for 1 week (phase-contrast microscopy is recommended) (12). Another option is to use reduced light with the condenser lowered to provide more contrast. If parasites are seen in the sediment, the material can be placed in a fecal preservative for staining and examination using the oil immersion objective (100× oil immersion objective).
Trichomonas vaginalis
Specimens from women may consist of vaginal exudate collected from the posterior fornix on cotton-tipped applicator sticks or genital secretions collected on polyester sponges. Specimens from men can include semen, urethral samples collected with polyester sponges, or urine. Urine samples collected from the patient should be the first voided specimen in the morning. It is critical that clinical specimens be inoculated into culture medium as soon as possible after collection (23–26). Although collection swabs can be used, there are often problems with specimens drying prior to culture; immediate processing is mandatory for maximum organism recovery. The culture method is considered to be the most sensitive for the diagnosis of trichomoniasis; however, because of the time and effort involved, many laboratories have changed to the monoclonal antibody detection kits or more automated molecular methods. Another approach would be to use the plastic envelope method for Trichomonas vaginalis (InPouch TV [BIOMED Diagnostics, San Jose, CA]), a simplified technique for the transport and culture that is illustrated in Fig. 8.3 and 8.4. Because of its long shelf life, relatively low expense, and high sensitivity, studies confirm that the pouch system provides a good diagnostic culture method for T. vaginalis (27, 28). It has been found to be more sensitive than either Diamond’s or Trichosel medium (27).
Figure 8.3 Illustration of the InPouch TV culture system for Trichomonas vaginalis. From top to bottom: (1) introduction of the specimen into the upper chamber containing a small amount of medium; (2) application of a plastic holder for microscope viewing prior to expressing medium into the lower chamber (optional); (3) transfer of a small amount of medium in the upper chamber to the lower chamber; (4) rolling down the upper chamber and sealing it with the tape; (5) plastic viewing frame used to immobilize the medium in the pouch for examination under the microscope. doi:10.1128/9781555819002.ch8.f3
Figure 8.4 InPouch TV culture system for Trichomonas vaginalis. Note the pouch, swabs, and plastic pouch holder for microscopic examination of the pouch contents. doi:10.1128/9781555819002.ch8.f4
Lash’s Casein Hydrolysate-Serum Medium
Lash’s casein hydrolysate-serum medium is a culture medium used to diagnose T. vaginalis.
Lash’s Casein Hydrolysate-Serum Medium
Dissolve in 500 ml of distilled water. To this solution, add:
1. Adjust the pH to 6.0 with concentrated phosphoric acid. Dispense in 5-ml aliquots, plug the tubes, and autoclave at 121°C for 15 min.
2. Prepare serum solution:
3. Sterilize the serum solution by filtration.
4. The complete medium contains 5 ml of serum solution and 5 ml of basic solution for each tube.
5. Incubate the cultures at 37.5°C, and examine 24 and 48 h after inoculation.
T. vaginalis has been studied for a number of years with agar plate cultures for both cloning and diagnosis (2–4). Although broth and agar culture for T. vaginalis are successful, many laboratories have transitioned to molecular based methods. Many media are available for the isolation of T. vaginalis; some of these can be purchased commercially and have a relatively long shelf life.
CPLM (Cysteine-Peptone-Liver-Maltose) Medium
Ringer’s solution
Dissolve the ingredients in the order listed, and bring the volume up to 100.0 ml with distilled water.
Liver infusion
1. Place the distilled water in a large beaker.
2. Add the liver infusion powder.
3. Infuse for 1 h at 50°C.
4. Raise the temperature to 80°C for 5 min to coagulate the protein.
5. Filter through a Whatman no. 1 paper with a Büchner funnel.
Methylene blue solution
Mix well until dissolved.
CPLM (Cysteine-Peptone-Liver-Maltose)
Complete Medium
1. With a magnetic stirrer, mix Ringer’s solution and the liver infusion in a large beaker.
2. Add peptone, maltose, cysteine HCl, and agar in that order, and heat the mixture until dissolved.
3. Add 0.7 ml of aqueous methylene blue.
4. Adjust the pH to 5.8 to 6.0 with 1 N NaOH or 1 N HCl.
5. Dispense 8-ml volumes into culture tubes.
6. Autoclave at 121°C for 15 min.
7. Aseptically add 2 ml of human serum, heat inactivated at 56°C for 30 min and cooled, per tube. Horse serum is recommended as a replacement for human serum, particularly when considering safety issues such as handling human blood and blood products.
8. Label as CPLM medium with the preparation date.
9. Store at room temperature. Use as long as the amber zone indicating an anaerobic condition persists.
Diamond’s TYM (Trypticase-Yeast Extract-Maltose) Complete Medium
1. Dissolve the buffer salts in the distilled water with a magnetic stirrer.
2. Add the remaining ingredients except the agar, in the order given, one at a time until dissolved.
3. Adjust the pH to 6.0 with 1 N HCl.
4. Add agar, and heat to dissolve.
5. Autoclave at 121°C for 15 min.
6. Cool to 45°C, and add 100 ml of bovine, sheep, or horse serum that has been inactivated for 30 min at 56°C.
7. Aseptically dispense 10-ml volumes to 16- by 125-mm screw-cap tubes.
8. Label as TYM medium with the preparation date and an expiration date of 10 days.
9. Store at 4°C.
Hollander’s Modification of TYM Complete Medium
Hollander’s modification of TYM complete medium differs from TYM medium by the replacement of cysteine with additional ascorbic acid and the addition of potassium chloride, potassium carbonate, and ferrous sulfate (2).
