A Simple Multistep Protocol for Differentiating Human Induced Pluripotent Stem Cells into Functional Macrophages

Human iPSCs are delicate and more difficult to maintain in culture compared to most other conventional human cell lines. They show a natural tendency to spontaneously differentiate into fibroblast like cells. Cytokines and growth factors present in the medium allow them to be kept in pluripotent state; hence, daily change of media is essential.

iPSCs grow in individual colonies. During handling, it is critical that iPSCs remain as small clumps and not broken down into single cells. Hence, iPSC colonies should be always treated gently; do not pipette too harshly or too many times as this will result in single-cell colonies. Use wide-bore stripettes, and centrifuge slowly and with shorter duration where possible.

It is important to ensure that iPSC colonies remain separate in culture and do not fuse to each other during expansion to prevent spontaneous differentiation. However, low levels of spontaneous differentiation are sometimes unavoidable especially on feeder-dependent culture. Culture can be cleaned up by carefully selecting only pluripotent colonies during passage steps—this will eventually clean up the culture after few passages

Prepare vitronectin-coated maintenance plate at least 2 h before plating the hiPSCs. Thaw 250µL at 4Room temperature and dilute it in 6mL. Add 1mL into each well of 6-well tissue culture plates or add whole 6mL into 100 mm tissue culture dish.

Make sure that the vitronectin solution covers the whole surface of the plastic vessel and incubate at 4Room temperature for at least 2h 0m 0s or at 4°C 2h 0m 0s tightly sealed with Parafilm. If necessary, sealed plates can be stored at 4°C for up to 5 days without removing the vitronectin solution ( see Note 7 ).

Prepare a 15 mL centrifuge tube with 9mL supplemented with 1micromolar (µM) for each vial of iPSC that will be defrosted.

Thaw the vial at 37°C in water bath. Gently transfer the defrosted cells into 9mL and slowly invert the tube to evenly mix the cells and to dilute out DMSO.

Centrifuge the tube at 290x g,20°C.

Resuspend the cell pellet into desired volume of E8 medium supplemented with 10micromolar (µM).

Plate 1mL into each well of a 6-well plate. Content of one frozen vial (equivalent to 1 well of ~70–80% confluent 6-well plate or 1/10th of a 100 mm dish) can be distributed into 2–3 wells of a 6-well plate with 1mL.

Place the plate into a humidified 37°C incubator with 5%. Once inside the incubator gently rock the plate in different directions to make sure that colonies are evenly distributed throughout the surface of the dish. Keep the plate undisturbed for at least 24h 0m 0s to allow colonies to attach to the vitronectin-coated surface.

After 24h 0m 0s the majority of the colonies will attach to the surface. Remove all media from the dish to discard any unattached colonies and dead cells, and add 2mL–3mL for each well of a 6-well plate or 10mL–12mL for each 100 mm dish ( see Note 10 ).

Change medium every day and replace with fresh E8 medium until cells are ~70–80% confluent and ready for passaging.

For passaging, start by preparing vitronectin-coated plates as before.

Prepare vitronectin-coated maintenance plate at least 2 h before plating the hiPSCs. Thaw 250µL at 4Room temperature and dilute it in 6mL. Add 1mL into each well of 6-well tissue culture plates or add whole 6mL into 100 mm tissue culture dish.

Make sure that the vitronectin solution covers the whole surface of the plastic vessel and incubate at 4Room temperature for at least 2h 0m 0s or at 4°C 2h 0m 0s tightly sealed with Parafilm. If necessary, sealed plates can be stored at 4°C for up to 5 days without removing the vitronectin solution ( see Note 7 ).

Once cells are sufficiently confluent, remove the spent media and gently wash the plate twice with an appropriate volume (~5mL) of D-PBS.

Remove the spent media.

Wash the plate with an appropriate volume (~5mL) of D-PBS. (1/2)

Wash the plate with an appropriate volume (~5mL) of D-PBS. (2/2)

Remove the D-PBS and add required volume of 5millimolar (mM) to cover the entire surface of the dish (1mL for each well of a 6-well plate, and 8mL for 100 mm dish).

