Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 18
ОглавлениеFixation of Parasite Specimens and Arthropods
Adequate fixation of parasites is important not only for diagnostic procedures but also as a means of preserving positive material for personnel and student training. There are many fixatives and preservatives available; however, only the more common ones are presented here.
Although this chapter does not include commentary related to the histopathology laboratory, some information about formalin fixatives may be helpful. Concerns about the toxicity of formalin, particularly in the quantities used in a routine histology laboratory, have led to trials of alternative methods of tissue fixation that do not require formalin. Alcohol-based fixatives have been proposed as optimal for immunohistochemical and nucleic acid methods and may be useful for diagnostic light microscopy. Some laboratories have converted to use of an alcoholic fixative (containing 56% ethanol and 20% polyethylene glycol). Although comparative scores between the two were slightly in favor of the formalin, there were no significant differences in terms of tissue architecture, cell borders, cytoplasm, nuclear contours, chromatin texture, red blood cell membranes, and uniformity of staining. Alcohol-polyethylene glycol appears to be a satisfactory alternative to formalin in routine diagnostic surgical pathology (1). In another study, Histochoice produced a staining intensity that was comparable, and in many cases superior, to that of formalin (2).
Formalin-fixed and paraffin-embedded tissues present particular challenges for proteomic analysis. However, most archived tissues in pathology departments and tissue banks worldwide are available in this form. Different approaches to removal of the embedding medium and protein digestion have been developed, thus releasing tryptic peptides, which are suitable for analysis by liquid chromatography-mass spectrometry. Peptide identifications made using this approach are comparable to those from matched fresh frozen tissue. Apparently, a high level of sequence coverage can be seen for proteins under study (3).
The effect of fixation on the degradation of nuclear and mitochondrial DNA in different tissues has been examined (4). Samples of different tissues were preserved in seven fixatives for periods extending from 1 to 336 days to determine which fixatives reduce the time-dependent degradation of DNA and preserve the histologic structure. For long-term storage in combination with amplification of nuclear and mitochondrial DNA, consistent results were obtained with Carnoy’s solution and glutaraldehyde. Variable results were observed for buffered formalin. In regard to comparison of the different tissues, the quantities recovered from skeletal muscles and kidneys were larger than from other tissues. These fixative studies will become even more important as molecular methods applied to fixed tissues become more common.
Molecular characterization of morphologic change requires precise tissue morphology and RNA preservation; however, traditional fixatives usually result in fragmented RNA. To optimize molecular analyses of fixed tissues, morphologic and RNA integrity in rat liver was assessed when sections were fixed in 70% neutral buffered formalin, modified Davidson’s II medium, 70% ethanol, UNIFIX, modified Carnoy’s solution, modified methacarn, Bouin’s medium, phosphate-buffered saline, or 30% sucrose. Each sample was treated with standard or microwave fixation and standard or microwave processing, and sections were evaluated microscopically. RNA was extracted and assessed for preservation of quality and quantity (5). Modified methacarn, 70% ethanol, and modified Carnoy’s solution resulted in tissue morphology providing a reasonable alternative to formalin. Modified methacarn and UNIFIX, best preserved RNA quality. Neither microwave fixation nor processing affected RNA integrity relative to standard methods, although morphology was somewhat improved. Modified methacarn, 70% ethanol, and modified Carnoy’s solution provided acceptable tissue morphology and RNA quality with both standard and microwave fixation and processing methods. Of these three fixatives, modified methacarn provided the best results and can be considered a fixative of choice where tissue morphology and RNA integrity are being assessed in the same specimens. In another study, the PAXgene Tissue System preserves histology similarly to formalin, but does not chemically modify RNA. RNA purified from PAXgene fixed tissues performs as well as RNA from fresh frozen tissue in real-time PCR regardless of amplicon length (6).
PCR has also been used to identify tissue-embedded ascarid nematode larvae. Two sequences of the internal transcribed spacer (ITS) regions of the rRNA gene of the ascarid parasites (ITS1 and ITS2) were compared with those registered in GenBank. PCR amplification of the ITS regions was sensitive enough to detect a single larva of Ascaris suum mixed with porcine liver tissue. These results suggest that even a single larva embedded in tissues from patients with larva migrans could be identified by sequencing the ITS regions (7).
