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Para-Fix™: PVA

Para-Fix™: PVA

Intended Use

The Para-Fix™ stool collection kits provide a standardized method for untrained personnel to properly collect and preserve stool specimens for the detection of helminth larvae and eggs, protozoan trophozoites and cysts, coccidian oocysts, and microsporidian spores. A ten language instruction sheet is provided to assist patients or healthcare professionals with the proper use of the kits at home or in the healthcare setting.

Summary and Explanation

The diagnosis of intestinal parasitic infection is confirmed by the recovery of helminth larvae and eggs, protozoan trophozoites and cysts, coccidian oocysts, and microsporidian spores. The ability to detect and identify intestinal parasites in fresh stool specimen depends on immediate collection, transportation, and examination by the laboratory; often, these time requirements from passage of the specimen to examination cannot be guaranteed. Brooke and Goldman, in 1949, described the use of PVA as a means of preserving intestinal protozoa. In 1981, Horen described substituting cupric sulfate for mercuric chloride in the preparation of Schaudin’s fixative for use in preparing PVA fixative. Results obtained with this method do not have the consistency required for routine laboratory work. In 1992, Medical Chemical Corp. substituted zinc sulfate for the mercuric chloride in the preparation of it’s Z-PVA fixative. Although mercuric chloride based PVA provides the best fixation, tests with Z-PVA have shown it to be a viable alternative, and far more effective than copper sulfate based PVA.

Kit Contents

  1. 1 vial (15ml) PVA or Z-PVA
  2. 1 vial clean or (15ml) 10% formalin
  3. 1 10-language instruction sheet

Materials Not Provided

  1. ethyl acetate, saline
  2. zinc sulfate solution
  3. applicator sticks and cotton tipped applicator
  4. transfer pipets
  5. centrifuge
  6. microscope, slides and cover slips
  7. trichrome reagents

Specimen Collection

    1. Collection of fecal specimens for intestinal parasites should always be performed prior to the use of any antacids, barium, bismuth, antidiarrheal medication, or oily laxatives.
    2. For routine examination for parasites prior to treatment, a minimum of three specimens, collected on alternate days, is recommended. Two of the specimens should be collected after normal movements, and one after a cathartic, such as magnesium sulfate or fleet Phospho-Soda. If the patient has diarrhea, do not use a laxative.
    3. Fecal specimens should be collected in a clean, dry wide mouthed container. A waxed, cardboard half-pint container with a tight-fitting lid is ideal, however, a clean, dry milk carton with the top two thirds removed is acceptable. Contamination with urine should be avoided.
    4. Small samples of the specimen should be placed into the vial, using the spork built into the lid of the vial. Pay particular attention to the areas that appear bloody or watery. Add samples until the fluid level reaches the red “fill line”. This will insure the appropriate three to one ratio of fixative to sample.
    5. Use the spork to mix the contents of the vial. Recap the vial, making sure the lid is securely fastened. Firmly shake the vial until contents are thoroughly mixed (the solution should appear homogeneous).
    6. Fill out the patient information on the side of each vial. Reseal the vials in the plastic bag.

Caution: Every sample should be treated as a potential source of contamination.

Specimen Examination

The use of the kits allows for a wide variety of examination procedures including gross examination (only if the kit contains a clean vial), direct microscopic examination, concentration procedures, and permanent staining.

Macroscopic examination
Examine the contents of the clean vial (unpreserved specimen) and record the consistency of the specimen, the presence of worms or proglottids, and blood if present.

Microscopic examination

  • Direct Wet Smears: The purpose of this procedure is to demonstrate trophozoite motility. Prepare the smear by mixing a small amount of fecal material (approximately 2 mg) with a drop of physiologic saline or D’ Antoni’s iodine on a glass slide. Cover with a 22 by 22 mm coverslip. Examine immediately using the low power objective (10X). Suspect objects may be examined using the high-dry objective (40X).
  • Permanent stained smears:
    Preparation of smears for permanent stains:

    1. Using two applicator sticks, mix the contents of the PVA, or Z-PVA vial.
    2. Pour approximately 0.5 to 1 ml of the vial contents onto a paper towel, or remove a small portion of fecal material with the applicator sticks or lid spork. Place sample on a paper towel for approximately 2 to 3 minutes to absorb the excess PVA. Smears made from solid (lumpy) portion of the preserved specimen will adhere better than smears made from the watery portion.
      Note: For liquid specimens or those containing an insufficient amount of fecal material, centrifuge the sample at approximately 500xg for about 10 minutes. Make the smear from the solid pellet at the bottom of the tube. This approach can also be used with specimens containing a lot of mucus.
    3. Using the applicator sticks, apply some of the stool material from the paper towel onto one or more glass slides and let dry for 2 hours to overnight at room temperature, or 30 minutes to 1 hour at 37 degrees C. For best adherence, spread the sample from edge to edge on the slide. If newsprint is not visible through the slide, the smear may be too thick.

Recommended procedure for staining with Wheatley’s Modified Gomori Trichrome:

    1. Place in 70% ethyl alcohol plus D’Antoni’s iodine (strong tea color) for 2 to 5 minutes.
    2. Place in two changes of 70% alcohol: 5 minutes in the first and 2 to 5 minutes in the second.
    3. Place in trichrome stain for 10 minutes.
    4. Dip twice in 90% ethanol with 0.5% acetic acid. Substitute with 90% ethanol if slide appears pale.
    5. Place in two changes of 100% alcohol for 2 to 5 minutes.
    6. Place in two changes of xylene or xylene substitute for 5 to 10 minutes.
    7. Mount with mounting medium using a #1 thickness cover glass.

