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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.