Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 11
ОглавлениеThis chapter discusses various collection methods that are available for specimens suspected of containing parasites or parasitic elements. When a laboratory selects its collection methods, the decision should be based on a thorough understanding of the value and limitations of each. One of the most important aspects of specimen collection is that the final laboratory results based on parasite recovery and identification will depend on the initial fixation of the organisms (1–3). Unless the appropriate specimens are properly collected and processed, these infections may not be detected (1). Considering the current era of cost containment and review of clinical relevance of laboratory information generated, specimen rejection criteria have become more important within the context of all diagnostic microbiology procedures. Diagnostic laboratory results based on improperly collected specimens may require excessive expenditures of time and supplies and may also mislead the physician. As a part of any overall total quality management or continuous quality improvement program for the laboratory, the generation of test results must begin with stringent criteria for specimen acceptance or rejection.
Clinically relevant diagnostic parasitology testing also depends on receiving appropriate test orders from the physician. Depending on the patient’s clinical condition and travel history, very specific diagnostic tests may be recommended. It is extremely important that physician clients are aware of the test order options available within the laboratory test menu and the pros and cons of each test when considered within the context of the patient’s history and symptoms. Without the proper test orders, diagnostic test results may be misleading or actually incorrect. Appropriate and complete communication regarding test orders between the laboratory and physicians is mandatory for high-quality patient care.
All fresh specimens should be handled carefully, since each specimen represents a potential source of infectious material (bacteria, viruses, fungi, and parasites). Safety precautions should include awareness of the following: proper labeling of fixatives; specific areas designated for specimen handling (biological safety cabinets may be necessary under certain circumstances); proper containers for centrifugation; acceptable discard policies; appropriate policies for no eating, drinking, or smoking, etc., within the working areas; and, if applicable, correct techniques for organism culture and/or animal inoculation.
Since diagnostic parasitology work is most often performed within the microbiology division of a clinical laboratory, all general guidelines for safety would also apply. Any special precautions which apply to a particular technique are discussed in the following chapters. In general, standard precautions as outlined by the Occupational Safety and Health Act must be followed when applicable, particularly when one is handling blood and other body fluids (4).
Procedures for the recovery of intestinal parasites should always be performed before barium is used for radiological examination. Stool specimens containing barium are unacceptable for examination, and intestinal protozoa may be undetectable for 5 to 10 days after barium is given to the patient. There are also certain substances and medications that interfere with the detection of intestinal protozoa: mineral oil, bismuth, antibiotics, antimalarial agents, and nonabsorbable antidiarrheal preparations. After administration of any of these compounds, parasitic organisms may not be recovered for a week to several weeks. The two most commonly used substances are barium and antibiotics, such as tetracycline, which modify the gastrointestinal tract flora. Specimen collection should be delayed for 5 to 10 days or at least 2 weeks after barium or antibiotics, respectively, are administered (1, 2, 5–24). The use of antibacterial therapy that affects the normal gastrointestinal tract flora will diminish the numbers of protozoa, since they feed on intestinal bacteria.
Fecal specimens should be collected in clean, wide-mouth containers; often a 0.5-pint (ca. 0.24-liter) waxed cardboard or plastic container with a tight-fitting lid is selected for this purpose. The fit of the lid is particularly important, both from the standpoint of accidental spillage and in order to maintain moisture within the specimen. The specimens should not be contaminated with water or urine, because water may contain free-living organisms that can be mistaken for human parasites and urine may destroy motile organisms. For safety reasons, stool specimen containers should be placed in plastic bags when transported to the laboratory for testing. Fresh specimens can also be submitted in collection vials (Fig. 2.1). All fresh specimens should be carefully handled, since they are potential sources of infectious organisms, including bacteria, viruses, and parasites. Every specimen should be identified with the following minimal information: patient’s name and identification number, physician’s name, and the date and time the specimen was collected (if the laboratory is computerized, the date and time may reflect arrival in the laboratory, not the actual collection time). The specimen must also be accompanied by a request form indicating which laboratory procedures are to be performed. It is also be very helpful to have information concerning the presumptive diagnosis or relevant travel history; however, this information is rarely available, and under certain circumstances, the physician will have to be contacted for additional patient history (Example: Fever of unknown origin [FUO]—possible malaria).
