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Thomson Lab Stem Cell Protocols
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The plasmids used to reprogram somatic cells and the resulting iPS cells (Yu et al., 2007) are now available. Please see our Reagents Page for more information.

Thomson Lab's Protocols

Human Embryonic Stem Cell Protocols

stem cell colony

Mouse Embryonic Fibroblasts Protcols

iPS Cell Protocols

Human Embryonic Stem (ES) Cell Protocols

Media and Reagents

PDF for mediaformulas

Serum Free Media for human ES cells on MEFs: can last for 7-10 days

Final Concentration
Amount for 250ml Stock solution
80% DMEM-F12
200ml
20% KO Serum Replacer
50ml
1% Non-essential Amino Acids
2.5ml
1mM L-glutamine (see recipe)
2.5ml
0.1mM b-mercaptoethanol
part of L-glutamine
4ng/ml bFGF (see recipe)
0.5ml

L-glutamine stock solution: Prepare on a per-use basis

bFGF Stock: Store aliquots at –20 or –70

0.1% Fraction V BSA:

Collagenase IV Split Media: This media lasts 2-3 weeks

Cell Freezing Media:

Cryopreservative Medium:
Final Concentration
Amount for 10ml Stock Solution
60% DMEM-F12
6ml
20% defined FBS
2ml
20% DMSO (do not filter sterilize)
2ml

 

Resuspension Medium:
Final Concentration
Amount for 10ml Stock Solution
80% DMEM-F12
8ml
20% defined FBS
2ml

 

Thawing Human ES cells:

PDF for thawing_es
  1. Remove Human ES cells from liquid nitrogen storage tank. Fill out a freeze/thaw form.
  2. Thaw cryovial by gently swirling in waterbath until only a small ice pellet remains, being careful not to completely submerge the cryovial under water.
  3. Completely submerge cryovial in 95% ethanol.
  4. Very gently, pipet cells from the vial into a 15 ml conical centrifuge tube.
  5. Slowly, add 9.5ml media dropwise to reduce osmotic shock. While adding media, gently mix the cells in the tube (by gently tapping the tube with a finger.)
  6. Centrifuge 1000rpm for 5 minutes.
  7. Wash cells by resuspending with 3ml media.
  8. Centrifuge 1000rpm for 5 mintutes.
  9. Resuspend in 2ml and add 0.5ml per well of a 4 well plate that has MEFs already plated on it.
  10. Change media daily, however it may take 2 weeks before cells are ready to be expanded.

Note: Wear safety glasses when removing cells from liquid nitrogen until the cells are completely thawed.

 

Splitting Human ES cells on MEFs:

PDF for splitting_es

stem cell photos

based on splitting onto one plate

  1. Warm collagenase IV split media to 37 °C in a water bath.
  2. Aspirate media off of cell culture plate.
  3. Add the following amount of collagenase:
    • 0.5ml/well of 4 well plate
    • 1.0ml /well of 6 well plate
  4. Incubate at 37 °C with 5% CO2 for 5-10 minutes. The cells are ready when the edges of the colony are rounded up and curled away from the MEFs on the plate.
  5. Using a 5ml pipet, scrape and wash the colonies off of the plate.
  6. Transfer cell suspension to a 15ml conical tube.
  7. Break up the colonies by pipetting up and down against the bottom of the tube until there appears to be a fine suspension of cells (no clumps of cells remain).
  8. Spin cells at 1000rpm for 5 minutes.
  9. Aspirate collagenase off and wash cells with 3 ml human ES media.
  10. Spin 1000rpm 5 minutes.
  11. While the cells are spinning prepare the feeder layers by aspirating off the MEF media and washing one time with Ca/Mg free PBS and adding human ES media to the feeder layers (2 ml/well of 6 well plate).
  12. Once cells are done spinning, aspirate off wash media.
  13. Resuspend cells in an appropriate volume (see notes).
  14. Plate cells by adding 0.4ml per well of a 6 well plate until the last 0.5-0.6ml remain. Add the last remaining volume dropwise to each well.
  15. Make sure the cells are evenly distributed across the entire well (see notes).
  16. Place gently in incubator. Again make sure the cell are not disturbed.
  17. Let cells settle overnight in incubator.