Although formalin fixation is the most common storage, transportation, and preservation method for stool samples, it dramatically reduces the ability to extract DNA from stool samples for PCR-based diagnostic tests. Apparently, the deleterious effects of formalin are both time and concentration dependent and may result from fragmentation of fixed DNA during its purification. This has been seen in studies of the effect of formalin fixation on PCR of Entamoeba histolytica (8). 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. 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 7/24/13). 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 (9, 10).
Parasites found in feces can be preserved in one of two ways (9, 10). One procedure involves thorough mixing of the fecal specimen directly in the fixative. This mixture can then be stored at room temperature without any additional processing. There are several disadvantages of this method: (i) if mixing is not complete, some of the organisms will not be well preserved and will disintegrate during storage; (ii) if the number of organisms is small, a random aliquot may not reveal the parasites; and (iii) the proper ratio of fixative to specimen may not be correct when one is working with larger volumes of material.
The second approach is to clean and separate the organisms from excess debris and concentrate them before fixation. The specimens can be mixed with a large volume of water, passed through a series of screens (large to small pore size), and finally allowed to sediment in a conical container (pilsner glass). After approximately 1 h, the sediment can be washed several times, resedimented, and finally fixed with hot (60 to 63°C) fixative to quickly preserve and stop any further development of certain helminth (Ascaris and Trichuris) eggs. The main disadvantage of this method is extensive washing and manipulation, which may lead to some organism distortion or destruction before fixation. Also, this approach is not very practical unless specimens are being processed for teaching/holding purposes.
The adult forms of most helminths must be relaxed prior to fixation. If this preliminary step is not performed, the worms often contract and curl up when they come in contact with the fixative, thus preventing visual examination of some of the key morphologic characteristics. Several methods, including those involving tap water, physiologic saline, and dilute menthol, are available for this purpose.
Definitive morphology needed for identification of protozoan trophozoites and cysts is best seen on the permanent stained smear, which can be prepared from fresh fecal specimens or from stool that has been submitted to the laboratory in fixative. No washing techniques should be used before bulk fixation, because the trophozoites will be destroyed. Immediately after collection or submission to the laboratory, the specimens should be thoroughly mixed with the fixative of choice. The ratio of fixative to stool should be at least 3 parts fixative to 1 part stool, and the fixation time should be at least 4 h (or less, depending on the amount of specimen used). The normal collection vial systems usually require a fixation time of at least 30 min. If an entire fecal specimen is used, 4 h is recommended. Individual slides can then be prepared from bulk-fixed material by a technique described by Scholten and Yang (11). This method involves centrifugation and smear preparation from the sediment. Although formalin-preserved specimens (except for sodium acetate-acetic acid-formalin-preserved fecal material) are not recommended for the preparation of permanent stained smears, formalin fixation for cyst preservation is recommended for the preservation of teaching specimens that will be examined as wet mounts only. The morphology of cysts can often be seen in formalinized wet mounts sufficiently well to identify organisms to the species level or certainly to be highly suggestive.
Streck tissue fixative has also been tested as a substitute for formalin and polyvinyl alcohol in fecal preservation. Stool samples were examined microscopically as follows: (i) in wet mounts (by bright-field and epifluorescence microscopy), (ii) in modified acid-fast-, trichrome-, and safranin-stained smears, and (iii) with two commercial test kits. Specific results showed that Cyclospora oocysts retained full fluorescence, modified acid-fast- and safranin-stained smears of Cryptosporidium and Cyclospora oocysts were equal in staining quality, and results were comparable in the immunofluorescence assay and enzyme immunoassay commercial kits. However, stool fixed in Streck tissue fixative and stained with trichrome showed unacceptable staining quality compared with stool fixed in preservatives, many of which contained PVA (fecal adhesive). Thus, Streck fixative is an excellent substitute for formalin; however, modifications to the trichrome procedure will be required to improve the staining characteristics of protozoan parasites (12). With the introduction of the Universal Fixative, TOTAL-FIX, formalin is no longer a consideration.
Formalin-Saline Solution
Although formalin is generally used at room temperature for routine diagnostic work, hot (60 to 63°C) 5% formalin in a ratio of 3 parts fixative to 1 part stool is recommended for bulk specimens containing intestinal protozoa. Long-term storage of protozoa is enhanced with buffered formalin; the solution should be replaced every 6 months. Acceptable morphology for teaching purposes can be maintained for 6 months to several years, depending on the organism and the lag time between specimen collection and fixation. The organism cytoplasm will tend to become glassy or granular with very poor nuclear definition. Cysts of Entamoeba coli and Giardia lamblia (G. duodenalis, G. intestinalis) tend to maintain their morphologic characteristics for years, whereas cysts of E. histolytica and E. hartmanni do not preserve well (Fig. 9.1). All of these cysts retain their ability to take up iodine in a wet, direct smear.
