Laboratory Techniques and Procedures





Introduction


Although most routine microbiology diagnostic laboratories process specimens for the diagnosis of parasitic infections, there are no best practice guidelines either for processing or for referral to specialist centers. Microscopy for parasites is most often requested on fecal samples, but other biological fluids may be tested. Certain parasitic infections may require other techniques for diagnosis such as serology or polymerase chain reaction (PCR). In this chapter will be described some of parasitological techniques used for the diagnosis of the main parasitic infections associated with disasters, emphasizing malaria, Schistosoma , Chagas disease, and intestinal parasitic infections. However, in many other parasitology handbooks or textbooks ( Box 4.1 ) and at many websites, more detailed descriptions of parasitological techniques for these and other parasitic infections may be found.



Box 4.1

Recommended Books and Webpages on the Diagnosis of Parasitic Infections


Books and Handbooks


Ash LR, Orihel TC, Bosman A. Bench Aids for the Diagnosis of Malaria Infections . Geneva: World Health Organization; 2000.


Ash LR, Orihel TC, Salvioli L. Bench Aids for the Diagnosis of Intestinal Parasites . Geneva: World Health Organization; 1994.


Ash LR, Orihel TC. Atlas of Human Parasitology . 4th ed. Chicago, IL: American Society of Clinical Pathologists; 1997.


Ash LR, Orihel TC. Parasites: A Guide to Laboratory Procedures and Identification . Chicago, IL: American Society of Clinical Pathologists; 1991.


Chiodini PL, Engbaek K, Heuek CC, et al. Basic Laboratory Methods in Medical Parasitology. Geneva: World Health Organization; 1991.


Garcia LS. Diagnostic Medical Parasitology . 4th ed. Washington, DC: American Society for Microbiology; 2001.


Garcia LS. Practical Guide to Diagnostic Parasitology . Washington, DC: American Society for Microbiology; 1999.


Melvin DM, Brooke MM. Laboratory Procedures for the Diagnosis of Intestinal Parasites . 3rd ed. US Department of Health and Human Services publication no. (CDC) 82-8282. Atlanta, GA: Centers for Disease Control and Prevention; 1982.


Melvin DM, Brooke MM. Morphology of Diagnostic Stages of Intestinal Parasites of Humans . 2nd ed. US Department of Health and Human Services publication no. (CDC) 89-8116. Atlanta, GA: Centers for Disease Control and Prevention; 1989.


National Committee for Clinical Laboratory Standards. Laboratory Diagnosis of Blood-Borne Parasitic Diseases. Approved Guideline M15-A . Wayne, PA: National Committee for Clinical Laboratory Standards; 2000.


National Committee for Clinical Laboratory Standards. Procedures for the Recovery and Identification of Parasites from the Intestinal Tract. Approved Guideline M28-A . Wayne, PA: National Committee for Clinical Laboratory Standards; 1997.


Orihel TC, Ash LR, Ramachandran CP, Ottesen E. Bench Aids for the Diagnosis of Filarial Infections . Geneva: World Health Organization; 1997.


Peters W, Gilles HM. Color Atlas of Tropical Medicine and Parasitology . 4th ed. London: Mosby-Wolfe; 1995.


Wilcox A. Manual for the Microscopical Diagnosis of Malaria in Man . Bethesda, MD: US Department of Health, Education, and Welfare; 1960.


Webpages


www.cdc.gov/dpdx/


www.diplectanum.dsl.pipex.com/purls/


www.southampton.ac.uk/∼ceb/


www.cdfound.to.it/HTML/atlas.htm



Parasitological Methods




  • A.

    Parasitological methods for intestinal parasitic infections



    • 1.

      Equipment: microscope:



      • a.

        The use of a good quality microscope with an oil immersion lens is fundamental. This should have a previously calibrated ocular micrometer to help accurately measure objects such as eggs, larvae, or protozoan cysts. Before using, place slide with stage micrometer under lens, then calibrate for each objective lens. Do this by lining up “0” lines of stage and ocular micrometers, and then read to the right until you find another set of perfectly superimposed lines. The number of microns corresponding with one ocular unit for each lens is found by: stage units/ocular units × 1000 = 1 ocular unit.



