(or ethyl acetate). The sediment should be well mixed, and a drop of sediment should be examined using the 10× low-power objective and the 40× high dry power objective. (B) Zinc sulfate (the surface film should be within 2 to 3 mm of the tube rim). Material from both the surface film and the sediment must be examined before the specimen is indicated as negative. The amount of sediment should not be excessive in either the sedimentation or flotation procedure. Heavy or operculated helminth eggs do not float. (Illustration by Sharon Belkin.) doi:10.1128/9781555819002.ch3.f5
Sedimentation methods (by centrifugation) lead to the recovery of all protozoa, oocysts, eggs, and larvae present; however, the concentration sediment that will be examined contains more debris. Although some workers recommend using both flotation and sedimentation procedures for every stool specimen submitted for examination, this approach is impractical for most laboratories. If one technique is selected for routine use, the sedimentation procedure is recommended as being the easiest to perform and the least subject to technical error (Fig. 3.6).
Figure 3.6 Sedimentation concentration. (Left) Unfertilized Ascaris lumbricoides egg. (Right) Hymenolepis diminuta egg. doi:10.1128/9781555819002.ch3.f6
A flotation procedure permits the separation of protozoan cysts, coccidian oocysts, and certain helminth eggs and larvae through the use of a liquid with a high specific gravity. The parasitic elements are recovered in the surface film, and the debris remains in the bottom of the tube. This technique yields a cleaner preparation than does the sedimentation procedure; however, some helminth eggs (operculated eggs and/or very dense eggs such as unfertilized Ascaris eggs) do not concentrate well in the flotation method (Fig. 3.7). The specific gravity may be increased, although this may produce more distortion in the eggs and protozoa. Laboratories that use only flotation procedures may fail to recover all of the parasites present; to ensure detection of all organisms in the sample, both the surface film and the sediment should be carefully examined. Directions for any flotation technique must be followed exactly to produce reliable results.
Figure 3.7 Flotation concentration. (Upper) Fasciolopsis buski egg (left), Diphyllobothrium latum egg (right). Note that both of these eggs in the top row are operculated and WILL NOT float in the zinc sulfate flotation concentration method; the opercula pop open, and the eggs fill with fluid and sink to the bottom of the tube. (Lower) Hookworm egg (left), Trichuris trichiura egg (right). These eggs concentrate using the flotation method and can be seen in the surface film. However, remember that both the surface film and the sediment must be examined by this method before reporting the final ova and parasite examination results. doi:10.1128/9781555819002.ch3.f7
Formalin-Ethyl Acetate Sedimentation Concentration
By centrifugation, the formalin-ethyl acetate sedimentation concentration procedure leads to the recovery of all protozoa, eggs, larvae, coccidia, and microsporidia present; however, the preparation contains more debris than is found in the flotation procedure. Ethyl acetate is used as an extractor of debris and fat from the feces and leaves the parasites at the bottom of the suspension in the sediment. The formalin-ethyl acetate sedimentation concentration procedure is recommended as being the easiest to perform, allowing recovery of the broadest range of organisms, and being the least subject to technical error.
The specimen must be fresh or formalinized stool (5 or 10% buffered or nonbuffered formalin or SAF or the Universal Fixative). Many of the single-vial preservative systems are also acceptable; however, the formulas are proprietary (e.g., UNIFIX; Medical Chemical Corp., Torrance, CA). Specimens preserved in fixatives containing polyvinyl alcohol (PVA) can also be used. However, PVA preservative formulations are rarely used for concentration methods in most laboratories but are recommended for the preparation of permanent stained smears.
5 or 10% Formalin
Formaldehyde (USP)
..................100 ml (for 10%) or
....................50 ml (for 5%)
Saline solution 900 ml (for 10%) or
0.85% NaCl 950 ml (for 5%)
Note Formaldehyde is normally purchased as a 37 to 40% HCHO solution; however, for dilution, it should be considered to be 100%.
Dilute 100 ml of formaldehyde with 900 ml of 0.85% NaCl solution. (Distilled water may be used instead of saline solution.)
Quality Control for Sedimentation Concentration
1. Check the liquid reagents each time they are used; the formalin and saline 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 in place on the microscope when objects are measured. The calibration factors for all objectives should be posted on the microscope or close by for easy access. Some researchers feel that a microscope does not require calibration every 12 months; however, if the microscope is moved periodically, can be easily bumped, or does not receive adequate maintenance, it should be rechecked yearly for calibration accuracy.
3. Known positive specimens should be concentrated and organism recovery should be verified at least quarterly and 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, can mimic protozoan 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 Sedimentation 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, and 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 other non-PVA single-vial preservatives), stir the stool-preservative mixture.
2. Depending on the amount and viscosity 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) for step 3 below. Usually, 8 ml of the stool-formalin mixture prepared in step 1 is sufficient. If the specimen is received in a vial of preservative (5 or 10% formalin, SAF, or other single-vial preservatives), approximately 3 to 4 ml of the preservative-stool mixture is sufficient for testing. If the vial contains very little specimen, then the entire amount may be used in the procedure. If the specimen contains a lot of mucus, do not strain through gauze but immediately fix in 5 or 10% formalin for 30 min and centrifuge for 10 min at 500 × g. Proceed directly to step 10.
3. Add 0.85% NaCl or 5 or 10% formalin (some workers prefer to use formalin for all rinses) almost to the top of the tube, and centrifuge for 10 min at 500 × g. The amount of sediment obtained should be approximately 0.5 to 1 ml.
4. Decant and discard the supernatant fluid, and resuspend the sediment in saline or formalin; add saline or formalin almost to the top of the tube, and centrifuge again for 10 min at 500 × g. This second wash may be