Figure 9.1 (Left) Entamoeba coli cyst (unstained). (Right) Giardia lamblia cyst (unstained). doi:10.1128/9781555819002.ch9.f1
The standard 10% formalin will fix protozoan cysts; however, it does not preserve morphology as well as does the buffered 5% formalin.
Formalin-Saline Solution
Formaldehyde is normally purchased as a 37% HCHO solution; however, for dilution, it should be considered to be 100%.
Buffered Formalin-Saline Solution
10% Formalin Solution
Solutions To Induce Relaxation in Adult Helminths ( 13 )
Tap Water or Physiologic Saline
1. Place living worms into a dish containing tap water or 0.85% NaCl solution.
2. Refrigerate the dish for 2 to 4 h.
Note Tap water generally works better than saline; also, the trematodes expel their eggs, thus allowing the internal morphology to be seen more clearly.
Dilute Menthol (14)
1. Dissolve 24 g of menthol crystals in 10 ml of 95% ethyl alcohol, and mix well.
2. Store the solution until needed.
3. Place living worms into a dish containing 100 ml of tap water or 0.85% NaCl.
4. Add 1 drop of menthol solution to the dish containing the worms.
5. Refrigerate the dish for 2 to 3 h.
Note The use of menthol accelerates relaxation of the worms. When this approach is used, the rostellum on the scolex of adult cestodes remains extruded during fixation.
Roundworms can be preserved with several different fixatives (Fig. 9.2). Formalin is usually not recommended, since it tends to harden the tissues. Nematodes can be killed with hot water (60 to 63°C) and then transferred to a preservative, such as alcohol-glycerin or alcohol-formalin-acetic acid (AFA). It is recommended that nematode larval stages be fixed in hot water. Direct fixation in preservatives may cause the cuticle to become “sticky,” and the larvae will be damaged when they adhere to the glass container (15).
Figure 9.2 Adult Ascaris lumbricoides nematode (male worm). doi:10.1128/9781555819002.ch9.f2
Alcohol-Glycerin
Alcohol (70%) containing 5% glycerin is an excellent fixative for most nematodes and should be used hot (60 to 63°C). Specimens can be left in this original fixing solution indefinitely. The glycerin will protect the specimen if the alcohol evaporates. Fixative evaporation is another reason why fixed specimens should be routinely checked every 6 months for possible fixative replacement.
Alcohol-Glycerin
AFA Solution
A solution of alcohol, formalin, and acetic acid can be routinely used for nematodes, trematodes, and cestodes. The fixative should be used hot (60 to 63°C); after fixation for 24 h, parasites can be stored in the alcohol-glycerin mixture.
AFA Solution
Glacial Acetic Acid
Undiluted glacial acetic acid is recommended for fixation of the smaller nematodes (Fig. 9.3). They are killed instantly in an extended position. This fixative is not recommended for the larger worms, such as Ascaris lumbricoides. The acetic acid will clear the worm tissue so that the internal structure becomes visible. Morphologic characteristics can be seen under the microscope if the worm is placed in water on a slide. Worms fixed with acetic acid can be placed in AFA for 24 h and then into alcohol-glycerin for long-term storage.
Figure 9.3 Adult Trichuris trichiura nematode (note the “whiplike” appearance; this is a much smaller adult worm than Ascaris lumbricoides). The small end attaches to the intestinal wall, while the large end is free in the bowel lumen. doi:10.1128/9781555819002.ch9.f3
Dilute Formalin
Although formalin is not recommended as a general fixative for nematodes, a dilute solution (1 to 2%) can be used to kill A. lumbricoides. Higher concentrations should not be used since the differences in osmotic pressure may cause the worms to rupture. After storage in dilute formalin for at least 24 h, the worms can be transferred to 10% formalin for long-term storage.
Note Ascaris, Toxocara, and Baylisascaris eggs continue to develop in formalin (10% or less) to the infective stage and remain viable and infective for a number of months (Fig. 9.4). Worms should be handled with caution, particularly if they are to be dissected.