                    Important Note: For slides prepared in non-mercury fixatives (Cu-PVA or Z-PVA), steps 1 and 2 may be omitted.

Concentration Procedures

There are a number of concentration techniques available designed to separate the parasitic components from the excess fecal debris through differences in specific gravity.

Sedimentation Procedure – ethyl acetate technique:
Mix contents of either the 10% formalin, Z-PVA, or PVA vial thoroughly.

1. Place one layer of pressed gauze or two layers of woven gauze in a funnel. Strain approximately half of the vial contents through the gauze into a 15 ml centrifuge tube. Add saline until the level in the tube is almost at the top and centrifuge at 500xg for 10 minutes.

2. There should be approximately 1 ml of sediment in the tube. If not, resuspend sediment and add or remove as necessary and centrifuge. Decant. If supernatant is not light tan or clear, a second wash with saline is optional. Over manipulation of the specimen can cause a loss of organisms.

3. Resuspend the sediment on the bottom of the tube with saline*, filling the tube half full. Add approximately 3 ml of ethyl acetate or ethyl ether and stopper. Hold the tube so that the stopper is directed away from your face and shake vigorously for 30 seconds.

4. Carefully remove the stopper. The resulting solution should have four layers:

Top: ethyl acetate or ethyl ether
Second: debris plug
Third: saline *
Fourth: sediment

5. Loosen the plug of debris with an applicator stick and decant all the fluid. While the tube is still inverted, ring debris from the sides of the tube with one or two cotton tip applicators. This will remove ethyl acetate or debris left behind.

6. Resuspend the remaining sediment with a few drops of saline*. Use either saline or iodine mounts for microscopic examination.

Flotation technique – zinc sulfate:

  1. Transfer a portion of the formalin preserved or fresh specimen to a 15 ml centrifuge tube. Fill to within 2 to 3 ml of the top with saline.
  2. Centrifuge for 10 minute at 500xg.
  3. The sediment should be approximately 1 ml in volume; if not, adjust with saline* or specimen and centrifuge. Decant the supernatant fluid.
  4. Half fill the tube with zinc sulfate solution sp. gr. 1.18 (for formalin preserved specimens adjust the specific gravity of the zinc sulfate to 1.20) and resuspend the sediment with an applicator stick.
  5. Add enough zinc sulfate solution to fill the tube to within 2 to 3 ml of the top.
  6. Centrifuge at 500xg for 1 minute.
  7. Without removing centrifuge tube from centrifuge, remove 1 or 2 drops from the center film with a freshly flamed and cooled wire loop. The loop should be bent to form a 90ç angle, so it is parallel to the surface of the fluid. Do not dip the wire loop below the surface when collecting the drops from the centrifuge tube. Transfer the contents of the loop to a microscope slide.
  8. Add iodine to the preparation if desired.
  9. Add cover glass and examine under microscope.

Alternate method:

    1. After step 6, remove centrifuge tube from centrifuge and place in test tube rack, avoiding agitation.
    2. Carefully fill tube to the top with zinc sulfate solution. Do not overfill.
    3. Place a clean cover slip on the centrifuge tube, if the cover slip does not contact the meniscus, carefully add zinc sulfate solution until it does.
    4. Allow tube to stand for 10 minutes.
    5. Remove the coverslip with a quick motion directly upward, so that a drop of liquid remains on the coverslip.
    6. Place the coverslip on a clean slide. If an iodine mount is desired, place the coverslip on a slide containing a drop of iodine.

* 5% or 10% formalin may be used in place of physiological saline.


  1. Ethyl acetate and diethyl ether are flammable. Use in a well ventilated area. Keep away from direct flame. Avoid contact of the solution with skin and eyes. Should contact occur flush with running water. Avoid breathing fumes.
  2. Every sample should be treated as a potential source of infection. Good laboratory practice should be followed at all times. The use of gloves and hand washing is recommended.


The expiration date of each kit is printed on the outer label. The expiration dates of each vial are printed on the individual vial label. The kits should be stored at room temperature. If the PVA and Z-PVA vials are exposed to freezing temperatures for an extended period of time they will gel. If the solutions are restored to a liquid state, there will be no change in performance. To restore the solutions, heat to between 40 – 50 degrees C for 30 minutes.


  1. Bartlett, Marilyn, et al, 1978. Comparative evaluation of a modified zinc sulfate floatation technique. J Clin Microbiol 7:524-528.
  2. Brook, M. M. and M. Goldman, 1944. Polyvinyl alcohol fixative as a preservative and adhesive for protozoa in dysenteric stools, and other liquid materials. J Lab and Clin Med 34:1554-1560.
  3. Erdman, Dean, 1981. Clinical comparison ethyl acetate and diethyl ether in the formalin-ether sedimentation technique. J Clin Microbiol 14:483-485
  4. Garcia, L.S., 2001. Diagnostic Medical Parasitology, 4th ed.; ASM Press: Washington D.C.
  5. Horen, W.P., 1981. Modification of Schaudin’s fixative. J Clin Microbiol 13:204-205.
  6. Young, Kirk H., et al, 1979. Ethyl acetate as a substitute for diethyl ether in the formalin-ether sedimentation technique. J Clin Microbiol 10:852-853.