ES / MEF cell culture and electroporation of targeting construct
Day 0
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.
Day 2
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.
Day 4
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.
Day 6
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.
Day 7
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.
Day 9
Feed the transfected ES cells with Selection Media, 13 ml dish. The clones should now be fairly large.
Day 10
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.
Day 11/12
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.
Day 13
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.
Day 14
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.
Day 14/15
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.
Day 15/16
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.
Day 16/17
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.