Figure 2.1 Stool collection vial; “clean vial” contains no fixatives. doi:10.1128/9781555819002.ch2.f2.1
Number of Specimens To Be Collected (Standard Recommendation)
It is recommended that a normal examination for stool parasites before therapy include three specimens, consisting of two specimens collected from normal movements and one collected after the use of a cathartic such as magnesium sulfate or Fleet’s Phospho-Soda. A cathartic with an oil base should not be used, and a stool softener (taken either orally or as a suppository) is usually inadequate for obtaining a purged specimen. The purpose of the laxative is to stimulate some “flushing” action within the gastrointestinal tract, possibly allowing one to obtain more organisms for recovery and identification. Obviously, if the patient already has diarrhea or dysentery, the use of any laxatives would be contraindicated. Since the majority of patients are symptomatic prior to submission of stool specimens for examination, the need for a laxative is relatively uncommon.
When a patient is suspected of having intestinal amebiasis, six specimens may be recommended. The examination of six specimens ensures detection of approximately 90% of amebic infections (5) (Fig. 2.2). However, because of cost-containment measures, the examination of six specimens is rarely requested.
Figure 2.2 Increased detection of Entamoeba histolytica by using various diagnostic techniques and serial stool specimens. (Adapted by Markell EK, Voge M, John DT, 1992, Medical Parasitology, 7th ed., The W. B. Saunders Co., Philadelphia, PA, from references 17 and 31, with permission.) doi:10.1128/9781555819002.ch2.f2.2
Three specimens are also recommended for posttherapy examinations, and they should be collected as outlined above. However, a patient who has 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. In some cases, the physician will assume a cure for tapeworm infection unless proglottids reappear in the stool; therefore, no posttherapy specimens are submitted for examination.
Number of Specimens To Be Collected (Pros and Cons of Various Options)
During the past few years, a number of issues have surfaced regarding the collection, processing, and testing of stool specimens for diagnostic parasitology. Many of the new suggestions and options have arisen as a result of continued cost-containment measures, limited reimbursement, and the elimination of mercury-based compounds for stool preservatives. The number of nonmercury preservative choices, collection systems, concentration devices, and immunoassays has increased dramatically. Many laboratories continue to review the options, and some may be having difficulties in selecting the proper approach (7–22).
It is important to realize that there are many acceptable options and that many laboratories will select different approaches. These differences should not be categorized as “right or wrong” or “acceptable or unacceptable”—they are merely different! To assume that there is only one correct approach for the examination of stool specimens is neither appropriate nor realistic. There are many parameters to consider before selecting the approach for your own laboratory. In no particular order, some of the considerations include client base, physician ordering patterns, number of specimens received per month, cost, presence or absence of appropriate equipment, current and possible methodologies (including immunoassays such as enzyme immunoassay [EIA], fluorescent-antibody assay [FA], and rapid membrane flow cartridge devices [rapids]), availability of expert microscopists, collection options, selection of preservative-stain combinations, reimbursement issues, client education, area of the world where laboratory is located, and emphasis on the most common infections (helminth or protozoa or both).
When considering available options and laboratory test menus, it is important to make sure that the pros and cons of the approaches selected are thoroughly understood and that diagnostic tests, potential results, and reporting formats are carefully explained to all clients. As an example, if the results of a stool examination are based on a concentration sediment examination only, this information must be conveyed to the physician. Many of the intestinal protozoa are missed by this diagnostic test approach, and it is important for the physician to recognize the limitations of such testing. Most physicians receive very little, if any, exposure to medical parasitology in medical school, and many newer physicians trained as generalists or family practitioners also have limited parasitology training or experience.
In most cases, it is probably realistic to assume that patients are symptomatic if they are submitting stool specimens for diagnostic parasitology testing. In an excellent article by Hiatt et al., the premise tested was that a single stool sample from a symptomatic patient would be sufficient to diagnose infections with intestinal protozoa (23). However, with additional stool examinations for symptomatic patients, the yield of intestinal protozoa increased dramatically (Entamoeba histolytica, 22.7% increase; Giardia lamblia, 11.3% increase; and Dientamoeba fragilis, 31.1% increase). This publication again demonstrates the problems with performing only a single stool examination (using the ova and parasite examination). If the patient becomes asymptomatic after the first stool examination, it may be acceptable to discontinue the series of stool examinations (this should be a clinician decision).
Available options are compared to the “gold standard,” which includes a series of three stool examinations (direct, concentrate, and permanent stained smear for a fresh specimen; concentrate and permanent stained smear for a preserved specimen). The single-specimen pros and cons are discussed in the previous paragraph (symptomatic versus asymptomatic patients). A suggestion has been made to pool three specimens and to perform a single concentration and a single permanent stained smear. Depending on the number of positives and negatives, this may be a viable option, but some organisms may be missed. Another testing option for three pooled specimens might be to perform a single concentration and three separate permanent stained smears. This approach would probably increase the yield of intestinal protozoa over the previous option. Another suggestion involves placing a sample of each of three stools into a single vial of preservative. This collection approach would require only a single vial, but it is very likely that the vial would be overfilled and that mixing and the ratio of fixative to stool would be inaccurate.
