Laboratory Diagnosis of Protozoan Infections

Many different technics have been used for the laboratory diagnosis of protozoan infections and for the study of parasitic protozoa. Only the commonest and those which have been found most useful in the author's laboratory are given here. Other routine and specialized technics are given by Craig (1948), Hoare (1949), Kirby (1950), and various textbooks of human parasitology.

Some of these technics are useful not only for protozoa but also for helminth eggs or larvae. If so, their value for these purposes is mentioned.

Direct Microscopic Examination of Wet Fecal Smears

Place a drop of physiological salt solution on a microscope slide. Take up a small amount of feces on the end of a toothpick and mix thoroughly with the salt solution. Do not make too heavy a suspension, or it will be impossible to see objects clearly under the microscope. An emulsion thru which newsprint can be read is about right. Place a coverslip on the drop. Examine under the low and high dry powers of the microscope.

Flagellates and ciliates can be seen moving about actively. Amoebae may move sluggishly or may remain still. Oocysts of coccidia and helminth eggs can be recognized from their shape and size. Many other objects will be seen, some of which may be mistaken for protozoan parasites. These include bacteria, yeasts, fungus spores, the fungus Blastocystis, pollen grains, undigested food particles such as starch grains and plant fibers, and ingested pseudoparasites such as grain mites or coccidian oocysts of animals which have been eaten by or have defecated in the feed of the animals under examination. In cases of enteritis, red or white blood cells or epithelial cells may be present.

In examining preparations under the microscope, move the slide systematically back and forth or up and down in order to bring every part of the preparation into view.

Iodine Staining. In order to bring out certain details which are not visible in the living protozoon, wet smears may be stained with iodine. Prepare a fecal suspension slightly heavier than that described above, and mix it with an equal amount of D'Antoni's aqueous iodine solution or of 1 part of Lugol's solution diluted with 4 parts of distilled water.

Direct Microscopic Examination of Intestinal Mucosa

This technic can be used only in animals which have been killed and have had their intestinal tracts opened. It permits a greater amount of material to be examined on a single slide than does the direct examination of diluted feces. It can be used to find the intracellular and extracellular stages of coccidia, other protozoa, small nematodes such as Strongyloides and Capillaria, small trematodes, cestodes or cestode scolices, and schistosome eggs.

Make a rather deep scraping of the suspected intestinal mucosa with a scalpel, toothpick or similar instrument, or even with the end .of a slide. Place the material thus obtained on a microscope slide and cover with a coverslip. Press the coverslip down if necessary to flatten out the preparation and make it thin enough to see thru.

To search for Trichomonas, Giardia, Hexamita and other motile flagellates, mix a little physiological salt solution with the scraping before placing the coverslip on it.

Microscopic Diagnosis of Tritrichomonas Foetus Infections

In heavy infections of female cattle, T. foetus can be found by direct microscopic examination of mucus or exudate from the vagina or uterus. In aborted fetuses it can be found in the amniotic or allantoic fluid, fetal membranes, placenta, fetus stomach contents, oral fluid or other fetal tissues; it occurs most commonly in the stomach contents and the material around the base of the tongue. In bulls, it can be found in the sheath cavity.

Clean the external genitalia thoroughly before taking samples in order to avoid contamination with intestinal or coprophilic protozoa. Take samples from the vagina by introducing about 10 ml of physiological salt with a bulbed dose syringe and washing it back and forth several times by squeezing the bulb repeatedly. Take samples from the preputial cavity of bulls in the same way, using a long, bulbed pipette or syringe, or introduce a cotton swab into the cavity and rub it around to obtain a sample of exudate; in the latter case, wash off the swab in physiological salt solution.

Allow the washings to stand 1 to 3 hours or centrifuge them before examination. Place a drop of the sediment on a slide, cover with a coverslip, and examine under the microscope.

If trichomonads cannot be found on direct microscopic examination, inoculate some of the washings into CPLM, PGPS or Diamond's medium, and examine after 1, 2 and 4 days' incubation at 37° C.

Sporulation of Coccidian Oocysts

In order to identify coccidia, it is often necessary to allow the oocysts to sporulate (i.e., to develop to the infective stage). To permit this, mix feces containing the coccidia with several volumes of 2.5% potassium bichromate solution and place the mixture in a thin layer in a petri dish. The potassium bichromate prevents bacteria from destroying the oocysts. Oxygen is necessary for the oocysts to develop, so the layer of fluid should never be more than a few millimeters thick. In most species, sporocysts and sporozoites form in a few days, but it is well to allow development to proceed for a week (or, for a few species, even longer). If it is not desired to study the sporulated oocysts immediately, the fecal suspension can be transferred to a bottle and stored in the refrigerator. The oocysts will remain alive for several months or, in some species, as long as a year.

It is best to sporulate coccidian oocysts before they have been subjected to refrigeration, since in some species (apparently a minority), refrigeration of the unsporulated oocysts prevents subsequent sporulation altho it does not harm sporulated oocysts.

MIF (Merthiolate-Iodine-Formaldehyde) Stain-Preservation Technic

This technic was first introduced by Sapero, Lawless and Strome (1951) and improved by Sapero and Lawless (1953). It was designed especially to permit identification of human protozoan trophozoites and cysts, but can also be used for helminth eggs and for parasites of domestic animals. It is simple and relatively cheap, permits rapid (almost immediate) wet-fixed staining of the smears, and preserves the parasites so that feces can be collected in the field or by untrained persons and shipped to the laboratory for later diagnosis. There is no appreciable loss or deterioration of parasites or cellular exudates for 6 months or more.

