UCR Stem Cell Core Protocols

• Stem Cell Culture

  Note: Refer to "Media and Cell Supporting Materials" section for media and cell supporting substrates preparation, and to "MEF Extraction and Culture" section for MEF feeders preparation.

  Passaging of hESCs/iPSC on Vitronectin XF or Lonza L7 hPSC matrix by using Lonza L7 hPSC Passaging Solution

  L7 hPSC passaging solution is a chemically-defined, non-animal origin, non-enzymatic cell detachment formulation based on a hypertonic sodium citrate solution manufactured by Lonza. L7 hPSC passaging solution gently dislodges hESC colonies from a substrate without the need of mechanical manipulation of cultures.

  NOTE: The following passaging procedure is written for cells grown in tissue culture treated 6-well plates, if using other culturing vessels then adjustments should be made to the volumes needed for the specific culturing vessels.

  1. Remove and discard cell culture medium from each well.

  2. Add 1ml/well of 1xPBS and gently wash cells by rocking and swirling the plate back and forth.

  3. Remove and discard 1xPBS.

  4. Add 1ml/well of L7 hPSC Passaging Solution.

  5. Incubate for 5 to 15 minutes at room temperature or for 5 to 10 minutes at 37^oC - 5 minutes in the incubator has shown to be optimal on stem cells grown in the UCR Stem Cell Core:

  NOTE: Treatment times may vary and are dependent on culture confluence, cell line and the type of culture medium and matrix. Monitor cultures until an accurate treatment time has been determined.

  • Monitor culture under the microscope until the following characteristics are observed:
    • Cells are rounding up and gaps begin to appear between groups of cells in the colonies.
    • Cells at the edge of the colonies begin to lift from the plate.
    • A small number of colony aggregates are observed floating in solution.

  6. After incubation, remove and discard the L7 hPSC Passaging Solution.

  ***The majority of the cells should remain attached to the culture surface***.

  Over-treatment of colonies can occur and can result in complete detachment of all cell colonies. If this occurs do the following:

  • Collect passaging solution + cells.
  • Centrifuge for 3 minutes at 800 rpm.
  • Remove supernatant and re-suspend in appropriate amount of media and directly seed the cells onto coated culture vessels.

  7. Tilt the plate and gently detach cellular colonies by rinsing the cell aggregates off the surface with fresh hPSC medium (Lonza

  L7 hPSC media (Catalog# FP-5007), Stem Cell Technologies mTeSR media (Catalog # 05850), or similar)

  NOTE: It is crucial to define the treatment time for each cell line being used. Cells should easily be rinsed off culture surface. Excessive pipetting will result in single cells and therefore reduced cell viability.

  8. Collect the detached cell aggregates and seed directly at desired cell density, onto coated culture vessels with hPSC Matrix (Lonza L7 matrix (Catalog# FP-5020), Stem Cell Technologies Vitronectin XF (Catalog# 07180), or similar).

  • At this point centrifugation is optional. If a cell pellet is desired perform the following:
    • After collecting the detached cell aggregated in desired media centrifuge for 3 minutes at 800 rpm.
    • Remove supernatant and re-suspend in appropriate amount of media and directly seed the cells onto coated tissue culture vessels.

  Passaging of hESCs/iPSC on Matrigel/Geltrex

  1. Aspirate out media.

  2. Add 1ml of DPBS without Mg²⁺/ Ca²⁺ per well to wash the cells briefly.

  3. Remove DPBS and add 750ul of accutase per well and incubate at 37^oC for 1 minute until most cells look like they are loosely attached.

  a. Observe cell detachment under microscope.

  4. Add glass beads to each well (~10-15 glass beads/well) and gently roll the colonies off.

  5. Add equal volume of mTeSR to a 15ml conical and transfer cell suspension (accutase + cells) from each well to the 15ml conical tube.

  6. Add 1ml more of mTeSR to each well to wash off any remaining cells in the well and add that to the 15ml conical tube.

  7. Spin at 800rpm for 3 min at RT.

  a. To the pre-coated 6-well plate: remove substrate completely and add 1ml of mTeSR to each well.

  8. Aspirate supernatant from the tube, leaving cell pellet intact.

  9. Using a 1000ul pipette tip, gently re-suspend the human ES cell pellet in appropriate volume of mTeSR and distribute evenly between wells (drop-wise in and around the wells). a. Passaging ratio should be determined by individual researcher.

  10. Look under microscope to confirm cell clumps are present and carefully place into 37oC/CO2 incubator. Swirl the plate carefully to ensure an even distribution of cells across wells.

  11. Feed cells daily until cells are ready to be split again (when cells reach 80% confluency).

  Freezing of hESCs/iPSC

  Human ES cell freezing medium: 90% FBS, 10% DMSO.

  1. Aspirate out media.

  2. Add 1ml of DPBS without Mg²⁺/ Ca²⁺ per well to wash the cells briefly.

  3. Remove DPBS and add 750ul of Accutase per well and incubate at 37^oC for 1 minute until most cells look like they are loosely attached.

  a. Observe cell detachment under the microscope.

  4. Add glass beads to each well (~10-15 glass beads/well) and gently roll the colonies off.

  5. Add equal volume of mTeSR to a 15ml conical and transfer cell suspension (Accutase + cells) from each well to the 15ml   conical tube.

