ES / MEF cell culture and electroporation of targeting construct
One frozen vial of Murine Embryonic Fibroblasts (MEFs) is thawed quickly in a 37oC water bath. When the last bit of ice is melted, spray the vial with 70% ethanol and transfer the contents of the vial into one 75 cm2 flask (T-75) containing 20 ml of MEF media. Place the MEFs in a 37oC, 5% CO2, 86% humidity incubator. Every frozen MEF preparation thaws a little differently. If on Day 1, the MEFs are only 50% cconfluent, thaw another vial into your ongoing T-75 MEF flask. It is important for the MEFs to be maintained at a relatively high density, or they will not continue to expand.
MEF passage 1: Expand the MEFs as follows: The MEFs should be 90% confluent (if they are not, feed them another day). Split as follows: Suction off old media, rinse flask x 1 with 10 ml phosphate buffered saline (PBS). Add 5 ml Trypsin/EDTA media (0.05%/0.02%) to the flask and incubate for 5 to 10 minutes at 37oC. After 10 minutes, the cells should be detached from the bottom. Now add 10 ml MEF media to this flask, pipette up and down to disaggregate the cells into a single cell suspension, and add the entire contents of the flask into one 150 cm2 flask (T-150). Add 20 ml MEF media to the flask, so that the final volume is 30 to 35 ml. Incubate at 37oC, 5% CO2, 86% humidity.
You should now have 1 T-150 flask of confluent MEF cells.
MEF passage 2: Mitotically inactivate cells as follows: Rinse with PBS (10 ml) and harvest with 7 ml Trypsin/EDTA for 10 minutes at 37oC as above. Add 10 ml of MEF media and pipette cells up and down several times. Take 10 ml of cells from the flask and put in a 50 ml centrifuge tube. Refeed the T-150 ml flask with 25 ml of fresh media. To the 50 ml centrifuge tube containing the MEF cells, add 5 ml MEF media (bringing the final volume up to 15 ml). Take this tube to the blood bank and irradiate (3000 rads). These inactivated MEFs will be used to make two 10cm dishes of MEF feeder layers for your ES cell thaw on Sunday. Using a hemacytometer, count the MEFs before adding them to your dish. A good monolayer will be formed if you add approximately 1x106 MEF cells to each 10 cm dish. Plate inactivated MEFs in 2-10 cm dishes having a final concentration of 1x106 Mef's per dish and a final volume of 12.0 ml. We prepare 1 extra 10cm dish in case of contamination or poor monolayer.
Sunday, Day 5
Thaw ES cells as follows: Suction the existing MEF media off one of the inactivated MEF 10 cm dishes. Refeed the dish with 12 ml ES media. Thaw one frozen vial of RW4 cells (1x106 cells) in a 37oC waterbath. When the last bit of ice has melted, spray the tube with 70% ethanol and transfer the contents of the vial to the inactivated MEF dish. Rock the dish to evenly disperse the cells. Incubate overnight at 37oC, 5% CO2, and 86% humidity. The T-150 flask of ongoing MEFs is ready to be expanded today. Rinse with PBS, add 7ml Trypsin/EDTA, for 10 minutes, then add 10 ml of MEF media, and pipette up and down several times. Pipette 8 ml of these cells into a new T-150 flask containing 25 mls of MEF media. Refeed the existing T-150 with 25 ml MEF media to create 2 T-150's of expanding MEFs.
Feed the ES cells with 12 ml ES media. Approximately 60% of the ES cells will form colonies in the dish. They are football shaped, shiny, and plump. You will need 4 x 10 cm dishes of irradiated MEFs for your electroporation on Day 7. You have 2 T-150's ongoing from which to make these 4 dishes. Follow the procedure for mitotically inactivating MEFs on Day 4. However, use both T-150s this time. Don't forget to refeed them, or you will not have MEFs for the end of the week. If the 2 T-150 flasks of MEF cells are not confluent today, feed them, and then inactivate them on the morning of Day 7. If you inactivate your MEFs on Day 7, you must either give them 2 to 3 hours to attach before changing the MEF media to ES media or initially inactivate and plate them in ES media.
