Protocols

Cell Thawing & Media Preparation

Biosafety Cabinet Preparation

1. Turn on the biosafety cabinet.
2. Allow it to run for at least 15 minutes before beginning work to purge the air and ensure sterile airflow.
3. Disinfect the surface. Wipe down all internal surfaces (sash, sidewalls, and work area) with 70% ethanol, using sterile wipes.
4. Take and spray all items with 70% ethanol before placing them in the hood:
   • 15 mL, 50 mL Falcon tubes
   • Sterile pipettes (1 mL, 5 mL, 10 mL) and pipette controller
   • Petri dishes (6 cm/10 cm)
   • Sterile waste container (for pipettes and tips)
   • 70% ethanol spray bottle

Media Preparation

1. Turn on the water bath and set the temperature to 37 °C.
2. While the water bath equilibrates, get your bottle of complete DMEM from the refrigerator.
3. Prepare the fresh DMEM medium in 50 mL Falcon tube according to this composition:
   • DMEM - 45 mL
   • FBS 10% - 5 mL
   • Penicillin-Streptomycin (100×) 1% - 500 μL
4. Place the bottle in the 37 °C water bath until warm (~10 min).

Thawing Cells

1. Wear gloves and loosen the vial cap slightly to avoid pressure buildup.
2. Re-tighten the cap and immerse the vial in the 37 °C water bath, swirling gently.
3. Thaw rapidly, until only a tiny ice crystal remains (~1 minute).
4. Immediately remove the vial, dry it with tissue, and spray with 70% ethanol.
5. Transfer the vial into the biosafety cabinet.
6. Inside the hood, open the vial carefully.
7. Transfer its contents (~1 mL) into a 15 mL conical tube.
8. Slowly add 9 mL of warm complete DMEM dropwise while gently swirling to avoid osmotic shock.
9. Mix gently by flicking the tube or pipetting slowly.
10. Centrifuge at 200 × g for 5 minutes at room temperature.
11. Carefully aspirate the supernatant.
12. Resuspend the cell pellet in:
    • 10 mL complete DMEM for one 10 cm dish, or
    • 4-5 mL if using two 6 cm dishes.

Plating

1. Add the resuspended cells to the Petri dishes.
2. Gently swirl the dish front-to-back and side-to-side to distribute cells evenly.
3. Label each dish with:
   • Cell line name "HEK293T"
   • Date
   • Passage number
   • Your initials
4. Place the dishes in a 37 °C, 5% CO₂ incubator.
5. Allow cells to attach overnight.

Post-Thaw Care

1. After attachment, replace the medium with fresh, warm complete DMEM to remove residual DMSO and dead cells.
2. Feed cells every 2-3 days or when the medium starts to turn yellow.
3. Passage when 70-90% confluent using TrypLE.
4. Always record passage number and date.

Cell Culturing

Prepare the Biosafety Cabinet

1. Turn on the hood 15 minutes before starting.
2. Wipe all inner surfaces with 70% ethanol.
3. Spray and place all sterile materials inside:
   • Warmed DMEM complete medium (RT)
   • TrypLE Express (RT)
   • PBS (Ca²⁺/Mg²⁺-free) for washing (RT)
   • New tissue-culture dishes or flasks
   • Sterile pipettes (5 mL, 10 mL, 25 mL)
   • Waste container and ethanol spray bottle

Cell Assessment

1. Take out the cells from the incubator and observe your cells under the microscope.
2. Cells should cover ~80–90% of the dish and look healthy.
3. Place the dish in the hood.

Washing

1. Aspirate the spent medium using a sterile aspirator or vacuum pipette.
2. Add enough PBS to cover the cell layer (2-3 mL for a 6 cm dish, 5-10 mL for a 10 cm dish).
3. Aspirate the PBS completely.

Trypsinization

1. Add just enough TrypLE to cover the cell layer:
   • 6 cm dish: 1 mL
   • 10 cm dish: 2-3 mL
2. Tilt the plate to ensure the entire surface is coated.
3. Incubate at 37 °C for 1-3 minutes. Do not over-trypsinize - HEK293 detaches quickly.
4. When cells start rounding up and detaching, tap the side of the dish gently to release them.

