3.2.1. Vector Preparation
1. In a volume of 0.3 mL, linearize 150 pg of vector DNA with the appropriate restriction enzyme. (Plasmid DNA for electroporation is prepared by alkaline hydrolysis and banded on a cesium chloride gradient as described in ref. 20).
2. Precipitate the digested DNA sample in 2 vol of absolute ethanol on ice for 5 min. Spin in microfuge, wash pellet several times with 70% ethanol, and drain off as much of the ethanol as possible.
3. Evaporate off the remaining ethanol in the hood by keeping the lid of the tube open (approx 1 h). Resuspend DNA pellet in 100 pL of sterile PBS, and vortex sample occasionally over a period of at least 4 h to ensure that the DNA is completely dissolved.
3.2.2. Electroporation of ES Cells and Picking G418-Resistant Colonies
1. Trypsinize three 175-cm2 flasks using 5 mL trypsin/flask as described above. Add 8 mL of medium to each flask, and combine cells in a 50-mL disposable centrifuge tube. Spin for 5 min at 260g, and resuspend in 20 mL of PBS.
2. Count cells, and resuspend at a concentration of 108 cells in 0.7 mL of PBS.
3. Add cells to the tube containing the 150 |g of linearized plasmid, and transfer immediately to a 0.4-cm electroporation cuvet. Electroporate in Bio-Rad Gene Pulsar unit set at 3 mF/800 V (time constant = 0.1 ms).
4. Leave cells to recover in the cuvet for 20 min, and then transfer them to 200 mL of medium. Plate 10 mL (5 x 106 cells) of the cell suspension onto 20 10-cm diameter gelatinized tissue-culture dishes.
5. After 24 h, aspirate medium, and replace with medium containing 200 |g/mL of G418. For the first 5 d, change medium daily. Once G418-resistant colonies appear, the medium may be changed every other day (see Note 2).
6. After about 10-12 d of growth, colonies should be about 1 mm in diameter. Circle colonies with a marker pen on the bottom of the dish.
7. Gelatinize the required number of 24-well plates.
8. Aspirate medium, and add 10 mL of PBS to each dish. Ideally, colonies should be picked in a tissue-culture hood. However, it is possible to pick colonies on the bench, resulting in minimal or no contamination as long as procedures are performed swiftly. Using a P200 Pipetman set at 100 |L and sterile tips, break up the colony, pipet the cells up in a volume of 100 |L PBS, and transfer to a 24-well dish. Once 24 colonies have been picked, add 100 |L of trypsin to each well, and incubate at 37°C for 10 min. Tap the dish several times to disperse the cells, and then add 2 mL of medium to each well.
9. To identify Pgal-positive colonies, we normally split four-fifths of a nearly confluent well into a new 24-well plate (Experimental) for staining with X-gal, and split the remaining one-fifth into a second 24-well plate (Master) for maintenance and subsequent expansion of selected cell lines (see Note 3). Since the colonies grow at different rates, the cells are split in batches over a period of several days. Cells in the Experimental plates are stained with X-gal the following day, and Pgal activity is scored after an overnight incubation. By this time, the cells on the Master plate should be ready to expand for further analysis.
3.2.4. Staining Cells for P-Galactosidase Activity
1. Aspirate medium and wash cells once with PBS.
2. Fix cells for 10 min at room temp.
4. Stain with X-gal overnight in a humidified chamber at 37°C. Colonies expressing low levels of P-gal may appear slightly discolored compared to P-gal negative cell lines. Pgal staining is difficult to see using phase contrast. We recommend using either bright field illumination or Nomarski optics.
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