  6. Add 1ml more of mTeSR to each well to wash off any remaining cells in the well and add that to the 15ml conical tube.

  7. Spin at 800rpm for 3 min at RT.

  8. Aspirate supernatant from the tube, leaving human ES cell pellet intact.

  9. Using a 1000ul pipette tip, gently re-suspend the human ES cell pellet in 500ul of freezing medium/well.

  10.Transfer 500ul of freezing media + cells into a cryogenic vial pre-label each tube.

  11. Place in freezing container and transfer to an -80^oC freezer overnight. Next day transfer to a liquid nitrogen tank for longer cell storage.

  Thawing of hESCs/iPSCs

  1.Remove cryogenic vial from freezer/LN2.

  2.Thaw by immersing the bottom half of the vial in a 37^oC water bath.

  a.Do not leave vial in water bath for more than 1.5 minutes.

  b.Do not immerse the whole tube in the water bath; this could lead to contamination problems.

  3.Using a 5ml serological pipette, take 5 ml of pre-warmed mTeSR and transfer to a 15ml conical tube.

  4.Transfer the 500ul of thawed cells from the cryogenic vial to the 15ml conical tube.

  5.Using a 1000ul pipette add 1ml of mTeSR to wash any remaining cells off the cryogenic vial and add to 15ml conical tube.

  6.Spin at 800 rpm for 3 minutes.

  a.To the pre-coated 6-well plate: remove substrate completely and add 1ml of mTeSR to each well.

  7.Aspirate out supernatant from the tube, leaving the human ES cell pellet intact.

  8.Using a 1000ul pipette tip, gently re-suspend the human ES cell pellet in appropriate volume of mTeSR and distribute evenly between wells (drop-wise in and around the wells).

  9.Look under microscope to confirm cell clumps are present and carefully place into 37^oC/CO2 incubator. Swirl the plate carefully to ensure an even distribution of cells across wells.

  Passaging of hESC/iPSCs on MEF feeders

  1.Coat culture vessel (6 well plate, T25, T75 etc.) with 0.1% gelatin for a minimum of 20 minutes.

  2.Make sure to remove gelatin solution before seeding MEF feeder cells onto culture vessel.

  3.Aspirate out media from the culture vessel that is to be passaged.

  4.Add 1ml of DPBS without Mg²⁺/ Ca²⁺ per well to wash the cells briefly.

  5.Remove DPBS and add 750ul of Accutase per well and incubate at 37^oC for 1 minute until most cells look like they are loosely attached.

  a.Longer incubation with Accutase is not advisable.

  6.Add glass beads to each well (~10-15 glass beads/well) and gently roll the colonies with MEF feeder cells off.

  7.Add equal volume of Human Embryonic Stem cell (HES) media to a 15ml conical and transfer cell suspension (Accutase + cells) from each well to the 15ml conical tube.

  8.Add 1ml more of HES media to each well to wash off any remaining cells in the well and add that to the 15ml conical tube.

  9.Spin at 800rpm for 3 min at RT.

  a.To the pre-coated 6-well plate – pre-coated with 0.1% gelatin: remove substrate completely and add 1ml of HES to each well.

  10.Aspirate supernatant from the tube, leaving cell pellet intact.

  11.Using a 1000ul pipette tip, gently re-suspend the cell pellet in appropriate volume of HES media and distribute evenly between wells (drop-wise in and around the wells).

  12.Look under microscope to confirm cell clumps are present and carefully place into 37^oC/CO2 incubator. Swirl the plate carefully to ensure an even distribution of cells across wells.

  13.Feed cells daily until cells are ready to be split again (when cells reach 80% confluency).

  a.Cells from 1 well on a 6-well plate typically yield 1.5-2 million cells.

Differentiation

• Embryoid Bodies Formation

  Differentiation in suspension culture: Embryoid bodies (EBs)

  EB medum

  The components of the medium used during the differentiation of hESC/iPSC cells can profoundly alter the cell types generated. Thus, the individual researcher should tailor the exact supplements and media type. EBs can be collected at certain time points for mRNA extraction and RT-PCR analysis, or replated on monolayer to let differentiated cells spread out.

  With Collagenase IV

  Collagenase IV solution (1 mg/ml)

  • Weigh out 50 mg of collagenase type IV (Invitrogen, 7104019).
  • Solubilize lyophilized powder in 50 ml DMEM/F12.
  • If 50ml is too much then adjust collagenase IV and DMEM accordingly – adjust to get a final concentration of 1mg/ml.
  • Filter-sterilize with a 0.2 micron cellulose acetate filter.
  • Store at 4oC. Use within 1-2 weeks.

  Note: From batch to batch the activity of the collagenase will be different. Always adjust solution concentration or time of substrate exposure to the enzyme for the change in activity.

  1.Remove media and briefly wash with DPBS without Mg²⁺/Ca²⁺.

  2.Add 1 ml of collagenase IV per well of a 6-well plate (3ml if a T25 flask – adjust accordingly) and place in a 37°C incubator for 60-90 minutes.

  a.Monitor the detachment of cells under the microscope.

  b.Detached cells should look slightly curled on the edges of the colony.

  3.Gently tap the plate to dissociate colonies and to promote complete detachment of cell colonies from the plate.

  4.Add 5 ml of HES media to a 15ml conical tube and gently transfer cell suspension into the 15ml conical tube.

  5.With another 1ml of HES media gently wash the wells to attain any remaining cells and transfer to a 15ml conical tube tube.

  6.Spin at 800rpm for 3 minutes at RT.

  7.Carefully remove the supernatant and gently re-suspend cell colonies in 10 ml of HES medium (without bFGF).

  a.For lineage specific differentiation – growth factors can be supplemented to the HES media.