Notes:

 

Freezing Human ES Cells:

PDF for freezing_es
  1. Collagenase cells for approximately 7 minutes at 37 °C (until edges of colonies are curling up).
  2. With a 5 ml pipet, gently pipet and scrape colonies from plate. Add cell suspension to a 15 ml centrifuge tube and GENTLY break up colonies. It is important to be gentle in this step as “chunkier” colonies will thaw out better than single cells. Ideally, colonies meant for freezing are left slightly larger than they would be for splitting.
  3. Spin 5 minutes at 1000 rpm.
  4. Resuspend pellet (gently) in 3 ml ES media to wash away collagenase.
  5. Spin 5 minutes at 1000 rpm.
  6. Resuspend pellet (again, gently!) in 0.25 ml Resuspension Medium per vial (This is one half the final volume required for freezing).
  7. Dropwise, add an equivalent volume (0.25 ml per vial) of Cryopreservative Medium and mix. Your DMSO concentration is now 10%.
  8. Place 0.5 ml of cells in each freezing vial.
  9. Rapidly transfer the cells to a freezing container and place at -70 °C overnight (cells don’t like to be in DMSO at room temperature for long periods of time).
  10. Transfer cells to liquid nitrogen the next day for long-term storage.

Notes:

 

Matrigel Aliquoting and Plating:

PDF for matrigel

Aliquoting Matrigel:

Day one:

Day two:

How to Make Matrigel Plates:

There are at least two possible methods for this. The main objective is not to let the undiluted matrigel sit at room temperature for too long (or it will become chunky and solidify).

A second possible method is a little quicker, the only modification being that you take a tube directly from the freezer and IMMEDIATELY resuspend the pellet of Matrigel in 6 ml ice cold media. Keep pipetting vigorously until all chunks are gone, and add to plate.

If after either of these methods, your plate still looks like it has chunks of Matrigel instead of a smooth even layer, try placing the plate at 4 °C overnight. This should "re-melt" the matrigel into an even layer. Some chunks are acceptable, but make sure that the whole surface of the plate is coated with matrigel.

Caring for Cells Growing on Matrigel:

hES cells grown on Matrigel-coated plates require MEF-conditioned medium. This medium is identical to regular hES medium, but is made without bFGF. Twenty-four hours before using the medium, it is conditioned on MEFs (at a density of 2.12 x 105 cells/ml, 2.5ml/well of 6 well plate). bFGF is added fresh every time to each individual well (5 ml/well).

*When ordering Matrigel, it is best to request "high-density" Matrigel (ie. over 12 mg/ml). There are some applications in the lab which require a higher density to have successful experiments. Additionally, one bottle of high density Matrigel will go farther than one bottle of low density Matrigel.

**The main objective and constant throughout working with Matrigel is to keep it COLD. As Matrigel warms to room temperature, it begins to solidify, making it harder to work with (ie. sticking inside of the pipet) and making it chunkier when attempting to use and plate.

Splitting Human ES cells on Matrigel:

PDF for splitting_es_matrigel

based on splitting onto on plate

  1. Warm collagenase media to 37°C in a water bath.
  2. Aspirate media off of cell culture plate.
  3. Add the following amount of collagenase
    • 0.5ml/well of 4 well plate
    • 1.0ml /well of 6 well plate
  4. Incubate at 37 °C for 5-10 minutes; stop incubation when edges of colonies begin to pull away from the plate.
  5. Aspirate the collagenase and add appropriate volume (3ml) of conditioned media (CM) to the plate (see notes).
  6. Collect cells off of the plate by scraping and washing with the CM, transfer to a 15ml tube.
  7. Aspirate matrigel off of 6 well plate (see matrigel aliquoting and plating procedure)
  8. Add 2ml of CM per well of 6 well plate.
  9. Add 5ul bFGF.
  10. Plate 0.4ml/well into each well until there is approximately 0.6 ml remaining.
  11. Add the remaining 0.6ml dropwise to each well until done.
  12. Make sure the cells are evenly distributed across the plate.
  13. Place gently into incubator.
  14. Let settle overnight.