One could also test patient specimens by using immunoassay reagents or kits for Giardia lamblia (G. duodenalis, G. intestinalis), the Entamoeba histolytica/E. dispar group, Entamoeba histolytica, or Cryptosporidium spp. However, patient history information is rarely received with the clinical specimen; fecal testing based on patient risk group or situation is impossible without sufficient information. Testing of patient specimens by immunoassay procedures should be performed on request; however, client education is critical for successful implementation of this approach. Collection and testing options and the pros and cons of each are listed in Tables 2.1 and 2.2.
The Clinical and Laboratory Standards Institute (formerly the National Committee for Clinical Laboratory Standards) document Procedures for the Recovery and Identification of Parasites from the Intestinal Tract (Approved Guideline M28-2A) was updated in 2005. Various stool collection, processing, and testing options are also included in that publication (1, 25).
In summary, laboratories performing diagnostic parasitology testing must decide on the appropriate test methods that are relevant for their own operations based on a number of variables mentioned above. It is unrealistic to assume or state that one approach is applicable for every laboratory; however, it is important to thoroughly understand the options within your test menu and to convey this information to your clients once your approach has been selected for implementation. Prior discussion with clients, written educational memos, meetings, and examples of revised report formats are highly recommended prior to implementation.
A series of three specimens as indicated above should be submitted on separate days; if possible, the specimens should be submitted every other day; otherwise, the series of three specimens should be submitted within no more than 10 days. If a series of six specimens is requested, the specimens should also be collected on separate days or within no more than 14 days. Many organisms, particularly the intestinal protozoa, do not appear in the stool in consistent numbers on a daily basis, and the series of three specimens is considered a minimum for an adequate examination. It is inappropriate for multiple specimens to be submitted from the same patient on the same day. One possible exception would be stool collections from a patient who has severe, watery diarrhea such that any organisms present might be missed because of the tremendous dilution factor related to fluid loss. Even under these circumstances, acceptance of more than one specimen per patient per day should not be routine but should be done only after consultation with the physician. It is also not recommended for the three specimens to be submitted one each day for three consecutive days; however, use of this collection time frame would not be sufficient cause to reject the specimens. Adequate spacing between specimens helps to provide parasite recovery within the recommended time frames.
Although the recommended number of stool specimens is three, laboratories have been more willing to accept two specimens, primarily because of cost savings and the assumption that if the patient is symptomatic, confirmation of any organisms present is just as likely to be possible from two specimens as from three specimens. However, it is important that clients understand the pros and cons of two compared with three stools. Both collection approaches are being used by diagnostic laboratories.
Specimen Type, Specimen Stability, and Need for Preservation
Fresh specimens are mandatory for the recovery of motile trophozoites (amebae, flagellates, or ciliates). The protozoan trophozoite stage is normally found in cases of 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. The time limit recommendations listed below are most relevant for the intestinal protozoa; most helminth eggs and larvae, coccidian oocysts, and microsporidian spores survive for extended periods. However, no one can predict which organisms will be present in the stool specimens; therefore, it is important to use the most conservative time frames for parasite 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 at any time within 24 h after passage, the protozoan cysts should still be intact (5, 16).
In review, remember that trophozoites only are usually found in liquid specimens, both protozoan trophozoites and cysts can be recovered in soft specimens, and generally cysts only are recovered in formed specimens. The time limits mentioned above are merely guidelines; however, if fresh specimens remain unpreserved for longer times before examination, many if not all organisms may disintegrate or become distorted. Fecal specimens should never be incubated or frozen prior to examination using routine microscopy. When the acceptance criteria for specimen collection are not met, the laboratory should reject the specimen and request additional specimens.
Because there is often a time lag from the time of specimen passage until receipt in the laboratory, many clinicians, clinics, and inpatient wards use a specimen collection system that includes stool preservatives (Table 2.3). A number of commercial systems are available with many preservative choices; the use of such systems has become routine for many institutions, and some request a custom collection kit that may contain several types of preservatives for stool specimens, depending on the tests normally ordered by the clinicians that they service. Specific information concerning collection kit components is provided in Appendix 1.
KEY POINTS—STOOL SPECIMEN COLLECTION
1. Occupational Safety and Health Act regulations (including standard precautions) should be used for handling all specimens.
