through the use of a liquid (zinc sulfate) with a high specific gravity. The parasitic elements are recovered in the surface film, and the debris and some heavy parasitic elements remain in the bottom of the tube. This technique yields a cleaner preparation than does the sedimentation procedure; however, some helminth eggs (operculated and/or very dense eggs, such as unfertilized Ascaris eggs) do not concentrate well in the flotation method; a sedimentation technique is recommended to detect these infections.
When the zinc sulfate solution is prepared, the specific gravity should be 1.18 for fresh stool specimens; it must be checked with a hydrometer. This procedure may be used on formalin-preserved specimens if the specific gravity of the zinc sulfate is increased to 1.20; however, this usually causes more distortion in the organisms present and is not recommended for routine clinical use. To ensure detection of all possible organisms, both the surface film and the sediment must be examined. For most laboratories, this is not a practical approach.
The specimen must be fresh or formalinized stool (5 or 10% buffered or nonbuffered formalin, SAF, or other non-PVA single-vial preservatives). PVA-preserved specimens can also be used; however, this approach is not commonly used or recommended.
Zinc Sulfate (33% Aqueous Solution)
1. Using a magnetic stirrer, dissolve the zinc sulfate in distilled water in an appropriate flask or beaker.
2. Adjust the specific gravity to 1.20 by the addition of more zinc sulfate or distilled water. Use a specific gravity of 1.18 with fresh stool (nonformalinized).
3. Store in a glass-stoppered bottle with an expiration date of 24 months.
Quality Control for Flotation Concentration
1. Check the reagents each time they are used. The formalin, saline, and zinc sulfate should appear clear, without any visible contamination.
2. The microscope should be calibrated (within the last 12 months), and the objectives and oculars used for the calibration procedure should be used for all measurements on the microscope. The calibration factors for all objectives should be posted on the microscope or close by for easy access. As mentioned above, some workers feel that recalibration of the microscope is not necessary each year; however, this would depend on the use and maintenance of that particular piece of equipment.
3. Known positive specimens should be concentrated and organism recovery should be verified at least quarterly, particularly after the centrifuge has been recalibrated. Human WBCs (buffy coat cells) mixed with negative stool can be used as a QC specimen. These human cells, when mixed with negative stool, mimic human parasites. The human cells concentrate just like human parasites, such as protozoa and helminth eggs and larvae.
4. All QC results should be appropriately recorded; the laboratory should also have an action plan for “out-of-control” results.
Procedure for Flotation Concentration
1. Transfer 1/2 teaspoon (about 4 g) of fresh stool into 10 ml of 5 or 10% formalin in a shell vial, unwaxed paper cup, or round-bottom tube (the container may be modified to suit individual laboratory preferences). Mix the stool and formalin thoroughly. Let the mixture stand for a minimum of 30 min for fixation. If the specimen is already in 5 or 10% formalin (or SAF or Universal Fixative), stir the stool-formalin (or SAF) mixture.
2. Depending on the size and density of the specimen, strain a sufficient quantity through wet gauze (no more than two layers of gauze or one layer if the new “pressed” gauze [e.g., Johnson & Johnson nonsterile three-ply gauze, product 7636] is used) into a conical 15-ml centrifuge tube to give the desired amount of sediment (0.5 to 1 ml) in step 3 below. Usually, 8 ml of the stool-formalin mixture prepared in step 1 is sufficient. If the specimen is received in vials of preservative (5 or 10% formalin, SAF, or other single-vial preservatives), approximately 3 to 4 ml of the mixture is sufficient unless the specimen has very little stool in the vial. If the specimen contains a lot of mucus, do not strain through gauze but immediately fix in 5 or 10% formalin or other fixative (does not contain PVA) for 30 min and centrifuge for 10 min at 500 × g. Proceed directly to step 5.
3. Add 0.85% NaCl almost to the top of the tube, and centrifuge for 10 min at 500 × g. Approximately 0.5 to 1 ml of sediment should be obtained. Too much or too little sediment results in an ineffective concentration examination. See the section on Commercial Fecal Concentration Devices later in this chapter.
4. Decant and discard the supernatant fluid, resuspend the sediment in 0.85% NaCl almost to the top of the tube, and centrifuge for 10 min at 500 × g. This second wash may be eliminated if the supernatant fluid after the first wash is light tan or clear. Some prefer to limit the washing to one step (regardless of the color and clarity of the supernatant fluid) to eliminate additional manipulation of the specimen prior to centrifugation. The more the specimen is manipulated and/or rinsed, the more likely it is for parasitic elements to be lost.
5. Decant and discard the supernatant fluid, and resuspend the sediment on the bottom of the tube in 1 to 2 ml of zinc sulfate. Fill the tube within 2 to 3 mm of the rim with additional zinc sulfate.
6. Centrifuge for 2 min at 500 × g. Allow the centrifuge to come to a stop without interference or vibration. Two layers should result: a small amount of sediment in the bottom of the tube, and a layer of zinc sulfate (Fig. 3.5). The protozoan cysts and some helminth eggs are found in the surface film; some operculated and/or heavy eggs are found in the sediment.
7. Without removing the tube from the centrifuge, remove 1 or 2 drops of the surface film with a Pasteur pipette or a freshly flamed (and allowed to cool) wire loop and place them on a slide. Do not use the loop as a “dipper”; simply touch the surface (bend the loop portion of the wire 90° so that the loop is parallel with the surface of the fluid). Make sure the pipette tip or wire loop is not below the surface film (Fig. 3.8).
8. Add a coverslip (22 by 22 mm, no. 1) to the preparation. Iodine may be added to the preparation (optional).
9. Systematically scan with the 10× objective. The entire coverslip area should be examined under low power (total magnification, ×100).
10. If something suspicious is seen, the 40× objective can be used for more detailed study. At least one-third to one-half of the coverslip should be examined with high dry power (total magnification, ×400), even if nothing suspicious has been seen. As in the direct wet smear, iodine can be added to enhance morphological detail, and the coverslip can be gently tapped to observe objects moving and turning over.
Figure 3.8 Method used to remove the surface film in the zinc sulfate flotation concentration procedure. (A) A wire loop is gently placed on (not under) the surface film. (B) The loop is then placed on a glass slide. (Illustration by Nobuko Kitamura.) doi:10.1128/9781555819002.ch3.f8
Results and Patient Reports from Flotation Concentration
Protozoan trophozoites and/or cysts and some helminth eggs and larvae may be seen and identified. Heavy helminth eggs and operculated eggs do not float in zinc sulfate; they are seen in the sediment within the tube. The high specific gravity of the zinc sulfate causes the opercula to pop open; the eggs fill with fluid and sink to the bottom. Protozoan trophozoites are less likely to be seen. In a heavy infection with Cryptosporidium spp. or C. cayetanensis, oocysts may be seen in the concentrate sediment; oocysts of C. belli can also be seen. Spores of the