Electroporation: Use three of the 10 cm dishes of inactivated MEF cells prepared on Day 6. From each dish, remove the old media and add 10 ml of fresh ES media. Place these dishes back in the incubator. Next, take the 10 cm dish containing the MEFs and ES cells and remove the old media. Rinse the dish x 1 with PBS, then add 2 ml Trypsin/EDTA and incubate for 5-10 minutes at 37oC. After 5 minutes, the cells will look like "bunches of grapes" under the inverted scope. Add 2 ml more Trypsin/EDTA to the dish, pipette up and down to break up the clumps and incubate for 3 more minutes. [You must have single cells for the electroporation.] Look at the cells again under the inverted microscope. The MEFs are the larger cells, and the ES cells are small and shiny; most if not all should now be single cells. To the 10 cm dish add 7 ml ES media, pipette up and down, and transfer all the cells to a 15 ml centrifuge tube. Pellet the cells by centrifuging gently (1000 RPM in a Sorvall tabletop) for 5 minutes at 10oC. Take off the supernatant and resuspend the pellet in 1.0 ml ice cold 1x Hebs (see Reagents, Transfection Buffer 1 X Hebs). Prepare a 5 ml tube of ES media for the cells after electroporation. Get out a sterile "flat pack" 1.8 mm gap cuvette (BTX order #485) and insert the cuvette between the safety stand contacts. Make sure there is a good contact between the cuvette and the safety stand contact. Having the safety stand connected to the rear of the unit using the cables supplied, turn on the power switch. Set electroporator (BTX 600 or equivalent) as follows: 500V/Capacitance and resistance, 500uF capacitance timing, 360 ohms R8 Resistance timing, Charging voltage 185V. Pipette the ES cells up and down with a 5 ml pipette and add to a microfuge tube containing the targeting construct DNA ( 40 ug of clean linear DNA in 1 X TE @ 1 ug/ul for each electroporation). Pipette cells and construct up and down with a pasteur pipette carefully. Slowly add the cells to the cuvette, taking care not to introduce any bubbles. Slide the cuvette into the electroporation chamber, dial the charging voltage to 185V and push the pulse button. Wait until the charging is over, then push the reset button, dial down the voltage, and turn the power off. With a sterile pasteur pipette, take the electroporated cells out of the cuvette and place them into the 5.0 ml of fresh ES media in a centrifuge tube (final vol. = 6 ml total). Take the three 10 cm dishes of inactivated MEFs (freshly fed with ES media) and add 2 ml of the transfected ES cells per dish. Label the dishes with the targeting construct's name and date. Rock the dishes slowly to evenly disperse cells.
Alternate Electroporation Procedure using Safety Chamber 630 A, BTX cuvette, 2 mm gap:
Prepare a 5.2 ml. tube of ES media for cells after the electroporation. Prepare ES cells for electroporation as described in original text on Day 7, except add only 800 ul cold 1 X Hebs to your pelleted cells. Place a sterile BTX cuvette (2 mm gap) in a 630 A Safety Chamber and attach the electrodes securely to the Electro Cell Manipulator 600. Pipette the cells up and down with a 5 ml pipette, and add 800 ul of cells to a microfuge tube containing the targeting construct (40ug of clean linear DNA in 1 X TE @ 1 ug/ul for each electroporation). Mix the cells and DNA with a 200-1000 ul barrier tip trying not to create bubbles. Add 400 ul of the cell/DNA mixture to a BTX cuvette. Set Electroporator as follows:
500Vcapacitance and resistance, 500uF capacitance timing, 360 ohms R8 Resistance timing, Charging Voltage 160V. When capacitors are charged hit the pulse button. When charging is complete, with the pipette provided, harvest the electroporated cells and place them into 5.2 ml of fresh ES media. Repeat the electroporation with the other 400 ul cells/construct in a new BTX cuvette. Add this to the 5.2 ml yielding a final volume of 6.0 ml. Take the 3-10 cm. dishes containing inactivated MEFs freshly fed with ES media and add 2 ml. of the electroporated cells per dish. Rock the dishes slowly to evenly disperse the cells. Label the dishes and place in the incubator.
Feed the transfected ES cells with Selection Media, 13 ml dish. The clones should now be fairly large.
Feed transfected ES cells with Selection Media as above. You should begin to see some selection in your dishes. Dead cells should be suspended in the media above your ES clones. Using your ongoing T-150 MEF flasks (2 flasks), you will prepare five, 24 well dishes for the isolation and expansion of your individual clones on Day 13. MEF passage 3: split ongoing MEFs (90% confluent) and irradiate as follows: Take off the old media and rinse x 1 with PBS (10 ml). Add 7 ml Trypsin/EDTA and incubate 5 to 10 minutes at 37oC. Add 10 ml MEF media, and pipette up and down. Transfer all of the MEF cells to a 50 ml centrifuge tube. Repeat with the other flask of MEFs. If you wish to keep an ongoing flask of MEF cells at this time, you may leave 5 ml. of cells in one of the flasks and refeed this flask. However after preparing your 24 well dishes today you will not need MEFs again for the completion of this electroporation. Take the MEF cells in the centrifuge tube and irradiate the cells as before (3000 rads) at the blood bank. Using a hemacytometer, count your MEFs. You will need 7x 104 MEFs per well. To attain this, you should add 9x106 irradiated cells in a final volume of 125 ml. MEF media into a sterile plastic bottle. Now you will have enough cells to prepare 5 - 24 well dishes. Mix the cells gently so that they are evenly dispersed and add 1.0 ml of irradiated MEFs to each well of a 24 well dish in a total of 5 dishes. Incubate at 37oC until Day 13.
