Comprehensive Culture Protocols for Human iPSCs:
Feeder-Dependent, Essential-8,
Conditioned Medium, & KSR Systems

Maintaining human induced pluripotent stem cell (hiPSC) cultures requires reliable, well-defined systems suited to diverse research needs. This comprehensive guide brings together key protocols for culturing hiPSCs across multiple conditions—including feeder-dependent systems on mouse embryonic fibroblasts (MEFs), feeder-free culture in Essential 8 Medium, and KnockOut Serum Replacement (KSR)-based media. Each protocol provides detailed guidance for medium preparation, plating, passaging, and long-term maintenance to help ensure consistent pluripotency, healthy morphology, and reproducible experimental outcomes.

hESCs and iPSCs are biologically very similar, but there are subtle yet important differences in how they behave and respond when cultured on feeder layers such as mitotically inactivated MEFs. The table below describes the comparison of growing these cells in culture.

1. Cover the whole surface of each new culture vessel with Attachment Factor (AF) solution and incubate the vessels for 30 minutes at 37°C or for 1 hour at room temperature.
2. Using sterile technique in a laminar flow culture hood, completely remove the AF solution from the culture vessel by aspiration just prior to use. Coated vessels may be used immediately or stored at room temperature for up to 24 hours.

3. One to two days before initiating or passaging hESC culture, plate 30,000/cm² of mitotically inactivated MEFs on an Attachment Factor-coated culture vessel in MEF medium (Table 1).
4. Place MEF dishes into a 37°C, 5% CO₂ incubator.

1. Aspirate the MEF medium from a dish containing inactivated MEFs and add pre-warmed PSC Culture Medium to the dish, 3–4 hours before plating iPSCs.
2. Label the dish containing inactivated MEF cells with the passage number from the vial, the date, and user initials.
3. Remove the vial of hESCs from liquid nitrogen storage using metal forceps.

4. Roll the vial between your gloved hands until the outside is free of frost. This should take ~10–15 seconds.
5. Immerse the vial in a 37°C water bath without submerging the cap. Swirl the vial gently.
6. When only an ice crystal remains, remove the vial from the water bath.
7. Spray the outside of the vial with 70% ethanol and place it in hood.
8. Pipet cells gently into a sterile 50-mL conical tube using a 5-mL sterile pipette.
9. Slowly add 10 mL of PSC Culture Medium drop-wise to cells in the 50-mL conical tube. While adding the medium, gently move the tube back and forth to mix the hESCs. This reduces osmotic shock to the cells.
10. Rinse the vial with 1 mL of PSC Culture Medium and add to the 50 mL conical with cells.
11. Transfer cell suspension to a 15 mL conical tube and centrifuge the cells at 200 × g for 5 minutes.
12. Aspirate and discard the supernatant.
13. Resuspend the cell pellet in sufficient volume of PSC Culture Medium according to Table 2 by gently pipetting the cells up and down in the tube a few times.
14. Aspirate the spent PSC Culture Medium from the MEF dish and slowly add the thawed colonies onto the dish. Place dish gently into the 37°C, 5% CO₂ incubator and move the dish in several quick, short, back-and-forth and side-to-side motions to disperse cells across the surface of the dishes.
15. Incubate the cells overnight.
16. The next day, remove the spent medium with debris using a sterile serological pipette and transfer it into a prepared MEF dish. You can use this dish as a backup in case there is a problem with the main dish.
17. Add fresh PSC Culture Medium to each dish according to the volumes in Table 2. Place both plates gently into a 37°C, 5% CO₂ incubator overnight.
18. Examine cells under the microscope and replace spent medium daily from both plates. If feeding more than one plate, use a different pipette for each well to reduce risk of contamination. Colonies may not be visible for up to a week.

In general, split cells when one of the following occurs:

• The MEF feeder layer is 2 weeks old.
• iPSC colonies are becoming too dense or too large.
• Increased differentiation occurs.

• The split ratio can vary, though it is generally between 1:2 and 1:4. Occasionally, cells will grow at a different rate and the split ratio will need to be adjusted. A general rule is to observe the last split ratio and adjust the ratio according to the appearance of the iPSC colonies.
• If the cells look healthy and colonies have enough space, split using the same ratio. If they are overly dense and crowding, increase the ratio. If the cells are sparse, decrease the ratio. Cells will need to be split every 4–10 days based upon appearance.
• iPSCs do well in iMEF plates that have been conditioned with PSC Culture Medium. It is common practice to condition new feeder plates before passaging iPSCs into them. 