2. Interfering substances (oil-based laxatives, barium, antibiotics) should be avoided when stool specimens are collected.
3. Contamination with urine or water should be avoided.
4. Recommendation for collection: three specimens collected, one every other day or within a 10-day time frame.
Note: There are some exceptions; see Table 2.1.
5. Liquid stool: examine or preserve within 30 min of passage (trophozoites). Soft stool: examine or preserve within 1 h of passage (trophozoites and cysts). Formed stool: examine or preserve within 24 h of passage. Note that Dientamoeba fragilis trophozoites can be found in formed stool specimens as well as liquid specimens.
There are a number of reasons why a lag time may occur from the time of specimen passage until examination in the laboratory (e.g., the workload in the laboratory or the transit distance or time for the specimen to reach the facility). To preserve protozoan morphology and to prevent the continued development of some helminth eggs and larvae, the stool specimens can be placed in preservative immediately after passage (by the patient using a collection kit) or once the specimen is received by the laboratory. There are several fixatives available; the more common ones, including formalin, Merthiolate (thimerosal)-iodine-formalin (MIF), sodium acetate-acetic acid-formalin (SAF), Schaudinn’s fluid (with or without polyvinyl alcohol [PVA]), the single-vial systems, and the Universal Fixative, are discussed in detail (Table 2.4). Regardless of the fixative selected, adequate mixing of the specimen and preservative is mandatory. A flow diagram for preservation and processing is shown in Fig. 2.3.
Figure 2.3 Flow diagram for preservation and processing of stool specimens. As mentioned in the text, the examination of fecal specimens using the ova and parasite examination is not considered complete unless a concentration and a permanent stained smear are examined for every specimen submitted to the laboratory. For a fresh specimen, a direct wet mount should be performed if the specimen is very soft to liquid; the complete ova and parasite examination would include the direct wet mount, the concentration, and the permanent stained smear. If the specimen is submitted in preservative, the direct wet mount should be eliminated (no motility is possible); the complete ova and parasite examination would include the concentration and the permanent stained smear (1, 2). doi:10.1128/9781555819002.ch2.f2.3
Note When selecting an appropriate fixative, keep in mind that a permanent stained smear is mandatory for a complete examination for parasites (1, 2, 26). It is also important to remember that disposal regulations for compounds containing mercury are becoming more restrictive; each laboratory will have to check applicable state and federal regulations to help determine fixative options.
Note It is important to remember that when using fecal immunoassays for the Entamoeba histolytica/E. dispar group or Entamoeba histolytica, fresh or frozen stool is required (in some cases Cary-Blair may be acceptable, but check the package insert prior to use). To date, stool submitted in routine fecal fixatives is not acceptable for these particular fecal immunoassays.
Formalin
Formalin has been used for many years as an all-purpose fixative that is appropriate for helminth eggs and larvae and for protozoan cysts, oocysts, and spores. 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 (Fig. 2.4). 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 will permit the examination of the specimen as a wet mount only, a technique much less accurate than a permanent stained smear for the identification of intestinal protozoa. However, the fecal immunoassays for Giardia lamblia (G. duodenalis, G. intestinalis) and Cryptosporidium spp. can be performed from the aqueous formalin vial. Fecal mmunoassays for the Entamoeba histolytica/E. dispar group and Entamoeba histolytica are limited to fresh or frozen fecal specimens (some kits may work with stool submitted in Cary-Blair; check the package insert). After centrifugation, special stains for the coccidia (modified acid-fast stains) and the microsporidia (modified trichrome stains) can be performed from the concentrate sediment obtained from formalin-preserved stool material. The most common formalin preparation is 10% formalin, prepared as follows:
Figure 2.4 When the stool specimen is added to the vial, the final ratio of stool to preservative is about 1:3. doi:10.1128/9781555819002.ch2.f2.4
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 (1, 3) 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. Prepare 1 liter of either 10 or 5% formalin, and add 0.8 g of the buffer salt mixture.
Protozoan cysts (not trophozoites), coccidian oocysts, microsporidian spores, helminth eggs, and larvae are well preserved for long periods in 10% aqueous formalin. Hot (60°C) formalin can be used for specimens containing helminth eggs, since in room-temperature formalin, some thick-shelled eggs (e.g., Ascaris lumbricoides) continue to develop, become infective, and remain viable for long periods. Several grams of fecal material should be thoroughly mixed in 5 or 10% formalin.
