DNA Content for Cell Cycle Analysis of Fixed Cells With Propidium Iodide

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DNA Content for Cell Cycle Analysis of Fixed Cells With Propidium Iodide

Adapted from Current Protocols in Cytometry
 
This protocol uses ethanol to fix and permeabilize cells for staining of DNA in intact cells with propidium iodide (PI). PI staining solutions provided are a reasonable starting point for concentrations of fluorochrome, however, this will vary with cell type and cellular state (accessibility of DNA binding sites). 
 
Since the binding of the fluorochrome is not covalent and is reversible, it is highly recommended that you establish the optimal concentration to reach equilibrium for saturation of DNA binding sites of your cells. You have reached saturating conditions when you do not see any additional increases in the mean channel of your G0/G1 population and you have achieved the lowest CV possible of your G0/G1 population (ideally < 5%). It is also critically important to keep the ratio of dye:cells relatively constant, especially if you plan to compare the fluorescence histogram between samples. Yes, this means you have to count your cells every time prior to PI staining!! (For a more detailed explanation of why this is important click here).
 

Materials

70% ethanol
Cells to be stained
Phosphate-buffered saline (PBS)
Propidium iodide (PI) staining solution; freshly made (see recipe below)
12X75 mm centrifuge tubes (preferably polypropylene)

PI staining solution (with Triton X-100 and RNAase)

To 10 ml of 0.1% (v/v) Triton X-100 (Sigma) in PBS add 2 mg DNase-free Rnase A (Sigma) and 200uL of 1 mg/ml PI **(Sigma, BioSure, Molecular Probes, etc.). Prepare freshly! A stock solution of PI, made by dissolving 1 mg PI in 1 ml DH2O, can be stored for several months at 0 - 4°C.
**-These concentrations are a recommended starting point. Saturating concentrations should be determined for specific cell types/numbers.
If the Rnase is not DNase-free, boil a solution of 2 mg Rnase A in 1 ml water for 5 min.

Fixing cells with ethanol

  1. Prepare the fixative by filling 12X75 mm tubes with 4.5 ml of 70% ethanol. Keep tubes on ice.
  2. Collect cells and suspend 106 to 107 cells in 5 ml PBS in a centrifuge tube.
  3. Centrifuge cells for 6 min at ~300 x g.
  4. Using a Pasteur pipette, thoroughly resuspend cells in 0.5 ml PBS. It is very important to achieve a single cell suspension. Fixation of cells that are in aggregates while suspended in PBS stabilizes the aggregates, which are then impossible to disaggregate. It is essential, therefore, to make sure cells are in a single cell suspension prior to the time of mixing the cells with ethanol!
  5. Transfer the cell suspension into the tubes containing the 70% ethanol. Keep cell in fixative for at least 2 hours. Cells suspended in ethanol can be stored at 0-40°C for several months to a year.

 

Staining cells with PI

  1. Centrifuge the ethanol fixed cells 5 min at 300 x g and decant ethanol thoroughly (be careful not to lose your cells!).
  2. Suspend the cell pellet in 5 ml PBS, wait 60 sec, and centrifuge 5 min at 200 x g.
  3. Suspend the cell pellet in 1 ml PI staining solution that has been optimized for your cell type and concentration. Keep at either 37°C for 15 min or 30 min at room temperature.
  4. Bring cells in PI solution to the flow lab for analysis.