A. Direct Examination Technic for Fresh Fecal Specimens

1. The MIF stain is composed of tincture of 1:1000 merthiolate No. 99 (Lilly), Lugol's solution (5%) and 40% formaldehyde solution (USP). Since LugoFs solution is unstable, it should be freshly prepared every 2 weeks, and the amount used should be varied with its age. The following amounts (in ml) are recommended:

Table 1

2. Place 1 ml of the stain (sufficient for 25 to 30 fecal smears) in a Kahn tube. Place some distilled water in a second tube. Put a small caliber medicine dropper in each tube.

3. Place 1 drop of distilled water at 1 end of a slide. Add a drop of MIF stain. Mix.

4. Add a small amount of feces and mix. Do not use too much feces, or fixation and staining will be poor. The finished wet smear should be thin enough so that the slide can be tipped on edge without the coverslip sliding.

5. Add a coverslip and examine at once. If it is desired to examine the slide later, ring it with petrolatum to keep the preparation from drying out.

B. Collection and Preservation of Fecal Specimens in the Field for Subsequent Examination in the Laboratory

1. Prepare the following stock MF solution:

Table 2

Store in a brown bottle

2. Measure 2.35 ml MF solution into a standard Kahn tube and stopper with a cork.

3. Measure 0.15 ml of 5% Lugo's solution into another Kahn tube and close with a rubber stopper. (Or keep the Lugol's solution in a bottle, and add the proper amount to the MF solution just before adding the feces in step 4 below. )

4. At the time the fecal sample is collected, pour the MF solution into the Lugol's solution. Within a few seconds, add an amount of feces equal to 2 medium-sized peas (about 0.25 g), and mix thoroughly with an applicator stick. Do not use too much feces. Stopper the tube and set aside for future examination.

5. To examine, draw off a drop of mixed supernatant fluid and feces from the top of the sedimented layer with a medicine dropper and place on a slide. Mix thoroughly, crushing any large particles. Add a coverslip and examine.

Permanent Fixing and Staining Technics

It is often desirable to make permanent preparations of fecal smears or to make hematoxylin-stained slides for detailed study. For this purpose, smears must first be fixed, i.e., the protozoa must be killed by the action of a chemical or mixture of chemicals which will preserve their structures as nearly as possible in the same form as in life.

Many different technics are used for fixing and staining tissues, cells and small organisms. Those given below are especially suitable for protozoa. The standard hematoxylin and eosin stain used routinely for tissue sections is also valuable for protozoa in tissues, but it is so well known that it is not described here. For further information on fixing, sectioning, staining and mounting technics, any text or reference book on microscopic technic may be consulted.

Fixation

Schaudinn's fluid is probably the best all-round fixative for intestinal protozoa, and it also serves well for other forms. Smears may be made on slides and stained in Coplin jars, or they may be made on coverslips and stained in Columbia jars. The latter method has the advantages that smaller amounts of reagents are necessary, a neater preparation is obtained (since there is no possibility of a portion of the smear extending beyond the coverslip), and in the completed slide the mounting medium is beneath the smear rather than above it, so that the microscope objective can come closer to the smear. This factor may be of importance when the oil immersion objective is used. Coverslips are fragile, however, and greater care must be exercised in handling them than in handling slides.

Clean the coverslips by dipping them in 95% alcohol, and dry them with a clean cloth before use. Be careful to handle them only by the edges in order not to leave fingerprints.

Place a tiny drop of albumen fixative in the center of the coverslip (or slide) and smear it over the surface with the little finger. (The finger should previously have been cleaned and rid of its oil by dipping it in 95% alcohol and wiping it with a clean cloth.) Albumen fixative is used to make the feces adhere to the glass.

Take up a small amount of feces on a toothpick (preferably a round, smooth one) and spread as evenly as possible in a very thin layer over the surface of the coverslip. Do not allow it to dry. Drop immediately into a Columbia jar containing Schaudinn's fluid at room temperature or 37° C. Allow to remain about 10 minutes and then transfer to 70% alcohol.

In some cases it may be necessary to mix the feces with a little physiological salt solution in order to make it thin enough to spread well. In other cases the feces are so fluid that if the coverslips are dropped edgewise into the fixative, all the material will come off. To prevent this, place the fixing solution in a small, flat vessel such as a petri dish, and place the coverslip face down on its surface. After a few seconds it can be transferred to a Columbia jar.

After fixation, wash the smear in 2 changes of 70% alcohol for at least 5 minutes each. Then transfer to 70% alcohol containing enough iodine to give it a port wine color. Allow to remain at least 10 minutes (preferably longer). This treatment takes out the excess mercuric chloride which may otherwise form crystals in the preparation. Then transfer to fresh 70% alcohol. Fixed material may be kept in 70% alcohol indefinitely without injury.

Staining with Heidenhain's Hematoxylin

In order to bring out many structures of organisms it is necessary to color them with a dye or dyes. The best and most commonly used dye employed in parasitologic and histologic work is hematoxylin, which is extracted from logwood. Hematoxylin alone has very poor staining properties, and a mordant must be employed to make it effective. Many different formulas have been used for hematoxylin staining solutions. In some, the mordant is mixed with the hematoxylin, while in others it is used separately. Many different compounds are used as mordants, the great majority being salts of heavy metals such as iron, lead, copper, cobalt, tungsten and molybdenum. One of the best hematoxylins is Heidenhain's iron-hematoxylin. A modification of this technic is given below. Starting with the smears in 70% alcohol after passing thru iodine, the staining schedule is:

Table 3

Gradual changes in alcohol concentration are used in all staining and dehydration procedures to avoid distortion of tissues. Hematoxylin-stained smears and sections can be kept in 70% alcohol indefinitely.

