Index
Live/Dead
Fluorescence Assay
Objectives
· Assess the impact of a toxic material on living cells.
· Become familiar with fluorescent microscope.
Materials
TC-treated 24-well plates
Fibroblasts cells (HDF)
DMEM with 10% serum and 1% antibiotic
PBS
Ethanol
Materials as required by cell passaging procedure.
Live/Dead reagent stock solutions: Calcein AM, Ethidium Homodimer (EthD-1)
Safety
1. Wear disposable latex gloves at all times
2. Wear safety glasses at all times.
3. Ethanol is flammable. Do not place near open flame or heat.
Test Conditions
A. Live cells, 3 wells
B. Dead cells, 3 wells
C. Mix of live and dead cells, 3 wells
Seed Cells (Day 1)
NOTE: One partner can do steps 1. - 2. for both partners.
1. Before harvesting cells for experiment, ensure that cells are confluent.
2. Proceed through cell passaging protocol until step #15. After centrifugation, resuspend the cells in 10 mL DMEM with 10% serum and 1% antibiotic.
NOTE: For steps 3. - 5., each partner should do her/his own work on her/his own TC-treated 24-well plate.
3. Dilute part of your cell suspension 1:3 in complete media by adding 3.5 mL cells to 7 mL complete media.
4. Seed 1.0 mL of this diluted cell suspension into each of 9 test wells on the TC-treated plates. When seeding, mix the cells gently and often (every 4-5 wells) to prevent settling of the cells.
5. Place cells in incubator for two nights. Cells will firmly attach to the surface during this time.
1. Remove Live/Dead reagent stock solutions from the freezer and allow them to warm to room temperature.
2. Add 20 mL of the supplied 2 mM EthD-1 stock solution to 10 mL sterile PBS in 15 mL centrifuge tube. Vortex to ensure mixing. (Resulting solution is 4 mM EthD-1.)
3. Add 5 mL of the supplied 4 mM calcein AM stock solution to the 10 mL EthD-1 solution. Vortex to ensure mixing. (Resulting solution is 2 mM calcein AM.)
4. Store with foil wrap. Use within one day.
Stain with Live/Dead Reagent (Day 3)
NOTE: For steps 1. - 10., each partner should do her/his own work.
1. (5 pts) Observe cells under light microscope. Note confluency and morphology.
2. Aspirate media from the wells.
3. Rinse wells with PBS using the following procedure:
· Add 1 mL of PBS to each well using a micropipette.
· Swish lightly.
· Aspirate PBS using glass Pasteur pipet.
· Repeat.
NOTE: When adding PBS to each well, squirt the liquid on side wall of well (where there are no attached cells) rather than in center of well (where there are cells)!
NOTE: Tilt plate slightly to assist in pooling liquid for aspiration.
NOTE: To streamline procedure, aspirate all wells, then add PBS to all wells, etc.
4. For Condition A, add 250 mL PBS and 100 mL dye to each of three wells.
5. For Condition B, add 250 mL ethanol and 100 mL dye to each of three wells.
6. For Condition C, add 250 mL PBS, 2 drops of ethanol, and 100 mL dye (in that order) to each of three wells.
7. Cover plate completely with foil. Incubate cells for 30 min at room temperature.
8. (10 pts) Observe cells under light microscope. Record pattern and morphology of cells.
9. (15 pts) Observe cells under fluorescence microscope. See instructions on Fluorescent Microscopes (p. 43-44). Record color, pattern, and morphology of cells. Live cells are seen as green through the filters. Dead cells are seen as red through the filters.
10. Dispose of biohazardous material properly. Aspirate the remaining liquid using a glass Pasteur pipet in the hood. NOTE: SAVE YOUR 24-WELL PLATE FOR USE IN THE PCNA STAINING STUDY. PUT IT BACK IN THE INCUBATOR!
Questions
1. (10 pts) How does ethanol effect HDF cells?
2. (10 pts) What advantages do viewing cells using a live/dead assay have over viewing cells under standard light microscopy?
Observations and answers to Questions should be recorded in your laboratory notebook. The point values are listed above.