Neutralization and Collection

1. Once cells are detached, add warm complete DMEM to stop trypsin:
   • Add 4-5 mL for a 6 cm dish, or 8-10 mL for a 10 cm dish.
2. Pipette up and down gently to break up clumps and make a single-cell suspension.
3. Transfer the suspension into a 15 mL conical tube.
4. (Optional) Take 10 µL to check viability or cell count with trypan blue.

Preparation for Reseeding

1. Prepare new labeled dishes with pre-warmed medium:
   • 6 cm dish: 2-3 mL
   • 10 cm dish: 8-10 mL
2. Centrifuge at 300 × g for 5 minutes at RT. Do not forget to balance!
3. Remove the supernatant and add fresh pre-warmed medium (1 mL) to the cell pellet.
4. Resuspend the pellet carefully and transfer to the new dishes.

Seeding

1. Depending on growth rate, split at ratios of:
   • 1:3 (if very confluent)
   • 1:5 (for routine maintenance)
   • 1:10 (if you want slower growth or preparing frozen stocks soon)
2. Gently rock the dish to distribute cells evenly.
3. Place the new dishes in the 37 °C, 5% CO₂ incubator.
4. Move gently front-to-back and side-to-side to ensure even seeding.
5. Check after 24 h for attachment and healthy morphology.

Western Blot

Harvesting HEK293T Cells

1. Prepare the required materials:
   • Labeled micro-centrifuge tubes, rack, ice bucket filled with ice, pipettes, PBS, Trypsin-EDTA, 4× Laemmli Sample Buffer, DMEM complete growth media.
2. Remove the conditioned growth media (DMEM+/+) by aspirating.
3. Gently wash the cell monolayer with 1 mL of cold PBS. Carefully add PBS to the side of the flask/dish so as not to forcefully dislodge adherent cells. Keep PBS in the ice!
4. Remove the PBS by suction.
5. Add 200 μL of pre-warmed Trypsin-EDTA to the flask and place in an incubator for 1-2 minutes. Check flask frequently to ensure all cells have dissociated from the flask surface.
6. When all cells are detached, neutralize/deactivate the dissociation reagent with 1 mL DMEM+/+.
7. Transfer cell suspension to the corresponding cold microcentrifuge tube on ice. Ensure that all cells have been harvested from the flask.
8. Centrifuge the cell suspension for 1 min at 12,000 RPM at 4°C. Remove the supernatant.
9. Add 1 mL of PBS to the pellet.
10. Centrifuge the cell suspension for 1 min at 12,000 RPM at 4°C. Remove the supernatant.
11. Centrifuge the cell suspension for 1 min at 12,000 RPM at 4°C. Remove the residual PBS***.
12. Add 40 μL of PBS and suspend the cells in the ice.
13. Add 40 μL of 4× Laemmli Sample Buffer. Mix the suspension by vortexing.
14. Insert the protective cap into microcentrifuge tubes, then boil for 13 minutes.
15. Cool down the suspension.
16. Centrifuge the cell suspension for 1 min at 7,000 RPM at room temperature.
17. Mix the suspension.
18. Centrifuge the cell suspension for 1 min at 7,000 RPM at room temperature.
19. Proceed with the SDS-PAGE.

***Note: It is possible to freeze the cells at this stage. You have to melt the cells in the ice for 10 minutes prior to preparing the sample.

SDS-PAGE

1. Take the gel cassette, supporting rubber and holder.
2. Prepare the 12% resolving gel:
   • Add distilled water.
   • Add 30% acrylamide.
   • Add 1.5M Tris (pH 8.8).
   • Add 10% SDS.
   • Add 10% APS.
   • Add TEMED.
3. For preparing 5% stacking gel:
   • Add distilled water.
   • Add 30% acrylamide.
   • Add 1.0M Tris (pH 6.8).
   • Add 10% SDS.
   • Add 10% APS.
   • Add TEMED.
4. Wait for 15 minutes to allow the resolving gel to harden.
5. Pour 1 mL isopropanol to even out the resolving gel surface.
6. Remove isopropanol, then wash with distilled water.
7. Pour the 5% stacking gel on top of the resolving gel.
8. Introduce clean combs.
9. Wait for the stacking gel to harden, then remove the combs.
10. Wash the gel cassette carefully, again place them inside the gel box.
11. Add a new running buffer inside and a used running buffer outside of the cassette.
12. Prepare 8 μL protein marker:
    • 1.5 μL protein marker + 6.5 μL 2× sample buffer.
13. Carefully load the samples (buffer, S1, S2, …, 8 μL protein marker, buffer).
14. Close the lid.
15. Connect the cables to the power supply. Make sure to match the colors on the cables to those on the power supply inputs.
16. Program the power supply and start the run:
    • 200 Volt - 40 minutes, or
    • 120 Volt - 60+ minutes, or
    • 50 Volt - 180 minutes
17. Check whether the bubbles are appearing or not.