  8.Transfer to a sterile 10 cm bacterial-grade Petri dish (the surface of bacterial Petri dishes does not facilitate the attachment of the EBs).

  a.Three wells or one T25 of cells should be transferred to one 10 cm bacterial grade Petri dish. If cell pellet is too small, cells from one well of 6-well plate can be transferred to one well of non-attachment 6-well plate – adjust media accordingly.

  9.Place in 37°C incubator and feed EBs every other day (see protocol for feeding EBs).

  a.Monitor differentiation for several days (depending of the lineage of interest).

  With 0.25% Trypsin/EDTA

  1.Pre-treat cell colonies with Y27632 ROCK inhibitor (10uM) for 1hr minimum.

  2.Remove media and briefly wash with DPBS without Mg²⁺/Ca²⁺.

  3.Add 1 ml of 0.25% Trypsin/EDTA per well of a 6-well plate (3ml if a T25 flask – adjust accordingly) and place in a 37°C incubator for 3 minutes maximum.

  a.Monitor the detachment of cells under the microscope.

  4.Gently tap the plate to promote complete detachment of cell colonies from the plate.

  5.Add 9 ml of MEF media to a 15ml conical tube and gently transfer the cell suspension into the 15ml conical tube.

  a.9ml of MEF media is sufficient enough to inactivate the trypsin.

  b.FBS can also be used to inactivate trypsin at 1ml FBS to 1ml 0.25% Trypsin/EDTA

  6.Spin at 800rpm for 3 min at RT.

  7.Carefully remove the supernatant and gently re-suspend cell colonies in 10 ml of hES medium (without bFGF and supplemented with Y27632 ROCK inhibitor (10uM)).

  a.For lineage specific differentiation – growth factors can be supplemented to the HES media.

  b.Y27632 ROCK inhibitor is needed to promote cell survival for the first and second day of EB growth. Y27632 ROCK inhibitor can be removed on the third day of culture.

  8.Transfer to a sterile 10 cm bacterial-grade Petri dish (the surface of bacterial Petri dishes does not facilitate the attachment of the EBs).

  a.Three wells or one T25 of cells should be transferred to one 10 cm bacterial grade Petri dish. If cell pellet is too small, cells from one well of 6-well plate can be transferred to one well of non-attachment 6-well plate – adjust media accordingly.

  9.Place in 37°C incubator and feed EBs every other day (see protocol for feeding EBs).

  a.Monitor differentiation for several days (depending of the lineage of interest).

   

  Feeding Embryoid Bodies (EBs)

  EBs require feeding every 2-3 days, depending upon density and size.

  1.Gently transfer the EBs along with old medium to a sterile 15ml conical tube.

  2.Allow the EBs to settle down under gravity at room temperature (takes 5-10 minutes – depending on size of EB, it may take longer).

  3.Aspirate old medium and add new medium gently.

  a.Take care not to break up the EBs.

  4.Transfer to a new sterile 10 cm bacterial-grade Petri dish or any other low attachment plate.

  5.Place in 37°C incubator and feed EBs and continue feeding EBs every other day.

  Re-plating Embryoid Bodies (EBs) in monolayer

  EBs can be re-plated onto uncoated TC treated plastic and various substrates (i.e. Poly-D-Lysine, Geltrex, Matrigel, fibronectin, etc.)

  1.Gently transfer the EBs along with old medium to a sterile 15ml conical tube.

  2.Allow the EBs to settle down under gravity at room temperature (takes 5-10 minutes – depending on size of EB, it may take longer).

  3.Aspirate old medium and add new medium gently to which the EBs will be plated with.

  a.Take care not to break up the EBs.

  4.Transfer to desired coated or uncoated culture vessel.

  a.The amount of media should only be enough to coat the plating surface – this allows the EBs to be closer to the plating surface and encourages EB attachment to surface.

  Note: Some helpful websites for 3D Culture: 3D Biomatrix

• Direct Differentiation

  Direct differentiations were carried out in RPMI (Mediatech Inc., Manassas, VA) with varying concentrations of defined FBS (HyClone, Logan, UT):

  • 0.0% at days 0–1
  • 0.2% at days 1–3
  • 2.0% at days 3–4

  For definitive endoderm differentiation,

  cells were treated with 100 ng/mL Activin A for 4 days, and 25 ng/mL Wnt3a from days 0 to 1 only.

  For extraembryonic differentiation,

  cells were treated with 100 ng/mL BMP4 and 3 nM FGFR inhibitor PD173074 (EMD Chemicals Inc., Gibbstown, NJ) for 4 days.

  For ectodermal differentiation,

  cells were treated with 100 ng/mL Noggin and 5 μM ACTR inhibitor (SB431542; Sigma Aldrich, St. Louis, MO) for 4 days.

  Note: Start with confluent ES cell culture for best differentiation efficiency.

• MEFs Extraction and Culture

  Deriving Mouse Embryonic Fibroblasts (MEFs)

  An established working protocol will be provided up on request.

  Passaging of Mouse Embryonic Fibroblasts

  At 90% -- 95% confluency:

  1.Remove media and briefly wash with DPBS without Mg²⁺/ Ca²⁺.

  2.Remove DPBS without Mg²⁺/ Ca²⁺ and add 3ml of Trypsin 0.25%/EDTA into a T75 flask (1ml of Trypsin 0.25%/EDTA into a T25 flask).

  3.Incubate at 37^oC for 3 minutes.