In the classical Heidenhain's hematoxylin staining procedure, the stained smears are destained with iron alum. In the above procedure, saturated aqueous picric acid is used instead; this requires a minimum of observation (usually none) during the destaining process, and the resultant stain is dark blue instead of brownish black as with iron alum.

If desired, longer mordanting and staining times can be used. The smears can be mordanted for 2 hours and stained for 4 hours, or they can be mordanted for 3 hours and stained overnight. These give a little more precise staining, but not enough to make them worthwhile for routine purposes.

Counterstaining

If desired, the smears can be counterstained with eosin Y. However, this has a tendency to obscure fine nuclear detail somewhat. To counterstain the smears, transfer them from 70% alcohol to 0.5% solution of eosin Y in 90% alcohol. The pH of this solution should be brought to 5.4 to 5.6 by adding 4.0 ml of 0.1 N HCl per 100 ml. The acidified solution will not keep more than 10 days to 2 weeks. After that its pH will become too high for satisfactory use. Stain for 45 seconds to 2 minutes. Transfer to 95% alcohol to wash out excess dye and then proceed as directed below.

Mounting

Permanent slides are mounted in a medium which, quite fluid at first, later becomes hard. Most mounting media are immiscible with water, and many with alcohol. Hence, before mounting, all water and alcohol must be removed from the smears. This cannot be done simply by allowing the smears to dry, for such dehydration in air would ruin the preparations by distorting the protozoa. Mounting media which have been employed include natural resins such as Canada balsam and damar, and synthetic resins such as euparal, naphrax, permount and clarite.

Starting with stained coverslips in 70% alcohol, pass them thru the following solutions:

Table 4

Mount in permount: Place a drop of permount on a clean slide, place the coverslip slantingly, smear side down, alongside the drop, and gently lay it down on the drop, taking care to prevent air bubbles from forming.

Neutral xylene may be used in place of toluene, altho it hardens the tissues more. Neutral balsam or other resinous mounting media may be used in place of permount. Neutralize the xylene and balsam by placing marble chips in their containers. If this is not done, the stains will fade more or less after months to years.

Feulgen Stain

The Feulgen nucleal stain, which is used for the detection of deoxyribonucleic acid (DNA), is essentially a modification of the Schiff reaction for aldehydes. When DNA is hydrolyzed by hydrochloric acid, aldehyde-like substances are formed which, when treated with colorless fuchsin sulfite, stain a purplish red. Whether the reaction is limited to DNA is doubtful, but at any rate, when properly carried out, the Feulgen technic produces a preparation in which only chromatin is stained.

Not all samples of basic fuchsin are satisfactory for the Feulgen stain. Hence, care must be taken to use dye from a batch which has been found satisfactory and which has been certified as such by the Biological Stain Commission.

1. Fix material to be stained by this method for 24 hours in a saturated solution of mercuric chloride containing 2% acetic acid.

2. Wash in running water, and pass thru 30%, 50%, and 70% alcohol. Do not treat with iodine.

3. Cut sections in the usual manner.

4. Before staining, leave smears and sections in 95% alcohol 48 hours to remove "plasmalogen" substances which may take the stain.

5. To stain, run down thru the alcohols to distilled water, and place the smears or sections in 1 N HCl at 60° C for 4 minutes.

6. Wash in cold 1 N HCl, then rinse with distilled water.

7. Transfer to the decolorized fuehsin solution, and stain 1 to 3 hours.

8. Wash thoroughly in water containing a little sodium bisulfite plus a few drops of HCl.

9. Wash in distilled water.

10. Dehydrate by passing up thru the alcohols as described above, clear, and mount in permount.

Bodian Silver Impregnation Technic

This method is superior to ordinary stains for demonstration of flagella and other diagnostic structures of flagellates. The technic given below is essentially that described by Honigberg (1947). Not all batches of protargol are equally good for this stain, and care must be taken to use a sample which has been tested and found satisfactory.

1. Fix in Hollande's or Bouin's solutions for 10 minutes.

2. Wash in 50% alcohol.

3. Transfer to 30% alcohol and then to distilled water.

4. Bleach in 0.5% aqueous potassium permanganate for 5 minutes.

5. Wash in distilled water.

6. Bleach in 5% aqueous oxalic acid for 5 minutes.

7. Wash several times in distilled water.

8. Place in freshly prepared 1% aqueous protargol solution. (To prepare this solution, place the proper amount of distilled water in a beaker and scatter the protargol powder on its surface; do not stir, heat or disturb the vessel until the protargol has dissolved.)

9. Keep copper wire or thin copper sheeting in the vessel thruout the staining process. Use 5 g copper per 100 ml protargol solution. Columbia jars contain 10 ml of solution. If they are used, it is convenient to place a coil of copper wire weighing 0.5 g in the bottom of each jar before adding the protargol.

10. Stain for 1 to 2 days at room temperature or 37° C in the protargol-copper solution. The staining time and temperature will depend on the material being stained and the final intensity desired. If staining is continued for more than a day, transfer to fresh protargol solution containing fresh copper for the second day.

