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Chapter 12: Cell Cultures

Exercise 12.10 - Establishment of a Primary Culture



Procedure 8

  1. Candle an 8 day old egg to ensure that it is alive. This is easily accomplished by holding the egg in front of a bright light source; the embryo can be seen as a shadow. Circle the embryo with a pencil.

  2. Place the egg in a beaker with the blunt end up, and wash the top with a mild detergent, followed by swabbing with ethanol.

  3. Carefully puncture the top of the egg with the point of a pair of sterile scissors and cut away a circle of shell, thus exposing the underlying membrane (the chorioallantois).

  4. With a second pair of sterile scissors, carefully cut away and remove the chorioallantoic membrane, exposing the embryo.

  5. Identify and carefully remove the embryo by the neck, using a sterile metal hook or a bent glass rod, and place the embryo in a 100mm petri dish containing phosphate buffered saline (PBS). Wash several times with PBS by transferring the embryo to fresh petri plates. After removal of all yolk and/or blood, move the embryo to a clean dish with PBS.

  6. Using two sterile forceps, remove the head, limbs, and viscera. Be sure to remove the entire limb by pulling at the proximal end. Move the remaining tissues of the embryo to yet another dish and wash with PBS.

  7. Mince the embryo finely with scissors and transfer the minced tissue to a flask containing PBS. Allow the tissue pieces to settle.

  8. Remove the PBS with a sterile pipette and add 25 ml of trypsin, a proteolytic enzyme. Stir the solution gently at 37° C for 15-20 minutes.

  9. Allow the larger, undigested tissue pieces to settle and decant the supernatant into an equal volume of Minimal Essential Medium (MEM) + 10% Fetal Calf Serum (FCS). FCS contains protease inhibitors which will inactivate the trypsin.

  10. Centrifuge the cells in MEM at 1000 rpm for 10 minutes in a standard clinical centrifuge. Remove the supernatant and resuspend the pellet in 25 ml of fresh MEM + 10% FCS.

  11. Remove 0.1 ml of the culture and determine cell concentration and viability as directed in the previous section.

  12. Seed two 25 cm^2 plastic culture flasks containing 25 ml of MEM + 10% FCS to a final concentration of 10^5 cells/ml.

  13. Label and place your cultures in the tissue culture incubator at 37° C and examine daily for cell density and morphology.

  14. Note any changes in the color of the media. Tissue Culture media has a pH indicator (Phenol Red) added in order to check on the growth of cells. The media initially is a cherry red (with slight blue haze) and turns orange and then yellow as the cells grow, thereby reducing the media. Should this color change occur within 24 hours, the culture is most likely contaminated and should be disposed of.

  15. Examine the cultures using an inverted phase contrast microscope. This will allow observation of the cells without opening or disturbing the growth.

  16. Make cell density determinations at 10 X magnification using a square ocular grid, as explained in Chapter One for the determination of area.

  17. Plot the cell density on a log scale vs. time of culture.

  18. Diagram the shape of the cells at each phase.


The cultures will develop differently than the suspension cultures. The viable cells will grow out of the trypsinized pieces of tissue and will remain in contact with the bottom of the culture flask. They will continue to divide and migrate until the entire bottom of the flask is covered with a single layer of cells (contact inhibition and the formation of a monolayer).

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Cell Biology Laboratory Manual
Dr. William H. Heidcamp, Biology Department, Gustavus Adolphus College,
St. Peter, MN 56082 --