You may passage cells via the enzymatic method as described below, or mechanically as described in the following section.

1. Aspirate the MEF medium from a dish containing inactivated MEFs and add pre-warmed PSC Culture Medium to the dish, 3–4 hours before plating iPSCs.
2. Label the new MEF dish with the cell line name, the new passage number, the date, the split ratio, and user initials. Return the plate to the incubator.
3. Under a dissecting microscope, remove differentiated colonies from the dish to be passaged.
4. Aspirate the spent medium from the dish with a Pasteur pipette.
5. Add Collagenase Type IV (1 mg/mL) solution to the dish containing iPSCs. Adjust the volume of Collagenase Type IV for various dish sizes (e.g., 35-mm dishes require 1 mL of Collagenase IV).
6. Incubate the dish(es) for 30–60 minutes in a 37°C, 5% CO₂ incubator. Note that the incubation times may vary among different batches of collagenase; therefore, examination of the colonies is needed to determine the appropriate incubation time.

7. Stop the incubation when the edges of the colonies are starting to pull away from the plate (Figure 4).

8. Aspirate the Collagenase Type IV Solution with a Pasteur pipette. Remove the collagenase carefully without disturbing the attached cell layer.
9. Add PSC Culture Medium to each dish. Use a 5-mL pipette to gently blow the cells off the surface of the dish while pipetting up and down. Make sure to pipet gently to minimize the formation of bubbles.
10. After the iPSCs have been removed from the surface of the well, pool the contents of the wells into a 15-mL conical tube.
11. Using a 5-mL pipette, add PSC Culture Medium to the dish to wash and collect any residual cells. Pipet up the medium and cells and then add the collected cells to the 15-mL tube.
12. Pipet cells up and down gently a few times in the 15-mL tube to further break up cell colonies. Pipet carefully to reduce foaming.

13. Centrifuge at 200 × g for 5 minutes and then aspirate the supernatant from the iPSC pellet.
14. Resuspend the pellet with an appropriate amount of PSC Culture Medium (Table 2). This is dependent on the split ratio and the number of dishes used.
15. Mix the cell suspension well with a 10-mL pipette. Be careful not to break up the colonies too much or cause bubbles in the media.
16. Add appropriate volume of cell suspension to each dish. Return the dish to the incubator.
17. Move the dish(es) in several quick, short, back-and-forth and side-to-side motions to disperse cells across the surface of the dishes.
18. Incubate cells overnight to allow colonies to attach. Replace spent medium daily.

1. Replace the medium in the dish containing the cells with fresh PSC Culture Medium.
2. Under a laminar flow hood, open the package containing the EZPassage tool and remove the tool.
3. Hold the culture vessel in one hand and pull (roll) the EZPassage tool across the entire dish in one direction. Apply gentle but firm pressure so that the entire roller blade touches the dish and maintains uniform pressure during the rolling action.
4. Keep rolling the EZPassage tool parallel to the first pass until the entire dish has been covered.
5. Rotate the culture dish 90° and then repeat rolling the cell layer as described above.
6. When you are finished, discard the EZPassage tool and do not reuse. Use a cell scraper to lift cell clusters off the plate, if necessary.
7. Using a serological pipette, rinse the dish with medium so that the cut colonies are suspended in the medium.
8. Transfer the medium containing the colonies to a 15-mL sterile tube.
9. Seed the cell colonies on dishes plated with mitotically inactivated MEFs at an appropriate density.
10. Place the plates into a 37°C, 5% CO₂ incubator. Shake the plates gently to evenly disperse cells.

Preparing MEF medium (for 100 mL of complete medium)

1. To prepare 100 mL of complete MEF medium, aseptically mix the following components:

Preparing PSC culture medium (for 100 mL complete medium)

1. To prepare 100 mL of complete PSC culture medium, aseptically mix the following components:

      2.   Complete MEF medium can be stored at 2–8°C for up to 4 weeks.

Preparing MEF-conditioned medium (MEF-CM)

1. Cover the whole surface of each new culture vessel with Attachment Factor (AF) solution and incubate the vessels for 30 minutes at 37°C or for 1 hour at room temperature. For MEF-CM generation, a T-175 flask is recommended.
2. Using sterile technique in a laminar flow culture hood, completely remove the AF solution from the culture vessel by aspiration just prior to use. Coated vessels may be used immediately or stored at room temperature for up to 24 hours.