Proceed as described above for Diamond’s TYM complete medium.
Note Hollander also includes agar to 0.05%.
Diamond’s Complete Medium (Modified by Klass)
1. Dissolve the ingredients one at a time in the order given.
2. Adjust the pH to 6.0 with 1 N HCl or 1 N NaOH.
3. Dispense in 12.5-ml aliquots to 16- by 125-mm screw-cap tubes.
4. Autoclave at 121°C for 15 min.
5. When cool (50°C), add 1 ml of sterile inactivated horse serum and 0.5 ml of antibiotic mixture to each tube.
6. Label as modified TYM medium with the date of preparation and an expiration date of 3 weeks.
Antibiotic Mixture
1. Mix thoroughly.
2. Dispense 1 ml of the antibiotic mixture into sterile screw-cap vials or sterile cryovials.
3. Label as antibiotic solution with the date of preparation and an expiration date of 1 year.
4. Store at 220°C in cryoboxes.
Serum Substitutions
Bovine, sheep, or horse serum can be substituted in the step involving CPLM complete medium.
Axenic Culture
1. Inoculation of culture medium.
A. Remove tubes containing culture medium from 4°C, and incubate at 37°C for 1 to 2 h.
B. Vigorously shake the cotton-tipped portion of the applicator stick containing the patient specimen in the medium, and then break off the tip with sterile forceps and drop it into the medium.
C. If the material is collected on polyester sponges, drop the sponges into the medium and shake the tube.
D. Centrifuge urine samples for 10 min at 250 × g, aspirate the supernatant, and inoculate the sediment into the medium.
E. Examine the tubes daily for several days, and subculture if necessary. To subculture, first shake the tube to disperse the organisms uniformly, remove about 1 to 2 ml, and inoculate into a warmed, fresh tube.
F. Incubate the tubes in a slanted position (45° angle).
G. Incubate the control tubes and those containing patient material for at least 72 to 96 h.
H. Examine the entire length of the tube. If the specimen is positive, T. vaginalis will be found freely swimming or attached to the tube walls.
I. Do not report negative results until 96 h.
2. To maintain stock cultures:
A.With an inverted microscope, examine stock culture tubes of T. vaginalis for any signs of bacterial contamination. Select one or more tubes showing good growth. Since the tubes are incubated in a slanted position, usually at an angle of 45°, a thick button of organisms is seen at the bottom of the tube. Gently invert the tube once or twice to disperse the trichomonads uniformly, and examine the tubes again. A large number of the organisms should be freely swimming, and a few will be attached to the tube walls.
B. Immerse the tubes in a bucket of ice-cold water for about 5 to 10 min to dislodge the trichomonads from the tube walls. Invert the tubes several times to distribute the organisms.
C. With a sterile Pasteur pipette, remove about 1.0 to 1.5 ml; inoculate 0.5 to 1.0 ml into a fresh tube. Inoculate the rest of the fluid into nutrient agar, brain heart infusion, and thioglycolate broth for routine monitoring of bacterial contamination. Inoculate several additional tubes this way, and incubate the cultures slanted at a 45° angle at 37°C as before.
D. If growth is poor and only few organisms are present along with a lot of debris, centrifuge the tube at 250 × g for 10 min, aspirate the supernatant, transfer the sediment to a fresh tube, and incubate as before.
Quality Control for Pathogenic Flagellates (T. vaginalis)
ATCC 30001 (T. vaginalis) should be available as a control strain when these cultures are used for clinical specimens.
1. Check all reagents and media (at least once a week). All media, including Ringer’s solution, should be free of any signs of precipitation and bacterial and/or fungal contamination.
2. Depending on use, the microscope(s) may have had to be calibrated within the last 12 months, and the original optics used for the calibration should be in place on the microscope(s). The calibration factors for all objectives should be posted on the microscope for easy access.
3. Maintain stock cultures of ATCC 30001 (T. vaginalis).
A. Transfer stock cultures weekly.
a. Stock organisms should always be cultured at the same time a patient specimen is inoculated into culture medium.
b. If the stock organisms multiply and remain viable during the 96 h, patient results can be reported.
B. Stain
a. A slide prepared from a stock strain of T. vaginalis is run in parallel with the patient slide.
b. Staining results are acceptable when the control organisms stain well.
4. Control organisms must be cultured each time a patient specimen is inoculated into the culture medium.
Note If no trophozoites are seen after 4 days of incubation, discard the tubes and report the culture as negative. Results of patient specimens should not be reported as positive unless control cultures are positive. Cultivation is the most sensitive method for the diagnosis of trichomoniasis. Every effort, therefore, must be made to inoculate patient materials into culture medium. However, since this method may take as long as 3 to 4 days and the patient materials occasionally contain nonviable organisms, it is imperative that microscopic examination of wet smears and/or stained smears (Giemsa) also be performed.
Axenic Cultivation of T. vaginalis in a Serum-Free Medium
Mammalian serum or bovine serum albumin is required for T. vaginalis grown under axenic conditions. However, these components inhibit several biological properties of these parasites. PACSR is a serum replacement, free of bovine serum albumin, which is used for axenic cultivation of E. histolytica. Studies indicate that PACSR added to several of the standard media used for the cultivation of T. vaginalis can support growth in the absence of serum (29).
