Preservation of specimens for Parasitology

PRESERVATION OF SPECIMENS

Principle

Fecal specimens that cannot be processed and examined in the recommended time should be placed in an appropriate preservative or combination of preservatives for examination later. Preservatives will prevent the deterioration of any parasites that are present. A number of fixatives for preserving protozoa and helminthes are available. Each preservative has specific limitations, and no single solution enables all techniques to be performed with optimal results. The choice of preservative should give the laboratory the capability to perform a concentration technique and prepare a permanent stained smear for every specimen submitted for fecal examination.

Procedure

1. Wear gloves when performing this procedure.
2. Add a portion of fecal material to the preservative vial to give a 3:1 or 5:1 ratio of preservative to fecal material (a grape-sized formed specimen or about 5 ml of liquid specimen).
3. Mix well by stirring with an applicator stick or the “Spork” insert that is attached to the fixative vial lid to give a homogeneous solution.
4. Allow to stand for 30 min at room temperature to allow adequate fixation.
5. While using commercial collection system follows the manufacturer’s directions concerning shaking the vials, etc.

A. Schaudinn’s fixative

This preservative is used with fresh stool specimens or samples from the intestinal mucosal surface. Many laboratories that receive specimens from in-house patients (no problem with delivery times) may select this approach. Permanent stained smears are then prepared from fixed material.

Advantages

1.       Designed to be used for the fixation of slides prepared from fresh fecal specimens or       samples from the intestinal      mucosal surfaces

2.    Prepared slides can be stored in the fixative for up to a week without distortion of      protozoan organisms.

3.      Easily prepared in the laboratory

4.      Available from a number of commercial suppliers

Disadvantages

1.      Not recommended for use in concentration techniques

2.      Has poor adhesive properties with liquid or mucoid specimens

3.      Contains mercury compounds (mercuric chloride), which may cause disposal problems.

B. PVA (Polyvinyl Alcohol)

PVA is a plastic resin that is normally incorporated into Schaudinn’s fixative. The PVA powder serves as an adhesive for the stool material; i.e., when the stool-PVA mixture is spread onto the glass slide, it adheres because of the PVA component. Fixation is still accomplished by the Schaudinn’s fluid itself. Perhaps the greatest advantage in the use of PVA is the fact that a permanent stained smear can be prepared. PVA fixative solution is highly recommended as a means of preserving cysts and trophozoites for examination at a later time. The use of PVA also permits specimens to be shipped (by regular mail service) from any location in the world to a laboratory for subsequent examination. PVA is particularly useful for liquid specimens and should be used at a ratio of 3 parts PVA to 1 part fecal specimen.

Advantages

a.       Ability to prepare permanent stained smears and perform concentration techniques

b.      Good preservation of protozoan trophozoites and cyst stages

c.       Long shelf life (months to years) in tightly sealed containers at room temperature

d.      Commercially available from a number of sources

e.       Allows shipment of specimens

Disadvantages

a.       Some organisms (Trichuris trichiura eggs, Giardia lamblia cysts, Isospora belli oocysts) are not concentrated as well from PVA as from formalin-based fixatives, and morphology of some ova and larvae may be distorted.

b.      Contains mercury compounds (Schaudinn’s fixative), which may cause disposal problems

c.       May turn white and gelatinous when aliquotted into small amounts (begins to dehydrate) or if refrigerated

d.      Difficult to prepare in the laboratory.

C. SAF (Sodium acetate formalin)

SAF lends itself to both the concentration technique and the permanent stained smear and has the advantage of not containing mercuric chloride, as is found in Schaudinn’s fluid and PVA. It is a liquid fixative much like 10% formalin. The sediment is used to prepare the permanent smear, and it is recommended that the stool material be placed on an albumin-coated slide to improve adherence to the glass. SAF is considered a “softer” fixative than mercuric chloride. The morphology of organisms will not be quite as sharp after staining as that of organisms originally fixed in solutions containing mercuric chloride. Staining SAF-fixed material with iron-hematoxylin appears to reveal organism morphology more clearly than staining SAF-fixed material with trichrome.