11. Wash in distilled water.

12. Place in a solution of 1% hydroquinone in 5% aqueous sodium sulfite for 5 to 10 minutes to reduce the silver.

13. Wash several times in distilled water o 140 Place in 1% (or more dilute) aqueous gold chloride for 4 to 5 minutes.

15. Wash in distilled water.

16. Place in 2% aqueous oxalic acid for 2 to 5 minutes until a purplish color appears.

17. Wash several times in distilled water.

18. Place in 5% sodium thiosulfate for 5 to 10 minutes.

19. Wash several times in distilled water.

20. Pass up thru a graded series of alcohols to dehydrate, clear in toluene or xylene, and mount in permount or balsam.

Giemsa Stain for Tissue Sections

The following technic is based on that described by Hewitt (1940) for staining tissue sections with Giemsa stain.

1. Fix small pieces of tissue in formol-Zenker's fluid for 18 to 24 hours.

2. Wash in running tap water overnight.

3. Place in 30% alcohol, 50% alcohol and 70% alcohol for 2 hours each.

4. Treat overnight with 70% alcohol containing enough iodine to give it a port-wine color. This removes the excess mercuric chloride.

5. Place in fresh 70% alcohol for 2 to 3 hours or longer to remove the iodine.

6. Finish dehydration, and infiltrate, embed, section and mount in the usual manner.

7. Run the sections down thru xylene and the alcohols into distilled water, in the usual manner.

8. Mordant in 2.5% aqueous potassium bichromate solution 1/2 to 1 hour.

9. Wash quickly in distilled water.

10. Stain for 24 hours in the following solution:

Table 5

11. Wash in distilled water colored lemon yellow with 2.5% potassium bichromate to remove the excess stain.

12. Differentiate in 70% alcohol. This is the most critical step in the whole procedure. It usually takes 30 seconds to 2 minutes, but the time varies with the type of tissue and the thickness of the sections. Liver usually takes less time than enlarged, engorged spleen, which takes less time than normal spleen. Thick sections take longer than thin. Stop differentiating as soon as the stain is being removed in noticeable quantities. Tissues which contain a large amount of blood will show sharply differentiated red and blue areas macroscopically when they are properly differentiated.

13. Stop the differentiation by washing quickly in distilled water.

14. Dehydrate and mount. Alcohol cannot be used for the dehydrating process, since it will remove too much dye. The simplest and best method of dehydration is to pass the sections thru 3 changes of anhydrous tertiary butyl alcohol for 4 to 10 minutes each (Levine, 1939). (Ordinary samples of tertiary butyl alcohol contain water and cannot be used. A simple way of determining whether a sample is anhydrous is to place it in the refrigerator; its melting point is 25° C, and it will crystallize.)

Transfer the sections from the third tertiary butyl alcohol to 2 changes of xylene and then mount in permount or another resinous mounting medium. It is important that the mounting medium be neutral; if it is acid it will soon decolorize the preparations.

The following dehydration procedure, recommended by Hewitt, can be used if tertiary butyl alcohol is not available:

Table 6

Microscopic Examination of Blood

In searching for blood protozoa, thick or thin smears of the blood are prepared and stained with one or another of the Romanowsky (methylene blue-eosin combination) stains. Thick smears are preferable to thin ones for mammalian blood because their use permits one to examine a relatively large amount of blood in a relatively short time. However, they cannot be used for avian blood because of its nucleated erythrocytes. The protozoa may be distorted in thick smears enough so that some practice is needed to differentiate species, especially of the malaria parasites.

Romanowsky stains may be either rapid (such as Wright's and Field's stains) or slow (such as Giemsa's stain). The rapid stains are satisfactory if speed is necessary, but they stain unevenly, particularly in thick smears, and they are not as precise as the slow stains. Giemsa's stain is best for most purposes. Mammalian blood should be stained at pH 7.0 to 7.2, and avian blood at pH 6.75. These pH's can be obtained by using Clark and Lubs phosphate buffers.

Trypanosomes, microfilariae and most protozoa can be found in fresh, wet, unstained smears, but for critical study they must be stained.

Preparation of Thin Blood Smears

Clean 2 slides by rinsing in 95% alcohol and wiping with a clean cloth. Handle the slides only by their edges to avoid leaving finger marks. Place a small drop of fresh blood at the end of one slide, place the other slide at a 30° angle to the first slide, touch the drop of blood with the end of the slanted slide so that the blood runs into the space beneath it, and then draw the slanted slide rather quickly over the length of the other slide. The blood should be pulled behind the slide and not pushed ahead of it as the smear is being made. A thin, even film of blood should result. Wave the slide in the air until it dries (a matter of a few seconds if the smear is thin enough). If the smear is to be stained in Giemsa's stain, fix it by dipping in absolute methyl alcohol (CP). If the smear is to be stained in Wright's stain, fixation is not necessary, since it will take place during the staining process. If the smear is to be stored for more than a day or so before staining, it should be fixed.

Preparation of Thick Blood Smears

Prepare slides as for thin smears. Place a medium-sized drop of blood or several tiny ones on the slide, and mix with a toothpick or the corner of another slide. Allow to dry in air or in an incubator at 37° C. A hair dryer can be used to speed up the drying process. Thick smears must be laked (i.e., the hemoglobin must be extracted) before being stained. This can be done by placing them in water until the color has disappeared. If Giemsa's stain is used and the smears are fresh, laking will take place during the staining process. If the smears are to be stored for more than a day or so before staining, they should be laked and then fixed with absolute methyl alcohol (CP) before storage, since it is often extremely difficult to remove the hemoglobin from smears which have been stored for some time.

