Culturing Mouse Embryonic Fibroblasts

互联网2013-09-06

689

Materials
Trypsin (Gibco 25200-023)
3T3 Medium: 500 mL DME (Invitrogen) + 50 mL FBS (Hyclone) + 5 mL 100x Pen/Strep
2x Freezing Medium: 3T3 Medium + 20% DMSO: Filter sterilized.
Sterile PBS
P100 and P60 tissue culture dishes
Dissecting tools: Scissors, forceps, scalpel
95% ethanol in a squeeze bottle and in a beaker
Mouse breeders

Note:
This protocol describes the isolation of primary MEFs and passage according to the 3T3 protocol. Passaging primary cells on the 3T3 protocol will maximize the growth prior to the development of cellular senescence. The 3T3 protocol was originally described as the passage of 3 x 10⁵ cells every 3 days on 50 mm. dishes which is equivalent to 1.2 x 10⁶ cells on a P100 dish (Nilausen K, Green H. Exp Cell Res . 1965 ). If cells are passaged continuously every three days under normoxic conditions they will experience a period of rapid growth (passage 1 - 5), progressively slower growth (passage 5 - 10) and senescence (little or no growth, passage 10 - 25). After this time the a subset of cells will emerge from crisis, through the selection for a more rapidly growing, immortalized subclone. This is typically accompanied by the development of an aneuploid karyotype. Early passage primary MEFs and immortalized MEFs are usually spindle shaped and form densely packed monolayers, whereas senescent cells tend to flatten and spread out. Some lines may become transformed and display neuron-like or refractile morphology, growth despite serum starvation, growth in suspension or in soft agar, and the ability to form fibrosarcoma tumors in mice. Senesence in murine fibroblasts is a result of oxidative stress as growth in physiologic (3%) O2 allows continual growth of murine cells similar to human fibroblasts (Parrinello S., et al. Nature Cell Biol, 2003 ).

A. Isolating Primary MEFs:

B. Passaging primary MEFS:

PROCEDURE

1. Setup mouse breeder pairs in the afternoon. Ideally one male should be setup with one or two virgin 7-8 week old females.
2. Check females for copulatory plugs the following morning. If present this should be marked as day +0.5 (or round off to day +1). Remove plugged females from the males breeder cages. Check the remaining females on a daily basis.
3. Embryos should be harvested on day 11.5 to 13.5 as follows:
  1. Place clean forceps and scissors into a beaker with ethanol. Fill a second beaker with sterile water or PBS to wash instruments between embryos. Aliquot sterile PBS into P60 dishes (2 - 3 per embryo).
  2. Euthanize the pregnant female according to standard protocols. CO₂ inhalation should be avoided because this induces acidosis.
  3. Saturate fur with ethanol. Incise the skin vertically and pin to the side.
  4. Using a fresh pair of forceps and scissors open the body wall and pin this back as well.
  5. Extract the intact uterus by grasping one horn with forceps. Withdraw the uterine horn from the abdomen without letting it touching skin or non-sterile areas. Cut the cervix and blood vessels and and transfer the uterus to a sterile petri dish.
  6. The embryos will appear likes pearls on a string. Cut the uterus into sections between each embyo. Place a uterine section with a single embryo into a fresh petri dish with sterile PBS. Squeeze the uterus with forceps to expell the embryo through the cut portion of the uterus.
  7. Wash the embryo in a fresh dish with sterile PBS to wash away any remaining maternal blood cells.
  8. Transfer the embryo to a fresh Petri dish with sterile PBS. Remove the membranes and umbilical cord. With a scalpel vertically incise the abdominal wall and remove the pink hematopoietic tissue (liver and spleen) as well as the tubular intestine.
  9. Trim off the bulk of the CNS tissue by dissecing away the head above the level of the oral cavity.
  10. Save CNS or liver tissue in a 1.5 mL tube for DNA extraction and genotyping. Embryos should be signed a mouse ID and genotyped according to standard protocols. It may be necessary to dilute the DNA or limit the PCR cycle number because of the large amount of DNA and the possibility of maternal blood contamination.
4. Transfer the embryo body to the tissue culture hood in a P100 dish with 5 mL of 1x trypsin.
5. Mince each embryo for 2 - 3 min. with a sterile forceps and scissors or a scalpel to create bits smaller than 2 mm. Place the dish in a 37ºC incubator. Rinse the instruments in sterile water and then 95% ethanol to clean and air dry them between each embryo.
6. After a total of 15 minutes transfer the cells and tissue bits with a 10 mL pipet to a 15 mL conical tube. Pipet the cells several times to further dissociate tissue bits. Add 5 mL of media (DME + 10% FBS).
7. Spin at 1000 RPM in a table top centrifuge for 5 min.
8. Aspirate the supernatant. Resuspend the embryonic cell pellet in 10 mL fresh media and transfer to a clean sterile P100 dish. Small bits of tissue in the cell suspension are OK. MEFs will migrate out of tissue bits onto the dish.
9. Mark the plates as "passage 0", with the date, and the embryo ID. Evenly distribute the cells and tissue bits by horizontal agitation and then incubate at 37ºC in 5% CO2. (Optional: To minimize senescence grow cells in 3% O₂ , see above).
1. Check the plates daily until they are nearly confluent (usually 1-3 days).
2. To trypsinize the cells:
  1. Aspirate away the media. Add 5 mL of PBS. Swirl to rinse and then aspirate off the PBS.
  2. Add 3-4 mL of 1x trypsin (or enough to cover the plate). Incubate at 37ºC for 2 min.
  3. Tap the plate to dislodge cells. Continue incubating until start to come off in large sheets. (For the 1st passage it is sufficient to loosen up the cells in the monolayer. Do not dislodge larger chunks of tissue, if present.)
  4. Add 5 mL of media (DME + 10% FBS) and transfer to a 15 mL conical tube. Pipet to create a single cell suspension. (If larger chunks of tissue are present in the first passage - allow them to settle out of suspension for 1 minute. Transfer the suspended cells to a fresh conical tube.)
3. Determine the viable cell count:
  1. Add 10 µL of cells suspension to a multiwell dish. Add 10 µL of 2x Trypan Blue stain.
  2. Mix by pipetting and transfer 10 µL to each side of a hemocytometer.
  3. Count white (live) and blue (dead) cells on each side of the hemocytometer. Count a total of 100 - 200 cells to be reasonably accurate.
  4. Multiply the average number of live cells in each large square x10⁴ to calculate the number of cells per mL.
  5. Keep a record the number of cells present and amount that they will be diluted to be able to calculate doubling times.
4. Meanwhile spin the remaining cells for 5 min. at 1000 RPM.
5. Aspirate off the supernatant leaving ~ 0.2 mL of media behind. Resuspend pellet by flicking bottom of tube. Add fresh media to bring the cells to a density of 10⁷ cells / mL.
6. Add 9 mL of media to each 100mm dish + 1 mL of cell suspension to plate 1.2 x 10⁶ cells per dish.
7. Mix the cells on the dish by swirling the plates. Horizontally agitate the plates in the incubator to ensure an even distribution of cells. Incubate 37ºC + 5% CO2 . (Optional: To minimize senescence grow cells in 3% O₂ .)
8. Passage the cells every 3 days as described above (B.2-B.7) even if there is little or no growth since the last passage.