    • 2.

      Preserving samples and wet preparations:



      • a.

        Preservatives: formalin (5% or 10%), polyvinyl alcohol (PVA), or sodium-acetate formalin (SAF). The 5% formalin is ideal for protozoan cysts, and 10% is good for eggs and larvae. Maintain 3:1 fixative:specimen ratio. Formalin is cheap, with a long shelf-life, but not good for trophozoites, and cyst/egg details fade with time. PVA is effective (if combined with Schaudinn’s solution, called “gold standard”). Good on trophozoites, cysts, eggs. Can do permanent stains (trichrome or iron hematoxylin) afterwards. However, large mercury content, not easy to prepare in the laboratory. SAF is an alternative to gold standard, and it is good for intestinal protozoans. Very good if followed by permanent smears, stained with modified acid-fast. No mercury, cheap, easy to prepare, long shelf-life are some advantages. Can use in concentration procedures, followed by iron hematoxylin staining. With SAF, remember it dilutes specimen, and albumin may be needed to add to slides to aid adhesion.


      • b.

        Buffered water: commonly used for variety of laboratory purposes. pH 6.8: 48.6 cc Na 2 HPO 4 + 50.4 cc NaH 2 PO 4 in 900 cc water.pH 7.0: 61.0 cc Na 2 HPO 4 + 39.0 cc NaH 2 PO 4 in 900 cc water. pH 7.2: 72.0 cc Na 2 HPO 4 + 28.0 cc NaH 2 PO 4 in 900 cc water.


      • c.

        Conditions for a good fecal sample:



        • i.

          Three stool samples collected on alternate days will increase the sensitivity of parasitological methods.


        • ii.

          Drinking of barium, bismuth, or mineral oil should be avoided because of the ingestion of these liquids can modify the parasitological test results.


        • iii.

          Do not contaminate with water, urine, or other biological fluids.


        • iv.

          Mushy/liquid feces usually have protozoan trophozoites. Cysts have predilection for formed stools. Helminth eggs/larvae variable. First examine exterior for adults, then break up with applicator stick. If adult worms are found can be filtered through a screen.




    • 3.

      Parasitological techniques for fecal samples:



      • a.

        Direct wet mount:



        • i.

          Saline: small drop of 0.85% saline mix (0.85% NaCl) with small sample should be able to read newsprint through, then place coverslip. Good for helminth eggs and larvae as well as motile protozoan trophozoites.


        • ii.

          1% Eosin: allows the observation of many protozoa, in general the background is stained pink color in contrast to protozoa, which stain with a white color if they are alive.


        • iii.

          Iodine: use Lugol’s iodine in same fashion as saline preparation. Good for protozoan cysts because a good stain of nuclear structures allows better identification.




          • Lugol’s iodine:


            Iodine: 1.0 g


            Potassium iodide (KI): 2.0 g


            Distilled water: up to 100 cc


            Dissolve KI in water, then add iodine, store in amber glass bottle out of direct sunlight. Interpretation: positive result—protozoan nuclei take up the iodine and stain pale brown while cytoplasm remains colorless.



          • Some workers prefer to make saline and iodine mounts on separate slides. There is less chance of getting fluids on the microscope stage if separate slides are used. The microscope light should be reduced for low power observations as most organisms will be overlooked by bright light. Illumination should be regulated so that some of the cellular elements in the feces show refraction. Most protozoan cysts will refract light under these conditions. For this method to work effectively, the 1% Lugol’s iodine solution should be a fresh preparation (10–14 days).




      • b.

        Formol ether concentration for ova and cysts



        • i.

          To prepare 10% formol saline solution:




          • NaCl: 8.5 g



          • Formaldehyde (37%–41% solution: 100 cc distilled)



          • Water: 900 cc



          • Dissolve NaCl in water; add formaldehyde solution.



          • Add 2 g feces to the bottle, place lid, shake bottle well.



          • Pass this solution through plastic tea strainer into a 15-cc glass centrifuge tube to within 2 cm of the top.



          • Centrifuge at 3000 rpm for 5 minutes. If centrifuge not available, leave to stand out of direct sunlight for 30 minutes.



