Fuente: Frances V. Sjaastad1,2, Whitney Swanson2,3y Thomas S. Griffith1,2,3,4
1 Programa de Posgrado en Microbiología, Inmunología y Biología del Cáncer, Universidad de Minnesota, Minneapolis, MN 55455
2 Centro de Inmunología, Universidad de Minnesota, Minneapolis, MN 55455
3 Departamento de Urología, Universidad de Minnesota, Minneapolis, MN 55455
4 Centro De Cáncer Masónico, Universidad de Minnesota, Minneapolis, MN 55455
Los anticuerpos policlonales se definen como una colección de anticuerpos dirigidos contra diferentes determinantes antigénicos de un antígeno o varios antígenos (1). Si bien los anticuerpos policlonales son herramientas poderosas para identificar moléculas biológicas, hay una limitación importante: son incapaces de distinguir entre antígenos que comparten determinantes antigénicos. Por ejemplo, cuando se utiliza albúmina sérica bovina para inmunizar a un animal, las células B con diferentes superficies Ig responderán a diferentes determinantes antigénicos en la albúmina sérica bovina. El resultado es una mezcla de anticuerpos en el antisuero. Debido a que la albúmina sérica bovina comparte algunos epítopos con albúmina sérica humana en regiones evolutivamente conservadas de la proteína, este antisuero de albúmina sérica antibovina también reaccionará con albúmina sérica humana. Por lo tanto, este antisuero no será útil para distinguir entre albúminas de suero bovino y humano.
Se han adoptado varios enfoques para superar el problema de especificidad de los antisueros policlonales. Una es absorbiendo los anticuerpos no deseados pasando el antisuero a través de una columna de cromatografía de antígenos inmovilizados (2). Este método es tedioso y con frecuencia incapaz de eliminar completamente los anticuerpos no deseados. Otro enfoque es aislar las células B productoras de anticuerpos individuales y expandirlas en cultivo. Sin embargo, como la mayoría de las células no transformadas normales, las células B no sobreviven en el cultivo a largo plazo.
Para superar la incapacidad de las células B para sobrevivir en el cultivo, un enfoque es preparar un hibrioma de células mieloma-B. En 1847, Henry Bence-Jones descubrió que los pacientes con mieloma múltiple, un tumor linfoide, producían una gran cantidad de anticuerpos (3). Las células B de estos pacientes se han vuelto malignas y crecen incontrolablemente. Dado que las células B malignas se derivan de un solo clon, son idénticas y producen un solo tipo de anticuerpo(es decir,un anticuerpo monoclonal o mAb). Sin embargo, la mayoría de estas células del mieloma producen anticuerpos de especificidades desconocidas. En 1975, al fusionar una célula de mieloma a una célula B, Cesar Milstein y Georges Kohler lograron producir un hibrioma que se puede cultivar indefinidamente in vitro y produce un número ilimitado de anticuerpos monoclonales de especificidad antigénica conocida (4). La razón de trás de su enfoque es combinar las propiedades inmortales de la célula del mieloma y el anticuerpo que produce propiedades de la célula B. Su técnica revolucionó la producción de anticuerpos y proporciona un poderoso medio para la identificación y purificación de moléculas biológicas utilizando anticuerpos monoclonales.
Generalmente, preparar un anticuerpo monoclonal requiere varios meses. El procedimiento general incluye los siguientes pasos:
Este protocolo se centra en el último paso : el crecimiento del hibrioma y la preparación del anticuerpo monoclonal. El anticuerpo se purifica del sobrenadante de cultivo mediante precipitación de sulfato de amonio (a menudo conocida como salazón) – un método comúnmente utilizado para eliminar proteínas de una solución. Las proteínas en solución forman enlaces de hidrógeno, junto con otras interacciones hidrófilas, con el agua a través de sus grupos polares e iónicos expuestos. Cuando se añaden concentraciones de iones pequeños y altamente cargados (como amonio o sulfato), estos grupos compiten con las proteínas para unirse al agua. Esto elimina las moléculas de agua de la proteína y disminuye su solubilidad, lo que resulta en la precipitación de la proteína.