1. Setup mouse breeder pairs in the afternoon. Ideally one male should be setup with one or two virgin 7-8 week old females.
2. Check females for copulatory plugs the following morning. If present this should be marked as day +0.5 (or round off to day +1). Remove plugged females from the males breeder cages. Check the remaining females on a daily basis.
3. Embryos should be harvested on day 11.5 to 13.5 as follows:
  1. Place clean forceps and scissors into a beaker with ethanol. Fill a second beaker with sterile water or PBS to wash instruments between embryos. Aliquot sterile PBS into P60 dishes (2 - 3 per embryo).
  2. Euthanize the pregnant female according to standard protocols. CO₂ inhalation should be avoided because this induces acidosis.
  3. Saturate fur with ethanol. Incise the skin vertically and pin to the side.
  4. Using a fresh pair of forceps and scissors open the body wall and pin this back as well.
  5. Extract the intact uterus by grasping one horn with forceps. Withdraw the uterine horn from the abdomen without letting it touching skin or non-sterile areas. Cut the cervix and blood vessels and and transfer the uterus to a sterile petri dish.
  6. The embryos will appear likes pearls on a string. Cut the uterus into sections between each embyo. Place a uterine section with a single embryo into a fresh petri dish with sterile PBS. Squeeze the uterus with forceps to expell the embryo through the cut portion of the uterus.
  7. Wash the embryo in a fresh dish with sterile PBS to wash away any remaining maternal blood cells.
  8. Transfer the embryo to a fresh Petri dish with sterile PBS. Remove the membranes and umbilical cord. With a scalpel vertically incise the abdominal wall and remove the pink hematopoietic tissue (liver and spleen) as well as the tubular intestine.
  9. Trim off the bulk of the CNS tissue by dissecing away the head above the level of the oral cavity.
  10. Save CNS or liver tissue in a 1.5 mL tube for DNA extraction and genotyping. Embryos should be signed a mouse ID and genotyped according to standard protocols. It may be necessary to dilute the DNA or limit the PCR cycle number because of the large amount of DNA and the possibility of maternal blood contamination.
4. Transfer the embryo body to the tissue culture hood in a P100 dish with 5 mL of 1x trypsin.
5. Mince each embryo for 2 - 3 min. with a sterile forceps and scissors or a scalpel to create bits smaller than 2 mm. Place the dish in a 37ºC incubator. Rinse the instruments in sterile water and then 95% ethanol to clean and air dry them between each embryo.
6. After a total of 15 minutes transfer the cells and tissue bits with a 10 mL pipet to a 15 mL conical tube. Pipet the cells several times to further dissociate tissue bits. Add 5 mL of media (DME + 10% FBS).
7. Spin at 1000 RPM in a table top centrifuge for 5 min.
8. Aspirate the supernatant. Resuspend the embryonic cell pellet in 10 mL fresh media and transfer to a clean sterile P100 dish. Small bits of tissue in the cell suspension are OK. MEFs will migrate out of tissue bits onto the dish.
9. Mark the plates as "passage 0", with the date, and the embryo ID. Evenly distribute the cells and tissue bits by horizontal agitation and then incubate at 37ºC in 5% CO2. (Optional: To minimize senescence grow cells in 3% O₂ , see above).
1. Check the plates daily until they are nearly confluent (usually 1-3 days).
2. To trypsinize the cells:
  1. Aspirate away the media. Add 5 mL of PBS. Swirl to rinse and then aspirate off the PBS.
  2. Add 3-4 mL of 1x trypsin (or enough to cover the plate). Incubate at 37ºC for 2 min.
  3. Tap the plate to dislodge cells. Continue incubating until start to come off in large sheets. (For the 1st passage it is sufficient to loosen up the cells in the monolayer. Do not dislodge larger chunks of tissue, if present.)
  4. Add 5 mL of media (DME + 10% FBS) and transfer to a 15 mL conical tube. Pipet to create a single cell suspension. (If larger chunks of tissue are present in the first passage - allow them to settle out of suspension for 1 minute. Transfer the suspended cells to a fresh conical tube.)
3. Determine the viable cell count:
  1. Add 10 µL of cells suspension to a multiwell dish. Add 10 µL of 2x Trypan Blue stain.
  2. Mix by pipetting and transfer 10 µL to each side of a hemocytometer.
  3. Count white (live) and blue (dead) cells on each side of the hemocytometer. Count a total of 100 - 200 cells to be reasonably accurate.
  4. Multiply the average number of live cells in each large square x10⁴ to calculate the number of cells per mL.
  5. Keep a record the number of cells present and amount that they will be diluted to be able to calculate doubling times.
4. Meanwhile spin the remaining cells for 5 min. at 1000 RPM.
5. Aspirate off the supernatant leaving ~ 0.2 mL of media behind. Resuspend pellet by flicking bottom of tube. Add fresh media to bring the cells to a density of 10⁷ cells / mL.
6. Add 9 mL of media to each 100mm dish + 1 mL of cell suspension to plate 1.2 x 10⁶ cells per dish.
7. Mix the cells on the dish by swirling the plates. Horizontally agitate the plates in the incubator to ensure an even distribution of cells. Incubate 37ºC + 5% CO2 . (Optional: To minimize senescence grow cells in 3% O₂ .)
8. Passage the cells every 3 days as described above (B.2-B.7) even if there is little or no growth since the last passage.