          • Remove fatty plug at top of solution and discard along with supernatant. Resuspend deposit with saline or iodine over glass slide and examine under microscope.




      • c.

        Cellophane (Kato) thick fecal smear technique: fecal egg counts may be useful techniques for research, epidemiology. The previous modified Stoll method described many years ago is more troublesome and time-consuming than Kato-Katz. In fact, the Stoll technique is not effective when the number of eggs is less than 200 ova per gram. This technique is useful for the diagnosis of the main soil transmitted helminthic and Schistosoma infections.



        • i.

          With a wood or plastic applicator stick, place 41.7 mg of feces on slide using the template. If sample has excessive amounts of fiber, pass through 105-mesh metal/nylon sieve. Cover sample with previously soaked cellophane coverslip, press against absorbent paper until smear covers area under coverslip, ensuring that the fecal sample does not extend past the edges of the cellophane.


        • ii.

          Leave smear until it clears, usually about 1 hour at room temperature.


        • iii.

          Examine first under 4× objective lens and 40× later.


        • iv.

          Some considerations:




          • Hookworm ova visible for up to 30 minutes after preparation. Schistosoma ova are best seen about 24 hours later. Trichuris and Ascaris eggs are visible at any time.



          • As the cellophane coverslips are soaked in a malachite green solution, the background will be green, but schistosome and hookworm eggs will remain colorless.



          • Cellophane: water wettable cellophane available as 22-mm wide rolls can be cut into rectangles of 22 × 30 mm.



        • v.

          Glycerin-malachite green solution: mix 100 cc glycerol, 100 cc water, and 1.0 cc of 3% aqueous malachite green solution. Place in screw-topped jar, add coverslips, and soak for at least 24 hours before use ( Fig. 4.1 ).




          Fig. 4.1


          Materials used for Kato-Katz technique: two plastic templates with a hole of 6 mm (for approximately 41.7 mg of feces); two plastic screens, one of them with a cardboard border; two spatulas, one plastic and the other one of aluminum; and strips of hydrophilic cellophane 40- to 50-µm thick, soaked in glycerol-malachite green solution for at least 24 hours before use.



      • d.

        Modified cold Ziehl-Neelsen’s stain (for Cryptosporidium and other coccidian species): This technique is used for the demonstration of oocysts of Cryptosporidium species in feces. Alternatively, the modified auramine-phenol stain may be used.



        • i.

          Prepare a medium to thick smear and air dry.


        • ii.

          Fix in methanol for 3 minutes and air dry.


        • iii.

          Flood the slide with modified Kinyoun’s acid-fast stain (3% carbol fuchsin) and leave for approximately 15 minutes.


        • iv.

          Rinse with tap water.


        • v.

          Flood the slide with 1% acid methanol to decolorize and leave for 15–20 seconds.


        • vi.

          Rinse with tap water.


        • vii.

          Counterstain with 0.4% malachite green or alternative and leave for 30 seconds.


        • viii.

          Rinse with tap water and air dry.


        • ix.

          Examine using 40× or 50× objective and 1× eyepiece lenses. Morphology may be examined closely with a high-power objective.


        • x.

          Positive result: Cryptosporidium species are 4–6 µm with a spherical shape; the affinity to fuchsine stain is high, despite some oocysts appearing less stained. Cyclospora cayetanensis oocysts stain pinkish red, are spherical, 8–10 µm, and contain a central morula; staining is variable and some oocysts may appear unstained.


        • xi.

          Cystoisospora belli species stain red, measure around 31 × 15 µm, and are elongated oval bodies tapered at both ends, containing a granular zygote or two sporoblasts. The oocysts observed in feces are usually unsporulated. Yeasts, other biota, and fecal debris may also take up the stain ( Fig. 4.2 ).




          Fig. 4.2


          Oocysts of Cryptosporidium spp. colored with acid-fast stain in a fecal sample.





  • B.

    Parasitological methods for diagnosis of malaria ,



    • 1.