Dientamoeba fragilis
Dientamoeba fragilis is a protozoan pathogen of the human gastrointestinal tract that was originally classified as an amoeba. Later studies confirmed its relationship with trichomonads. The approach to in vitro culture is not new; however, some excellent improvements have recently been developed (30, 31). Currently, no axenic cultures of D. fragilis exist. D. fragilis will grow happily with a support flora consisting mostly of E. coli. Slight variations in the species of prokaryotic support flora present within D. fragilis cultures are unlikely to exhibit any significant effect on growth. A modified Earle’s Balanced Salt Solution (EBSS) containing cholesterol, ferric ammonium citrate, and rice starch can be regarded as a superior liquid overlay that can be used along with the Loeffler’s serum slope for culture of D. fragilis under anaerobic conditions.
Loeffler’s Serum Slants (30)
1. Aliquot 5 ml into tubes, slant, and inspissate in 80°C drying oven.
Note If the slants are left too long at 80°C, Dientamoeba will not grow. Slants should be allowed to cool IMMEDIATELY after they solidify.
2. Add approximately 2 mg of rice starch into the bottom of each slant.
3. Overlay each slant with 5 ml of PBS.
4. After inoculation with fresh stool (pea size), incubate at 42°C under microaerophilic conditions (6% O2, 7.2% CO2, 3.6% H2, 83.3% N2).
Earle’s Balanced Salt Solution Overlay
Another overlay that has proven to be very effective is EBSS, pH 7.8. EBSS provides the highest trophozoite numbers and contains a variety of inorganic salts, as well as sodium hydrogen carbonate (NaHCO3) and glucose. The inorganic salts in the medium help to maintain the osmolarity as well as providing a range of divalent cations needed for cell metabolism. NaHCO3 maintains the buffering capacity in the anaerobic in vitro culture and glucose provides an additional energy source in addition to the rice starch. This saline solution has also been used with good results when supplemented with 40 mg/liter ferric ammonium citrate and 50 mg/liter cholesterol (31).
Autoclave and store at room temperature.
Cryoprxrvation (30)
1. Pool four culture tubes of Loeffler’s medium containing viable organisms.
2. Centrifuge at 500 × g for 5 min; discard supernatant fluid.
3. Add 1 ml of PBS to the pellet and invert tube to obtain even cell suspension.
4. Count the cells present.
5. Dilute quantified cell suspensions in PBS to a final concentration of 106 trophozoites/µl.
6. Plate tubes overnight at −80°C freezer.
7. Remove tubes from the −80°C freezer the following morning and place in a liquid nitrogen freezer.
Flagellates of Blood and Tissue
The first types of media used to culture the blood and tissue flagellates, which are still useful for establishment of cultures, were undefined and contained a complex mixture of ingredients. Improvements led to semidefined formulations that included tissue culture media as a base and, as a next step, addition of tissue culture cells as a feeder layer to promote parasite growth. Newer developed media are completely defined, having replaced the feeder cells with various supplements. Serum, a variable component of the media, can now be replaced by various serum substitutes. Fully defined formulations are available for the cultivation of many of these organisms (8).
For the hemoflagellates found in the bloodstream, probably the most direct means of diagnosis is by examination of a stained blood smear under the microscope (any blood stain is acceptable). However, if isolation of the parasite is required for confirmation, culture can be used. The easiest approach involves the use of Novy-MacNeal-Nicolle (NNN) medium. Because of its relatively short shelf life (2 to 4 weeks with refrigeration) and the infrequent need for culture, most clinical laboratories would not routinely stock NNN medium. However, the Centers for Disease Control and Prevention (Division of Parasitic Diseases, Atlanta, GA) can provide culture services, and in other parts of the world, major reference laboratories may provide a similar service. The procedure for isolating Leishmania spp. from patients with suspected cases of cutaneous, mucocutaneous, and visceral leishmaniases is basically similar to that for isolating Trypanosome spp. The amastigote found within host cells, or the promastigote that develops from the amastigote in culture, would be confirmatory. Punch biopsy specimens or needle aspirates serve as the inocula for NNN medium or Schneider’s Drosophila medium with 30% fetal bovine serum. Other procedures such as splenic puncture and liver biopsy can also yield material but are invasive and less likely to be performed (8).
Specimens for culturing Leishmania spp. may consist of aspirates, scrapings, or biopsy material from skin lesions of patients with cutaneous leishmaniasis; bone marrow aspirates or, more rarely, splenic aspirates from visceral leishmaniasis patients; or normal skin biopsy specimens, lymph node aspirates, or pieces of liver and spleen from suspected or potential wild- or domestic-animal reservoirs. Ensure that the skin surrounding the ulcer is thoroughly cleaned and swabbed with 70% alcohol (sterile saline is not acceptable as a cleansing agent) and allowed to dry before the sample is removed. Also ensure that alcohol does not get into the ulcerated area or broken skin. A punch biopsy specimen taken from the advancing margin of the lesion is often recommended. If skin biopsy fragments are placed in transport medium (RPMI medium supplemented with 10% fetal calf serum) maintained at ambient temperature, delays in transit through the mail will still not prevent recovery of the organisms. In one study, after being received in the laboratory (transit time as long as 3 to 17 days), the biopsy specimens were ground in sterile saline and inoculated into NNN culture tubes. The tubes were incubated at 25°C and subcultured every week until the fifth week. Cultures were positive in 9 of 16 cases in all seasons and for three different Leishmania species. Thus, delayed culture can still yield valuable results from biopsy specimens obtained under field conditions (32).