Notes:

Splitting Human ES cells in Defined Media

PDF for define media split

The Thomson lab currently uses TeSR media, however this protocol will work with other define medium

  1. Warm 2mg/ml dispase to 37C.
  2. Aspirate media off of cell culture plate.
  3. Add the following amount of dispase (2mg/ml in DMEM/F12):
    • 0.5ml/well of 4 well plate
    • 1.0ml /well of 6 well plate
  4. Incubate at 37 C with 5% CO2 for 5-7 minutes.
  5. During 5 minute incubation, replace media on matrigel coated plate (see matrigel coating procedure) with fresh, ES cell culture media.
  6. After 5 minutes of incubation, check cells to determine if they are ready to be passaged. (Cells are ready for passaging when the majority of colonies are beginning to round up away from the plate. If cells begin to float off the plate, you will need to collect the cells and dispase, spin them down by centrifuging at 1000 rpm for 5 minutes, washing (with repeat centrifuging) twice with DMEM/F12, and proceed to step 9)
  7. If cells are not ready to be passaged, return them to incubator for 1-3 minutes.
  8. Wash cells (on plate) 3 times with sterile DMEM/F12. Gently add media as dispase treated cells will easily wash off plate.
  9. Resuspend cells in appropriate volume of media for plating:
    • 3.5 ml for 6 well plate
    • 1 ml for a 10cm dish
  10. Gently break up cells by pipeting up and down 1-2 times.
  11. Plate cells onto new plates (for a 6 well plate, add 0.5ml/well and distribute the final 0.5ml dropwise to each well for even cell distribution.
  12. Place cells in incubator.

Notes:

Embryonic Bodies:

PDF for embyonic_bodies
  1. Let human ES cells grow until the colonies are large and the cells are pretty piled up - about the time when you would normally split or even a day past that.
  2. Treat cells with 0.2 - 0.5 mg/ml dispase. You want to use the lowest possible concentration of dispase, but it tends to vary a bit.
  3. Wait until the colonies completely detach from the plate. Do not blow colonies off with a pipet. This should take about 20-30 min. If nothing is happening by that point, add more dispase.
  4. Once the colonies come up, gently transfer them to a 15 ml conical tube with a 10ml pipet. You don't want to break up the colonies.
  5. The cells should sink to the bottom of the tube after a minute or two without any spinning. Aspirate off the media and wash once in hES media. If you are in a hurry and need to spin the colonies down, 1 min at 500 rpm is enough.
  6. Transfer cells to a flask containing ES media without bFGF. Put all of the EBs from one 6-well plate into a T80 flask with about 25 mls media.
  7. The cells will round up into actual embryoid bodies after about 12-24 hrs. They should then be fed every day by exchanging half the media with fresh media. The EBs should not attach, if they do, gently tap the flask to dislodge the EBs.

Notes:

General notes on ES cell culture:

PDF for general_es

Mouse Embryonic Fibrolasts (MEFs) Protocols

Harvesting of Mouse Embryonic Fibroblasts from Murine Embryos

PDF for harvesting MEFs
  1. Inject approximately 0.5 ml avertin IP into a 13 or 14 day pregnant mouse*. When mouse is anesthetized, perform a cervical dislocation.
  2. Saturate abdomen with 70% Ethanol and pull back the skin to expose the peritoneum. With sterile tools, cut open the peritoneal wall to expose the uterine horns. Remove the uterine horns and place them in a 10 cm dish. Wash three times with 10 ml PBS w/o CaMg.
  3. Cut open each embryonic sac with a scissors and release the embryos into the dish.
  4. Using two pairs of watchmakers forceps, remove the placenta and membranes from the embryo. Once they have been removed, dissect out the visceral tissue (ie. anything that is dark in color). Place the embryos in a clean petri dish and wash three times with 10 ml PBS.
  5. With a curved iris scissors, FINELY mince the tissue. When your hand is too tired to mince any more, add 2 ml Trypsin/EDTA and continue to mince. Add an additional 5 ml of Trypsin/EDTA and incubate at 37o°C (for about 20 minutes). At this time return to step 1 and start another mouse.
  6. Perform steps 1-4 and return to the embryos in Trypsin/EDTA.
  7. Pipet the embryos in Trypsin/EDTA vigorously, until few chunks remain. Return plate to the incubator for an additional 10 minutes.
  8. Neutralize the Trypsin/EDTA with about 20 ml culture medium**, and transfer the contents of the dish to a 50 ml conical tube.
  9. Mix the contents of the tube well, and evenly add to T75 culture flasks containing 20 ml culture medium. There should be approximately 3 embryos per T75.
  10. Place these flasks in a 37°C incubator overnight.
  11. Return to the embryos sitting in PBS, and begin process at step 5 again.
  12. The next day, change the medium to get rid of debris and toxic cell death products.
  13. When flasks are becoming about 80-90% confluent and still in the log growth phase, it is a good time to freeze them. In general, this happens about the second day after preparing the embryos. It may happen sooner or later, so keep watch over your flasks.