Look under the microscope every 2 min to confirm that colonies are detaching and have changed their morphologies but do not wait too long such that the colonies are fully detached or broken up into single cells.

Carefully remove PBS-EDTA solution without disturbing iPSC colonies and add same volume of fresh E8 media. Gently pipette up and down using a 10 mL stripette to dislodge the colonies from the surface. If necessary repeat this step two more times to recover as many iPSC colonies as possible.

Collect the media and detached iPSC colonies into a new 15 mL centrifuge tube and centrifuge at 290x g,20°C.

Resuspend the colonies in 2mL supplemented with 10micromolar (µM). If colonies are too big at this stage pipette up and down 2–3 times to break them into smaller clumps.

Transfer colonies in a desired splitting ratio into a fresh vitronectin-coated dish with appropriate volume of E8 media supplemented with 10micromolar (µM).

After24h 0m 0s remove medium and add fresh E8 medium (without Rock inhibitor); change medium every day until next passage.

To freeze iPS cells, freshly prepare the freezing mixture—10% in KSR (9mL + 1mL) at Room temperature. After lifting off iPSC colonies as described in Subheading "hiPSC Culture in Feeder-Free System", resuspend the cell pellet into desired volume of freezing mixture and quickly transfer 1mL into each cryovial. We recommend one 100 mm dish (70–80% confluent) to divide into ten cryovials and each well of a 6-well plate into two cryovials for freezing. Immediately transfer the cryovials into a Mr. Frosty or CoolCell container and place them into -80°C freezer . After 24h 0m 0s transfer the cryovials into liquid nitrogen for long-term storage.

At least 24 h before transferring the iPSCs from feeder-free culture, prepare feeder plates using inactivated mouse embryonic fibroblasts (MEFs).

First coat the tissue culture plates by adding 2mL into each well of 6-well plates or 8mL for 100 mm dishes; ensure that whole surface of the dish is covered and then incubate at 80Room temperature for at least 2h 0m 0s.

Thaw one vial (2 × 10⁶ cell) of irradiated mouse embryonic fibroblasts into 9mL in a 15 mL centrifuge tube and spin at 290x g,20°C,0h 0m 0s.

Resuspend the cell pellet in desired volume of human iPSC base medium.

Remove the gelatine solution from the plate, seed the MEFs, and place them in 37°C incubator 2h 0m 0s. One vial of MEF (2 × 10⁶ cell) is sufficient for a whole 6-well plate or one 100 mm dish.

Lift off the iPSC colonies growing under feeder-free conditions as described in Subheading "hiPSC Culture in Feeder-Free System".

Resuspend these colonies into the required volume of human iPSC base medium supplemented with 4ng/mL and 10micromolar (µM) (final concentrations).

Remove medium and any unattached MEFs from the feeder plate and transfer iPSC colonies onto the feeder layer. Usually a splitting ratio of 1:10 works well at this stage.

Place the dish inside the incubator, redistribute colonies evenly on feeder layer, and keep undisturbed for at least 24h 0m 0s.

After 24h 0m 0s a majority of colonies will have attached to the feeder layer. Remove medium and add fresh human iPS base medium supplemented with 4ng/mL but no Rock inhibitor. Change medium every day until colonies are ready to passage ( see Note 10 ).

For passaging, thaw collagenase, dispase aliquots at 37Room temperature, and mix them in 1:1 ratio in a tube.

Remove medium from iPSCs growing on feeder layer, wash once with D-PBS, and add collagenase-dispase mix on the dish to cover the whole surface.

Incubate at 37°C for 0h 30m 0s–1h 0m 0s until pluripotent colonies start to lift off but feeder layer and any differentiated cells remain attached.

Start checking under a microscope for floating colonies after 0h 20m 0s and every 0h 5m 0s–0h 10m 0s subsequently.

Gently mix the collagenase-dispase solution up and down few times using a 10 mL stripette to dislodge loosely adherent colonies from the feeder layer. Collect collagenase-dispase mixture along with all floating colonies into a new 50 mL and add at least same volume of human iPSC basal medium to neutralize enzyme activity.