It is imperative that only a few drops of bone marrow juice or spleen aspirate be inoculated into tubes. Inoculate several tubes with a few drops each rather than inoculating a single tube with a large volume (1 to 2 ml), since the serum in the specimen may contain leishmanicidal or inhibitory factors that prevent the growth of organisms. Alternatively, bone marrow juice may be centrifuged for 10 min at 250 × g and the sediment may be washed in 0.85% saline by centrifugation and then inoculated into culture tubes. Buffy coat from the blood sample rather than whole blood should be inoculated. Because the leishmanias are fastidious organisms and not all isolates may grow in any one medium, it is imperative that at least two media be used; for example, use NNN or modified Tobie’s medium and Schneider’s Drosophila medium.
Specimens for culturing Trypanosoma cruzi may consist of blood or the gut contents of the triatomid bug. It is advisable to use two different media such as liver infusion-tryptose (LIT) and NNN for the initial isolation of T. cruzi. Once growth is established, use the medium in which best growth is obtained for subculture. According to James Sullivan, LIT medium, when used as an overlay on Tobie’s slants, is excellent for isolation and diagnosis (8). The major culture form is the epimastigote; occasionally, however, trypomastigotes and amastigotes are also seen.
NNN Medium (Leishmaniasis or Chagas’ Disease) (33)
1. Mix the NaCl and agar in the distilled water in a 500-ml flask.
2. Heat the mixture until the agar melts.
3. Autoclave at 121°C for 15 min.
4. Cool to about 50°C.
5. Add 10 ml of aseptically collected, defibrinated rabbit blood.
Note Although blood collected with EDTA as the anticoagulant can be used for routine stock culture subcultures, it may not be quite as effective as defibrinated blood in isolating organisms from patient specimens. Certainly, if defibrinated blood is not available, blood collected with EDTA as the anticoagulant can be used. Make sure that the rabbit blood is fresh (no older than 10 days). It should be aseptically collected and stored at 4°C until used.
6. Dispense 4 ml into 16- by 125-mm sterile screw-cap culture tubes.
7. Place the tubes at a 10° angle (shallow slant position) until the agar sets.
8. Immediately transfer the tubes into test tube stands, and let stand upright at 4°C so that the bottom portion of the slants is covered with the water of condensation. Rapid cooling increases the water of condensation (Fig. 8.5).
9. Label as NNN medium with the preparation date and an expiration date 3 weeks from the date of preparation.
10. Store at 4°C.
Figure 8.5 Illustration of a tube of NNN medium, used for the culture and recovery of Leishmania spp. (A) Fluid condensate and tissue culture medium overlay containing the developing organisms. (B) Blood agar medium. (Illustration by Sharon Belkin.) doi:10.1128/9781555819002.ch8.f5
NNN Medium, Offutt’s Modification (Leishmaniasis)
1. Heat until the agar is dissolved in the distilled water in a 500-ml flask.
2. Autoclave at 121°C for 15 min.
3. Cool to about 50°C.
4. Add 15 ml of aseptically collected, defibrinated rabbit blood.
5. Dispense 4 ml into 16- by 125-mm sterile screw-cap culture tubes.
6. Place the tubes at a 10° angle (shallow slant position) until the agar sets.
7. Immediately transfer the tubes to test tube stands, and let stand upright at 4°C so that the bottom portion of the slants is covered with the water of condensation. Rapid cooling increases the water of condensation.
8. Label as Offutt’s medium with the preparation date and an expiration date 3 weeks from the date of preparation.
9. Store at 4°C.
Overlay Solution (To Be Used with NNN or NNN Modified)
1. Autoclave at 121°C.
2. Dispense 4 ml aseptically into 16- by 125-mm sterile culture tubes.
3. Label as 0.9% saline with the preparation date and an expiration date 3 weeks from the date of preparation.
4. Store at 4°C.
Evan’s Modified Tobie’s Medium (Leishmaniasis or Chagas’ Disease) (33)
1. Mix all the ingredients in the distilled water in a large beaker with a magnetic stirrer.
2. Heat the mixture until the agar melts.
3. Dispense 5 ml into 16- by 125 mm screw-cap culture tubes.
4. Autoclave at 121°C for 15 min.
5. Cool to about 50°C.
6. Add 1.2 ml of aseptically collected, defibrinated horse blood.
7. Hold the tubes upright in the palm of your hand, and roll the tubes gently to mix the blood and the agar well.
8. Place the tubes at a 10° angle (shallow slant position) until the agar sets.
9. Immediately transfer the tubes into test tube stands, and let stand upright at 4°C so that the bottom portion of the slants will be covered with the water of condensation. Rapid cooling increases the water of condensation.
10. Label as Evan’s modified Tobie’s medium with the preparation date and an expiration date 3 weeks from the date of preparation.
11. Store at 4°C.
Overlay Solution (To Be Used with Tobie’s Medium)
1. Place 750 ml of double-distilled water in a 1-liter beaker, and add the above ingredients one at a time in the order listed until dissolved. Use a magnetic stirrer.
2. Adjust the pH to 7.2 by adding slowly, while stirring, solid Tris (white, crystalline powder, C4H11NO3 used for buffering capacity).
