FREEZING AMPHIBIAN TISSUE BIOPSY SAMPLES FOR LATER
INITIATION OF CELL CULTURE
Contributed by San Diego Zoo's Institute for Conservation Research, Genetics Division
Materials and Reagents Needed
• sterile glass plate or petri dish
• biopsy vials (sterile)
• growth medium (supplemented with 10% FBS, 1% glutamine, 1% pen-strep
fungizone)
• dimethylsulfoxide (DMSO)
• 15- or 50-cc centrifuge tubes
• Hank's balanced salt solution (HBSS) supplemented with 1% pen-strep fungizone
• sterile pipets, 5- and 10-cc, pipettors or pipet bulbs
• sterile forceps and scalpel
• cryo freezing vials (~2cc capacity)
• liquid nitrogen
Protocol (use sterile technique throughout)
1. Take biopsy sample as described in Skin Biopsy Procedure). A 6mm 2 biopsy will
provide enough material for 1 frozen vial.
2. Prepare freezing medium with 10% DMSO in centrifuge tube: Make enough to
allow for 1 cc of freezing medium per vial of tissue to be frozen. Keep on ice or
refrigerated.
3. On a sterile glass plate (or petri dish) mince tissue biopsy into tiny fragments
(1mm2) with sterile forceps and scalpel. Do not let fragments dry out; keep
moistened with media.
Note: If biopsy sample was not adequately prepared at initial sampling, it may be
necessary to rinse piece in HBSS or medium and remove any excessive amount of
hair, fur, etc. This usually needs to be done if working from ear notch samples.
4. Gather tissue pieces with forceps and carefully place into sterile freezing vial.
5. Add 1 ml cold freezing medium to each vial. Cap carefully but securely, making
sure that silicone gasket does not bulge out. If capped too tightly, liquid nitrogen
will seep inside and cause contamination and/or explosion upon thaw. Gently
agitate cryovial to expose all tissue pieces to the freeze medium.
6. Label vials with appropriate information, using water- and alcohol-resistant
marker.
7. Place directly into liquid nitrogen for storage.
To Thaw (use sterile technique throughout)
1. Quick thaw sample in 37° C waterbath. Wipe the outside of the vial with alcohol
drenched gauze. Transfer contents of vial to a 15-cc centrifuge tube containing 10
cc of HBSS.
2. Centrifuges at 1000 RPM for 10 min.
3. Remove as much supernatant as possible. If the pieces look adequately diced,
proceed with step 5. If the tissue pieces look larger than reasonable for adequate
digestion, remove the washed tissue to a sterile petri dish and dice the pieces
smaller.
4. Using scalpel and forceps, cut the tissue into small squares (approximately
0.5mm3). Smaller pieces will digest more rapidly than larger pieces. Use clear
cuts. Do not shred or tear the tissue.
5. Gather approximately 10-20 pieces together in a clump. Using the forceps, push
the pieces to the bottom of a 15-cc conical tube. It is easiest to do this with the
pieces in a clump rather than individually.
6. Add approx. 0.3 - 0.5 cc of 0.5% collagenase to the pieces so they are completely
covered.
7. Leave tube at room temperature.
8. About once each hour, gently agitate the tissue pieces in the tube being careful
not to let any pieces stick to the side of the tube above the collagenase.
9. After 3-6 hrs the tissue pieces should look "chewed" (crisp edges become fuzzy
looking) and the collagenase liquid becomes cloudy with cells and tissue debris.
10. When the tissue looks sufficiently digested, add 5 cc of media to the tube and
pipette the digested tissue pieces and cell pellet in and out of a 5-cc pipet to
dislodge any cells that are loosely attached to the tissue. Place entire contents
of the tube into a T25 flask.
11. Incubate the T25 flask at 27° C, 6% CO2, humidified.
12. Leave the flask undisturbed for 2-3 days, then examine under the microscope for
small cell patches (there may be only a few patches of 3-10 cells). If a few or no
cell patches are seen, replace half the media and observe again in a few days. If
many patches are seen, feed as usual.
13. When outgrowth is sufficient the cells must be "passed" to a larger flask
(secondary culture). Remove the medium. Rinse with 4 cc Hank's balanced salt
solution (HBSS). Remove HBSS. Add 4 cc Trypsin-EDTA and swirl around flask
to coat. Remove all but 1 cc Trypsin-EDTA. Place flask on warming tray set at
37° C until cells have detached.
14. Add 10 cc medium to the flask and swirl around to get all the cells into the
medium. Transfer to a T75 culture flask. Refeed the 1° cultures. Incubate both
1° and 2° cultures. Secondary cultures should be checked every other day and
fed accordingly.
15. Cultures may be passed indefinitely, frozen when sufficient numbers of 2° flasks
have reached confluency, or harvested when active growth is most evident.
NOTE: Tissue that has been frozen in this manner tends to take longer to
initiate growth and may also in general be a slow growing culture.