Protein Silver Staining of Intestinal Flagellates
OBJECT
This staining method will allow the student to differentiate the rather exotic intracellular morphology of flagellates found in the frog rectum or termite intestine. The student will become familiar with chemical preparation and the discipline necessary to perform this rather laborious stain technique. Another object of this project is to familiarize the student with the morphology and identification of the different organisms found in the frog rectum and termite intestine. I was taught this technique by Felix H. Lauter, many years ago in graduate school.
METHODS AND MATERIALS FOR PROTEIN-SILVER IMPREGNATION
Materials
live Rana pipiens and termites
dissecting tools and trays
microscopes, slides and cover slips
Coupling jars of Columbia staining dishes
incubator
albumen slide fixative
Hollande's fixative
ethanol-absolute, 95%,70%, 50% and 30%
distilled water
xylene
0.25% potassium permanganate
2.0 % oxalic acid
1.5% protargol
copper wire
1% hydroquinone
0.5 % gold chloride
5% sodium thiosulfate
permount
slide labels
applicator sticks
glacial acetic acid
PROCEDURE
1. Coat a meticulously-clean glass
slide or cover slip with a minute amount of albumen fixative (Mayer). I
seem to have better success working with cover slips but they obviously are more
fragile and prone to breakage.
2. Add a concentrate of rectal contents or intestinal
contents of either frog flagellates or termite flagellates to the coated slide
and allow to partially air dry. This is a critical step because the
majority of the flagellates will eventually wash off the slide in the numerous
solution transfers. The drying will occur from the outer edge of the smear
inwardly. Allow the outer edge to dry before you proceed. You should have
enough Coupling/Columbia jars prepared and labeled for each of the transfers.
Each of the following steps will involve moving the slides/cover slips from jar
to jar and therefore solution to solution. The alternative of dumping the
previous solution and adding the next solution in the same jar is too disruptive
and will eventually wash all surface content off the slide/cover slip.
3. Fix in Hollande's fixative for 20 minutes at room
temperature in a Coupling jar (if your smear is on a slide) or a Columbia
staining dish (if your smear is on a cover slip). Each can hold four slides or
cover slips. Make sure to mark one side of the jar or dish so you can
remember which side of the slide/cover slip the smear is on. I have had
students get to the end of this laborious technique, only to wipe the smear off
rather than the opposite side.
4. Wash in two changes of distilled water to remove the
yellowish tinge (excess picric acid). The picric acid used in Hollande's
is potentially explosive if allowed to dry, so be careful.
5. Transfer the slide/cover slip to 50% ethanol for 2
minutes.
6. Transfer the slide/cover slip to 30% ethanol for 2
minutes.
7. Transfer sequentially through 3 changes of distilled
for water for a total of 2 minutes.
8. Bleach in 2.0% aqueous oxalic acid for 10 minutes.
9. Wash in tap water for 2 minutes (3 changes).
10. Place in fresh 1.5% protargol; this is made by
sprinkling the protargol powder on the surface of distilled water in a clean
beaker and allowing to dissolve overnight. Do not heat or stir and place
the beak in the dark.
11. Add 5 gm of copper wire per 100 ml of protargol to
the staining jar/dish before use.
12. Stain for 33 hours, changing the staining solution
after 24 hours (including the addition of fresh protargol and copper). The
staining time has been determined by trial and error.
13. Wash in distilled water for 30 seconds.
14. Place in a solution of 1% hydroquinone in 5%
aqueous sodium sulfite for 10 minutes.
15. Wash several times in distilled water (3 changes of
3, 3, and 2 minutes).
16. Place in 0.5% aqueous gold chloride for 5 minutes.
17. Wash in distilled water (2 changes, totalling three
minutes).
18. Place in 2% oxalic acid for 5 minutes until a
purplish hue appears on the smear.
19. Wash several times in distilled water (4 changes,
total of 10 minutes).
20. Place in 5 % sodium thiosulfate for 10 minutes.
21. Wash several times in distilled water (4 changes
for a total of 10 minutes).
22. Dehydrate through a graded series of ethanols, 30%.
50%, 70%, 95% and absolute for 2 minutes each.
23. Clear in xylene for 1 minute. The xylene
should be used in a hood since it is toxic and carcinogenic.
24. Mount thinly with permount. Make sure you
mount the smear down. I have known students to mistakenly have turned the
slide or cover slip around during the various transfers and ended up mounting
the smear on top, where it is quickly rubbed off.
DISCUSSION
Obviously, this stain technique is not for the inexperienced student, nor the easily-frustrated person. There are quite a few pitfalls along the way but the presentation of flagella, parabasal bodies, kinetoplasts, axostyles and other exotic morphology of these flagellates is without comparison. Proper precautions should be taken working with the above chemicals, some of which are toxic or dangerous. Students or lab workers should read all chemical bottle labels and take proper precautions.