Notes
* We have been using CF-1 mice for fibroblast preparations
** Culture medium is:

*** As with any new cell in the lab, a representative sample should be tested for mycoplasma

Freezing MEF cells:

PDF for freezing MEFs

Resuspension Medium

Cryopreservative Medium

  1. Wash cells once with PBS w/o CaMg.
  2. Add Trypsin/EDTA to cells for approximately 5 minutes at 37° C.
  3. Detach cells from the plate by pipetting off or tapping against the heel of your hand.
  4. Neutralize Trypsin/EDTA with an equal volume of culture medium.
  5. Pipet to break up chunks. If clumps remain, add suspension to a 50 ml tube and allow the chunks to settle out.
  6. Take the supernatant and divide it amongst conical tubes and spin 5 minutes at 1000 rpm.
  7. Resuspend pellet in 0.5 ml Resuspension Medium per vial (This is one half the final volume required for freezing).
  8. Dropwise, add an equivalent volume (0.5 ml per vial) of Cryopreservative Medium and mix. Your DMSO concentration is now 10%.
  9. Place 1 ml of cells in each freezing vial.
  10. Rapidly transfer the cells to a freezing container and place at -70°C overnight (cells don’t like to be in DMSO at room temperature for long periods of time).
  11. Transfer cells to liquid nitrogen the next day for long-term storage.
Notes:

Thawing MEF cells:

PDF for thawing MEFs
  1. Remove vial from liquid nitrogen.
  2. Do a quick thaw in a 37°C water bath, being careful not to immerse the vial above the level of the cap.
  3. When just a small crystal of ice remains, sterilize the outside of the vial with 95% EtOH and place it in the hood.
  4. Gently pipet the cells up and down once, and place them in a 15 ml conical tube.
  5. Dropwise, slowly add 10 ml culture medium to the tube. This should not take less than 2 minutes. To do this faster shocks the cells and you may end up with a lower viability than you otherwise would.
  6. Spin the cells at 1000 rpm for 5 minutes.
  7. Resuspend the cells in 10 ml culture medium and add to T75.
  8. Place in 37°C incubator.
  9. Fill out a freeze/thaw form to keep the database current.

Splitting MEF cells:

PDF for splitting MEFs
  1. Remove MEF medium*.
  2. Wash cells with 5 ml of PBS w/o CaMg (to get rid of trypsin inhibitors).
  3. Add 1.5 ml Trypsin/EDTA (0.05% Trypsin) to each flask and allow to sit for about 5 minutes.
  4. To loosen cells, either tap flask against the heel of your hand or pipet the cells off.
  5. For every 1 ml of Trypsin/EDTA added, add at least 1 ml of MEF medium to neutralize the trypsin reaction.
  6. Add the cell suspension to a 15 ml conical tube and pipet several times to individualize the cells.
  7. Add 10 ml MEF medium to new T75 flasks.
  8. Divide the cell suspension appropriately amongst the new T75 flasks and place at 37°C**.

Notes:
* MEF medium is:

** MEFs will grow slower with each passage. It is possible that you may begin by splitting the first passage 1:5, but end up splitting the last usable passages (p4 and p5) only 1:2.

