Nucleofection of mouse iPSCs, colony picking, and genotyping

Notes:
  • Warm all the 4C reagents (STOP buffer, IMDM, culture media) to 37C before using. TrypLE Express and DPBS are kept at room temp.
  • Lonza kit = Amaxa P3 Primary Cell 4D-Nucleofector X Kit L (cat# V4XP-3024)
Day 1. Harvest miPSCs & setup nucleofection
  1. Coat a 6-well culture plate with 0.1% gelatin (2mL/well). Incubate for 30 min at 37C.
  2. Prepare Lonza nucleofection P3 solution + supplement in a sterile Eppendorf tube. Allow to warm to room temp in the hood.
    • Create a master mix based on a 1x volume of 82uL P3 primary solution + 18uL NS supplement per reaction
    • Once mixed, can be stored up to 3 months at 4C
  3. Obtain plasmids from -20C and thaw on ice.
    • AAV-Flpe-mCherry + pCol-FRT plasmid containing transgene of interest
    • Will need 1.5ug of each plasmid per reaction (3ug total)
  4. Harvest miPSCs from culture plate using TrypLE express and STOP buffer.
  5. Spin down cells and resuspend in 10mL IMDM, then plate onto a sterile 10cm dish. Swirl gently to allow even coating of the dish. Place in incubator at 37C for 15 min to deplete feeders from suspension.
    • If the pellet is large after spin, can split the cells to 2x 10cm dishes (in 10mL IMDM each)
  6. Remove 10cm dish from incubator and gently swirl a few times. Tilt the dish at a 45 degree angle to collect iPSC cell suspension at the bottom.
    • Feeders will mostly remain attached to the plastic (visible as a thin film)
  7. Keep the dish at an angle and gently rinse the base with the cell suspension before transferring the full volume back to the falcon tube.
  8. Take 10uL cells to an Eppendorf and count. Determine the volume needed for 1-3M cells per nucleofection reaction.
  9. Transfer appropriate volume of cells for nucleofection to a new 15mL falcon tube.
  10. Spin down cells at 1500 rpm for 4 min at room temp.
    • During spin, aspirate the gelatin from the 6-well plate and add 2mL fresh, warm 2i media to each well—replace in the incubator to keep warm
  11. Resuspend cells in prepared nucleofection solution. Will need 100uL per nucleofection.
    • Work quickly from this point to minimize time in nucleofection solution as it can decrease viability if left for prolonged time
  12. Transfer 100uL cells in nucleofection solution to sterile Eppendorf tubes.
    • Each tube will correspond with a nucleofection reaction in the Lonza cuvette
  13. Add appropriate amounts of pCol-FRT-transgene and AAV-Flpe-mCherry plasmids to each reaction tube. Pipette gently to mix.
    • Reaction tubes (both plasmids) get 1.5ug of each plasmid
    • Negative control tubes (pCol-FRT-transgene only, no AAV-Flpe) will only get 3ug of FRT-insert plasmid
  14. Gently transfer 100uL cell/plasmid mixture from each Eppendorf tube to a large nucleofection cuvette. If bubbles form, try to gently pop with the pipette.
  15. Take cuvettes to Amaxa Nucleofector machine with attached X unit.
  16. Select the CG-104 program, then select the icon for large cuvettes.
    • Load 2 cuvettes at a time and press Start
    • Continue loading pairs of cuvettes until all reactions have completed
  17. Bring cuvettes back upstairs and carefully transfer nucleofected cells dropwise to prepared culture plate using the plastic transfer pipettes included in the kit.
    • Using one pipette per cuvette, carefully pull up all cells and drip into one well of the gelatinized plate containing 2mL of 2i media
  18. Label plate, check cell density under microscope, and incubate overnight at 37C. Cells should become very dense in all wells and reach near confluence over the next couple of days.
Day 2

Carefully aspirate spent media and refresh with 2mL/well of fresh, pre-warmed 2i media.

