Call Us

Local: (310) 787 - 6800
Outside The Area: (800) 424 - 9394
(800) 252 -1125

Para-Fix™: SAF

Para-Fix™: SAF

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.

Junod, in 1972, described the use of SAF (sodium acetate-acetic acid-formalin) as a multipurpose means of preserving intestinal parasites. Scholten and Yang, in a study of more than 900 specimens, confirmed the suitability of SAF for routine use in the clinical parasitology laboratory.

Proper use of the SAF kits assures the preservation and proper identification of all stages of intestinal parasites. SAF allows permanent stains and concentration procedures to be performed on a single sample. The clean vial is used for the examination of stool fat, occult blood, enteric or amoeba culture in an unpreserved specimen.

Kit Contents

  • 1 vial (15ml) SAF
  • 1 vial clean or (15ml) 10% formalin
  • 1 10-language instruction sheet

Materials Not Provided

  • ethyl acetate, saline
  • applicator sticks and cotton tipped applicator
  • transfer pipets
  • centrifuge
  • microscope slides and cover slips
  • microscope

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).

Concentration and permanent stain procedure
Mix contents of the SAF 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. Mix the sediment and prepare a slide as follows:

4. Place a small drop of Mayer’s egg albumin (supplied with each case of SAF) on a glass slide and wipe so that a thin coating remains. Note: excess albumin on the slide will cause a reddish tinge after decolorization.

5. Place a small sample of the suspended sediment on the albumin coated slide. Spread the sample over the slide to prepare a thin smear that varies in thickness. Allow to dry at room temperature (smear will appear opaque when dry).

6. Proceed with staining method of choice. We recommend the iron hematoxylin method, although the Gomori trichrome method is also used. Proceed with the concentration procedure using the remaining sediment.

7. Resuspend the sediment on the bottom of the tube with 10% formalin, 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.

8. Centrifuge at 500 xg for 10 minutes.

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

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

10. 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.

11. Resuspend the remaining sediment with a few drops of saline or 10% formalin. Use either saline or iodine mounts for microscopic examination.

If using a Para-Fix™ Para-Sed™ use the following procedure. If using Sed-Connect™ or Micro-Sed™ follow the procedures provided with those items.

      1. Thoroughly mix the contents of the SAF vial. Proceed with the specimen processing instructions in the Para-Sed™ kit instruction sheet. Insert the following procedure between steps 5 and 6:
      2. Bring the liquid level in the vial to fill line on Para-Sed™ tube with physiological saline.
      3. Place the screw cap provided with the Para-Sed™ on the conical centrifuge vial and centrifuge at 500X for 10 minutes.
      4. Carefully pour off supernatant.
      5. Add a small drop of saline and mix the sediment with an applicator stick. Prepare a slide as outlined in steps 4 to 6 in the previous section.
  • Permanent stained smears with iron hematoxylin:
      1. Place slide in 70% alcohol for 5 minutes.
      2. Wash in tap water for two minutes.
      3. Place in Kinyoun stain (MCC Cat#483A) for 5 minutes.*
      4. Wash in running tap water for 1 minute.*
      5. Place slide in acid-alcohol decolorizer for 4 minutes.**
      6. Wash slide in running tap water for 1 minute.*
      7. Place in iron hematoxylin (MCC Cat# 6185A and 6188A) working solution for 8 minutes.
      8. Wash slide in distilled water for 1 minute.
      9. Place slide in picric acid 0.6% (MCC Cat.# 733A) solution for 3 to 5 minutes.
      10. Wash slide in running tap water for 10 minutes.
      11. Place slides in 70% alcohol plus ammonia for 3 minutes.
      12. Place slides in 95% alcohol for 5 minutes.
      13. Place slides in 100% alcohol for 5 minutes.
      14. Place slides in two changes of Xylene (or Xylene substitute) for 5 minutes.
      15. Mount with mounting media using #1 thickness coverglass.

* If the lab is not looking for cryptosporidium these steps may be omitted.

** This step can be performed as follows:

  1. Place the slide in acid-alcohol decolorizer for 2 minutes.
  2. Wash the slide in running tap water for 1 min.
  3. Place the slide in acid-alcohol decolorizer for 2 minutes.
  4. Wash the slide in running tap water for 1 minute.
  5. Continue with step 7 of staining procedure.
  • Permanent stained smears with Wheatley’s Gomori Trichrome:
    1. Place the prepared slide in 70% ethyl alcohol for 2 to 5 minutes.
    2. Place in trichrome stain for 10 minutes.
    3. Dip twice in 90% ethanol with 0.5% acetic acid or with 90% ethanol if slide appears pale.
    4. Place in two changes of 100% alcohol for 2 to 5 minutes.
    5. Place in two changes of xylene or xylene substitute for 5 to 10 minutes.
    6. Mount with mounting medium using a #1 thickness cover glass.


  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. Avoid contact of SAF solution with skin or eyes. If contact occurs, flush effected area with water. If irritation develops contact a physician immediately.
  3. SAF solution is poisonous. If ingestion occurs, drink milk or water. Contact a physician immediately.
  4. 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 SAF vials are exposed to freezing temperatures for an extended period of time they will freeze. If the vials are restored to room temperature, there will be no change in performance.


  1. Brooke, M.M., 1974. “Intestinal and Urogenital Protozoa”, Manual of Clinical Microbiology, ASM, Washington, D.C., Second Edition, 582-601.
  2. Garcia, L.S., 2001. Diagnostic Medical Parasitology; 4th Edition. ASM Press: Washington D.C.
  3. Junod, L. 1972. Technique Coprologique Novelle Essentiellement Destinee a la Concentration des Trophozoites d’Amibes. Bul. Soc. Pathol. Exot., 65:390-398.
  4. Scholten, T., 1972. An Improved Technique for the Recovery of Intestinal Protozoa. J. Parasitol. 58:603-634.
  5. Yang, J., and Scholten, T., 1977. A Fixative for Intestinal Parasites Permitting the Use of Concentration and Permanent Staining Procedures. Am. J. Clin. Pathol., 67:300-304.