Thomson Lab 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

  • 0.146g L-glutamine
  • 10ml Ca/Mg free PBS
  • 7 ul b-mercaptoethanol

bFGF Stock: Store aliquots at –20 or –70

  • 10ug vial of bFGF
  • 5ml 0.1% Fraction V BSA
  • Aliquot 0.5ml into sterile tubes

0.1% Fraction V BSA:

  • 0.1g Fraction V BSA
  • 100ml PBS with Ca and Mg
  • Filter Sterilize

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

  • Final concentration of 1mg/ml in DMEM-F12 media ie. 0.05g collagenase IV in 50ml DMEM-F12

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
back to top ^

Matrigel Aliquoting and Plating:

PDF for matrigel

Aliquoting Matrigel:

Day one:
  • Put the sterilized tip box (either 200 ml or 1000 ml tips), sterilized microfuge tube container, and appropriate pipettor in -20 °C freezer.
  • Thaw the Matrigel bottle on ice in the 4 °C fridge overnight (until it liquifies).

Day two:
  • Each bottle of Matrigel is at a different density, and it is aliquoted out at 2mg/tube (which is enough to coat one six well plate).
  • Fill an ice bucket and place the Matrigel, along with pipets and tubes on ice to keep them cold*.
    • Example calculation with Matrigel arriving at 13.841 mg/ml:
    • 13.841mg/1ml = 2mg/x ml
    • x = 0.145ml/tube
  • Using good sterile technique, aliquot Matrigel to tubes, switching tips whenever Matrigel seems to be clogging the tip and/or causing the pipet to measure inaccurately. Place tubes on ice as they are finished.
  • When the entire bottle has been aliquoted the tubes are stored in either the –20 °C or –70 °C until ready to be used.

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).

  • Thaw tube overnight on ice at 4 °C.
  • Dilute with 6ml cold basal media and mix well.
  • Add 1ml per well of 6 well plate.
  • Allow plate to sit at room temperature for one hour or overnight at 4 °C.
  • Plate may either be used immediately or stored at 4 °C (plate will be good for at least one week).
  • When ready to use the plate(s), remove the excess liquid and wash once with basal medium.

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 6ml 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 (5ml/well).

*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.
back to top ^

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:
  • High density MEFs for CM last about 2 weeks after plating. Be sure to check them regularly for cell death.
  • bFGF is stable at 4 °C for one month.
back to top ^

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 37 ºC.
  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 ºC 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:
  • Dispase splitting solution lasts approximately 2 weeks and the fresher the solution, the less time it takes for colonies to up off the plates.
  • There is an "art" of cell culture spitting. Cells should be Dispased to effect. Cells shouldn't need to be harshly scraped off of the plates (as is acceptable in feeder layer culture conditions). Cells should be enzymatically treated until the colonies easily slough off the plate. This will give the user the ability to control the colony size. Too little treatment and the resulting harsh scraping will kill many cells.
  • Do not skip wash steps! Any amount of residual Dispase will inhibit cell attachment.
  • Cells should be passaged as larger colonies using this media system than using MEFs. Therefore, do not break up colonies as much by pipeting up and down as many times.
back to top ^

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 15ml 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 25mls 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:
  • Dispase is dissolved in DMEM-F12 basal media.
back to top ^

General Notes on ES Cell Culture:

PDF for general_es
  • hES media has a two week shelf life.
  • hES cells should be cultured in 4, 6, 24, 48, 96 well plates. Growing cells in flasks is not recommended because it is very difficult to scrape cells in flasks.
  • hES cells need to be fed every day with hES media. You can add 2 volumes of media and then not feed for 48 hours (ie. over the weekend.)
  • You should pick differentiated cells off of the plate if more than 5% of your culture is differentiated.
  • Alternatively, you can pick to keep the good colonies if there is massive differentiation within your culture.
back to top ^

Mouse Embryonic Fibrolasts (MEFs) Protocols


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.

back to top ^