Irradiationg & Plating MEF cells:

PDF for plating MEFs hemocytometer counting chamber
  1. Remove MEF medium.
  2. Wash cells with 5 ml of PBS w/o CaMg (to get rid of trypsin inhibitors).
  3. Add 1.5 ml Trypsin/EDTA (0.05% Trypsin) to each flask and allow to sit for about 5 minutes.
  4. To loosen cells, either tap flask against the heel of your hand or pipet them off.
  5. For every 1 ml of Trypsin/EDTA added, add at least 1 ml of MEF medium to neutralize the trypsin reaction.
  6. Add the cell suspension to a 15 ml conical tube and pipet several times to individualize the cells.
  7. Perform a cell count:
    • Mix cell suspension thoroughly and remove 10 μl
    • Add this to 10 μl Trypan Blue and mix well
    • Add ≤10 μl cell suspension/Trypan Blue mix to hemacytometer
    • Count bright cells in two of the 4x4 squares--I usually do opposite corners (ie. A and C, or B and D)
    • Don't include dead cells in the count--these pick up the Trypan Blue
    • Total cell number = (Total cells counted in two 4x4 squares) ÷ (two 4x4 squares) x (two--Trypan Blue dilution factor) x (1 x 10^4--cell dilution factor) x (total ml of cell suspension)
    • Example: You have just finished counting your sample of 23 ml cell suspension and saw 432 live cells in the two 4x4 squares. Your calculation will look like this:
    • 432 cells ÷ 2 x 2 x (1x104) = 4.32 x 106 cells/ml x (23 ml) = 99.36 x 106 total cells
  8. Irradiate cells for 8000 rads. This number is highly variable between MEF batches--the idea is to irradiate them enough to stop them from growing, but not enough to kill them. We have used between 5000 and 8000 rads in the past.
  9. Spin cells at 1000 rpm for 5 minutes.
  10. Determine how to dilute cells
    • Total cell number = (Desired concentration) x (# ml needed to dilute)
    • Example: You have 99.36 x 106 total cells, and wish to get your sample to 1.5 x 105 cells/ml. Your calculation will look like this:
    • (99.36 x 106 total cells) = (1.5 x 105 cells/ml)(x ml)
    • x = 662.4 ml needed
  11. 11. Resuspend cells appropriately with MEF medium
  12. Remove 0.1% gelatin from wells and plate with the following volumes of MEF cell suspension:
    • 0.5ml/well 4 well plate
    • 2.5ml/well 6 well plate and 35mm dish

Note - The general densities for plating MEFs are:

iPS Cell Media & Reagents

PDF for iPS media

Freezing andMedia for iPS cells is almost identical to regular ES media, except for the concentration of bFGF which is 100 ng/ml final concentration, 25x as much as regular ES media. Cryopreservation and Resuspension media are the same as for ES cells.

Serum Free Media for human ES cells on MEFs: can last for 14 days

Final Concentration
Amount for 250ml Stock solution
80% DMEM-F12
200ml
20% KO Serum Replacer
50ml
1% Non-essential Amino Acids
2.5ml
1mM L-glutamine (see recipe)
2.5ml
0.1mM b-mercaptoethanol
part of L-glutamine
100ng/ml bFGF
25 ug

iPS cells are very sensitive to quality of MEFs and the freshness of the KOSR. The KOSR you add to your media should not be older than 4-5 days (freezing down aliquots is fine). Bad MEFs with fresh KOSR (or vice versa) and your cells will be okay, but bad MEFs with old KOSR and your cells will look awful.

Splitting iPS cells on MEFs:

PDF for iPS cell splitting

based on splitting onto one plate

  1. Warm collagenase IV split media to 37 °C in a water bath.
  2. Aspirate media off of cell culture plate.
  3. Add 1ml of collagenase split media to 1 well of 6 well plate
  4. Incubate at 37 °C with 5% CO2 for 10 minutes. The cells are ready when the edges of the colony are rounded up and curled away from the MEFs on the plate.
  5. Gently wash cells 3 times with sterile DMEM/F12 while colonies are still attached to plate.
  6. Using a 5ml pipet, scrape and wash the colonies off of the plate.
  7. Break up the colonies by pipetting up and down against the bottom of the tube until just before there appears to be a fine suspension of cells. iPS cells do best with slighly bigger colonies than ES cells.
  8. Plate onto fresh feeder layers (previously washed with PBS). Plate cells by adding 0.4ml per well of a 6 well plate until the last 0.5-0.6ml remain. Add the last remaining volume dropwise to each well.
  9. Make sure the cells are evenly distributed across the entire well.
  10. Place gently in incubator. Again make sure the cell are not disturbed.
  11. Let cells settle overnight in incubator.

Notes:

iPS Cell Certification

The following are test results performed on the iPS cells described in Yu et al., 2007. For information about obtaining these cells please see our Reagents page.