  4.To promote cell detachment tap the flask against the palm of your hand or against the bench top. Look through a microscope to make sure that the MEF cells are completely detached. a.Avoid over incubating MEF cells with Trypsin 0.25%/EDTA, doing so can cause an increase in cell death.

  5.Inactivate trypsin:

  a.With MEF medium (9 ml MEF to 1 ml trypsin).

  b.FBS (1ml of FBS to 1ml of Trypsin 0.25%/EDTA).

  6.Transfer cell suspension into a 15ml conical tube and centrifuge at 1000rpm for 5 minutes.

  7.Aspirate supernatant from the tube, leaving MEF cell pellet intact – gently flick the tube to disperse the cell pellet.

  8.Re-suspend the cell pellet with 3ml of MEF media and distribute the cells evenly to 3 T75 flasks containing 14ml of MEF media.

  a.This is a 1:3 passage dilution, higher passaging ratios are not recommended.

  b.Culture vessels do not need a substrate, the MEF cells will attach to the plastic just fine.

  9.MEF cells can be grown out for 3 passages before inactivation into feeder cells.

   

  Freezing of Mouse Embryonic Fibroblasts (MEFs)

  Materials -freezing Media - 90% FBS,10% DMSO.

  At 90% -- 95% confluency:

  1.Remove media and briefly wash with DPBS without Mg²⁺/Ca²⁺.

  2.Remove DPBS without Mg²⁺/ Ca²⁺and add 3ml of Trypsin 0.25%/EDTA into a T75 flask (1ml of Trypsin 0.25%/EDTA into a T25 flask)

  3.Incubate at 37^oC for 3 minutes.

  4.To promote cell detachment tap the flask against the palm of your hand or against the bench top. Look through a microscope to make sure that the MEF cells are completely detached.

  a.Avoid over incubating MEF cells with Trypsin 0.25% EDTA, doing so can cause an increase in cell death.

  5.Inactivate trypsin:

  a.With MEF medium (9 ml MEF to 1 ml trypsin).

  b.FBS (1ml of FBS to 1ml of Trypsin 0.25%/EDTA.

  6.Transfer cell suspension into a 15ml conical tube and centrifuge at 1000rpm for 5 minutes.

  7.Aspirate supernatant from the tube, leaving MEF cell pellet intact.

  8.Re-suspend the cell pellet in 500ul of freezing media.

  9.Transfer 500ul of freezing media + cells into a pre-labeled cryogenic vial.

  10.Place in freezing container and transfer to an -80^oC freezer overnight.Next day place the frozen tubes with cells in liquid nitrogen tank for longer storage.

   

  Thawing of Mouse Embryonic Fibroblasts

  1.Remove cryogenic vial from freezer/LN2.

  2.Thaw by immersing the bottom half of the vial in a 37oC water bath. Do not leave vial in water bath for more than 1.5 min. Do not immerse the whole tube in the water bath, this could lead to contamination problems.

  a.Using a 5ml serological pipette, take up 5 ml of pre-warmed MEF media and transfer to a 15ml conical tube. Transfer the 500ul of thawed cells from the cryogenic vial to the 15ml conical tube.

  3.Using a 1000ul pipette add 1ml of MEF media to wash any remaining cells off the cryogenic vial and add to 15ml conical tube.

  4.Spin at 1000 rpm for 5 minutes.

  5.Aspirate out supernatant from the tube, leaving the MEF cell pellet intact.

  6.Using a 1000ul pipette tip, gently re-suspend the MEF cell pellet in appropriate volume of MEF media and distribute evenly culture vessels (T25 – T75 flasks).

  a.Culture vessels do not need a substrate, the MEF cells will attach to the plastic just fine.

  7.Look under microscope to confirm cells are present and carefully place into 37^oC/CO2 incubator. Swirl the plate carefully to ensure an even distribution of cells across culture vessels.

  Preparation of Mouse Embryonic Fibroblast (MEF) feeder layers

  Note: Feeder cells should be prepared from P3 MEF cells. To obtain decent feeder cells, it is essential that the MEF cells be growing in log phase before growth innactivation. MEF cells can be mitotically inactivated by either mitomycin C treatment or by irradiation.

  Mitomycin C treatment

  To a 90% confluent P3 MEF cell culture:

  1.Aspirate out media and add MEF media supplemented with mitomycin C. Final concentration of mitomycin C should be 10ug/ml.

  2.Place into 37^oC/CO2 incubator and incubate MEF cells with MEF media supplemented with 10ug/ml mitomycin C for 1.5 hours.

  3.After incubation, remove MEF media supplemented with 10ug/ml mitomycin C and wash with DPBS without Mg²⁺/ Ca²⁺ (15ml of DPBS without Mg²⁺/Ca²⁺ for a T75 flask/ 5ml for a T25 flask) – swirl the flask and remove the DPBS without Mg²⁺/Ca²⁺). Repeat washing 3 additional times.

  a.IMPORTANT – sufficient washing with DPBS without Mg²⁺/Ca²⁺ is required to remove any residual amount of mitomycin C. Inadequate washing can yield unsuitable feeder cells.

  10.After washing, add 3ml of Trypsin 0.25%/EDTA into a T75 flask (1ml of Trypsin 0.25% EDTA into a T25 flask) and incubate at 37^oC for 3 minutes.

  11.To promote cell detachment tap the flask against the palm of your hand or against the bench top. Look through a microscope to make sure that the MEF cells are completely detached.

  a.Avoid over incubating MEF cells with Trypsin 0.25%/EDTA, doing so can cause an increase in cell death.