Centrifuge harvested colonies at 290x g,20°C and discard the supernatant.

Wash at least two times by resuspending colonies in iPSC base medium followed by centrifugation.

Resuspend colonies in iPSC base medium followed by centrifugation. (1/2)

Resuspend colonies in iPSC base medium followed by centrifugation. (2/2)

After the last wash, if required, break the colonies into smaller sizes, gently resuspend in desired volume of iPSC base medium supplemented with bFGF and Rock inhibitor, transfer to new feeder plates in a desired splitting ratio, and continue as before.

Once iPSC colonies are 70–80% confluent on feeder-dependent culture detach them from the feeder layer using collagenase-dispase method. Transfer all the floating iPSC colonies with enzyme mix into a fresh 50 mL tube and add at least twice the volume of iPSC base medium to neutralize enzyme function.

Keep the tube upright in a tube rack undisturbed for 0h 3m 0s–0h 5m 0s to allow colonies to settle.

Aspirate medium from the top without disturbing iPSC colonies in the bottom of the tube.

Wash 2–3 times by adding excess volumes of iPSC base media and by allowing the colonies to settle.

Wash by adding excess volumes of iPSC base media. (1/3)

Wash by adding excess volumes of iPSC base media. (2/3)

Wash by adding excess volumes of iPSC base media. (3/3)

Resuspend iPSC colonies in iPSC base medium without any cytokine supplement, and distribute them to desired numbers of 100 mm low-adherent bacteriological dishes.

Add extra medium as necessary to make the final volume of each 100 mm low-adherent bacteriological dish ~30mL.

Transfer the dishes to a humidified 37°C incubator and leave undisturbed for 96h 0m 0s (4 days) to allow EB formation.

On day 5, the EBs should have formed and be visible with the naked eye. Harvest them by collecting all the media into 50 mL tubes using a 10 mL stripette and leaving the tubes in an upright position for 0h 3m 0s–0h 5m 0s to let the EBs settle by gravity.

Carefully aspirate the medium along with single cell and cell debris and gently resuspend EBs into desired volume of myeloid precursor base medium supplemented with IL-3 and M-CSF.

Distribute the EBs onto gelatine-coated 100 mm tissue culture-treated dishes with final volume of ~12mL in each dish.

After 4–5 days, the majority of EBs will attach to the gelatine-coated surface and start to spread out as a stromal layer around the EB. It is possible that some EBs are still not attached to the surface at this stage; in such a case add additional 12mL–15mL and leave the EBs in culture for another 4–5 days ( see Note 12 ).

Change the medium every 4–5 days for next 2–3 weeks by harvesting all media from the EB dish and pass through a 70 μm cell strainer. Discard the flow through as this will mostly contain contaminating small cells and debris. Any floating EBs will be retained by the cell strainer and should be transferred back into the culture dish; invert the cell strainer and directly pipette the medium onto the bottom surface of the strainer back into the same dish.

Approximately 3–4 weeks after transferring the EBs to gelatinized plates, the smaller apoptotic looking cells will disappear from the culture to be replaced by larger blast-like cells with dendrite-like structure. These are the myeloid precursor cells that could be further differentiated into mature macrophages

as described below.

From onwards this is a continuous culture; myeloid precursors can be harvested every 4–5 days and as long as cytokine-supplemented fresh medium are added to EBs they will produce further precursors. This process can be continued for 6–8 months after which precursor number drops significantly ( see Notes 13–15 ).

Harvest the myeloid precursor cells by removing spent medium from EB plates.

Filter through a 70 μm cell strainer and collect the flow through in 50 mL tubes as it will contain the precursor cells.

Centrifuge at 290x g,20°C and resuspend the pellet in macrophage differentiation base medium supplemented with 100ng/mL.

Count cells and plate ~150,000 precursor cells in each well of a 6-well plates or 500,000 cells in a 100 mm tissue culture dish ( see Note 16 ).

Culture cells at 37°C incubator for another 144h 0m 0s (6 days) to differentiate into mature macrophages.

From day 7 onwards fully mature macrophages will be ready for phenotypic assessment or downstream functional assays.