Resolving Gel Recipes (1.5M Tris, pH 8.8)

Stacking Gel Recipe (1.0M Tris, pH 6.8)

Transfer

1. Prepare all the necessary instruments: sponges, knife, nitrocellulose membrane, 3M paper, ice box, ice bucket, transfer buffer, power supply, transfer tank.
2. Gently remove the gel from the cassette.
3. Label the nitrocellulose membranes according to the given samples.
4. Place the transfer tank inside the ice bucket.
5. Put ice inside the transfer tank and around it.
6. Pour the transfer buffer into the washing box.
7. Soak the 3M papers, sponge & nitrocellulose membrane before the assembly of the sandwich.
8. Prepare the sandwich:
   • Sponge
   • 3M paper
   • Membrane (anode (+)) (RED)
   • Gel (cathode (-)) (BLACK)
   • 3M paper
   • Sponge
9. Gently remove the bubbles between the gel and the membrane by using your hand. Otherwise, they will inhibit the transfer process.
10. Place the sandwich into the transfer cassette. Remember that the gel side of the cassette faces the cathode (-) while the membrane side faces the anode (+).
11. Close the square lid.
12. Connect the cables to the power supply. Transfer time is fixed to 90 minutes.
13. Start with 180 mA (-90 Volts).
14. Place the brick on the lid to press it.

Immunodetection

1. Block the membrane with skim milk in TBST for 30 minutes at room temperature on a shaker.
2. Add primary antibody and incubate overnight on a shaker.
3. Wash the membrane with TBST for 10 minutes. Do this 3 times! (All washing and antibody incubation steps should be done on a shaker at room temperature to ensure even agitation)
4. Dilute the secondary antibody:
   • Add 6 mL of skim milk per blot.
   • Add secondary antibody 1/2000 (1/5000 for housekeeping genes).
5. Add a secondary antibody and incubate for 1 hour on a shaker.
6. Discard the secondary antibody.
7. Wash the membrane with TBST for 10 minutes. Do this 3 times! (All washing and antibody incubation steps should be done on a shaker at room temperature to ensure even agitation)
8. Add 1 mL of ECL and 3 μL of H₂O₂.
9. Mix well and spread over the membrane surface.
10. Visualize the result using the Fusion Solo software.

MTT Assay

Perform the experiment according to the manufacturer's protocol (CyQUANT™ MTT Cell Proliferation Assay Kit). On the day of the experiment, prepare the following reagents.

Reagent Preparation

1. Prepare a 12-mM MTT stock solution by adding 1 mL of sterile PBS to one 5-mg vial of MTT (Component A). Vortex to mix or sonicate the solution until it is dissolved. If some particulate material does not dissolve, remove by filtration or centrifugation.
   Note: MTT stock solution can be stored at 4°C for up to 4 weeks protected from light.
2. Prepare the SDS-HCl solution by adding 10 mL of 0.01 M HCl to one tube containing 1 g of SDS (Component B). Mix the solution gently by inversion or sonication until the SDS dissolves. Once prepared, use the SDS-HCl solution promptly.

Assay Procedure

1. Replace the culture medium according to your cell type. For adherent cells, remove the medium, then add 100 μL of fresh culture medium.
2. Add 10 μL of the 12-mM MTT stock solution to each well. Include a negative control by adding 10 μL of the MTT stock solution to 100 μL of medium alone.
3. Incubate at 37°C for 4 hours or overnight. For cell densities >100,000 cells per well, the incubation time can be shortened to 2 hours.
4. Add 100 μL of the SDS-HCl solution to each well, then pipet up and down thoroughly to mix.
5. Incubate the microplate at 37°C for 4-18 hours in a humidified chamber. Longer incubations will decrease the sensitivity of the assay (Niks, 1990).
6. Pipet up and down to mix each sample again, then read the absorbance at 570 nm.