  12.Inactivate trypsin:

  a.With MEF medium (9 ml MEF to 1 ml trypsin.

  b.FBS (1ml of FBS to 1ml of Trypsin 0.25% EDTA.

  13.Transfer cell suspension into a 15ml conical tube.

  14.Perform a cell count:

  a.Cell number to be frozen down per cryogenic vial must be determined by individual researcher based on their use of the cells.

  15.Spin at 1000rpm for 5 minutes.

  16.Aspirate supernatant from the tube, leaving MEF cell pellet intact.

  17.Re-suspend the cell pellet in appropriate amount of freezing media so that each cryogenic vial contains 500ul of freezing medial + desired cell number.

  18.Transfer 500ul of freezing media + cells into a cryogenic vial and label each tube accordingly and place in freezing container and transfer to an -80oC freezer overnight followed by transfer to liquid nitrogen tank for longer storage.

  Preparation of mouse embryonic fibroblast (MEF) feeder layers for hESC/iPSC culture

  Note: Once MEF cells are mitotically inactivated, a substrate is needed to promote attachment. The substrate used to promote MEF feeder cell attachment is 0.1% gelatin. MEF feeder cells can be seeded onto culture vessels from:

  a) a previously frozen MEF feeder cell vial.

  b) directly after inactivation.

  If from a previously frozen vial:

  1.Coat culture vessel (6-well plate, T25, T75 etc.) with 0.1% gelatin for a minimum of 20 minutes.

  a.Make sure to remove gelatin solution before seeding MEF feeder cells onto culture vessel.

  2.Remove cryogenic vial from freezer/LN2.

  3.Thaw by immersing the bottom half of the vial in a 37^oC water bath. Do not leave vial in water bath for more than 1.5 minutes. Do not immerse the whole tube in the water bath, this could lead to contamination problems.

  a.Using a 5ml serological pipette, take up 5 ml of pre-warmed MEF media and transfer to a 15ml conical tube. Transfer the 500ul of thawed cells from the cryogenic vial to the 15ml conical tube.

  4.Using a 1000ul pipette add 1ml of MEF media to wash any remaining cells off the cryogenic vial and add to 15ml conical tube.

  5.Perform a cell count.

  6.Spin at 1000 rpm for 5 minutes.

  7.Aspirate supernatant and seed MEF feeder cells to pre-coated culture vessels.

  a.Seed MEF cells at 1x105 cells per well of a 6-well plate; 2 X 105 cells per T25 flask, etc.

  8.Gently re-suspend the MEF cell pellet in appropriate volume of MEF media so that each culture vessels has optimal number of cells and distribute evenly culture vessels (6-well plate, T25 – T75 flasks etc).

  a.Make sure that each culture vessel has appropriate amount of media (1ml for a well of a 6 well plate, 3ml for a T25 flask, etc).

  9.Look under microscope to confirm cells are present and carefully place into 37oC/CO2 incubator. Swirl the plate carefully to ensure an even distribution of cells across culture vessels.

  10.Feeders are ready to be used after overnight incubation (minimum of four hour incubation).

  a.Good feeder layers should be 70-80% confluent with spaces available around the cells. Human ES cells attach to the space between the feeders and push the feeder cells when they proliferate and expand as colonies.

  If directly after inactivation:

  1.Coat culture vessel (6 well plate, T25, T75 etc.) with 0.1% gelatin for a minimum of 20 minutes.

  a.Make sure to remove gelatin solution before seeding MEF feeder cells onto culture vessel

  After inactivation perform a cell count.

  a.Seed MEF cells at 1x105 cells per well of a 6-well plate; 2 X 105 cells per T25 flask, etc).

  3.Gently re-suspend the MEF cell pellet (MEF cell pellet acquired after inactivation process) in appropriate volume of MEF media so that each culture vessels has optimal number of cells and distribute evenly culture vessels (6-well plate, T25, T75 flasks etc.).

  a.Make sure that each culture vessel has appropriate amount of media (1ml for a well of a 6- well plate, 3ml for a T25 flask, etc.

  4.Look under microscope to confirm cells are present and carefully place into 37^oC/CO2 incubator. Swirl the plate carefully to ensure an even distribution of cells across culture vessels.

  5.Feeders are ready to be used after overnight incubation (minimum of four hour incubation).

• Media and Cell Supporting Materials Preparaton

  MEF Media:

  DMEM (high glucose)
  10% fetal calf serum (GIBCO, Origin: USA)
  1X L-glutamine or Glutamax
  1X non essential amino acid
  1X sodium pyruvate

  Human Embryonic Stem Cell (HES) Growth media

  For 500 ml HES medium with 20% KOSR:

  400ml KnockOut DMEM/ F12 (Life Technologies, Cat#12660-012)
  100ml KOSR
  5ml 1X Glutamax I (100x)
  5ml Non-Essential Amino Acid solution (NEAA) (100x)
  500ul 2-β mercaptoethanol
  bFGF 8-10 ng/ml

  Human Embryonic Stem Cell (HES) Differentiation media

  For 500 ml Differentiation medium with 10% FBS:

  450ml Knockout DMEM/ F12 (Life Technologies, Cat#12660-012)
  50ml FBS
  5ml 1X Glutamax I (100x)
  5ml Non-Essential Amino Acid solution (NEAA) (100x)
  500ul 2-β mercaptoethanol

  BD Matrigel™ – hESC-qualified Matrix,LDEV-Free (BD Catalog # 354234)

  Note: To avoid premature gelling of substrate thaw the bottle of matrigel overnight at 4^oC. Once matrigel is completely thawed at 4^oC: Aliquot as 1ml aliquots and dilute 1:30 with DMEM or add equal volume of DMEM, aliquot 1ml aliquots, and dilute 1:15.  Final dilution of matrigel should be 1:30.