C. Freezing/thawing MEFS:

Primary MEFs do not do freezer and thaw well and are likely senesce at a lower passage than are cells kept in continuous culture. If they are to be frozen it should probably be at passage 1 - 2.
To freeze MEFs:
1. Trypsinize cells as above B.2 - B.4. Resuspend the cells at a density of 4 x 10⁶ live cells /mL. Add an equal volume of 2x Freezing Media (media + 20% DMSO).
2. Aliquot 1 mL of cell suspension in freezing media into cryo vials. Cap securely, being careful not to contaminate the edges of the vial or cap. Label tubes as MEFs, passage #, date, and your initials.
3. Place vials in a styrofoam container or a cell freezing jar with isopropanol. Place at -80ºC overnight.
4. Transfer vials to a cryofreezer. Record the location in the Freezer database.
To thaw MEFs:
1. Pre-warm medium at 37ºC.
2. Remove a vial of cells from the cryo freezer and place in 37ºC bath.
3. Delete the vial entry from the Freezer database.
4. Wipe the vial with ethanol and place in the tissue culture hood.
5. Transfer the cells to a sterile conical tube with 10 mL of medium.
6. Count the viable cells with trypan blue as above (B.3).
7. Plate the cells at density of 1.2 x 10⁶ cells per P100 (or 1.5 x 10⁴ cells / cm² in a smaller dish).
8. Incubate 37ºC + 5% CO2 . (Optional: To minimize senescence grow cells in 3% O₂ .)