      Introduction: The observation of malaria parasites in blood films is the still the gold standard for the diagnosis of malaria. In thick blood, the red blood cells (RBCs) are lysed; consequently, diagnosis is based on the appearance of the parasite. In thick films, organisms tend to be more compact and denser than in thin films. The use of thin films is always required because RBCs are fixed so the morphology of the parasitized cells can be seen, allowing the identification of species and stages. However, malaria parasites may be missed on a thin blood film when there is a low parasitemia. With a thick blood film, the red cells are approximately 6 to 20 layers thick, which results in a larger volume of blood to be observed. Although some technicians stain thick and thin films on separate slides, it is recommended to use both stains in only one slide.


    • 2.

      Field’s staining method for thick blood films: The method is based in the use of two main components. Field’s A is a buffered solution of azure dye and Field’s B is a buffered solution of eosin. Both Field’s A and B are supplied ready for use by the manufacturers.



      • a.

        Place a drop of blood on a microscope slide and spread to make an area of approximately 1 cm 2 . It should just be possible to read small print through a thick film.


      • b.

        The film is air dried and NEVER fixed in methanol.


      • c.

        The slide is dipped into Field’s stain A for 3 seconds.


      • d.

        The slide is then dipped into tap water for 3 seconds and gently agitated.


      • e.

        The slide is dipped into Field’s stain B for 3 seconds and washed gently in tap water for a few seconds until the excess stain is removed.


      • f.

        The slide is drained vertically and left to dry.


      • g.

        Dry films upright.


      • h.

        Examine the film at 40× before going to oil immersion (1000× magnification).


      • i.

        Microscopic features of the Field’s stained thick blood film:



        • i.

          The end of the film at the top of the slide when it was draining should be looked at. The edges of the film will also be better than the center, where the film may be too thick or cracked.


        • ii.

          In a well-stained film, the malaria parasites show deep-red chromatin and pale-blue cytoplasm.


        • iii.

          White cells, platelets, and malaria pigment can also be seen on a thick film. The leukocyte nuclei stain purple and the background is pale blue. The red cells are lysed and only background stroma remain. The occasional red cell may fail to lyse.


        • iv.

          Schizonts and gametocytes, if present, are also easily recognizable.


        • v.

          A thick film should be examined for at least 10 minutes, which corresponds to approximately 200 oil immersion fields, before declaring the slide negative.


        • vi.

          As a result of hemolysis of the RBC due to staining of an unfixed film, the only elements seen are leukocytes and parasites, the appearance of the latter being altered.


        • vii.

          Consequently:




          • The young trophozoites appear as incomplete rings or spots of blue cytoplasm with detached chromatin dots.



          • The stippling of Plasmodium vivax and Plasmodium ovale may be less obvious although occasionally ghost stippling may be seen.



          • The cytoplasm of late trophozoites of P. vivax and P. ovale may be fragmented.



        • viii.

          Caution should be exercised when examining thick blood films, as artifacts and blood platelets may be confused with malaria parasites.




    • 3.

      Thin blood films: When examining thin blood films for malaria you must look at the infected RBC and the parasites inside the cells.



      • a.

        Rapid Field’s stain for thin films: This is a modification of the original Field’s stain to enable rapid staining of fixed thin films. This method is suitable for malaria parasites, Babesia spp., Borrelia spp., and Leishmania spp.



        • i.

          Air dry the film.


        • ii.

          Fix in methanol for 1 minute.


        • iii.

          Flood the slide with 1 mL of Field’s stain B, diluted 1 in 4 with distilled water.


        • iv.

          Immediately add an equal volume of undiluted Field’s stain A, mix well, and allow to stain for 1 minute.


        • v.

          Rinse well in tap water and drain dry.


        • vi.

          Dry films upright.


        • vii.

          Examine the film at 40× before going to oil immersion (1000× magnification).


        • viii.

          Uses: this is a useful method for fast presumptive species identification of malarial parasites. It shows adequate staining of all stages including stippling (mainly Maurer’s dots). However, staining with Giemsa is always the method of choice for definitive species differentiation.




    • 4.

      Giemsa stain for thin films:



      • a.

        Air dry thin films.


      • b.

        Fix in methanol for 1 minute.


      • c.

        Wash in tap water and flood the slide with Giemsa diluted 1 in 10 with buffered distilled water pH 7.2. The diluted stain must be freshly prepared each time.