To collect large numbers of cysts, eggs, or larvae relatively free from other debris, the whole stool specimen is mixed in water and then strained through several layers of gauze. The suspension is allowed to sediment in a cone-shaped glass or flask for 1 h or more, and the supernatant fluid is discarded. The specimen may be washed several times in this manner before the sediment is finally fixed in hot 10% formalin, as mentioned above. When working with watery diarrhea specimens from patients with suspected cases of coccidiosis or microsporidiosis, the specimen should not be strained through gauze (oocysts and small bits of mucus may cling to the gauze); centrifugation (500 × g for 10 min) is necessary to sediment the oocysts and/or spores.
MIF
MIF (5) is a good stain preservative for most kinds and stages of parasites found in feces; it is especially useful for field surveys. It is used with all common types of stools and aspirates; protozoa, eggs, and larvae can be diagnosed without further staining in temporary wet mounts, either made immediately after fixation or prepared several weeks later. Although some laboratories maintain that a permanent stained smear can be prepared from specimens preserved in MIF, most laboratories using such a fixative examine the material only as a wet preparation (direct smear and/or concentration sediment). For a good discussion of this technique, see reference 27.
The MIF preservative is prepared in two stock solutions, stored separately, and mixed immediately before use.
Solution I (stored in a brown bottle)
Solution II (Lugol’s Solution) (good for several weeks in a tightly stoppered brown bottle)
Combine 9.4 ml of solution I with 0.6 ml of solution II just before use.
1. Add about one-quarter teaspoon (1 g) of fresh feces to the solution, and mix with an applicator stick. Fecal material should be formed or soft if egg counts are to be made later; liquid stool does not work very well for worm burden estimates.
2. Within 24 h, if undisturbed, the specimen forms three well-defined layers. The top layer, a clear orange fluid, consists mainly of formalin, Merthiolate, and water; it does not trap eggs or protozoa. The interface is a thick, pale orange or creamy yellow layer, usually 1 to 2 mm thick; this layer may trap some protozoa and helminth eggs. The bottom layer consists of deeper-staining particulate matter; eggs and protozoa are found throughout this layer.
3. With a glass pipette, MIF direct smears can be made from both the interface and bottom layers. Best results are obtained by making smears from both layers.
4. It has been suggested by some workers that a concentration technique applied to the MIF method (referred to as the MIFC or TFC method) gives satisfactory results. This contention is debatable, and Dunn (27) suggests that it is not nearly as reliable as the MIF direct smear method.
SAF
SAF lends itself to the concentration technique, the permanent stained smear, and fecal immunoassays for Giardia and Cryptosporidium and has the advantage of not containing mercuric chloride, as is found in Schaudinn’s fluid and some of the other fixatives containing PVA (20, 21). It is a liquid fixative, much like the 10% formalin described above (Fig. 2.5). The sediment is used to prepare the permanent smear, and it is frequently recommended that the stool material be placed on an albumin-coated slide to improve adherence to the glass (Fig. 2.6).
Figure 2.5 Stool collection vials, one containing SAF fixative and the other containing Z-PVA, one of the non-mercury-based fixatives. This combination of collection vials is an excellent option; concentrations and fecal immunoassays can be performed from the SAF vial, while the permanent stained smear can be performed from the Z-PVA vial. doi:10.1128/9781555819002.ch2.f2.5
Figure 2.6 Albumin used to precoat the slide prior to the application of SAF-fixed stool concentration sediment. Once the smear is dry, it is ready for permanent staining. doi:10.1128/9781555819002.ch2.f2.6
SAF is considered to be a “softer” fixative than mercuric chloride. The organism morphology is not quite as sharp after permanent staining as that of organisms originally fixed in solutions containing mercuric chloride. The pairing of SAF-fixed material with iron hematoxylin staining provides better organism morphology than does staining SAF-fixed material with trichrome (personal observation). However, a change in the trichrome formulation tends to improve the trichrome staining using SAF (see chapter 3 on examination of fecal specimens). Although SAF has a long shelf life and is easy to prepare, the smear preparation technique may be a bit more difficult for less experienced laboratory personnel who are not familiar with fecal specimen techniques. Laboratories that have considered using only a single preservative have selected this option (concentration, permanent stain, fecal immunoassays for Giardia and Cryptosporidium). Helminth eggs and larvae, protozoan trophozoites and cysts, coccidian oocysts, and microsporidian spores are preserved by this method. After centrifugation, special stains for the coccidia (modified acid-fast stains) and the microsporidia (modified trichrome stains) can be used with the concentrate sediment obtained from SAF-preserved stool material.