While Leucocytozoon, microfilariae and sometimes Trypanosoma can be found with the low power of the microscope, the stained blood smears should be examined with the oil immersion objective for other protozoa. The faster thin smears have dried, the less distortion is produced. Hence, the most natural appearing protozoa will be found at the thin end and around the edges of the smear.

Cleaning Immersion Oil Off of Slides

Stained blood smears are customarily not covered with a coverslip, and immersion oil is placed on them for examination. The immersion oil should be removed after the examination has been completed if the slides are not to be thrown away. Many people do this by rubbing the slide with lens paper as tho they were polishing silver, a procedure which removes not only the oil but also many of the blood cells. The following technic, which I first saw demonstrated by Dr. Joseph A. Long, permits one to remove the oil quickly and neatly without disturbing the blood cells. It can also be used for slides which have been covered by a coverslip; by its use, one can remove the oil from a newly mounted slide without also removing either the coverslip or the wet mounting medium beneath it.

Fold a small piece (about 5 cm square) of lens paper twice so that it is 4 layers thick. Place the lens paper on top of the immersion oil and allow it to take up the oil. Pull if off the slide sideways in a single motion; do not rub.

Fold a second piece of lens paper like the first. Place a drop of xylene on it. Place the wet lens paper on what remains of the oil. Leave it for a second or two, and then pull it off the slide sideways in a single motion; do not rub. When the xylene has evaporated, the slide will be clean and dry. (Sometimes it is necessary to repeat this second step with a fresh piece of lens paper.)

Concentration of Protozoan Cysts from Feces

A number of technics have been developed for the concentration of protozoan cysts and helminth eggs from feces. They are of 2 general types, flotation and sedimentation. Each has certain advantages over the other.

Flotation Technics

These technics make use of solutions of higher specific gravity than protozoan cysts or helminth eggs, but of lower specific gravity than most of the fecal debris. When feces are mixed with them, the cysts and eggs will float to the top while most of the fecal material remains at the bottom. Flotation technics are most useful for coccidian oocysts, other protozoan cysts, nematode eggs and some tapeworm eggs. They are not satisfactory for trematode, acanthocephalan and other tapeworm eggs.

Many different solutions have been used, and many variations in technic have been proposed. The methods described here all work satisfactorily.

Sugar Flotation

This technic is preferable for general use, but is not satisfactory for protozoan cysts other than those of coccidia. Sugar solution is preferable to sodium chloride, sodium nitrate or other salt solutions except zinc sulfate. It does not crystallize as readily, and causes less distortion than salt solutions, and it is just as efficient (Levine et al. , 1960). The following technic is a modification of the DCF (direct centrifugal flotation) technic introduced by Lane (1923).

1. Make a rather heavy suspension of feces in physiological salt solution in a shell vial or other container.

2. Strain thru 2 layers of cheesecloth into a test tube or centrifuge tube, filling the tube almost half full. The lip of the tube must be smooth, or an air bubble will form under the coverslip following centrifugation (#6 below).

3. Add an equal volume of Sheather's sugar solution, leaving a small air space at the top. Cover with a plastic coverslip or small piece of card, and invert repeatedly to mix.

4. Add enough additional Sheather's sugar solution to bring the surface of the liquid barely above the top of the tube.

5. Cover with a round coverslip.

6. Centrifuge for 5 minutes. (If a centrifuge is not available, let stand for 45 minutes to 1 hour.)

7. Remove the coverslip, place it on a slide, and examine under the microscope.

(If desired, Steps 2 to 4 can be modified by straining the fecal suspension into a second shell vial, mixing with an equal volume of Sheather's sugar solution, and then filling the centrifuge tube with the mixture.)

Zinc Sulfate Flotation

Zinc sulfate solution has the advantage of concentrating the cysts of protozoa such as Entamoeba and Giardia without distortion. The following technic is a modification of that introduced by Faust et at. (1938).

1. Make a suspension of feces in physiological salt solution in a shell vial or other container.

2. Strain 4 ml of the suspension thru 2 layers of cheesecloth into a test tube or centrifuge tube. The lip of the tube must be smooth.

3. Add tap water to within 1 cm of the top of the tube.

4. Mix thoroughly and centrifuge for 4 minutes.

5. Pour off the supernatant fluid.

6. Add a small amount of zinc sulfate solution and mix with an applicator stick. Add more zinc sulfate solution until the tube is almost full, cover with a plastic coverslip or a small piece of card, and invert repeatedly to mix.

7. Add enough additional zinc sulfate solution to bring the surface of the liquid barely above the top of the tube.

8. Cover with a round coverslip.

9. Centrifuge for 5 minutes.

10. Remove the coverslip, place it on a slide, and examine under the microscope.

Sedimentation Technics

Sedimentation technics can be used for concentration of protozoan cysts, and are necessary for the concentration of trematode, acanthocephalan and some tapeworm eggs, which sink to the bottom of the solutions used in the flotation technics. A few protozoan cysts such as those of Eimeria leuckarti also sink to the bottom.

Since they are essentially washing processes, sedimentation technics may not concentrate cysts and eggs as much as flotation technics. Many different sedimentation technics have been developed. The two described below appear to be among the best.

Formalin-Triton-Ether (FTE) Sedimentation Technic

This technic was introduced by Ritchie (1948) and modified by Maldonado, Acosta-Matienzo and Velez-Herrera (1954). The latter considered it the nearest to an all-round diagnostic procedure, since it is highly effective for the detection not only of schistosome, hookworm, whipworm and ascarid eggs but also of protozoan cysts.