      • d.

        Stain for 25 to 30 minutes.


      • e.

        Run tap water on to the slide to float off the stain and to prevent deposition of precipitate onto the film.


      • f.

        Dry films upright.


      • g.

        Examine the film at 40× before going to oil immersion (1000× magnification).


      • h.

        Microscopic features of the thin blood film:



        • i.

          Examine the tail end of the slide where the red cells are separated into a one-cell-layer thickness.


        • ii.

          An alkaline buffer pH 7.2 is vital for clear differentiation of nuclear and cytoplasmic material and to visualize inclusions such as Schüffner’s/James’s dots in the red cells. Acidic buffer is unsuitable.


        • iii.

          The red cells are fixed in the thin film so the morphology of the parasitized cells and the parasites can be seen.


        • iv.

          On a well-stained film the chromatin stains red/purple and the cytoplasm blue. Leukocytes have purple nuclei. The red stippling, if present, should be clearly visible.



      • i.

        Infected RBCs:



        • i.

          Look at the size of the infected RBC.


        • ii.

          Are there any Schüffner’s dots present or not?




    • 5.

      Estimation of percentage parasitemia of Plasmodium falciparum : Counting of RBCs infected with parasites of P. falciparum is essential and the percentage parasitemia should always be reported as this has implications for prognosis and the pattern of treatment employed.



      • a.

        The recommended procedure for estimating the percentage parasitemia in a thin blood film is by expressing the number of infected cells as a percentage of the RBC, for example, 3 parasitized red cells/100 RBC, or 3% parasitemia.


      • b.

        An RBC infected with multiple parasites counts as one parasitized red cell. The percentage parasitemia should be calculated by counting the number of parasitized RBCs in 1000 cells in a thin blood film.


      • c.

        Alternatively, the World Health Organization recommends a method that compares the number of parasites in a thick blood film with the white blood cell count.


      • d.

        The parasitemia is estimated by first counting the number of parasites per 200 white blood cells in a thick blood film and then calculating the parasite count/μL from the total white blood cell count/μL.


      • e.

        Knowledge of either % parasitemia or total parasite count is essential for the correct clinical management of P. falciparum malaria.



    • 6.

      Effects of anticoagulant on the microscopic diagnosis of malarial parasites



      • a.

        Thin blood films for malaria diagnosis are best prepared from venous or capillary blood taken directly from the patient, without the addition of anticoagulant. However, this is not usually possible in a clinical laboratory, as many samples are received from general practices and other hospitals. All anticoagulants have some effect on the morphology of malaria parasites and the RBC they inhabit. This effect depends on the stage of the parasite, the time taken for the blood to arrive to the laboratory, and the type of anticoagulant used. If it is necessary to use an anticoagulant, the films should be prepared as soon as possible after the blood has been taken.


      • b.

        If the films cannot be made immediately, potassium ethylenediaminetetraacetic acid (EDTA) is the anticoagulant of choice. However, if the blood is left for several hours in EDTA, the following effects may be seen.



        • i.

          Sexual stages may continue to develop and male gametocytes can exflagellate, liberating gametes into the plasma. These can be mistaken for organisms such as Borrelia . Gametocytes of P. falciparum , which have a characteristic crescent shape, may round up and then resemble those of Plasmodium malariae .


        • ii.

          Accolé forms, which are characteristic of P. falciparum, may be seen in P. vivax because of attempted reinvasion of the RBC by merozoites.


        • iii.

          Mature trophozoites of P. vivax may condense when exposure becomes prolonged and in cases of extreme exposure, RBCs containing gametocytes and mature schizonts may be totally destroyed along with the contained parasites. The malaria pigment, hemozoin, always remains and can provide a clue to the presence and, to an expert eye, identity of the parasite.


        • iv.

          The morphology of the RBCs may be altered by shrinkage or crenation.




    • 7.

      Appearance of Plasmodium spp. in thin blood films ( Figs. 4.3– 4.6 , Tables 4.1– 4.5 )


Aug 20, 2021 | Posted by in GENERAL & FAMILY MEDICINE | Comments Off on Laboratory Techniques and Procedures
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