Figure 9.4 Ascaris lumbricoides fertilized eggs. Note the fully developed larva within each egg; these eggs were viable when photographed. In some cases, the moving larva can be seen, even in formalin-preserved specimens. doi:10.1128/9781555819002.ch9.f4
Most trematodes are very muscular and may contract when placed in fixatives. Living specimens should be placed in cold 0.85% saline for 30 min to several hours (depending on the size of the worm) before fixation. Specimens can be placed on a slide in a petri dish and then covered with another slide or coverglass to flatten the worm. Do not apply pressure, since doing so may distort internal organs (Fig. 9.5).
Figure 9.5 Adult Fasciola hepatica trematodes. (Upper) Fresh trematode (unfixed, unstained). (Middle) Unstained flukes. (Bottom) Stained fluke (note the morphologic details that can be seen after staining). doi:10.1128/9781555819002.ch9.f5
Note Formalin is never recommended for the fixation of trematodes (16).
AFA fixative (see above) should be used hot (60 to 63°C); the worms can be stored in AFA or transferred to 70% alcohol for long-term storage.
Acid fixatives dissolve the characteristic calcareous corpuscles found in tapeworm tissue and should not be used. Since these corpuscles may be used to diagnose tapeworm tissue in histologic sections, buffered formalin or non-acid-containing fixatives are recommended (16).
Formalin
Hot (60 to 63°C) buffered formalin is recommended for the fixation of cestodes. If the whole worms are immediately swirled around in the container of fixative, they will be rapidly fixed with minimal contraction of the proglottids. If placed in cold formalin, the proglottids tend to contract; subsequent India ink injection will be more difficult to accomplish.
AFA Solution
AFA fixes tapeworm tissue well; however, the acid dissolves the calcareous corpuscles. The worms can be transferred to 70% alcohol for long-term storage (Fig. 9.6).
Figure 9.6 Taenia saginata proglottids. (Upper) Note that morphologic details cannot be seen in this preserved but unstained preparation. (Lower) After India ink injection, the uterine branches are now visible and can be counted. doi:10.1128/9781555819002.ch9.f6
Note All tapeworms and proglottids should be handled with care, especially if the species has not been determined. The eggs of Taenia solium (cysticercosis) and Hymenolepis nana are infectious for humans (Fig. 9.7).
Figure 9.7 (Left) Taenia spp. egg (note the striated shell and hooklets seen within the embryo (oncosphere). (Right) Hymenolepis nana egg (note the six-hooked embryo and the polar filaments that are found between the oncosphere and the shell). doi:10.1128/9781555819002.ch9.f7
Whole fecal specimens can be mixed directly with an appropriate fixative, although it is recommended that the organisms be concentrated before fixation. The use of formalin-saline as described for the fixation of protozoa is recommended for helminth eggs and larvae. If well preserved, most helminth eggs maintain their morphologic characteristics indefinitely.
Clinical laboratory personnel may receive arthropods for identification from various patient sources (surface of the body, stool, sputum, etc.) (16, 17). Specimens may be submitted when the actual source is unknown (implicated in a bite, found in the house or yard, etc.). Small, wingless insects and other arthropods should be placed in alcohol (70 to 95%), where they can remain indefinitely. Formalin fixative is not recommended for such specimens. Most flying insects should be killed in a chloroform tube or cyanide bottle and then preserved as a dry mount. Large maggots and other larvae can be killed in hot water first to prevent body shrinkage and contraction when placed in alcohol (18). A number of books also contain detailed information on the collection, preservation, and mounting of arthropods (19–21).
DNA sequencing has become much more common in surveys of archival research collections, particularly in reviewing rare taxa of arthropods. As an example, marine invertebrates have historically been maintained in ethanol following initial fixation in formalin. These collections often represent rare or extinct species or populations, provide detailed time series samples, or come from presently inaccessible or difficult-to-sample localities. Results obtained from preserved crustaceans in archival research collections indicate that in the absence of fresh or frozen tissues, archived formalin-fixed, ethanol-preserved specimens will prove a useful source of material for gene sequence data analysis by PCR and direct sequencing (22). It has also been determined that the now widespread use of critical-point drying of wasps and other insects from alcohol is advocated as a potential source of DNA from rare taxa (23).
Another problem that has been documented involves morphology changes seen with the use of various fixatives. Thus, fixation and mounting can significantly influence the morphometric analysis of mites and other arthropods. It is recommended that morphometric studies be conducted using consistent methods to reduce experimental bias and that the methods used be reported in publications dealing with morphometric analyses (24).