Advantages

a.       Can be used for concentration techniques and stained smears

b.      Contains no mercury compounds

c.       Long shelf life

d.      Easily prepared or commercially available from a number of suppliers

Disadvantages

a.       Has a poor adhesive property. Albumin-coated slides are recommended for stained smears.

b.      Protozoan morphology with trichrome stain not as clear as with PVA smears. Hematoxylin staining gives better results.

c.       More difficult for inexperienced workers to use.

D. MIF

Merthiolate (thimerosal)-iodine-formalin (MIF) is a good stain preservative for most kinds and stages of parasites found in feces and is useful for field surveys. It is used with all common types of stools and aspirates; protozoa, eggs, and larvae can be diagnosed without further staining in temporary wet mounts. Many laboratories using this fixative examine the material only as a wet preparation (direct smear and/or concentration sediment). MIF is prepared in two stock solutions that are stored separately and mixed immediately before use.

Advantages

a.       Combination of preservative and stain (merthiolate), especially useful in field surveys

b.      Protozoan cysts and helminth eggs and larvae can be diagnosed from temporary wet-mount preparations.

Disadvantages

a.       Difficult to prepare permanent stained smears

b.      Iodine component unstable; needs to be added immediately prior to use

c.       Concentration techniques may give unsatisfactory results.

d.      Morphology of organisms becomes distorted after prolonged storage.

E. 5 or 10% formalin

Formalin is an all-purpose fixative that is appropriate for helminth eggs and larvae and protozoan cysts. Two concentrations are commonly used: 5% which is recommended for preservation of protozoan cysts, and 10%, which is recommended for helminth eggs and larvae. Most commercial manufacturers provide 10%, which is most likely to kill all helminth eggs. To help maintain organism morphology, formalin can be buffered with sodium phosphate buffers, i.e., neutral formalin.

Advantages

a.       Good routine preservative for protozoan cysts and helminth eggs and larvae. Materials can be preserved for several years.

b.      Can be used for concentration techniques (sedimentation techniques)

c.       Long shelf life and commercially available

d.      Neutral formalin (buffered with sodium phosphate) helps maintain organism morphology with prolonged storage.

Disadvantage

a.       Permanent stained smears cannot be prepared from formalin-preserved fecal specimens.

Shipment of Specimens

Principle

In outpatient situations, it may be necessary for a specimen to be shipped to the laboratory for examination. Only preserved fecal specimens should be shipped, as any delays in examination may result in deterioration of parasitic organisms. Prior fixation also reduces the risk of infection from any etiologic agents present in the specimen. The U.S. Postal Service regulates the shipment of clinical specimens through the mail. It is the responsibility of the sender to conform  to these regulations.

Specimens

1. Preserved fecal specimens in collection vials.
2. Fecal smears for staining and examination for parasitic organisms.
3. Blood smears for staining and examination for blood parasites (thin blood films should be fixed in methyl alcohol prior to shipment).

Procedure

1. Place the primary container of preserved fecal material into the secondary container (metal sleeve or a sealable bag), and seal.
2. Place into the mailing container, and seal.
3. Label appropriately for shipment.
4. Wrap glass slides in shock-absorbent material to protect from breakage, or place them in a sturdy slide container.
5. Slides need not be placed in double containers for shipping.
6.  Place padded slides in shipping container and label appropriately.                            Duodenal Contents: String Test (Entero-Test Capsule)

Principle

The Entero-Test capsule is usually administered and the string is retrieved by a physician. This test is used to procure specimens from the duodenum that are then examined for the presence of parasites. The Entero-Test is a gelatin capsule lined with silicone rubber that contains a spool of nylon string and a weight. The end of the string is taped to the back of the patient’s neck or the patient’s cheek just before the capsule is swallowed with water. After swallowing the capsule,

the patient is allowed to relax for 4 h. The patient is not allowed to eat during this time but is allowed to drink a small amount of water. As the capsule dissolves, the string unwinds and is carried by peristalsis to the duodenum, and the duodenal mucus adheres to the string. Any Strongyloides larvae, Giardia  trophozoites, or Cryptosporidium  or Isospora oocysts that are present will also adhere to the string and will be pulled up with the string when it is removed.