1. Mark off a test tube at the 5 ml and 5 ml levels.

2. Place 5 ml of 10% formalin containing a drop of Triton NE in the tube.

3. Add 1 ml of feces.

4. Break up the feces thoroughly with a wooden applicator.

5. Strain the suspension thru 4 layers of cheesecloth into a 15 ml conical centrifuge tube. Squeeze the cloth to get out as much liquid as possible.

6. Add 5 ml of commercial ether to the suspension in the centrifuge tube. Cover the tube with a plastic coverslip and shake vigorously.

7. Centrifuge (at 2000 r.p.m. in a horizontal centrifuge with a radius from the center to the tip of the tube of 8 inches; if another type of centrifuge is used, change the speed of centrifugation accordingly) for 1 minute after the centrifuge has reached its terminal speed.

8. Loosen the plug of detritus at the formalin solution-ether interface with an applicator stick, pour off all the supernatant fluid rapidly, and, holding the tube slightly inverted, clean its walls carefully with a piece of clean, dry gauze. This is done to prevent the liquid and debris on the walls of the tube from sliding down to the bottom and diluting the sediment.

9. Add a drop of physiological salt solution to the sediment to facilitate its removal.

10. Take up the sediment with a pipette (a Stoll pipette works well), place on a slide, add a coverslip, and examine under the microscope.

MIFC (Merthiolate-Iodine-Formaldehyde Concentration) Technic

This technic was introduced by Blagg et al. (1955) as a modification of the MXF preservative stain. They found that the MIFC technic was positive for protozoan trophozoites in 74% of 110 positive human fecal specimens as compared with 55% for the MXF direct smear; it was positive for 92% of 226 specimens containing protozoan cysts, as compared with 58% positive with the MXF direct smear.

1. Prepare an MXF preserved fecal specimen as described above (p. 379).

2. When ready to examine, shake the specimen vigorously for 5 seconds.

3. Strain thru 2 layers of wet surgical gauze into a 15 ml centrifuge tube.

4. Add 4 ml cold (refrigerated) ether to the centrifuge tube, insert a rubber stopper, and shake vigorously. If ether remains on top after shaking, add 1 ml tap water and shake again.

5. Remove the stopper and let stand for 2 minutes.

6. Centrifuge 1 minute at 1600 r.p.m. Four layers will appear in the tube: (a) an ether layer on top, (b) a plug of fecal detritus, (c) an MIF layer, (d) the sediment containing protozoa and helminth eggs on the bottom .

7. Loosen the fecal plug by ringing with an applicator stick.

8. Quickly but carefully pour off all but the bottom layer of sediment.

9. Mix the sediment thoroughly, pour a drop on a slide, cover with coverslip, and examine.

Protozoan Culture Media

NNN (Novy, MacNeal and Nicolle) Medium

This medium was developed for the cultivation of Leishmania, but it can also be used for trypanosomes of the lewisi group.

1. Measure or weigh out:

Table 7

2. Mix, bring to the boiling point, and place in bacteriologic culture tubes in 5 ml amounts. Sterilize in the autoclave. This is the medium base, and can be stored in the refrigerator.

3. To use, melt the agar in the tubes and cool to 48° C. Add to each tube 1/3 of its volume of sterile, defibrinated rabbit blood. Mix thoroughly by rolling the tube between the palms of the hands.

4. Place the tube on a slant without leaving a butt of medium at the bottom, and allow to solidify. This is best done in the refrigerator or in ice, since more water of condensation is obtained in this way. (The protozoa develop best in the water of condensation at the bottom of the slant.)

5. Seal the tubes to prevent the water of condensation from evaporating, and incubate at 37° C for 24 hours to test for sterility before inoculating.

6. Inoculate suspected material into the condensation water and incubate at 22 to 24° C. Transfer cultures every week or two.

Weinman's Trypanosome Medium

This medium was developed by Weinman (1946) for the cultivation of Trypanosoma gambiense and T. rhodesiense. It can also be used for other trypanosomes.

1. The base medium is Difco nutrient agar (1.5%), which consists of:

Table 8

Dissolve in 1 liter distilled water, bring to pH 7.3, sterilize by autoclaving.

2. To prepare the culture medium, heat the base medium to melt the agar. Before it has resolidified, add the following aseptically to each 75 ml of the base:

Table 9

3. Dispense in Kolle flasks or slanted in test tubes. Stopper with rubber corks or seal with Parafilm to retard drying. Store in the refrigerator until used.

4. Inoculate with suspected material and incubate at room temperature. The trypanosomes grow on the surface as small, rounded, colorless, transparent, slightly raised, glistening, moist-appearing colonies 1 to 2 mm in diameter; they are detectable in 5 to 10 days or, exceptionally, in 3 to 4 weeks.

Tobie, von Brand and Mehlman's Trypanosome Medium

This medium was developed by Tobie, von Brand and Mehlman (1950) for African trypanosomes. It consists of a solid slant with a liquid overlay.

1. Solid slant. Measure or weigh out:

Table 10

Mix the ingredients, dissolve by bringing to the boiling point, adjust to pH 7.2 to 7.4 with NaOH, and autoclave at 15 lbs. pressure for 20 minutes. Cool to 45° C, and add 1 part of inactivated, citrated rabbit blood to each 3 parts of the above base. Place 5 ml amounts in test tubes, slant, and allow to cool. If desired, 25 ml amounts may be placed in flasks.

2. Fluid overlay (Locke's solution). Measure or weigh out:

Table 11

Autoclave at 15 pounds pressure for 20 minutes.

3. Place 2 ml of the liquid overlay in each tube containing 5 ml of the base (or 10 to 15 ml in each flask), using aseptic technic.