Modified Berelese’s medium is an all-purpose medium that kills, fixes, and preserves many arthropod specimens. No dehydration in alcohol is necessary.
Modified Berelese’s Medium
Mounting and Staining of Parasite Specimens for Examination
Following preservation, many staining and mounting techniques for the preparation of permanent or semipermanent mounts of helminth eggs and larvae and of arthropods may be used.
Because the cuticle of nematodes prevents the uptake of stain , these worms are usually rendered transparent with glycerin or lactophenol. In this way, the internal morphology can be observed for identification purposes. Specific morphologic features that can be seen include the cuticle, alimentary tract, and reproductive structures. Small nematodes can be mounted in glycerin jelly. Standard mounting media containing resins and dehydrating agents are generally not satisfactory because the worms tend to collapse and become distorted during dehydration.
Glycerin Jelly Preparation
Specimens are transferred from pure glycerin directly into the glycerin jelly, a medium that will gel at room temperature, thus providing a semipermanent mount.
Glycerin Jelly (Refractive Index, 1.47)
Dissolve the gelatin in water in a beaker in a water bath. Add the glycerin and melted phenol. Store in small, wide-mouth bottles, and refrigerate. Remelt in a hot water bath before use. Dispense with a dropper onto the slide.
1. Nematodes should first be killed in glacial acetic acid, AFA, or alcohol-glycerin.
2. Place nematodes in a small dish containing 70% alcohol−5% glycerin solution (several millimeters deep).
3. Partially cover the dish to allow gradual evaporation of the alcohol and water for approximately 24 to 36 h. The larger the nematodes, the longer the time needed to complete the evaporation.
Note The evaporation procedure should be done slowly to prevent collapse of the worm; the larger the specimen, the longer the evaporation time.
4. Transfer the worms to another dish containing a few milliliters of glycerin. The specimen should be placed just below the surface; it will eventually (within a few hours) sink to the bottom of the dish. It is then ready to be mounted.
5. Place a drop of liquefied glycerin jelly on the slide, and transfer the specimen into the drop. When the coverglass is added, the glycerin jelly should flow out to the edges of the coverglass. Allow the glycerin jelly to begin to solidify, and apply the coverglass. If the specimen is large, place the coverglass onto small pieces of broken glass to provide more depth under the coverglass.
6. Allow the preparation to gel overnight (horizontal position). The following day, the coverglass can be sealed with Vaspar or enamel paint. These preparations usually last for several years, particularly if no unnecessary pressure is applied to the coverglass.
Note If personnel are not experienced in preparation of these permanent mounts, remember that the worms can be stored indefinitely in vials of pure glycerin. They can be studied in this medium as temporary mounts and then carefully returned to the vial of glycerin.
Glycerin
Nematode specimens can be examined as temporary mounts and subsequently stored in pure glycerin. This approach is particularly helpful for larger specimens (Fig. 9.8).
Figure 9.8 Semipermanent mounts of small nematodes. (Left) Small nematode from soil sample. (Right) Male Enterobius vermicularis (pinworm) nematode. doi:10.1128/9781555819002.ch9.f8
Lactophenol
Lactophenol is recommended when larger specimens are to be examined as temporary mounts. The worms can be placed into lactophenol from alcohol or formalin; clearing will occur, with the length of time depending on the size of the worm. The worms can then be washed in alcohol.
Lactophenol (Refractive Index, 1.44)
Trematodes are usually studied as stained whole mounts (Fig. 9.9). Two stains that are recommended are carmine and hematoxylin. Many modifications are found in the literature, and most give excellent results. The stains are usually available commercially, and they can also be easily prepared in the laboratory.
Figure 9.9 Stained trematodes. (Upper) Eurytrema sp. (Lower) Heterophyes sp. Note that the morphologic details are clearly visible after staining. doi:10.1128/9781555819002.ch9.f9
Semichon’s Acid Carmine
Add glacial acetic acid slowly to the water. Add the carmine powder, and heat to 95 to 100°C for 15 min. Cool and filter (stock solution). Add few drops of stock solution to 70% alcohol, making the working stain pink.
Van Cleave’s Combination Hematoxylin Stain
Van Cleave’s combination hematoxylin stain is a combination of Delafield’s hematoxylin and Ehrlich’s hematoxylin that rarely overstains trematode specimens.