The specimen can be examined as a wet preparation or as a permanent stained smear. In rare instances, Clonorchis sinensis eggs may be recovered. This test is a less invasive substitute for duodenal aspiration.

Procedure

1. Gloves must be worn when handling this specimen. Infectious Strongyloides larvae can penetrate the intact skin.
2. Record the color of the string. Yellow bile stain indicates that the string did reach the duodenum.
3. Place the specimen under the biosafety cabinet, hold the dry white end in one hand, and strip all the mucus off the string by gripping it between the thumb and index finger of the other hand and squeezing it all the way down to the end, so that the mucus goes into the screw-cap container.
4. Place 1 drop of mucus on a clean slide, and cover with a coverslip (22 by 22 mm). If the mucus is very viscous, add a drop of saline before adding the coverslip.
5. Store the remaining mucus in a transfer pipette placed in a labeled test tube (16 by 125 mm) so that it will not dehydrate.
6. Examine the entire coverslip under low power (100X) for larvae or motile trophozoites, looking especially carefully at the mucus, where Giardia lamblia may be entangled.
7. Examine the mucus under high dry power (400X), since G. lamblia may be detectable only by the flutter of the flagella rather than by motility.
8. If there is enough specimen, gently smear a drop or two of patient material on two slides, and immediately immerse the slides in Schaudinn’s fixative so that permanent stained slides may be made. If the specimen is not adequate for this, place the wet mount slide in a Coplin jar containing Schaudinn’s solution after it has been read. The coverslip will float off and sink to the bottom, allowing the remaining material to be stained. The fixation and staining times are identical with those for routine fecal smears.
9. If the material contains a lot of mucus or is a watery specimen, gently mix 1 or 2 drops of patient material with 3 or 4 drops of PVA fixative directly on the slide. Let the smear air dry for at least 2 h prior to staining. The fixation and staining times are identical to those for routine fecal smears.
10. Place a drop of the mucus on one or more slides to be stained for Cryptosporidium and Isospora species, and then repeat the wet-mount procedure.
11. Stain the Cryptosporidium and Isospora slide(s) with modified acid-fast stain, and examine as usual.
12. Examine the permanent stained smear with the oil immersion lens (100X) with maximum light. Examine at least 300 oil immersion fields on each smear.

Urogenital Specimens: Direct Saline Mount

Principle

Trichomonas vaginalis infections are primarily diagnosed by detecting live motile flagellates from direct saline (wet) mounts. Microscope slides made from  patient specimens can be examined under low and high power for the presence of actively moving organisms.

Specimens

Vaginal discharge, Urethral discharge, Penile discharge, Urethral-mucosa scrapings, First-voided urine with or without prostatic massage.

Procedure

1. Apply the patient’s specimen to a small area on a clean microscope slide.
2. Immediately before the specimen dries, add 1 or 2 drops of saline with a pipette. If urine sediment is used, the addition of saline may not be necessary.
3. Mix the saline and specimen together with the pipette tip or the corner of the coverslip.
4. Cover the specimen with the no. 1 coverslip.
5. Examine the wet mount with the low-power (10X) objective and low light.
6. Examine the entire coverslip for motile flagellates. Suspicious objects can be examined with the high-power (40X) objective.