Days 3-6. Perform hygromycin selection
  1. On day 3, prepare fresh hygromycin media by adding 2.8uL working stock hygromycin B per 1mL of 2i media.
    • Usually make several days’ worth of hygro media on day 3
  2. Aspirate spent media from the nucleofection plate and wash gently with 1mL/well of DPBS.
  3. Feed cells with pre-warmed 2i + hygro media (2mL/well). Incubate plate at 37C.
  4. On Days 4-6, repeat steps 2-3.
    • Hygro selection causes many cells to detach and float, so DPBS wash helps clear out any clumps that form
Days 7-14. Monitor wells for colony formation
  • On day 7, switch back to normal 2i media (without hygro) for daily feeds and begin to monitor the plate visually for formation of colonies. DPBS wash prior to media change may still be needed as cells continue to die from the selection.
  • “Good colonies” have a compact, tight border and should be fairly round in shape. They may have a slightly darker center and will grow quickly. Often, they form around the edges of the wells.
  • In general, colonies begin forming around day 8 but may also form in the negative control wells (no AAV-Flpe = should not have hygro resistance). If many colonies are present in both wells, perform a second 2-day hygro pulse on days 10-12.
  • After second selection step, any remaining colonies that still look good on day 12 can be marked for picking. Most “background” colonies in the negative control wells should be cleared out after this second treatment.
Day 12-14. Picking colonies for genotyping & expansion
Notes:
  • Once good colonies have been identified, it is helpful mark their location on the bottom of the plate with a fine-tip marker. If some colonies do not look healthy, mark these to avoid.
  • The best picking method is to wait until the colonies have grown large enough that they can be seen by eye and picked directly off the plate without a TC sterile scope. As long as the good ones are marked early, they can remain in culture a few extra days to grow larger. Note, however, that some background colonies may begin forming at day 14 that will likely not contain the insert and should not be picked.
  • The day before picking, plate feeders to a 24-well plate (one well for each colony). On picking day, check that the feeders have adhered at the correct density to support iPSCs.
  • Warm all the 4C reagents (STOP buffer, IMDM, culture media) to 37C before using. TrypLE Express and DPBS are kept at room temp.
  1. To prepare for picking, set out one sterile Eppendorf tube per colony to be picked, and add 50uL TrypLE Express to each tube.
  2. For picking, use a P20 pipette set to 10uL.
    • Carefully tilt the culture plate so that the colony is fully submerged in media (this makes it easier to see at an angle and is easier to capture).
    • Depress the plunger on the pipette, and in one smooth motion, slowly release the plunger while scooping/scraping at the base of the colony.
    • The colony should easily be pulled into the pipette tip and will be visible floating in the media.
  3. Transfer the picked colony to one Eppendorf tube and pipette up and down a few times to begin disaggregation. Close the lid and label the tube.
  4. Repeat steps 2-3 for all colonies.
  5. Once all colonies have been picked, allow to disaggregate for 10min at room temp.
  6. Vortex each tube briefly, then add 50uL STOP buffer to each tube and pipette up and down several times to complete disaggregation
  7. Add 200uL IMDM to each tube and spin down in the tabletop centrifuge at 6000 rpm for 5 min.
  8. Depending on the size of the colony, a small pellet should be visible. Return tubes to the TC hood and carefully aspirate the supernatant (using a P200 tip over the aspirator).
  9. Wash each colony pellet with 200uL IMDM and spin down at 8000 rpm for 5 min.
  10. In TC hood, aspirate tubes and resuspend each pellet in 750uL ESCM/2i culture media.
  11. Aspirate PMEF media from the 24-well feeders and plate 500uL of disaggregated colony cells per well (each well will be one colony picked).
  12. Check plate briefly under the microscope and place in the incubator at 37C.
  13. Take Eppendorf tubes with the remaining 250uL of colony cells to isolate DNA for genotyping. Spin down in tabletop centrifuge at 8000 rpm for 5 min.
  14. After aspirating (can continue on benchtop), begin DNA extraction by adding lysis buffer.
    • Use 50uL Viagen + 1uL Proteinase K per colony and pipette up and down to fully resuspend the pellet.
  15. Float tubes in the 56C water bath for 20 min, then 95C on the dry block for 10 min.
  16. Allow tubes to cool briefly then quick spin to collect condensation drops to the bottom of the tube. Colony DNA tubes can be stored at 4C.
Colony Genotyping
  • Perform genotyping using green HotStart protocol and “regular” PCR program. Use 2uL of DNA as template for each PCR.
    • Fusion transgene (primers spanning the fusion junction)
    • Vav-Cre
    • Delta ME retained (successfully recombined FRT site) & delta ME removed (FRT site still intact)
    • Any cooperating mutations present in the parent line (Flt3 ITD/wt or NrasG12D mut)
    • RIK wt/mut
  • For controls, include DNA from the parent iPSC line (non-nucleofected cells) and wt B6 tail, along with any positive controls required for each reaction and H2O negative control.
  • Run on a 1.5% agarose gel at 150V for about 30 min and image.
  • Correctly integrated colonies will give a bright band for del ME retained, matching the ME/ME control. The parent line and FRT/FRT controls will give a bright band for del ME removed, indicating the FRT site is still intact and has not been recombined. Wt B6 should not produce a bright band in either reaction.
    • Look for colonies that have Vav present as well as the transgene and cooperating mutations.
  • Any colonies without the insert (or lacking cooperating mutation) can be immediately marked to toss. Keep all correct genotype colonies in culture; they should begin attaching to feeders and growing as normal by the next day.
  • Monitor growth for spontaneous differentiation or changes in morphology. Normal colonies can be frozen or expanded for further analyses.