3. Bring up the volume to 1,000 ml with distilled water.
4. Dispense 100 ml into each of several screw-cap flasks or bottles.
5. Autoclave at 121°C for 15 min.
6. Label as overlay solution with the preparation date and an expiration date of 1 month.
7. Store at 4°C.
NIH Method for Trypanosomes and Leishmaniae
Solution 1
Solution 2 (Locke’s Solution)
1. Infuse the beef and distilled water in a water bath at 37°C for 1 h. Heat for 5 min at 80°C.
2. Filter through filter paper (Whatman no. 2V).
3. Add the rest of the above ingredients (solution 1), and adjust the pH to 7.0 to 7.4 with NaOH.
4. Autoclave at 121°C for 20 min.
5. Cool to 45°C, and aseptically add 10% defibrinated rabbit blood.
6. Dispense 5-ml quantities in sterile tubes. Slant and cool.
7. Just before inoculation, overlay with 2 ml of sterile Locke’s solution.
4 N Medium for Trypanosomes and Leishmaniae
The 4 N medium was adapted from the original formula described by Novy and McNeal (9, 34, 35).
4 N Medium for Trypanosomes and Leishmaniae
Agar base
1. Mix the agar and water, and dissolve by autoclaving or steaming.
2. Dispense liquid agar in 5-ml aliquots into 30-ml screw-cap glass bottles.
3. Autoclave again if necessary for sterility.
4. When the medium has cooled to 45°C, add aseptically to each bottle approximately 1 ml of fresh rabbit blood and allow the agar to solidify in a slant.
Overlay for 4 N medium
1. Add 1 ml of Locke’s (see NIH method for trypanosomes and leishmaniae) solution to each bottle containing 5 ml of agar.
2. Incubate the bottles at 37°C for 12 to 24 h to check for sterility.
3. Store the bottles at 4°C for 24 h or more before use.
Yaeger’s LIT Medium (for Chagas’ Disease) (33)
1. Add all the ingredients to the distilled water, and mix well with a magnetic stirrer until dissolved. Heat, if necessary, to dissolve all the ingredients.
2. Using a Whatman no. 42 filter paper in a Büchner funnel, filter with suction. Repeat this filtration one more time.
3. Adjust pH to 7.2 with 1 N NaOH or 1 N HCl.
4. Sterilize by filtration through a 0.22-µm-pore-size membrane filter.
5. Dispense 4.5 ml into each tube.
6. Label as LIT medium, with the date of preparation and an expiration date of 1 month.
Hemin Stock Solution
1. Mix triethanolamine with water, add the mixture to a tube containing hemin, shake well, and let dissolve.
2. To make the complete medium, just before inoculation, add 0.5 ml of inactivated fetal bovine serum and 0.25 ml of antibiotic solution. The final concentrations of the antibiotics are 100 U of penicillin per ml, 100 µg of streptomycin per ml, and 0.2 µg of Fungizone per ml.
USAMRU Blood Agar Medium for Leishmaniae (36)
1. Dissolve 40 g of agar in 1,000 ml of distilled water by heating.
2. Cool the solution, and add 150 ml of defibrinated rabbit blood.
3. Dispense 5-ml portions into 125-by-16-mm screw-cap tubes. Position the tubes at a slant, and allow to cool, preferably on ice, to produce moisture of condensation in the tubes.
4. Incubate the tubes at 35°C for 24 h to ensure sterility. Antibiotics (penicillin and streptomycin) can be added if necessary (see the antibiotic formula in this chapter). The final concentrations of antibiotics are 100 U of penicillin per ml and 100 µg of streptomycin per ml.
5. Inoculate the medium with aspirate material or triturated tissue from biopsy specimen.
Note This medium has been especially useful for primary isolation of the Leishmania braziliensis complex in Latin America.
Liquid Media for Cultivation of Hemoflagellates
Hendricks et al. have reported on the highly successful use of liquid culture media for the rapid cultivation of various species of Leishmania and Trypanosoma (33). Schneider’s Drosophila medium (GIBCO, Grand Island, NY), supplemented with 30% (vol/vol) fetal calf serum, has been used in making primary isolates from human and animal infections as well as for routine maintenance of a wide variety of leishmanial and trypanosomal species in the laboratory. Both Schneider’s medium and Grace’s insect tissue culture medium (GIBCO) promote better growth of organisms and are less costly than the widely used blood-agar-based media. In addition, Schneider’s medium may be freeze-dried for at least 2 years and reconstituted with distilled water in the field, and it will provide excellent culture results.
Stock Antibiotic Solution (To Be Used with All Media)
1. Mix the components thoroughly. The concentration of the stock solution is:
2. Dispense 1 ml of the antibiotic mixture into sterile screw-cap vials or sterile cryovials.
3. Label as antibiotic solution with the date of preparation and an expiration date of 1 year.
4. Store at −20°C in cryoboxes.
Axenic Culture
1. Remove tubes containing culture medium (one of NNN, modified NNN, or Tobie’s medium and one of Schneider’s for Leishmania; one of NNN or Tobie’s medium and one of LIT for T. cruzi) from 4°C, add fetal bovine serum and antibiotics if required, and incubate at 20 to 23°C for 1 to 2 h.