  Coating with BD Matrigel™

  • Add 1ml of diluted matrigel per well of a 6-well plate (if using another culture vessel – make sure to add appropriate volume to coat the bottom of the vessel).
  • Gently shake the plate to spread the coating solution across the entire surface.
  • Incubate at room temperature for a minimum of 1hr – 1.5hr. At 37^oC for maximum 1hr.
  • Culture vessels can be stored at 4^oC for one week.

  Geltrex™ LDEV-Free hESC-qualified Reduced Growth Factor Basement Membrane Matrix (Life technologies,Catalog # A1413302)

  Note: Geltrex is only used as a substrate for the RIV9 iPSC line (all other hESC and iPSC cell lines are grown on Matrigel).

  • To avoid premature gelling of substrate thaw the bottle of Geltrex overnight at 4^oC.
  • Once Geltrex is completely thawed at 4oC it can be used by diluting the substrate 1:100 with DMEM.

  Coating with Geltrex™

  • Add 1ml of diluted Geltrex per well of a 6-well plate (if using another culture vessel make sure to add appropriate volume to coat the bottom of the vessel).
  • Gently shake the plate to spread the coating solution across the entire surface.
  • Incubate at 37^oC for a minimum of 1hr.
  • Plates are ready to use after 1hr incubation at 37^oC. Plates can be stored in the incubator as long as the Geltrex does not evaporate and wells dry up.

  Stem Cell Technologies Vitronectin XF

  Vitronectin XF manufactured by Stem Cell Technologies (SCT) is xeno-free alternative to Matrigel and Geltrex. The use of Vitronectin XF  results in consistent cell growth of hESCs and iPSCs. It is currently our matrix of choice for growing human pluripotent stem cells at the Core.

  Coating Culture Vessels with Vitronectin XF

  Although SCT recommends the use of non-tissue culture treated vessels for the use of Vitronectin XF, we have used tissue culture treated vessels and have obtained satisfactory results. The following coating procedure is for tissue culture treated 6-well plates, if using other culturing vessels then adjustments should be made to the volumes needed to coat the specific culturing vessel.

  • Thaw Vitronectin XF at room temperature (15-25^oC) or at 4^oC overnight.

  To avoid additional freeze-thaw cycles aliquot appropriate amounts of Vitronectin XF into 1.5ml micro-centrifuge tubes and store at -20^oC or -80^oC. Vitronectin XF is stable for two weeks at 4^oC. Once is thawed, a final concentration of 10ug/ml is needed to coat culture vessels.

  • Dilute Vitronectin XF (250ug/ml) with 1xPBS (w/o Ca²⁺&Mg²⁺) to reach a final concentration of 10ug/ml (i.e. use 40 ul of Vitronectin XF per 1ml of PBS). Use an appropriate polypropylene conical tube (15ml or 50ml) to dilute the Vitronectin XF.
  • Gently mix the diluted Vitronectin XF inverting the tube 2-3 times and immediately use the diluted Vitronectin XF solution to coat tissue culture treated 6-well plates.
  • Add 1ml/well of the 6-well plate and gently rock and swirl the 6-well plate back and forth to spread the Vitronectin XF solution evenly across the surface.

  Note: if using non-tissue culture-treated vessels for coating with Vitronectin XF make sure the surface is fully coated by the Vitronectin XF solution - additional rocking and swirling may be required in order to coat the hydrophobic surface.

  • Incubate at room temperature for at least 1 hour before use. Do not let the Vitronectin XF solution evaporate.

  Note: if not used immediately, the culturing vessels can be stored at 4^oC as long as evaporation of the Vitronectin XF does not occur. Six-well plates can be sealed with parafilm to prevent evaporation of the Vitronectin XF. Allow coated culture vessels to come to room temperature for 30 minutes before use.

  • Remove excess Vitronectin XF solution.
  • Add 1ml/well of mTeSR media - cells can then be added to each well at desired passage ratio.

  Lonza L7 hPSC Matrix

  Lonza's recombinant hPSC Matrix is a xeno-free defined matrix suitable for teh culture of hPSCs. Lonza L7 hPSC Matrix arrives in powder form and is stable at -20^oc to -80^oC. Once is reconstituted, Lonza L7 hPSC Matrix  is stable for up to 3 months. To avoid repeated freeze-thaw cycles, prepare appropriate stocks (see below for recommended protocol) and store at -20^oC to --80^oC. 

  Preparation of L7 hPSC Matrix Stock Solution

  The following coating procedure is written for tissue culture treated 6-well plates, if using other culture vessels then adjustment should be made to the volumes needed for the specific culturing vessel.

  1. Remove the vial of Lonza L7 hPSC Matrix from -20^oc to -80^oC and briefly centrifuge to collect powder at the bottom of the vial.

  2. Add 1ml of 1xPBS (Lonza Catalog No. 17-516F, or similar) to the vial and and gently swirl to mix. 

  NOTE: This will be your stock solution of Lonza L7 hPSC Matrix, that can be further diluted prior to coating vessels. Avoid freeze-thaw cycles by aliquoting appropriate volumes and store at -20^oc to -80^oC for up to 3 months.