SAF fixative is prepared as follows:
To make Mayer’s albumin, mix equal parts of egg white and glycerin. Place 1 drop on a microscope slide, and add 1 drop of SAF-preserved fecal sediment (from the concentration procedure). After mixing, allow the smear to dry at room temperature for 30 min prior to staining. Mayer’s albumin is also available commercially.
Schaudinn’s Fluid
Schaudinn’s fluid is designed to be used with fresh stool specimens or samples from the intestinal mucosal surface. Many laboratories that receive specimens from in-house patients (no problem with delivery times) often select this approach. Permanent stained smears are then prepared from fixed material. A concentration technique for Schaudinn’s fluid-preserved material is also available but is not widely used.
Mercuric Chloride, Saturated Aqueous Solution
Use a beaker as a water bath; boil (use a hood if available) until the mercuric chloride is dissolved; let stand for several hours until crystals form.
Schaudinn’s Fixative (Stock Solution)
Immediately before use, add 5 ml of glacial acetic acid per 100 ml of stock solution.
Schaudinn’s fluid containing PVA (mercury base)
PVA is a plastic resin that is normally incorporated into Schaudinn’s fixative (28). The PVA powder is inert and is not a fixative but serves as an adhesive for the stool material; i.e., when the stool-PVA-fixative mixture is spread onto the glass slide, it adheres because of the PVA component. Fixation is still accomplished by the Schaudinn’s fluid itself. Perhaps the greatest advantage of the use of PVA is the fact that a permanent stained smear can be prepared. Although some laboratories may perform a fecal concentration from a PVA-preserved specimen, some parasites do not concentrate well, nor do some exhibit the typical morphology that would be seen in concentration sediment from a formalin-based fixative. Fixatives containing PVA are highly recommended as a means of preserving cysts and trophozoites for later examination. The use of fixatives containing PVA also permits specimens to be shipped (by regular mail service) from any location in the world to a laboratory for subsequent examination. Fixatives containing PVA are particularly useful for liquid specimens and should be used in the ratio of 3 parts PVA to 1 part fecal specimen. The formula is as follows:
Mix the liquid ingredients in a 500-ml beaker. Add the PVA powder (stirring is not recommended). Cover the beaker with a large petri dish, heavy wax paper, or foil, and allow the PVA to soak overnight. Heat the solution slowly to 75°C. When this temperature is reached, remove the beaker and swirl the mixture for 30 s until a homogeneous, slightly milky solution is obtained.
Schaudinn’s fluid containing PVA (copper base, zinc base)
There has been a great deal of interest in developing preservatives without the use of mercury compounds, and substitute compounds now provide the quality of preservation necessary for comparable protozoan morphology on the permanent stained smear. Copper sulfate has been tried (29, 30) but does not provide results equal to those seen with mercuric chloride (29). However, zinc sulfate has proven to be a good mercury substitute and is used with trichrome stain or iron hematoxylin (Fig. 2.5) (31, 32). Although zinc substitutes have become widely available, each manufacturer has a proprietary formula for the fixative.
Copper Sulfate Solution
Add the CuSO4 ∙ 5H2O to 1,000 ml of distilled water heated to 100°C. Mix until dissolved.
Modified PVA Fixative (Stock Solution)
Immediately before use, add 5 ml of glacial acetic acid per 100 ml of stock solution.
Single-Vial Collection Systems (Other than SAF)
Several manufacturers now have available single-vial stool collection systems, similar to SAF or modified PVA methods (11). From the single vial, both the concentration and permanent stained smear can be prepared (Fig. 2.7 through 2.9). It is also possible to perform fecal immunoassay procedures from some of these vials. Make sure to ask the manufacturer about all three capabilities (concentration, permanent stained smear, fecal immunoassay procedures) and for specific information indicating that there are no formula components that would interfere with any of the three methods. Like the zinc substitutes, these formulas are proprietary.