Delafield’s Hematoxylin Stain
Dissolve the hematoxylin in ethyl alcohol, add aluminum alum solution, and let stand for 1 week exposed to air and light (in a paper-capped container). Filter; add glycerin and methyl alcohol. Age for 6 to 8 weeks in a tightly capped bottle in the refrigerator. This is the working stain solution (no dilution is needed).
Ehrlich’s Hematoxylin Stain
Dissolve hematoxylin powder in alcohol; add the other ingredients. Expose to air and light for at least 2 weeks (in a paper-capped container). The solution can be ripened immediately by the addition of 0.4 g of sodium iodate (NaIO3). Store refrigerated in a tightly capped bottle. This is the working solution (no dilution is needed).
Van Cleave’s Hematoxylin Staining Solution
Dissolve the potassium alum in water and add the two hematoxylin stains. This is the working solution.
1. Staining should be done in dilute solutions of carmine or hematoxylin (at least overnight). Specimens stained in carmine are placed into the dilute stain from 70% ethyl alcohol. Those stained in hematoxylin are placed in the stain from water. It is best to overstain and then destain (regressive stain approach).
2. Rinse in 70% ethyl alcohol.
3. Destain in weak acid-alcohol (2 to 4 drops of concentrated HCl in 100 ml of 70% ethyl alcohol). Leach the color from tissues until they are clear but the internal organs remain well stained. Destaining may take several minutes to hours; however, the specimens must be periodically checked to avoid overdestaining.
4. Rinse in 70% alcohol.
5. Place the specimens for 30 min to 1 h in a solution of 70% ethyl alcohol containing 1 or 2 drops of saturated aqueous Na2CO3, NaHCO3, or LiCO3. This step neutralizes the acid step and prevents continued destaining.
6. Rinse in 70% ethyl alcohol.
7. Dehydrate through 80, 95, and 100% ethyl alcohol, with 11 to 15 min for each alcohol change.
8. Clear the specimens in xylene for at least 15 min.
9. Mount in Permount or other permanent mounting medium.
Cestodes can also be examined as stained whole mounts, although the Taenia tapeworms can be examined more rapidly with India ink in a temporary mount (see chapter 4 for India ink injection of tapeworm proglottids). The same carmine and hematoxylin stains as used for trematodes can be used for cestodes.
India Ink Procedure for Tapeworm Proglottids
Identification of adult worms usually involves examination of a tapeworm proglottid. A Taenia proglottid must be gravid, containing the fully developed uterine branches. Using a syringe (1 ml or less) and a 25-gauge needle, India ink is injected into the central uterine stem of the proglottid, filling the uterine branches with ink, or into the uterine pore. The proglottid can then be rinsed in water or saline, blotted dry on paper towels, pressed between two slides, and examined. After the identification has been made, the proglottid can be left between the two slides (place a rubber band around the slides), dehydrated through several changes of ethyl alcohol (50, 70, 90, and 100%), cleared in two changes of xylene, and mounted with Permount for a permanent record. After the xylene step, the proglottid will be stuck to one of the two slides; do not try to remove it (it is very brittle and will crack), but merely add the Permount to the proglottid and then add the coverglass (Fig. 9.10).
Figure 9.10 Taenia gravid proglottids. (Upper) Taenia solium stained with carmine stain. (Lower) Taenia saginata after India ink injection of the uterine branches. doi:10.1128/9781555819002.ch9.f10
Euparal Mounts of Tapeworm Proglottids
Most helminth eggs mounted in Euparal (Flatters and Garnett, Ltd., Manchester, UK) exhibit excellent optical and drying properties, and this mounting medium can also be used to mount tapeworm proglottids (25). Proglottids should be placed in 100% ethyl alcohol for 5 to 10 min and then pressed between two slides as described above. They should then be placed in Euparal; transparency will occur in 2 to 3 h. The best results have been obtained by keeping the slides in a 50°C incubator overnight.
Double-Coverglass Method for Microscopic Mounts of Cysts, Eggs, and Larvae
Slides prepared by the double-coverglass technique will last for several weeks to several years, depending on the organisms and the care taken in preparation. The procedure is as follows (16).
1. Place a small drop of 10% formalin or formalin-saline suspension (containing cysts, eggs, and/or larvae) in the center of a 22-mm round or square coverglass.
2. Using forceps, very carefully apply a smaller coverglass (12, 15, or 18 mm round or square) to the suspension so that the fluid flows to the edges of the small coverglass (with no bubbles). If excess fluid flows beyond the small coverglass, it should be blotted dry or allowed to evaporate. Do not allow it to dry too long; otherwise, bubbles will accumulate under the coverglass.