  3. When ready to coat a plate, thaw an aliquot of the L7 hPSC Matrix at room temperature and dilute the L7 hPSC Matrix stock solution 1:200 with 1xPBS.

  4. Add 1ml of diluted L7 solution and tilt teh 6-well plate from side to side to ensure complete coverage of surface area.

  5. Incubate at 37^oC for at least one hour.

  NOTE: Overnight incubation for next day use is acceptable, however, leaving the vessel at 37^oC longer then 24 hours is not recommended. Vessels that have been coated at 37^oC may be stored at 4^oC to room temperature prior to plating cells.

  6. Remove coating solution prior to addition of cells onto wells. There is no need to was wells with PBS prior to addition of media + cells.

  L7 hPSC BulletKit Medium

  Lonza L7 hPSC BulletKit Medium is a complete, xeno-free medium for everyday or every-other-day maintenance of hPSCs.

  Store basal media at -20^oc to -80^oC an Supplement at -20^oc to -80^oC. Once the supplement is thawed it should be stored at 2^o-8^oC and added to basal medium within 72 hours. After supplement is added to basal medium use within two weeks. 

  Preparation of Culture Media

  The following procedure is written for a 500ml bottle of media. Smaller amounts can be made by proper adjustment of media and supplement.

  1. Thaw L7 hPSC Supplement (Catalog No. FP-5107) at room temperature or overnight at 4^oC.

  2. Transfer the contents of the L7 hPSC Supplement to L7 hPSC Basal Medium with a pipette and rinse the vial with medium.

  NOTE: If sterility is a concern the prepared growth medium may be re-filtered with a 0.2um filter to assure sterility. Routine re-filtration is not recommended.

  Maintenance

  1. Change the growth medium  (warm media at room temperature) the day after seeding and every 2 days thereafter. Add 1ml per well medium for the day after re-plating the cells - adjust volumes accordingly with cell growth.

  NOTE: Avoid repeated warming and cooling of the medium. If the entire contents are not needed for a single procedure, transfer and warm only the required volume to a sterile , secondary container.

  2. Over time, cells within each colony will become smaller and more compact. Some cells with more epithelial morphology may be observed surrounding a colony, if the hPSCs have been directly transferred to grow in Lonza L7 hPSC medium from another media composition (for example, adaptation of hPSC, previously cultivated with mTeSR medium (SCT Catalog No.05850) may be required).

  3. Passage cells when they are 90% confluent.

  NOTE: For best results, change the medium the day before and the day after passaging the cells. Medium may also be changed daily by the user, if preferred.

• Immunocytochemistry

  Immunocytochemistry General Protocol

  1.Remove media and briefly wash cells (cell can be grown on chamber slides or on 24, 12, 6- well plates) with PBS w/ Mg²⁺ and Ca²⁺. Repeat twice.

  a.Cells are washed with PBS w/ Mg²⁺ and Ca²⁺ to minimize cell detachment.

  2.Remove PBS w/ Mg²⁺ and Ca²⁺ and add 1ml of 4% paraformaldehyde (PFA) to fix the cells.

  a.1ml of PFA is sufficient for one well of a 6-well plate – adjust PFA volume according to culture vessel being used.

  3.Incubate/fix the cells for 10 to 20 minutes at room temperature depend on the antigen.

  a.ALWAYS incubate at room temperate – DO NOT attempt to place cells back into incubator.

  b.Cells grown on same slides/plates must be fixed at the same time, cells cannot be grown after cells on a nearby well have been fixed.

  4.Remove 4% PFA.

  5.Wash the fixed cells with PBS w/o Mg²⁺ and Ca²⁺ for 5 minutes each. Repeat 2 additional times.

  6.Permiabilize with 0.1-0.2% TritonX-100 for 10-20 minutes.

  For example,

  For cell surface marker SSEA4

  -use wash buffer without 0.1% Triton-X100;

  For nuclear transcription factors

  -use wash buffer with 0.1-0.2%% Triton-X to permeabilize the cell membrane as this is an intracellular marker.

  7.Repeat step 5.

  8.Remove PBS w/o Mg²⁺ and Ca²⁺ and block the cells with blocking buffer (PBS w/o Mg²⁺ and Ca²⁺ + 1-2% Goat Serum or 1-2% Donkey Serum depend on the secondary antibody.

  9.After blocking – remove blocking buffer and wash the cells with DPBS w/o Mg²⁺ and Ca²⁺ and incubate for 10 minutes.

  10.Dilute antibody in DPBS w/o Mg²⁺ and Ca²⁺ and incubate fixed cells with primary antibody at 4^oC overnight. Alternatively the antibodies can be diluted in 0.1-0.2% blocking serum in PBS.

  Note: To obtain working dilutions for different antibodies used at the Core click here.

  11.The next day, wash the cells with PBS w/o Mg²⁺ and Ca²⁺ and incubate for 10 minutes. Repeat 3 additional times.

  12.Incubate the pre-stained cells with the appropriate secondary antibodies for 30 minutes to 1 hour based on the antibody manufacturer recommendations.

  13.Wash the cells with PBS w/o Mg²⁺ and Ca²⁺ for 10 minutes. Repeat 3 additional times.

  14.Mount the stained cells with DAPI mounting solution.

  15.Cells are ready to be observed under a microscope.