Feed transfected ES cells with 12 ml of Selection Media (G418). This may be done on Day 11 or Day 12., but does not have to be done both days.
View the 10 cm dishes containing the transfected ES cells through the inverted microscope. The clones are visible as small nests of rapidly growing cells. They have tight borders and are closely packed. Larger cells within the colony, with well defined membranes are the cells which are beginning to differentiate. Do not pick these clones! This is day 6 of the selection process. If the clones are big enough you may pick on this day, or wait until day 7. To pick the clones, you must view them through an inverted microscope in a laminar flow hood. Prepare your 24 well plates (made on Day 10) by taking out the old media and replacing it with 1.0 ml ES Selection Media per well. Place the 24 well dishes (all 5) back into the incubator. Replace the ES selection media in the first 10 cm dish with fresh ES Selection Media. Place the dish under the microscope and view the cells at 4X and 10X trying to be sure you are picking clones that have not yet started to differentiate. Isolate the clone that you wish to pick in the viewing field. Using a 0-160 ul barrier tip, gently push the ES clone forward from the surrounding MEFs. With the pipettor set between 30 and 50 ul, and the plunger button already depressed, pluck the clone using a forward scooping suction motion. If the pipettor is set on 30 ul, you should have enough suction to dislodge the clone from the plate. However, if the inactivated MEF layer is too dense, you may have trouble dislodging the ES clones. In this case, you must carefully tease the surrounding MEFs away from the clone without disrupting the clone with your tip. Then you should be able to harvest your clones as above. Place each clone into one of the 24 wells containing ES Selection Media. Continue to pick clones and place in the wells of the prepared 24 well plate until 12 clones are picked or the 10 cm. dish has been out of the incubator for 15 minutes (the clones are sensitive to pH and temperature changes). Now place the plate containing the picked clones in the incubator. Continue picking clones until all 24 wells have been filled in all of your prepared dishes. Leave the 24 well dishes in the incubator overnight.
If you were not able to pick all your clones on day 13, you may continue to pick today (selection day 7). Picking on Day 8 of selection is NOT recommended.
Examine the 24 well plates under the scope. You should see a single clone in each well. To disaggregate each clone, hold the dish at an angle and suction out the media. Now add 0.5 ml PBS to each well to rinse out the media and resuction each well. Next add 0.2 ml. Trypsin/EDTA to each well and place the plate back in the incubator for 20 minutes at 37oC. Finally add 1.0 ml ES selection media to each well and pipette the cells up and down 4 or 5 times with a 200 ul to 1000ul barrier tip to disaggregate the clone . After all clones are disaggregated, return your 24 well plate to the incubator. Look at your plates every day. You want your wells to have many small nests of colonies, evenly dispersed throughout the well. Note: ES selection media should be used until the clones are frozen to ensure that no Wild Type ES cells contaminate your clones.
Examine the clones with the inverted scope. They will look like many tiny clones of ES cells, evenly dispersed. Aspirate the existing media from the cells, then feed all your wells with 1.0ml ES selection media.
The individual wells of the 24-well plate should contain 50 to 100 small, healthy, undifferentiated colonies if they were properly disaggregated. Number your wells from 1 to 120. To freeze the individual clones, first rinse each well with 0.5 ml PBS, and then add 0.2 ml 0.05% Trypsin/EDTA to each well. Place the plate in the incubator for 15 minutes. Remove the plate from the incubator and add 0.5 ml ES media to each well. Individually disaggregate the cells with a 200 to 1000 ul barrier tip by pipetting up and down 4 to 5 times. Place 500 ul of the disaggregated colonies in the Nunc vial (1.8 ml) that is numbered the same as your well. Add 0.5 ml. of 2X ES freezing media to each vial. Place the numbered cryotubes in a freezer box labeled with your constructs name and put in a -70o freezer. After removing all the colonies to the cryotubes, refeed each well of the 24 well plate with 1.0 ml ES media. Allow the cells in these wells to grow to confluence (which will take 4-6 days) and use these wells to harvest cells for DNA analysis of the clones. Repeat this freezing procedure for all of your plates. When the DNA analysis confirms which numbered vials are homologous recombinants, these vials are then stored under liquid nitrogen for later expansion and injection. If all of your wells were not ready to be frozen by Day 17, you may need to disaggregate these wells again with Trypsin / EDTA . This will enable a well containing a small number of cells to be expanded further for a subsequent freeze on Day 19. Follow the freezing procedure above on Day 19 for these wells.
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