2. Inoculate the specimen (aspirate, scraping, or biopsy material from skin lesions from cutaneous leishmaniasis patients; bone marrow aspirates or splenic aspirates from visceral leishmaniasis patients; or normal skin biopsy specimens, lymph node aspirates, or pieces of liver and spleen from suspected or potential wild- or domestic-animal reservoirs) into the culture tubes. For Chagas’ disease, inoculate a few drops of buffy coat into the culture tubes.
3. Add 0.5 ml of overlay (either saline or other overlay, depending on the medium). The organisms will develop in the fluid condensate and overlay at the bottom of the slant (Fig. 8.5).
4. Incubate the tubes at 20 to 24°C.
5. Once every 2 to 3 days, remove a drop of medium and examine it under the low power (100×) of a microscope, preferably one equipped with phase-contrast optics.
6. If promastigotes are seen, inoculate a couple of drops of the medium into fresh culture tubes. Add a couple of drops of 0.85% saline or the overlay solution (depending on the culture medium used) to the old tube.
7. If visible contamination occurs, add antibiotics to the overlay (to contain 200 U of penicillin/ml and 200 µg of streptomycin/ml). The parasites will not proliferate if bacterial contamination is present.
8. Incubate the tubes containing the patient specimen for at least 2 weeks at 28°C or room temperature.
9. If no organisms are seen even after 2 weeks of incubation, examine several drops of fluid under the microscope for promastigotes. The culture should be observed for 1 month before being signed out as negative. Transfers to fresh medium should be made once or twice a week after the culture is established.
Quality Control for Flagellates of Blood and Tissue
The following control strains should be available when using these cultures for clinical specimens: ATCC 30883 (Leishmania mexicana) and ATCC 30160 (T. cruzi).
1. Check all reagents and media at least once a week. The media should be free of any signs of precipitation and bacterial and/or fungal contamination.
2. The microscope(s) should have been calibrated within the last 12 months, and the original optics used for the calibration should be in place on the microscope(s). The calibration factors for all objectives should be posted on the microscope for easy access.
3. Maintain stock cultures of Leishmania spp.
A. Transfer stock cultures weekly.
a. Stock organisms should always be cultured at the same time a patient specimen is inoculated into culture medium.
b. If the stock organisms multiply and remain viable during the 96 h, patient results can be reported.
B. Stain
a. Stain a slide prepared from stock culture in parallel with the patient slide.
b. Staining results are acceptable when the control organisms stain well.
Note Cultivating the organism from suspected materials provides a definitive diagnosis, but it may take as many as 3 to 7 days. Every effort, therefore, must be made to microscopically examine wet smears and/or permanent stained smears so that appropriate therapy can be instituted without delay if findings are positive (Fig. 8.6) (37, 38).
Figure 8.6 Leishmania spp. from culture systems. (Upper left) Stained smear of culture fluid sediment [U3] showing promastigotes of Leishmania sp. (Upper right) Stained smear of culture fluid sediment showing promastigotes of Leishmania sp. (higher magnification). (Lower left) Stained Leishmania promastigote. (Lower right) Leishmania major promastigotes during infection of primary fibroblast culture. Cells are stained with antitubulin (green) and antiactin (red). (Courtesy of the Pasteur Institute, Molecular Parasitology and Signaling Image Bank). doi:10.1128/9781555819002.ch8.f6
The diagnosis of toxoplasmosis may be difficult, because the clinical symptoms mimic a number of various infectious and noninfectious diseases. Serologic tests that are often used for diagnosis may be insensitive in patients lacking normal immune responses. Sometimes even examination of histologic material does not reveal the organisms. With the increase in the number of laboratories using tissue culture techniques for viral pathogens, these techniques have been used for the isolation and identification of T. gondii. The following procedure has been recommended for biopsy specimens, brain, liver, spleen tissue, CSF, amniotic fluid, and buffy coat preparations, and they may be particularly helpful in making the diagnosis in immunosuppressed patients (39). The procedure for buffy coat cells is as follows.
1. Collect 10 ml of blood anticoagulated with preservative-free heparin.
2. Allow the blood to sediment via gravity.
3. Remove the buffy coat (by an aseptic technique), and separate the cells from the plasma by centrifugation at 800 × g for 10 min.
4. Wash the buffy coat cells three times with Eagle’s minimal essential medium (GIBCO).
5. Inoculate the washed buffy coat material onto complete human foreskin fibroblast (HFF) monolayers (in tubes and shell vials). One HFF tube and two HFF vials should be inoculated for each patient specimen.
6. Observe the cultures weekly for cytopathic effect.
7. The shell vial coverslips can be fixed and stained at 7 and 14 days postinoculation for an indirect fluorescent-antibody (IFA) assay and observed for the tachyzoites.
Note CSF, placental tissue, or other tissues can also be used to inoculate tissue culture monolayers. Uncentrifuged CSF (0.1 to 1 ml) can be used. If more than 1 ml is submitted, the specimen should be centrifuged for 10 min at 500 × g. Positive and negative controls must be tested with each set of patient specimen vials.
Three continuous cell lines (HeLa, LLC, and Vero) and three cell culture methods (culture in conventional flasks, culture in membrane-based flasks, and an automated culture system) were investigated (Fig. 8.7). Overall, HeLa was the cell line of choice. Continuous passage in flasks was successful, and HeLa-derived tachyzoites can be used for the dye test, if applicable in your laboratory setting (40). Another study indicates that THP1 cells serve as a good model of invasion for T. gondii (41).