Figure 2.7 ECOFIX is an example of a fixative that represents the single-vial collection system. This fixative has been coupled with its own stain, the ECOSTAIN (11). Remember to inquire about the compatibility of all single-vial systems with the immunoassay procedures; not all single-vial preservatives are compatible with all fecal immunoassays. doi:10.1128/9781555819002.ch2.f2.7
Figure 2.8 ProtoFix is a single-vial collection option. It is always important to review peer-reviewed literature regarding the results of new products compared with those previously in use. doi:10.1128/9781555819002.ch2.f2.8
Figure 2.9 UNIFIX is a single-vial collection system; information on this product can be found at the company website (www.med-chem.com). This fixative works well with trichrome stain. doi:10.1128/9781555819002.ch2.f2.9
Universal Fixative (TOTAL-FIX)
The TOTAL-FIX stool collection kit is a single-vial system that provides a standardized method for untrained personnel to properly collect and preserve stool specimens for the detection of helminth larvae and eggs, protozoan trophozoites and cysts, coccidian oocysts, and microsporidian spores. Concentrations, permanent stains, most fecal immunoassays, and some molecular methods can be performed from a TOTAL-FIX preserved specimen (Fig 2.10). TOTAL-FIX is a mercury-, formalin-, and PVA-free fixative that preserves parasite morphology and helps with disposal and monitoring problems encountered by laboratories. www.med-chem.com (accessed 4/17/13). TOTAL-FIX is similar to Unifix and Zinc PVA (Z-PVA), commonly used fixatives that have been commercially available and used in many laboratories since 1992.
Figure 2.10 TOTAL-FIX is a single-vial collection system; the fixative contains no mercury, no formalin, and no PVA. It is considered a Universal Fixative and can be used for concentrations, permanent stained smears, fecal immunoassays, special stains, and some molecular methods. doi:10.1128/9781555819002.ch2.f2.10
Quality Control for Stool Fixatives
Fixatives for fecal specimens are checked for quality control by the manufacturer before sale, generally with the use of living protozoa. If you prepare your own fixatives, the following approach can be used for quality control. The specimen used for quality control presented below is designed to be used with fixatives from which permanent stained smears will be prepared (Schaudinn’s fluid, Schaudinn’s fluid containing PVA, copper- or zinc-based fixative containing PVA, SAF, MIF, and Universal Fixatives). However, the same quality control specimen can also be used in a concentration; the white blood cells (WBCs) can be seen in the concentrate sediment (sedimentation concentration) or in the surface film (flotation concentration).
1. Obtain a fresh, anticoagulated blood specimen, centrifuge, and obtain a buffy coat sample (try and find a specimen with a high WBC count).
2. Mix approximately 2 g of soft, fresh fecal specimen (normal stool, containing no parasites) with several drops of the buffy coat cells.
3. Prepare several fecal smears, and fix immediately in Schaudinn’s fluid to be quality controlled.
4. Mix the remaining feces-buffy coat mixture in 10 ml of fixatives with or without PVA, SAF, MIF, or Universal Fixative preservative to be quality controlled.
5. Allow 30 min for fixation, and then prepare several fecal smears. Allow to dry thoroughly (60 min at room temperature or 30 to 60 min in an incubator [approximately 35°C]). Do not use a heat block.
6. Stain the slides by the normal staining procedure (trichrome, iron hematoxylin).
7. After staining, if the WBCs appear well fixed and display typical morphology and color, one can assume that any intestinal protozoa placed in the same lot number of preservative would also be well fixed, provided that the fecal sample was fresh and fixed within the recommended time limits.
8. The bulk quality control specimen can be concentrated as for a normal patient specimen. If the fixative is performing correctly, the WBCs will be visible in the concentration sediment or surface film (depending on the method used).
9. Record all quality control results. If the WBC morphology does not confirm good fixation, describe the results and indicate what corrective actions were used (repeated the test, prepared new fixative).
Procedure Notes for Use of Preservatives
1. Most of the commercially available kits have a “fill to” line on the vial label to indicate how much fecal material should be added to ensure adequate preservation of the fecal material (ratio of 1 part stool to 3 parts fixative). However, patients often overfill the vials; remember to open the vials with the vials turned away from your face. There may be excess gas in the vials that may create aerosols once the vial lids are opened.