Expectorated Sputum: Direct-Mount and Stained Preparations

Principle

A direct smear can be used to detect large or motile organisms from the lung. Parasites which can be detected and may cause pneumonia, pneumonitis, or Loeffler’s syndrome include Entamoeba histolytica, Paragonimus spp., Strongyloides stercoralis, Ascaris lumbricoides, and hookworm. The smears can be examined with and without the addition of D’Antoni’s or Lugol’s iodine. Trichrome stains of material may aid in differentiating E. histolytica from Entamoeba gingivalis, and Giemsa stain may better define larvae and juvenile worms. Prepare a stain of material if organisms are found in examination of direct mounts which require additional differentiation. Although Cryptosporidium parvum will be difficult to see in a direct mount, examination of smears stained with modified acid-fast stains (hot or cold method) may provide confirmation of pulmonary cryptosporidiosis. In order to see microsporidial spores, centrifuged specimens stained with modified trichrome stains or optical brightening agents will be required (500 X g for 10 min).

Procedure

A. Wear gloves when performing this procedure.

B. Expectorated sputum (untreated with mucolytic agent)

1.   With a Pasteur pipette, place 1 or 2 drops (50 μL) on one side of a glass slide (2 by 3 in.), and cover with a no. 1 coverslip (22 by 22 mm).

2.   Place a second drop on the slide, add 1 drop of saline, and cover with a coverslip.

C. Material that has been treated with a mucolytic agent can be suspended in 100 μL of saline. Place 1 drop of the mixture on a slide (2 by 3 in.), and cover with a coverslip.

D. Reserve the specimen and remaining treated specimen for preparation of smears for staining should stains be required.

E. Examine the wet preparations field by field with low light and the 10X objective to detect eggs, larvae, oocysts, or amebic trophozoites.

F. If inconclusive, prepare smears of material for staining.

1.   Place 1 drop of sediment in the center of each of three glass slides (1 by 3 in.), and spread the material with the tip of the pipette.

2.   Place one slide in Schaudinn’s fixative while wet, and dry the other two thoroughly.

3.   Trichrome stain the slide fixed in Schaudinn’s fixative.

4.   Fix the air-dried smears in methanol. Stain one with Giemsa and the other with a modified acid-fast stain.

5.   Put immersion oil on stained smears, and examine Giemsa-stained smear with the 10X objective and trichrome-stained smear with the 50X oil objective if available. Otherwise, use the 100X oil objective.

Aspirates and Bronchoscopy Specimens

Principle

The examination of aspirated material for the diagnosis of parasitic infections is useful when routine specimens and methods have failed to demonstrate the organisms. Aspirates include liquid specimens collected from a variety of anatomic sites that delineate the types of organisms to be expected. Aspirates most commonly processed in the parasitology laboratory include fine-needle aspirates and duodenal aspirates. Fluid specimens collected by bronchoscopy include bronchoalveolar lavage (BAL) fluids and bronchial washings.

Specimens

A. Fine-needle aspirates

When specimens are collected and sent to the laboratory for processing, slides must be stained appropriately for suspected organisms and examined microscopically. Suggested stains are Giemsa and methenamine-silver nitrate for Pneumocystis carinii, Giemsa for Toxoplasma gondii, trichrome for amebae, and modified acid fast for Cryptosporidium parvum.

B. Aspirates of cysts and abscesses

Aspirates to be evaluated for amebae may require concentration by centrifugation, digestion, microscopic examination for motile organisms in direct preparations, and cultures and microscopic evaluation of stained preparations.

C. Duodenal aspirates

Aspirates to be evaluated for Strongyloides stercoralis, Giardia lamblia, or Cryptosporidium may require concentration by centrifugation prior to microscopic examination for motile organisms and permanent stains. In order to see microsporidial spores, centrifuged sediment stained with modified trichrome stains or optical brightening agents will be required.

D. Bone marrow aspirates

Aspirates to be evaluated for Leishmania amastigotes, Trypanosoma cruzi amastigotes, or Plasmodium spp. require Giemsa staining.

E. Fluid specimens collected by bronchoscopy

Specimens may be lavage fluids or washings, with BAL fluids preferred. Specimens are usually concentrated by centrifugation prior to microscopic examination of stained preparations. Organisms discussed here which may be detected are P. carinii, T. gondii, C. parvum, and the microsporidia.