Figure 8.7 Toxoplasma gondii RH tachyzoites replicated in Vero cell cultures. A rosette of many tachyzoites is seen at the left and several parasite pairs are at right (arrows). (Courtesy of I. Canedo-Solares, Abstract 42.009, 15th International Congress on Infectious Diseases, Bangkok, Thailand, 2012). doi:10.1128/9781555819002.ch8.f7
Techniques for the culture of Plasmodium falciparum were described in 1976 (42) and have been improved and modified since that time (43, 44). Life cycle stages of the five Plasmodium spp. that infect humans have been established in vitro. Of these five, P. falciparum and P. knowlesi are the only species for which all stages have been cultured in vitro. The life cycle includes the exoerythrocytic stage (within liver cells), the erythrocytic stage (within erythrocytes or precursor reticulocytes), and the sporogonic stage (within the vector). Culture media generally consist of a basic tissue culture medium to which serum and erythrocytes are added. Most of the culture methods have been directed toward the stage found in the erythrocyte. This stage has been cultivated in petri dishes or other containers in a candle jar to generate elevated CO2 levels or in a more controlled CO2 atmosphere. Later developments employed continuous-flow systems to reduce the labor-intensive requirement for replenishing the system with fresh media. The exoerythrocytic and sporogonic life cycle stages have also been cultivated in vitro. Although cultivation is of great help in understanding the biology of Plasmodium, it does not lend itself to use for routine diagnostic purposes (7).
The availability of the microaerophilous stationary phase (MASP) culture technique, in which the parasites proliferate in a settled layer of blood cells, has provided an opportunity to study Babesia, a formerly obscure disease agent regarded as within the purview of veterinary parasitology, in the laboratory. A number of Babesia spp. have been established in continuous culture using the MASP technique. It is possible to study the basic biology of the organism—as well as host-microbe interactions, immune factors triggered by the parasite, factors involved in innate resistance of young animals to infection, and antimicrobial susceptibility—to a degree not possible before the availability of cultures. These culture systems can produce quantities of parasite nucleic acid needed for defining phylogenetic relationships, developing diagnostic methods for parasite detection in asymptomatic individuals, and producing parasite antigens and attenuated strains of Babesia that could be used for immunization (5).
The in vitro cultivation of Cryptosporidium has improved significantly in recent years. These obligate intracellular parasites colonize the epithelium of the digestive and respiratory tracts, are often difficult to obtain in significant numbers, produce durable oocysts that defy conventional chemical disinfection methods, and are persistently infectious when stored at refrigerator temperatures (4 to 8°C). While continuous culture and oocyst production have not yet been achieved in vitro, routine methods for parasite preparation and cell culture infection and assays for parasite life cycle development have been established. Parasite yields tend to be limited, but in vitro growth is sufficient to support a variety of research studies, including assessing potential drug therapies, evaluating oocyst disinfection methods, and characterizing life cycle stage development and differentiation (1). Recent studies indicate that primary human intestinal epithelial cells (PECs) support Cryptosporidium better than other existing cell lines (45).
Although various microsporidia that infect humans can be identified from clinical specimens by serologic and/or molecular methods, none of these methods are commercially available. Unfortunately, in some cases microscopic examination of biopsy specimens does not yield conclusive results. It is also possible that microsporidial organisms may be present in very small numbers, which can be easily missed during routine histologic examinations. Some microsporidia such as Encephalitozoon, Enterocytozoon, Vittaforma, Trachipleistophora, and Anncaliia spp., even when they are present in small numbers, can become established in cell cultures, thus facilitating their identification at a later time. Therefore, attempts at culturing these organisms should be made whenever possible, since many clinical laboratory personnel are familiar with cell culture methodology (11). Anncaliia algerae has been successfully grown on the rabbit kidney cell line, RH-13 (46). Cell lines from goldfish skin (GFSK-S1) and brain (GFB3C-W1) and ZEB2J from zebrafish embryos, FHMT-W1 from fathead minnow testis, and Sf9 from ovaries of a fall armyworm moth were also found to be successful. All cultures were maintained at 27°C. Infection was judged to have taken place by the appearance of sporonts and/or spores in cells and occurred in all cell lines. These results suggest that cells of a wide range of vertebrates support A. algerae growth in vitro and fish cells can produce spores infectious to cells of mammals, fish, and insects (46). Expanded information on cell lines, media and supplements used for the culture of microsporidia causing human infections can be seen in Table 8.2 and Fig. 8.8 and 8.9.
Figure 8.8 (A) An E6 cell infected with E. intestinalis. Magnification, ×1,200. Note the well-defined multiple parasitophorous vacuoles (PV); N, host cell nucleus. (B) An HLF cell culture completely destroyed by E. hellem. Note the spores with everted polar tubules (at arrows), Magnification, ×600. All three species of Encephalitozoon destroy the cell culture, and often the cell cultures are completely covered by spores that either are intact or have discharged their polar tubules. Note It is very unusual to see the extruded polar tubules in routine clinical specimens such as urine or stool. (Courtesy of Govinda Visvesvara, from Visvesvara G, Clin Microbiol Rev 15:401–413, 2002). doi:10.1128/9781555819002.ch8.f8
Figure 8.9 (A) A host cell infected with Anncaliia (Brachiola) algerae. Note the arrangement of spores around the host (E6 cell) nucleus (N). A single spore is probably in the process of infecting an adjacent cell (arrowhead). Magnification, ×1,200. (B) A spore with an everted polar tubule. Magnification, ×1,200. (Courtesy of Govinda Visvesvara, from Visvesvara G, Clin Microbiol Rev 15:401–413, 2002). doi:10.1128/9781555819002.ch8.f9
Most routine clinical laboratories do not have the animal care facilities necessary to provide animal inoculation capabilities for the diagnosis of parasitic infections. Host specificity for many animal parasite species is a well-known fact which limits the types of animals available for these procedures. In certain suspect infections, animal inoculation may be requested and can be very helpful in making the diagnosis, although animal inoculation certainly does not take the place of other, more routine procedures.