2. Although the two-vial system (one vial of 5 or 10% buffered formalin [concentration] and one vial of PVA fixative [permanent stained smear]) has always been the “gold standard,” many laboratories are now using other options, including the single-vial collection systems and the Universal Fixative. Changes in the selection of fixatives are based on the following considerations:
A. Problems with disposal of mercury-based fixatives (availability of high-temperature incineration facilities and cost) and lack of multilaboratory contracts for disposal of such products
B. The cost of a two-vial system compared with the cost of a single collection vial
C. Selection of specific stains (trichrome, iron hematoxylin) to use with specific fixatives
D. Whether the newer fecal immunoassay kits can be used with stool specimens preserved in that particular fixative
Procedure Limitations for Use of Preservatives
1. Adequate fixation still depends on the following parameters:
A. Meeting recommended time limits for lag time between passage of the specimen and fixation
B. Use of the correct ratio of specimen to fixative (1:3)
C. Thorough mixing of the fixative and specimen (once the specimen is received in the laboratory, any additional mixing at that time will not counteract the earlier lack of fixative-specimen mixing and contact)
2. Unless the appropriate stain is used with each fixative, the final permanent stained smear may be difficult to examine (organisms hard to see and/or identify). Examples of appropriate combinations are as follows:
A. Schaudinn’s or PVA fixative with trichrome or iron hematoxylin stain
B. SAF fixative with iron hematoxylin stain or the modified trichrome formulation
Shipment of Diagnostic Specimens, Biological Products, Etiologic Agents, or Infectious Substances
Biological materials shipped domestically or internationally must be packaged in compliance with hazardous-materials transport regulations (33, 34). The United States has incorporated the United Nations Recommendations on the Transport of Dangerous Goods into law, making the U.S. regulations consistent with foreign shipment regulations. The U.S. regulations for packing diagnostic specimens and infectious agents for shipment were modified in 2006. The Public Health Service, Department of Transportation, and Postal Service specify requirements for packaging and shipping of biological materials. These regulations, plus those for packaging and shipping of materials via air, are similar; most carriers elect to follow the international shipping regulations within the International Air Transport Association (IATA) Dangerous Goods Regulations (Table 2.5).
Anyone packaging diagnostic specimens or infectious agents for shipment must receive training and be tested every 2 years. Those who complete the training and test will receive a certificate. The regulations are designed to protect all personnel who may come in contact with the package; compliance with these rules is the responsibility of the sender. Definitions of relevant terms can be found in Tables 2.5 and 2.6.
Double mailing containers should be used in shipping any parasitologic specimens other than microscope slides. The inner container is an aluminum screw-cap mailing tube that fits into an outer cardboard screw-cap mailing container. The specimen vials or tubes in the inner aluminum cylinder should be packed in cotton to absorb any moisture or material that might result from leakage or breakage. Instruction sheets, patient information sheets, etc., may be wrapped around the metal cylinder before it is placed in the outer cardboard mailer. For all packages containing infectious substances, an itemized list of contents must be enclosed between the secondary packaging and the outer packaging (33, 34). Training and training material for the transportation of dangerous goods and infectious substances are available at the American Society for Microbiology and IATA (training manuals).
Prepared slides, such as stained fecal smears or blood films, do not require double mailers for shipment. They may be packed in boxes, cardboard slide holders, or any other suitable container that will prevent damage or breakage. Slides should be individually wrapped in toilet tissue or facial tissue. If a number of slides are to be mailed, they can be wrapped in toilet paper as follows: place a slide on the tissue, wrap the slide several times, place the next slide on top of the first, and continue to wrap the slide several times. The series of slides will be padded and can be easily unwrapped on arrival. When you place slides in the flat cardboard containers, they need additional protection; some slides will arrive broken if this thin cardboard container is merely placed in an envelope for mailing. A plastic slide container with a snap top is an excellent option for shipping microscope slides (holds four to five slides).
Note If the slides are mounted with Permount or other mounting media, they should be completely dry before being packed.
All packages prepared for mailing should contain a complete information sheet about the specimen. An address label should be enclosed inside the package, as well as being on the outside container. All labels should be checked to ensure proper delivery. Also, remember that if you use an overnight carrier, the address must also contain a street address, not just a P.O. Box or general address such as department and institution.
Documentation
The Shipper’s Declaration for Dangerous Goods is a legal document that serves as a contract between the shipper and carrier; it must be accurate and legible and have no spelling errors. However, if minor discrepancies in typing or punctuation are present, they are no longer considered errors if they do not compromise safety. The document must be either handwritten by one person or typed. Two copies of the Shipper’s Declaration must be completed and signed for each shipment. One copy is retained by the shipper, and the other is forwarded with the package; one copy can be a carbon copy. This document is required for shipping infectious substances and toxic substances; it is not required for shipping diagnostic specimens or biological products. If there are no dangerous goods in a shipment with dry ice, the Shipper’s Declaration is not required. It is important for personnel within the laboratory to thoroughly understand these regulations and to have sample copies of appropriate documentation for referral. A checklist used by carriers to decide to accept or reject a package is widely used; a copy of this document should also be on file within the laboratory.
The current editions of the Code of Federal Regulations can be obtained from the Superintendent of Documents, U.S. Government Printing Office, Washington, DC 20402. Inquiries about Postal Service publications may be directed to U.S. Postal Service, 475 L’Enfant Plaza, Washington, DC 20260-5365. Dangerous Goods Regulations can be obtained from Publications Assistant, International Air Transport Association, 2000 Peel Street, Montreal, Quebec, Canada, H3A 2R4.
References
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