3. Plate 9.4 × 10⁶ Mitomycin C-treated or irradiated MEFs in a T-175 flask coated with AF and containing 30 mL of MEF medium.
4. The following day, replace the MEF medium with 90 mL of PSC Culture Medium.
5. Collect the PSC Culture Medium, now considered MEF-CM, from the flasks after 24 hours of conditioning for up to seven days in a row.
6. Each day, filter sterilize the collected MEF-CM with a 0.22 μM filter. Filtered MEF-CM can be stored at –20°C until use.
7. At the time of use, thaw the MEF-CM in a 37°C waterbath, and freshly supplement it with additional bFGF (20 ng/mL).

Coating culture vessels with Geltrex matrix

1. Thaw a 5-mL bottle of Geltrex matrix at 2–8°C overnight.
2. Dilute the thawed Geltrex solution 1:1 with cold sterile D-MEM/F-12 to prepare 1-mL aliquots in tubes
   chilled on ice. These aliquots can be frozen at –20°C or used immediately.

3. To create working stocks, dilute a Geltrex aliquot 1:50 with cold D-MEM on ice, for a total dilution of 1:100.

4. Quickly cover the whole surface of each culture dish with the Geltrex solution (refer to Table 3).
5. Incubate the dishes in a 37°C, 5% CO₂ incubator for 1 hour.

6. Aspirate the diluted Geltrex solution from the culture dish and discard. You do not need to rinse off the
   Geltrex solution from the culture dish after removal. Cells can now be passaged directly into MEF-CM onto the Geltrex matrix-coated culture dish.

Table 3. Volume of Geltrex hESC-qualified matrix required.

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Thawing and plating iPSCs

1. Label the Geltrex matrix-coated dish with the passage number from the vial, the date, and user initials.
2. Remove the vial of iPSCs from liquid nitrogen storage using metal forceps.

3. Roll the vial between your gloved hands until the outside is free of frost. This should take ~10–15 seconds.
4. Immerse the vial in a 37°C water bath without submerging the cap. Swirl the vial gently.
5. When only an ice crystal remains, remove the vial from the water bath, spray the outside of the vial with 70% ethanol to sterilize, and place it in hood.
6. Pipet the thawed cells gently into a sterile 50-mL conical tube using a 5-mL sterile pipette.
7. Slowly add 10 mL of MEF-CM drop-wise to cells in the 50-mL conical tube. While adding the medium, gently move the tube back and forth to mix the iPSCs. This reduces osmotic shock to the cells.
8. Rinse the vial with 1 mL of MEF-CM and add to the 50-mL conical tube with cells.
9. Transfer cell suspension into a 15-mL centrifuge tube. Centrifuge the cells at 200 × g for 5 minutes.
10. Aspirate and discard the supernatant.
11. Resuspend the cell pellet in sufficient volume of MEF-CM according to Table 4 by gently pipetting the cells up and down in the tube a few times.
12. Aspirate the excess Geltrex solution from the prepared dish and slowly add the thawed colonies onto the dish. Move the dish in several quick, short, back-and-forth and side-to-side motions to disperse cells across the surface the dish.
13. Place dish gently into the 37°C, 5% CO₂ incubator and incubate the cells overnight.
14. The next day, remove the spent medium with debris using a sterile serological pipet and transfer it into a prepared Geltrex matrix-coated dish. You can use this dish as a backup in case there is a problem with the main dish.
15. Add fresh MEF-CM to each dish according to the volumes in Table 4. Place both plates gently into a 37°C, 5% CO₂ incubator overnight.
16. Examine cells under the microscope and replace spent medium daily from both plates. If feeding more than one plate, use a different pipette for each well to reduce the risk of contamination. Colonies may not be visible for up to a week.

Table 4. Volume of MEF-CM required.

Figure 6. PSCs growing in MEF-CM on a Geltrex matrix-coated dish.