4. Inoculate with suspected material and incubate at 24 to 25° C for 10 to 14 days.

RES (Ringer's-Egg-Serum) Medium for Enteric Protozoa

This medium was first introduced by Boeck and Drbohlav (1925). Many different modifications have been proposed which are as useful as the one described below. The serum may be replaced by egg albumen, for instance, or the Ringer's solution by Locke's solution.

The medium is essentially a coagulated egg slant overlaid with a fluid nutrient solution.

A. Egg slant.

1. Mix 12.5 ml Ringer's solution with each egg used. For best results, mix in a Waring blendor for 30 seconds. If a blendor is not used, filter the mixture thru cheesecloth.

2. Place 2 ml amounts of the mixture in cotton-stoppered test tubes. (Other standard closures for bacteriologic work can also be used.)

3. Place the tubes upright in a vacuum desiccator. Evacuate the desiccator slowly. As evacuation proceeds, the egg mixture begins to bubble, and within 4 minutes a dense foam of egg begins to climb in the tubes. Stop the evacuation before the cotton plugs become wet, and allow the tubes to remain in the evacuated desiccator for an hour. The purpose of this treatment is to remove the dissolved air from the medium. If it is allowed to remain, it will bubble out during subsequent sterilization and coagulation, roughening and pitting the slant surface (Levine and Marquardt, 1954).

4. Release the vacuum, pack the tubes in baskets, slant them in the autoclave, and inspissate and sterilize them simultaneously at 15 pounds pressure for 20 minutes. Best results are obtained when no butt of medium is left in the tubes. When this is done, 2 ml of fluid makes a slant about 1. 5 inches long in an 18 x 150 mm tube.

B. Fluid overlay.

1. Mix the following aseptically:

Table 12

2. Add sufficient fluid overlay to each egg slant to cover the whole slant. Aseptic technic must be used thruout. Incubate at 37° C for 2 days prior to inoculation to test for sterility.

Balamuth's Amoeba Medium

This medium was developed by Balamuth (1946) for enteric amoebae, but it can be used for other enteric protozoa as well.

1. Mix 288 g dehydrated egg yolk with 288 ml distilled water and 1000 ml physiological salt solution. Mix with a Waring blendor or similar instrument until the suspension is smooth.

2. Heat over an open flame in the upper part of a double boiler, stirring constantly, for 5 to 10 minutes until coagulation begins.

3. Continue heating over boiling water in the double boiler for 20 minutes until coagulation is complete. Add 160 ml distilled water to replace water lost by evaporation.

4. Filter thru a muslin bag. When the bag cools, squeeze it gently to obtain the maximum amount of filtrate.

5. Add enough physiological salt solution to the filtrate to bring its volume to 1000 ml.

6. Place 500 ml of filtrate in each of 2 Erlenmeyer flasks. Autoclave at 15 pounds pressure for 20 minutes.

7. Chill the flasks by refrigeration overnight or in some other way.

8. Filter while cold thru 2 layers of Whatman qualitative filter paper in a Buchner funnel, using negative pressure. Pour the mixture thru the funnel in small amounts, replacing the filter paper frequently.

9. Add an equal volume of Balamuth's buffer solution to the filtrate.

10. Add 5 ml of crude liver extract (Lilly, No. 408) to each liter of medium.

11. Dispense in 5 to 7 ml amounts in tubes.

12. Autoclave at 15 pounds pressure for 20 minutes.

13. Add a small amount of sterile rice powder to each tube. Incubate for 24 hours at 37° C to test for sterility. (If desired, the medium can be stored in large flasks in the refrigerator after autoclaving; it can be kept for a month or more without deteriorating, but any sediment which forms should be removed by filtration before use.)

CPLM (Cysteine-Peptone-Liver Infusion-Maltose) Medium

This medium was developed by Johnson and Trussell (1943) for Trichomonas, but it can also be used for other enteric protozoa.

A. Liver infusion.

1. Mix the following thoroughly, using a Waring blendor if available:

Table 13

2. Infuse for 1 hour at about 50° C.

3. Heat with stirring at 80° C for 5 minutes to coagulate the protein.

4. Filter thru a Buchner funnel. About 320 ml of liver infusion are obtained.

B. Preparation of final medium.

1. Mix the following, using a Waring blendor if available:

Table 14

2. Add the liver infusion from A above.

3. Adjust the pH to 7.0 (approximately 20 ml of 1. 0 N NaOH are needed).

4. Heat to dissolve the agar.

5. Filter thru cotton into a 2000 ml flask.

6. Add 0.7 ml of 0.5% methylene blue solution.

7. Place 300 ml amounts in 500 ml Erlenmeyer flasks.

8. Autoclave for 15 minutes at 15 pounds pressure.

9. Add 75 ml sterile inactivated serum to each 300 ml flask.

10. Place 7 to 10 ml amounts aseptically in sterile, plugged test tubes.

11. Incubate for 2 days at 37° C to test for sterility before use.

BGPS (Beef Extract-Glucose-Peptone- Serum) Medium

This medium was introduced by Fitzgerald, Hammond and Shupe (1954) for use in the diagnosis of Tritrichomonas foetus infections, but it can also be used for other trichomonads.