• Flow cytometry

  General rules for cell analysis on a flow cytometer for avoiding clogs:

  • You must pass all cells trough a cell strainer  after labeling and before you bring them to the flow cytometer. Choose a cell strainer based on the size of the harvested cells. For an example of cell strainers please click here .
  • It is highly recommended to pass hESCs trough a cell strainer twice, before and after labeling.
  • Always vortex cells well before running them trough the flow cytometer. 

   

  General protocol for dissociating and labeling embryoid bodies for flow cytometry analysis

  1. Collect embryoid bodies in 15ml falcon tubes.

  2. Let them settle for 5-10 minutes.

  3. Aspirate supernatant .

  4. Add 1ml 0.25%Trypsin/EDTA in PBS.

  5. Incubate 5 minutes at 37⁰C. Occasionally vortex for at least three times during incubation.

  6. Quench trypsin activity with 2ml serum.

  7. Pass through 5ml syringe with 20G needle 6-8 times.

  8. Add 3ml PBS/0.5%BSA and pass through a 45-70um cell stainer, depend on the size of the cells.

  9. Centrifuge for 5 minutes at 1200rpm.

  10. Re-suspend in 3ml PBS/0.5%BSA and count the cells. Distribute the desired amount of cells in FACS tubes. Adjust the volume to 2ml with PBS/0.5%BSA.

  11. Centrifuge the cells for 5 minutes at 1200rpm. Aspirate the supernatant and perform immunostaining for membrane or intracellular antigens.

  NOTE: The same protocol could be used for harvesting clumping hESCs. Start the protocol from point N^o4. In point N^o5 omit the vortex. In point N^o7 pass the cells through the syringe for 3-5 times instead of 6-8 times.

   

  General protocol for labeling cell surface antigens

  1.Harvest cell culture as single cells by using 0.25% Trypsin/EDTA (Invitrogen) or milder cells dissociation reagent based on antigen characteristics.

  2. Pass cells trough a 45-70um strainer and count. Distribute equal number of cells per tube for labeling. Use conical tubes. Consider to not work with less then 1x10⁵ cells/tube. 1x10⁶ /tube is a safe number of cells for labeling when considering that some cells loss will occur during the washing steps.

  3.Incubate with primary antibody (5-20ug/ml) for 1hr. Mix the cells every 30 minutes by gently flipping the tube. Determine antibody concentration based on the antibody characteristics and manufacturer's recommendations.

  5.Wash with 0.5% BSA/PBS. Centrifuge at 1000rpm for 5-7 minutes to collect the cells. Remove the supernatant and gently re-suspend the pellet in 0.5% BSA/PBS. Repeat this step several times.

  6.Incubate with secondary (detection) antibody for 30min to 1hr. Use dilutions recommended by the manufacturer.

  7.Wash with 0.5% BSA/PBS several times as recommended in step 5. The last time wash with PBS only.

   

  General protocol for labeling intacellular antigens

  1.Harvest cell culture as single cells by using 0.25% Trypsin/EDTA (Invitrogen).

  2. Pass cells trough a 70um strainer and count. Distribute equal number of cells per tube for labeling. Use conical tubes. Consider to not work with less then 1x10⁵ cells/tube. 1x10⁶ /tube is a safe number of cells for labeling when considering that some cells loss will occur during the washing steps.

  3.Fix and permeablize cells by using IntraPrep permeabilization reagent kit (PN IM2388; Beckman Coulter) according to the manufacturer's instructions.

  a. Use 5% normal serum for blocking in the presence of the permeabilization reagent, for 20 minutes at room temperature.

  4.Incubate with primary antibody (5-20ug/ml) for 1–2 h at room temperature. Determine antibody concentration based on the antibody chracteristics and manufacturer's recommendations.

  5.Wash with 0.5% BSA/PBS. Centrifuge at 1000rpm for 5-7 minutes to collect the cells. Remove the supernatant and gently re-suspend the pellet in 0.5% BSA/PBS. Repeat this step several times.

  6.Incubate with secondary (detection) antibody for 1hr. Use dilutions recommended by the manufacturer.

  7.Wash with 0.5% BSA/PBS several times as recommended in step 5. The last time wash with PBS only.

• Beta Testing of Lonza L7 HPSCs Xeno-Free Medium System

  White paper Lonza L7 kit (Please click here to read) 

  RIV9 iPS cells white paper  images in ppt.format

  Videos #1 was taken at a pre-selected custom point in one well of 6-well plate on Nikon Biostation CT. Each of the 12 frames represents  a time point taken every 6 hours period. The magnification was set at 10x, phase. RIV9 iPS cells were maintained for 14 passages in Lonza 7 HPSC conditions. The first imaging frame is generated at passage 14 in 30 hours after the cells were passaged. The scale bar was placed on the bottom right side of the video.

  The videos #1 are attached one for PC users (click here) and one for MAC users (click here). The videos could be open by QuickTime Player software.

  Videos #2 was taken by applying Nikon full well scan protocol on one well of a 6-well plate on Nikon Biostation CT. Each of the 11 frames represents  a time point taken every 6 hours period. The magnification was set at 10x, phase. RIV9 iPS cells were maintained for 14 passages in Lonza 7 HPSC conditions. The first imaging frame is generated at passage 14 in 30 hours after the cells were passaged. The scale bar was placed on the bottom right side of the video.

  The videos #2  are attached one for PC users (click here) and one for MAC users (click here). The videos could be open by QuickTime Player software.

  Videos #3 is a snap shot of a cropped area from video#2.

  The videos #2  are attached one for PC users (click here) and one for MAC users (click here). The videos could be open by QuickTime Player software.