Antibody Generation: Producing Monoclonal Antibodies Using Hybridomas

Note: Sterile cell culture technique should be maintained when handling the hybridoma cells and the media in a sterile manner (e.g., in a biosafety cabinet) until the antibody purification steps.

1. Thawing frozen hybridoma cells

1. Incubate the vial containing the frozen hybridoma cells in a 37°C water bath until just thawed (approximately 2 minutes).
2. Add the thawed cells to a 15 mL conical tube containing 10 mL of complete RPMI (RPMI supplemented with 10% fetal bovine serum, 100 U/mL penicillin, 100 µg/mL streptomycin, 1mM sodium pyruvate, 1x non-essential amino acids, 50 µM 2-Mercaptoethanol, 10 mM HEPES).
3. Centrifuge for 5 min at 1200 RPM to wash out any contaminating freeze media.
4. After centrifugation, discard the liquid supernatant, and resuspend the cell pellet in 5 mL fresh complete RPMI. Then, add the cells to a T75 tissue culture flask containing 15 mL complete RPMI (final volume of RPMI is 20 mL).
5. Grow the cells in a standard incubator at 37°C with 5% CO₂. Cells are non-adherent while in culture.

2. Hybridoma expansion

1. Allow the cells to expand for approximately 3 days until the flask is ~80% confluent.
   Note: This amount of time can vary based on the starting number of cells, as well as inherent differences in growth rates of different cells. It is important for the cells to be in the exponential growth phase.
2. Once the initial flask reaches ~80% confluency, remove the cells from this initial T75 flask, place into a conical centrifuge tube and spin (1200 RPM for 5 min) to pellet the cells.
3. Resuspend the cells in 6 mL of complete RPMI, and then distribute the cell suspension into three new T75 flasks (2 mL/flask containing 18 mL RPMI). Incubate these three T75 flasks at 37°C with 5% CO₂ until they are ~80% confluent (This should take approximately 3 days).

3. Antibody Production in Serum-free Medium

Note: At this point, the cells are ready to continue their growth in a serum-free medium designed for the growth of hybridoma cell lines. In this protocol, we use the ready-to-use, commercially available HB Basal Liquid medium containing the HB101 Supplement Product.

1. The cells from each of the three T75 flasks (at ~80% confluency) are collected and centrifuged (1200 RPM for 5 min).
2. The cell pellet from each T75 flask is resuspended in 10 mL supplemented HB101 serum-free medium and then added to two T225 flasks supplemented HB101 serum-free medium (i.e. 5 mL cell suspension into each of 6 flasks containing ~220-240 mL HB101 in each flask).
3. Continue growing the cells in T225 flasks and HB101 media at 37°C with 5% CO₂ for three weeks, or until cells start to die. The cells are producing mAb during these 3 weeks and secreting it into the culture medium. Once the cells are starting to die, they are no longer producing any mAb.

4. Antibody Purification - Day 1

Note: At this point, sterility does not need to be maintained so handling the media in a sterile manner (e.g., in a biosafety cabinet) is not required. Furthermore, hybridomas are not considered a "BSL2 level" agent.

1. Pour media from the flasks into tubes for a fixed angle rotor. Spin the tubes in a centrifuge with a fixed angle rotor at 10,000 RPM for 8 minutes. This step is designed to remove the cellular debris from the culture supernatant.
   Note: Tube sizes can vary depending on the rotor used. The important thing to remember is to have the same volume in the tubes to ensure the rotor is properly balanced prior to centrifugation. It is likely the entire volume of the media will not fit in the centrifuge at one time. The remaining media will be centrifuged later (step 4.5) using the same tubes.
2. Prepare a 2L plastic beaker with a stir bar in a bucket surrounded by ice. Place on a stir plate.
3. Attach a 500 mL filter top on a 1L bottle. Attach bottle top filter unit to house vacuum via appropriate tubing.
4. Pour the supernatant from step 4.1 (which still contains the mAb produced by the hybridoma cells) into the filter top.
5. Continue using the same set of tubes to pellet cell debris from the media (the pellet will continue to build). Wait to run the vacuum until the filter top is full and another batch of supernatant is ready to pour in. Do not let the filter dry out or filtering will become very slow.
6. When the 1L bottle is close to full, remove the filter top and pour supernatant into 2L beaker prepared in step 4.2. Reattach the filter top. Keep track of the volume.
7. Repeat the centrifugation and filtration steps until all the media is processed.
8. Measure out 295g of ammonium sulfate per 1L of filtered supernatant collected. While stirring, slowly add the ammonium sulfate to the supernatant over the next couple of hours (~25 g every 15 minutes) to prevent a localized high concentration of ammonium sulfate salt that may cause unwanted proteins to precipitate.
9. Once all the ammonium sulfate has been added, cover the beaker with foil and move with the stir plate to 4°C (e.g., walk-in refrigerator or cold room). Stir overnight. Constant stirring of the solution is required to solubilize the salt, but prolonged stirring can lead to denaturation of proteins in the solution at the surface/air interface.
10. Wash the tubes using the fixed angle rotor.

5. Antibody Purification - Day 2

1. Pour the ammonium-sulfate containing supernatant from the 2L beaker into tubes for a fixed angle rotor. Spin in centrifuge with fixed angle rotor at 6500 RPM for 20 minutes without brake.
2. Vacuum aspirate the supernatant, taking care not to suck up the pellet as it will be soft. Continue using the same set of tubes to collect pellet from the ammonium-sulfate containing supernatant.
3. After the last aspiration, resuspend each pellet (which contains antibodies) in ~1 ml PBS.
4. Remove the ammonium sulfate from the precipitated mAb solution by dialysis against large volumes of PBS. To do this, first cut approximately 1 inch of dialysis tubing (e.g., Membra-Cel MD25-14 MWCO 12,000-14,000 Dalton cut-off cellulose dialysis tubing) for each mL of mAb solution. Wet the tubing with dH₂O. Tie a knot in one end of the tubing and fill with dH₂O to check that knot does not leak. Empty dH₂O from the tubing.
5. Pipette PBS/antibody solution into the tubing. Rinse the tubes with an additional 0.25 ml PBS, and transfer to the tubing.
6. Secure the top of the tubing as close to the solution as possible with an orange dialysis clip.
7. Tape the top of the tubing to the outside top of a 4L beaker. Allow the filled portion of the tubing to hang into the beaker. Fill the beaker with PBS and add a stir bar.
8. Stir overnight for ~8 hours at 4°C.
9. Replace the PBS in the beaker with fresh PBS and stir for ~8 hours two more times.
10. Transfer the antibody from the tubing to 15- or 50-ml conical tubes. Centrifuge for 5 minutes at 1200 RPM to remove any precipitate that may have formed. Transfer the supernatant to a fresh tube.
11. Dilute an aliquot of antibody 1:20 with PBS (5 µl antibody + 95 µl PBS). Quantitate the antibody concentration with a spectrophotometer (blank with PBS) at 280 nm. Use an extinction coefficient (ε) of 1.43. The concentration is calculated as
    
    
12. Aliquot the antibody into screw cap vials and store at -80°C.