1. Mix the following in a 3 liter flask:

Table 15

2. Dissolve by boiling. After cooling, adjust the pH to 7.4 with 1.0 N NaOH solution.

3. Cover the mouth of the flask with heavy paper and autoclave for 30 minutes at 15 pounds pressure.

4. After cooling, add 20 ml inactivated (at 56° C for 30 minutes) beef serum aseptically, and mix thoroughly.

5. Dispense in 10 ml amounts into 15 ml culture tubes. Test for sterility by incubating at 37° C for 2 days.

6. Just before inoculation, add 500 to 1000 units of penicillin and 0.5 to 1.0 mg of streptomycin to each ml of medium, and mix thoroughly.

7. Pipette the inoculum on the top of the medium in such a way as to minimize mixing. The trichomonads migrate to the bottom of the tube, while yeasts and molds tend to remain near the top. Incubate at 39° C for 3 to 5 days. To examine, remove a sample from the bottom of the tube with a pipette.

Diamond's Trichomonad Medium

This medium was introduced by Diamond (1957) for the axenic cultivation of trichomonads. It can be used successfully for more species than other media.

1. Mix the following:

Table 16

2. Adjust the pH to 6.8-7.0 with 1 N NaOH for all trichomonads except T. vaginalis; for this species, adjust the pH to 6.0 with 1 N HCl.

3. Add 0.05 g agar.

4. Autoclave for 10 minutes at 15 pounds pressure.

5. Cool to 48° C, and add the following:

Table 17

(The penicillin and streptomycin can be made up in 1 ml distilled water beforehand.)

6. Place 5 ml amounts of the medium aseptically in sterile, stoppered test tubes. Store in the refrigerator up to 14 days or longer.

7. Prior to inoculation, incubate the tubes at 35.5° C for 1 hour.

All the trichomonads which Diamond (1955) cultivated except T. gallinarum and T. gallinarum-like species grew well at 35.5° C; the latter grew better at 38.5° C.

It has been found in the author's laboratory that the phosphates are not necessary for the growth of T. foetus, T. suis, T. gallinae, T. gallinarum and several other species.

RSS (Ringer's-Serum-Starch) Medium for Balantidium

The following medium is slightly modified from that introduced by Rees (1927) for the cultivation of Balantidium coli.

1. Add 25 ml of horse, rabbit or bovine serum aseptically to 500 ml of sterile Ringer's solution.

2. Tube in 8 ml amounts, using aseptic technic.

3. To each tube add a 5 mm loop of rice starch which has been sterilized in a large test tube for 30 minutes at 15 lb pressure.

4. Incubate at 37° C for 48 hours to test for sterility. Store in the refrigerator.

5. Before inoculation, warm the tubes to 37° C by placing them in the incubator. Incubate at 37°. The protozoa grow in the bottom of the tube.

Formulae

Physiological Salt Solution

Table 18

D'Antoni's Iodine Solution

Table 19

Allow to stand 4 days before use. This is the stock solution and contains an excess of iodine. Filter small amounts before use. If tightly stoppered, the filtered solution will keep 4 weeks before too much iodine has volatilized for use.

Lugol's Iodine Solution

Table 20

Dissolve the potassium iodide in the water before adding the iodine.

Mayer's Albumen Fixative

Put the whites of several new-laid eggs in a shallow dish. Whip them a little with a fork or wire egg beater, 2 or 3 dozen strokes being sufficient. Do not beat them until they are white and stiff. Allow them to stand for about an hour, and then skim the foam from the top and pour the remaining liquid into a graduated cylinder. Pour in an equal amount of glycerol, and add 1 g of sodium salicylate for each 100 ml of the mixture. Shake thoroughly and filter thru paper into a clean bottle. Filtration will require 1 to several weeks. It may be accelerated somewhat by pouring a rather small amount at a time into the filter and replacing the paper every few days. Keep a small quantity in a vial or bottle provided with a glass rod stuck into the cork and projecting into the albumen. A drop can easily be placed on a slide with this rod.

Hollander Fixative

Table 21

Schaudinn's Fixative

Table 22

Add 5% acetic acid immediately before use.

Iron Alum Solution

Table 23

Filter immediately before use.

Heidenhain's Hematoxylin (Stock Solution)

Table 24

Allow to remain 1 month in a loosely stoppered bottle before use. To make the staining solution, add 0.5 ml of the stock solution to 9.5 ml of distilled water.

Feulgen Stain

1. Dissolve 1 g basic fuchsin (certified as suitable for the Feulgen stain) in 200 ml boiling distilled water.

2. Cool to 50° C.

3. Filter.

4. Add 20 ml 1 N HCl to the filtrate.

5. Cool to 25° C.

6. Add 1 g dried sodium bisulfite (NaHSO3); this liberates sulfurous acid.

7. Allow to stand at room temperature 24 hours until decolorized.

8. Store in the refrigerator in small glass-stoppered bottles filled to the top to exclude air. The solution will keep several weeks. It should be straw-colored; if it is red, it should be discarded.

Sorensen's Phosphate Buffers

To make M/15 Na2HPO4 solution, dissolve 9.5 g anhydrous Na2HPO4 or 11.9 g Na2HPO4-2H2O in 1 liter distilled water. To make M/15 KH2PO4, dissolve 9.08 g KH2PO4 in 1 liter distilled water. Store separately in pyrex, glass-stoppered bottles.

To prepare buffered water for the Giemsa stain, mix the following amounts of the solutions (in ml):

Table 25

Balamuth's Buffer Solution

Table 26

This is the stock solution. To prepare the final solution used in Balamuth's medium, add 14 parts of distilled water to 1 part of the stock solution.

Ringer's Solution

Table 27

Sheather's Sugar Solution

Table 28

Zinc Sulfate Flotation Solution

Table 29

The specific gravity of this solution is 1.180.