Preparing Essential 8 medium (500 mL of complete medium)

1. Thaw Essential 8 Supplement (50X) at room temperature for ~1 hr. Do not thaw frozen supplement at 37°C.
2. To prepare 500 mL of complete Essential 8 Medium, aseptically mix the following components:
   

   Component	Volume
   Essential 8 Basal Medium	490 mL
   Essential 8 Supplement (50X)	10 mL

3. Complete Essential 8 Medium can be stored at 2–8°C for up to 2 weeks.

Preparing 0.5 mM EDTA in DPBS (50 mL)

1. To prepare 50 mL of 0.5 mM EDTA in DPBS, aseptically mix the following components in a 50-mL conical tube in a biological safety hood:

   Component	Volume
   DPBS without calcium and magnesium	50 mL
   0.5 M EDTA	50 μL

2. Filter sterilize the solution. The solution can be stored at room temperature for up to six months.  

Coating culture vessels with vitronectin (VTN-N)

1. Upon receipt, thaw the vial of vitronectin at room temperature and prepare 60-μL aliquots of vitronectin in polypropylene tubes. Freeze the aliquots at –80°C or use immediately.
2. Prior to coating culture vessels, calculate the working concentration of vitronectin using the formula below and dilute the stock appropriately. Refer to Table 5 for culture surface area and volume required.
   
   The optimal working concentration of vitronectin is cell line dependent. We recommend using a final coating concentration of 0.1–1.0 μg/cm² on the culture surface, depending on your cell line. We routinely use vitronectin at 0.5 μg/cm² for human PSC culture.  
   
   
   
   Example: To coat a 6-well plate at a coating concentration of 0.5 μg/cm², you will need to prepare 6 mL of diluted vitronectin solution (10 cm²/well surface area and 1 mL of diluted vitronectin/well; see Table 5) at the following working concentration:
   
   
3. To coat the wells of a 6-well plate, remove a 60-μL aliquot of vitronectin from –80°C storage and thaw at room temperature. You will need one 60-μL aliquot per 6-well plate.
4. Add 60 μL of thawed vitronectin into a 15-mL conical tube containing 6 mL of sterile DPBS without Calcium and Magnesium at room temperature. Gently resuspend by pipetting the vitronectin dilution up and down.

5. Aliquot 1 mL of diluted vitronectin solution to each well of a 6-well plate (refer to Table 5 for recommended volumes for other culture vessels).

6. Incubate at room temperature for 1 hour.

7. Aspirate the diluted vitronectin solution from the culture vessel and discard. It is not necessary to rinse off the culture vessel after removal of vitronectin. Cells can be passaged directly onto the vitronectin-coated culture dish.

Table 5. Volume of diluted vitronectin required.

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Preparing CELLstart-coated culture dishes

1. Dilute CELLstart substrate (1 mL) 1:50 in Dulbecco’s Phosphate Buffered Saline (D-PBS) containing calcium and magnesium. Pipette the solution gently to mix. Do not vortex.

2. Cover the whole surface of each culture dish with the CELLstart solution (1 mL for a 35-mm dish, 1.5 mL for a 60-mm dish).
3. Incubate the dishes for 1–2 hours at 37°C.
4. Transfer each dish to a laminar flow hood and allow it to equilibrate to room temperate (about 1 hour) before use.

5. Immediately before use, aspirate all CELLstart solution from the culture dishes. It is not necessary to rinse the vessels after removing CELLstart substrate.

Preparing complete KnockOut SR feeder-free medium

1. To prepare 1 mL of 10 μg/mL Basic FGF solution, aseptically mix the components listed below. Aliquot the solution and store at –20°C for up to 6 months.

2. To prepare 100 mL of complete KnockOut SR XenoFree Feeder-Free (KSR XenoFree FF) medium aseptically combine the components listed in the table below.
   
   

   Component	Stock concentration	Final concentration	Volume
   Knockout DMEM/F12	–	1X	76.8 mL
   GlutaMAX Supplement	200 mM	2 mM	1 mL
   KnockOut SR XenoFree	–	20%	20 mL
   KnockOut SR-GFC	50X	1X	2 mL
   bFGF	10 μg/mL	20 ng/mL	200 μL

   You may store the KSR XenoFree FF medium at 2–8°C for one week.
3. Just before pre-equilibrating the complete medium to temperature and gases, aseptically add the required volume of 2-mercaptoethanol (55 mM stock concentration) for a 0.1 mM final concentration. For example, to prepare 100 mL of KSR-FF medium add 182 μL of 55 mM 2-mercaptoethanol (1:550 dilution) Alternatively, the 2-mercaptoethanol may be added to the 1X completed medium and stored at 2–8°C for up to one week.