Procedure

A. Wear gloves when performing this procedure.

B. Specimens that contain mucus may be treated with a mucolytic agent by adding a volume of agent equal to or one-half to two-thirds of the volume of specimen and incubating at room temperature for 15 min. Centrifuge at 1,000 X g for 5 min, and use sediment to prepare wet mounts and smears for staining.

C. Specimens that contain cell debris and proteinaceous material and that require digestion should be treated with streptokinase (1 part enzyme solution to 5 parts specimen) for 1 h at 35degree C. Shake at intervals (every 15 min). Centrifuge at 1,000X g for 5 min, and use sediment to prepare wet mounts and smears for staining.

D.  Specimens that include significant amounts of blood require treatment with an agent to lyse RBCs. Add 1 volume of lysing agent per volume of specimen, and incubate at room temperature for 5 min.

E. Place representative samples of untreated and Lyse-treated specimens in 15-ml conical centrifuge tubes, and centrifuge at 1,000X g for 5 min. For BAL or bronchial-washing specimens, which usually are 50 ml or more, place 20 to 24 ml of each specimen in a 50-ml conical centrifuge tube, and centrifuge as described above.

F.   Decant supernatants from centrifuged samples into a disposal container containing disinfectant.

G. With a Pasteur pipette, remove drops of sediment for wet mounts, stain preparations, and culture.

H. For duodenal aspirates and aspirates from cysts or abscesses, place 1 drop of sediment on a glass slide (2 by 3 in.), add a drop of 0.85% NaCl, and cover with a no. 1 coverslip (22 by 22 mm).

I.    Examine preparation field by field with low light until the entire mount has been examined.

J.   If the wet mount is equivocal for a protozoan, place a drop of sediment on a glass slide (1 by 3 in.), and add a drop of polyvinyl alcohol (PVA) fixative. Mix the drops with a pipette, and spread the mixture into an even film (about 22 by 22 mm). Dry the preparation thoroughly, and stain with trichrome stain.

K. For material from cysts or abscesses, prepare cultures by adding 0.5 ml of material to a tube of culture medium for the recovery of amebae.

L. Aspirates of bone marrow may be submitted for diagnosis of leishmaniasis, trypanosomiasis, and occasionally for malaria. Material should be stained with Giemsa stain and examined with the 100X oil immersion objective.

M. Sediments from BAL or bronchial washings are examined in stained preparations. Three slides should be stained.

1. Use a Pasteur pipette to place drops of sediment from each specimen on at least four slides. With the pipette, spread the sediment into a thin, even film. For specimens treated with an agent to lyse RBCs, use sediment from both treated and untreated samples for smears.
2. Air dry slides, and fix in methanol. If slides are to be stained with immunospecific stains, fix according to package instructions.
3. Stain one slide with rapid Giemsa stain. Rapid Giemsa stain procedure

a.       Stain solutions should be kept in dropper bottles to avoid bacterial contamination. Place 1 or 2 drops of red stain solution 1 on specimen smear and control slide, hold for 10 s, and drain.

b.      Add 1 or 2 drops of blue solution 2, hold for 10 s, drain, and rinse very briefly with deionized water.

c.       Stand slides on end to drain and air dry.

d.      Slides must be examined with oil or mounted with mounting medium.

   4.      Stain one slide with modified acid-fast stain.
   5.      Stain one slide with methenamine-silver nitrate or other cyst wall stain.

Biopsy Specimens

Principle

Biopsy specimens are recommended for the diagnosis of tissue parasites. The procedures that may be used for this purpose in addition to standard histological preparations are impression smears and teased and squash preparations of biopsy tissue from skin, muscle, cornea, intestine, liver, lung, and brain. Tissue to be examined by permanent sections or electron microscopy should be fixed as specified by the laboratories which will process the tissue.

Specimens

Tissue submitted in a sterile container on a sterile sponge dampened with saline may be used for protozoan cultures after mounts for direct examination or impression smears for staining have been prepared. If cultures for parasites will be made, use sterile slides for smear and mount preparation.