be submitted one each day for three consecutive days; however, use of this collection time frame would not be sufficient to reject the specimens.
Although three stool specimens are recommended, laboratories have been more willing to accept two specimens, primarily because of cost savings and the assumption that if the patient is symptomatic, the presence of any organisms is likely to be confirmed by testing two specimens. However, it is important that clients understand the pros and cons of the two approaches. Both collection approaches are being used by diagnostic laboratories.
Posttherapy Collection
Patients who have received treatment for a protozoan infection should be checked 3 to 4 weeks after therapy, and those treated for Taenia infections should be checked 5 to 6 weeks after therapy; these recommendations have been used for many years. If the patient remains asymptomatic, the posttherapy specimens may not be collected, often as a cost containment measure. If the patient becomes symptomatic again, additional specimens can be submitted. If fecal immunoassays are ordered and parasites are present, the patient should be tested 7 to 10 days posttherapy. It usually takes approximately a week for antigen to be eliminated from the stool. It is important to remember that if the post-therapy specimens are collected too long after therapy, the presence of parasites or parasite antigen may represent a reinfection.
Specimen Type, Stability, and Need for Preservation
Fresh specimens are required for the recovery of motile trophozoites (amebae, flagellates, or ciliates). The protozoan trophozoite stage is found in patients with diarrhea; the gastrointestinal tract contents are moving through the system too rapidly for cyst formation to occur. Once the stool specimen is passed from the body, trophozoites do not encyst but may disintegrate if not examined or preserved within a short time after passage. However, most helminth eggs and larvae, coccidian oocysts, and microsporidian spores can survive for extended periods. Since it is impossible to know which organisms might be present, it is recommended that the most conservative time limits be used for parasite preservation and recovery. Liquid specimens should be examined within 30 min of passage, not 30 min from the time they reach the laboratory. If this general time recommendation of 30 min is not possible, the specimen should be placed in one of the available fixatives. Soft (semiformed) specimens may contain a mixture of protozoan trophozoites and cysts and should be examined within 1 h of passage; again, if this time frame is not possible, preservatives should be used. Immediate examination of formed specimens is not as critical; in fact, if the specimen is examined any time within 24 h after passage, the protozoan cysts should still be intact (Table 3.3).
Currently, fresh or frozen fecal specimens are required for the following fecal immunoassays (either as a single-organism test or when combined with other organisms such as G. lamblia or Cryptosporidium spp.): E. histolytica/E. dispar group and E. histolytica.
Summary: Collection of Fresh Stool Specimens
1. Occupational Safety and Health Act regulations (Standard Precautions) should be used for handling all specimens.
2. Interfering substances (e.g., barium, mineral oil, or antibiotics) should be avoided when stool specimens are collected.
3. Contamination with urine or water should be avoided.
4. Recommendation for collection: two (minimum) or three specimens collected, one every other day or within a 10-day time frame; see Table 3.1 for options and pros/cons.
5. Liquid stool should be examined or preserved within 30 min of passage (trophozoites). Soft stool should be examined or preserved within 1 h of passage (trophozoites and cysts*). Formed stool should be examined or preserved within 24 h of passage.
6. Fresh or frozen fecal specimens are required for the following fecal immunoassays (either as a single-organism test or combined with other organisms such as G. lamblia or Cryptosporidium spp.): E. histolytica/E. dispar group and E. histolytica. Fresh, frozen, or preserved specimens can be used for G. lamblia and Cryptosporidium spp.; specimens submitted in Cary-Blair transport medium are also acceptable. Note: Due to the freeze-thaw cycle, frozen specimens cannot be used for the FA procedures for G. lamblia and/or Cryptosporidium spp. (destruction of the actual cysts and/or oocysts that are visual proof of a positive specimen).
* Dientamoeba fragilis trophozoites can be found in formed stool specimens.
Preservation of Stool Specimens
Overview of Preservatives
If there are likely to be delays from the time of specimen passage until examination in the laboratory, the use of preservatives is recommended. To preserve protozoan morphology and to prevent the continued development of some helminth eggs and larvae, the stool specimens can be placed in preservative either immediately after passage (by the patient using a collection kit) or once the specimen is received by the laboratory. Several fixatives are available: formalin, sodium acetate-acetic acid-formalin (SAF), Schaudinn’s fluid, polyvinyl alcohol (PVA), and single-vial systems (Table 3.3). Regardless of the fixative selected, use of the appropriate ratio of fixative to stool (3 parts fixative to 1 part stool) and adequate mixing of the specimen and preservative are mandatory. Although many products are commercially available, the most commonly used preservatives are discussed below. They are all available from various scientific supply houses.
When selecting an appropriate fixative, keep in mind that a permanent stained smear is required for a complete examination for parasites. If the physician orders a fecal immunoassay such as FA, EIA, or the rapid-flow method, you will need to confirm that the fixative you are using is compatible with the immunoassay you have selected. It is also important to remember that disposal regulations for compounds containing mercury are becoming more strict; each laboratory will have to check applicable state and federal regulations to help determine fixative options.
Formalin
Formalin is an all-purpose fixative that is appropriate for helminth eggs and larvae and for protozoan cysts. Two concentrations are commonly used: 5%, which is recommended for preservation of protozoan cysts, and 10%, which is recommended for helminth eggs and larvae. Although 5% is often recommended for all-purpose use, most commercial manufacturers provide 10%, which is more likely to kill all helminth eggs. To help maintain organism morphology, the formalin can be buffered with sodium phosphate buffers, i.e., neutral formalin. Selection of specific formalin formulations is at the user’s discretion. Aqueous formalin permits examination of the specimen as a wet mount only, a technique much less accurate than a stained smear for the identification of intestinal protozoa. The most common preparation is 10% formalin, prepared as follows:
Formaldehyde (USP) . . . . . . . . . . . . . 100 ml (or 50 ml for 5%)
Saline solution, 0.85% NaCl . . . . . 900 ml (or 950 ml for 5%)
Dilute 100 ml of formaldehyde with 900 ml of 0.85% NaCl solution. (Distilled water may be used instead of saline solution.)
Note: Formaldehyde is normally purchased as a 37 to 40% HCHO solution; however, for dilution it should be considered to be 100%.
If you want to use buffered formalin, the recommended approach is to mix thoroughly 6.10 g of Na2HPO4 and 0.15 g of NaH2PO4 and store the dry mixture in a tightly closed bottle. For 1 liter of either 10 or 5% formalin, 0.8 g of the buffer salt mixture should be added.
Protozoan cysts (not trophozoites),