Adapting human iPSCs to KSR XenoFree FF medium

1. Culture the human iPSCs on human foreskin fibroblasts or MEF feeder cells until they are 70–80% confluent.
2. Pre-warm the required volume of TrypLE enzyme in a 37°C water bath. Refer to Table 6 below for details on the volumes required.
3. Pre-equilibrate the required volume of KSR XenoFree FF medium in a 37°C water bath for 15 min. Refer to Table 6 below for details on the volumes required.
4. Aspirate the medium from the culture dishes and add an appropriate amount of TrypLE enzyme. Incubate the dishes at 37°C for 3–5 minutes.
5. Aspirate the TrypLE enzyme from each culture vessel and wash off the MEF feeder cells gently with D-PBS (2 to 3 times).
6. Add an appropriate amount of complete KSR XenoFree FF medium to each culture vessel. Use a cell scraper or a 5 mL pipette to gently scrape the cells off the surface of culture vessel.

7. Collect the cell suspension from each culture dish into separate 15 mL conical tubes. Rinse each culture vessel with an appropriate amount of complete KSR XenoFree FF medium, and add the D-PBS rinse medium into the 15 mL conical tubes containing the cell suspension. Be cautious not to break the cell clumps into single cells
8. Centrifuge the 15 mL conical tubes at 200 × g for 5 minutes to pellet the iPSCs.
9. Aspirate the supernatant from the iPSC pellet. Resuspend the pellet in an appropriate amount of KSR XenoFree FF medium according to the split ratio (Table 6). Do not break the cell clumps to a smaller size, because the smaller clumps do not attach well to the surface.

10. Aspirate the CELLstart solution from the pre-coated culture vessel and slowly add an appropriate amount of cell suspension to each culture vessel.
11. Move the culture dish back and forth and side to side several times to disperse the cells across the surface of the dish. Gently place the culture dish in a 37°C incubator with a humidified atmosphere of 4 to 6% CO₂ in air. Replace the spent medium with KSR XenoFree FF medium every day.

Passaging human iPS cells using KSR XenoFree FF medium

1. Observe the human iPSCs growing in complete KSR XenoFree FF under the microscope to confirm that the cells are 70–80% confluent and ready to be subcultured.

2. Cut out and remove any differentiated iPSC colonies prior to passaging the culture.
3. Pre-warm the required volume of TrypLE enzyme in a 37 °C water bath. Refer to Table 6 below for details on the volumes required.
4. Pre-equilibrate the required volume of KSR XenoFree FF in a 37°C water bath for 15 min. Refer to Table 6 below for details on the volumes required.
5. Aspirate the spent medium from the culture vessel using a pipette and rinse the cells twice with D-PBS.
6. Gently add pre-warmed TrypLE enzyme to the culture vessel (e.g., 1 mL of TrypLE solution per 60-mm culture dish). Swirl the culture vessel to coat the entire cell surface.
7. Incubate the culture vessel at 37°C for 3 minutes.
8. Remove the vessel from the incubator, aspirate the TrypLE enzyme, and gently wash the cells with D-PBS.
9. Gently scrape the cells off the surface of the culture dish using a cell scraper, and transfer the cells to a sterile 15 mL centrifuge tube.
10. Rinse the culture dish twice with KSR XenoFree FF, gently “spraying off” any cells that have not detached. Pool the rinse medium with the cells in the 15 mL tube.
11. Centrifuge the tube at 200 × g for 5 minutes at room temperature to pellet the cells.
12. Carefully aspirate the supernatant without disturbing the cell pellet and discard it.
13. Gently flick the tube to fully dislodge the cell pellet from the tube bottom.
14. Gently resuspend the cells in pre-equilibrated KSR XenoFree FF using a 5mL serological pipette. Do not triturate.
15. Transfer the cells to a fresh 60-mm CELLstart-coated dish at the desired split ratio and move the culture dish back and forth and side to side several times to disperse the cells across its surface.
16. Place the culture dish in a 37°C incubator with a humidified atmosphere of 4 to 6% CO₂ in air.

The next day, gently replace the spent medium with KSR XenoFree FF medium to remove cell debris. Replace the spent medium everyday thereafter. Observe the iPSCs daily and passage them as needed (approximately every 4–5 days). Passaging is recommended when the cells reach 70–80% confluence.

Table 6. Recommended volumes for KSR.

In the PSC Culture Medium, DMEM/F12 containing GlutaMAX Supplement can be substituted with the following alternatives:

• KnockOut DMEM/F-12

To prepare 100 mL of complete PSC Culture Medium using KnockOut DMEM/F-12, aseptically combine the components listed in the table below.

• KnockOut DMEM

To prepare 100 mL of complete PSC Culture Medium using KnockOut DMEM, aseptically combine the components listed in the table below.