The hamster is the laboratory animal of choice for the isolation of any form of Leishmania spp. A generalized infection results after intraperitoneal inoculation; spleen impression smears should be examined for the presence of organisms.
1. Aspirates or biopsy material obtained under sterile conditions from cutaneous ulcers, lymph nodes, spleen, liver, bone marrow, buffy coat cells, or CSF may be used for inoculation.
2. Material (0.25 to 1 ml) obtained from these sources should be inoculated under sterile conditions by the intraperitoneal route. Material from patients with mucocutaneous leishmaniasis should be inoculated by the intranasal route or into the feet.
3. Young (either sex) hamsters (2 to 4 months old) should be used for this procedure.
4. If the animal dies several days after inoculation, splenic aspirates should be examined for the presence of organisms. The material should be prepared as thick blood films and stained with Giemsa or other blood stains.
Note The infection develops slowly in hamsters; several months may be required to produce a detectable infection. For this reason, culture procedures are usually selected as more rapid means of parasite recovery. Intranasal lesions or those in the feet may develop very slowly; experimental animals should be kept for 9 to 12 months before a negative report is sent out.
Several laboratory animals can be used for the recovery of trypanosomes. Organisms are usually found in the blood of the animals within the first week after inoculation; however, if no organisms are found, the animals should be checked at the end of 2 and 4 weeks before results are reported as negative. Trypanosomes are infectious, so extreme care should be used when examining any blood or tissue suspected to contain these organisms.
1. Blood, lymph node aspirates, tissue, or CSF obtained under sterile conditions may be used for inoculation.
2. Material (up to 2 ml) should be inoculated intraperitoneally into guinea pigs or white rats for Trypanosoma gambiense and T. rhodesiense and into white mice for T. cruzi.
3. The number of organisms in the blood may vary; therefore, smears should be prepared frequently (every few days) over a 4-week period after inoculation.
Note Rats should be checked for the presence of T. lewisi (common parasite in rats) before inoculation to prevent a possible false-positive result.
All common laboratory animals can be infected by T. gondii. White rats and mice are generally used. White rats develop a chronic infection that can be a useful means of maintaining a strain of organisms. Mice that have been inoculated by the intraperitoneal route develop a fulminating infection that leads to death within a few days. Tremendous numbers of organisms can be recovered from the ascitic fluid. These specimens should be handled carefully to avoid an accidental laboratory infection.
General Procedure
1. The Toxoplasma organisms are found throughout the body after dissemination via the bloodstream. Any body tissue or fluid can be used for animal inoculation; the most common specimens are blood, lymph node fluid, and CSF.
2. The material used for inoculation (the amount may be very small [less than 0.25 ml]) should be obtained under sterile conditions and should be inoculated via the intraperitoneal route.
3. White mice of either sex and any age can be used. The animals should be checked daily for symptoms of illness.
4. After several days, the organisms can be recovered at necropsy from the peritoneal fluid of the mouse. The material should be prepared as thick blood films and stained with Giemsa or other blood stains.
5. Blind passage of peritoneal fluid to additional mice is recommended if the original mice appear to be negative.
6. If mice survive for up to 6 months, serum can be tested for the presence of antibody.
Procedure for Tissue
1. Grind the tissue in sterile 0.85% NaCl.
2. Prepare a 10% suspension (10% tissue, 90% NaCl).
3. Inoculate six mice via the intraperitoneal route. Each mouse should receive 1 ml.
4. The mice can be reinoculated the following day with an additional 1-ml dose.
5. Check the mice daily for symptoms.
6. If the animals are not sick at the end of 6 days, examine 1 drop of peritoneal fluid from each mouse directly and then stain it with Giemsa stain.
Xenodiagnosis is a technique that uses the arthropod host as an indicator of infection. Uninfected reduviid bugs are allowed to feed on the blood of a patient who is suspected of having Chagas’ disease (T. cruzi infection) (47) (Fig. 8.10). After 30 to 60 days, feces from the bugs are examined over a 3-month time frame for the presence of developmental stages of the parasite, which are found in the hindgut of the vector. This type of procedure is used primarily in South America for field work, and the appropriate bugs are raised in various laboratories specifically for this purpose. The term “xenodiagnosis” has also been applied to the diagnosis of trichinosis (Trichinella spiralis). Muscle tissue from a patient suspected of having the disease is fed to uninfected rats; the rats are then checked after the appropriate time for the presence of T. spiralis larvae, particularly in the diaphragm. This procedure is rarely requested and is not available in most clinical laboratories.
Figure 8.10 Illustration of the process of xenodiagnosis used for the diagnosis of Chagas’ disease. (Illustration by Sharon Belkin.)doi:10.1128/9781555819002.ch8.f10
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