3. Place a large drop of Permount or other permanent mounting medium in the center of a 1- by 3-in. (1 in. = 2.54 cm) microscope slide. Using forceps, place the double-coverglass preparation (small coverglass side down) onto the mounting medium so that the medium flows to the edges of the large coverglass.
4. Allow the preparation to thoroughly dry in a horizontal position (this may take several days).
Mounting of Arthropods for Examination (20, 26–28)
Before being mounted, specimens in which xylene is used as a solvent require dehydration through 50, 70, and 95% ethyl alcohol. Clearing can be done in clove oil, carbol-xylene (3 parts xylene to 1 part phenol crystals), or absolute ethyl alcohol followed by xylene. The specimens should remain in each solution for at least 15 to 20 min. Specimens mounted in balsam or Permount will take several days to harden; however, those mounted in isobutyl methacrylate will dry very rapidly (within a few hours). With this type of permanent mount, ringing or sealing of the coverglass is not necessary. Some specific recommendations are presented below.
Temporary mounts can be made with a drop of 50% ethyl alcohol, which is gently heated. Clearing and extension of the specimen occur, revealing the typical morphology (Fig. 9.11). Permanent mounts of living specimens can be made by placing the specimen in a drop of chloral-gum medium, adding a coverglass, and then gently heating until bubbling begins. Specimens originally preserved in alcohol must first be washed in distilled water to remove the alcohol.
Figure 9.11 Sarcoptes scabiei itch mites in wet mounts. (Upper) Adult mite. (Lower) Skin scraping containing adult mite (black arrow), eggs (red arrow), and mite feces (green arrow). These specimens could be seen using the high dry objective (magnification, ×400). (Lower image courtesy of National Institutes of Health, http://www.nlm.nih.gov/medlineplus/ency/images/ency/fullsize/2471.jpg). doi:10.1128/9781555819002.ch9.f11
Chloral-Gum Medium
Specimens may be preserved in 70% alcohol or mounted on slides for identification (Fig. 9.12).
Figure 9.12 (Left) Body louse, Pediculus humanus. (Right) Flea, Pulex irritans. (Courtesy of the CDC Public Health Image Library.) doi:10.1128/9781555819002.ch9.f12
1. Drop living fleas or preserved specimens into 10% KOH for a few days until sufficiently cleared.
2. Transfer for 30 min to a small volume of water containing a few drops of concentrated HCl.
3. Dehydrate in 50% ethyl alcohol for 30 min.
4. Dehydrate in 95% ethyl alcohol for 30 min.
5. Clear in beechwood creosote for 1 h, or place in several changes of absolute ethyl alcohol and clear in clove oil or xylene.
6. Mount on slides in balsam, Permount, or isobutyl methacrylate.
Specimens should be preserved in 70% alcohol or cleared and mounted on slides. They can be fixed in an extended position by being gently pressed between two slides while immersed in hot water. Clearing in KOH, dehydration in alcohols, and mounting with balsam, Permount, or isobutyl methacrylate can be done as recommended for fleas (Fig. 9.13).
Figure 9.13 Ticks. (Left) Example of a hard tick, Ixodes sp. (Right) Example of a soft tick, Argas sp. (Courtesy of the CDC Public Health Image Library.) doi:10.1128/9781555819002.ch9.f13
Spiders, scorpions, centipedes, lice, bedbugs, maggots and other larvae, nymphs, and soft-bodied insects can be preserved in 70% ethyl alcohol containing a small amount of glycerin to prevent drying and shrinkage. Containers should remain tightly sealed and should be checked periodically. Larger, hard-bodied insects can be pinned, labeled, and stored in boxes containing naphthalene flakes or paradichlorobenzene to prevent damage from mold and living insects (Fig. 9.14) (29).
Figure 9.14 Example of large hard-body insects, which are triatomid bugs. These large insects can be pinned, labeled, and stored in boxes containing naphthalene flakes or paradichlorobenzene. (Courtesy of the CDC Public Health Image Library.) doi:10.1128/9781555819002.ch9.f14
Note Identification of the many species of arthropods can best be handled by an entomologist or specialist working with a particular arthropod group. Additional help with identification may be obtained at a local university entomology department, a military base (over 100 entomologists are employed by the U.S. Army and Navy), or the entomology department of natural history museums or the Smithsonian Institution (30).
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