Using this protocol, we have obtained the following results with several different hybridomas:
Hybridoma: RB6-BC5 (rat anti-mouse Ly6C/Ly6G (Gr1) IgG2b, κ mAb)
OD280 – 1.103
(1.103/1.43)(20) = 15.42 mg/mL
Hybridoma: GK1.5 (rat anti-mouse CD4 IgG2b, κ mAb)
OD280 – 0.485
(0.485/1.43)(20) = 6.78 mg/mL
Hybridoma: 2.43 (rat anti-mouse CD8 IgG2b, κ mAb)
OD280 – 0.209
(0.209/1.43)(20) = 2.92 mg/mL
These are all example results, and it is important to note that each production run with the same hybridoma can be slightly different in the amount of mAb available at the end.
The procedure outlined above is a simple, straight-forward way to purify monoclonal antibodies from hybridoma culture supernatant. It is important to remember, though, that the ammonium sulfate will precipitate other proteins that may be in the culture supernatant. Consequently, the antibody concentrations determined from the absorbance measurements are estimates. The user may wish to assess the purity of the dialyzed sample by running a small amount on an SDS-polyacrylamide gel. The mAb produced by a hybridoma, once purified using this methodology, can be used in a variety of ways. The above-described RB6-BC5, GK1.5, and 2.43 mAb are commonly used for in vivo depletion of neutrophils, CD4 T cell, and CD8 T cells, respectively, in mice. Other mAb produced and purified using this protocol can be used for flow cytometry (when conjugated to a fluorophore or in conjunction with a secondary Ab), ELISA, or Western blotting.
Antibodies are a powerful tool for research and diagnosis, which means producing them in large quantities is often necessary.
The first step to generating antibodies is to inject the antigen of interest into a host animal. The antigen activates the host’s B-cells which then produce and release antibodies specific to that antigen. Then, regular screening of the host animal’s antisera for the presence of the target antibody is carried out, using ELISA or another detection method. Once it’s detected, the host animal’s spleen, which contains the B-cells, is removed. If all of the B-cells from the spleen are now isolated, this should include a population which are secreting antibodies to the antigen of interest. We refer to this population as polyclonal, because each cell likely bound to a different epitope of the antigen, and therefore, generated its own individual and unique antibody.
To generate monoclonal antibodies, antibodies raised to recognize one specific epitope, the individual B-cell that produces the desired antibody must first be isolated and cultured. Unfortunately, B-cells do not survive well in culture. So to overcome this hurdle, scientists fuse B-cells with immortal myeloma cells, resulting in hybridomas. These cells are then grown in a selective medium that only allows the hybridomas to grow and release antibodies. Again, the medium is screened using a method such as ELISA for the desired antibody. Once it is detected, the hybridomas are cloned via a process called limiting dilution, a serial dilution of the parent culture, which should result in single cells being seeded into the wells of a screening plate. This allows growth of hybridomas from a single parent cell, yielding a monoclonal cell line that only releases the desired antibody. These monoclonal lines can be expanded in tissue culture flasks to produce large quantities of monoclonal antibody. After this, as the cells begin to die off, the antibodies can be precipitated from the medium with ammonium sulfate. Normally, in solution, antibodies interact with water through hydrophilic interactions. However, ammonium and sulfate are highly-charged ions that separate the water molecules from the antibodies, decreasing the solubility of the antibodies and causing them to precipitate.
To begin, first check the list of materials and prepare all the media, supplies, and work surfaces for the protocol.
Then, turn on a water bath and set it to 37 degrees Celsius. Next, add 10 milliliters of complete RPMI to a 15-milliliter conical tube and 15 milliliters of complete RPMI to a T75 cell culture flask and set them aside. Using caution and wearing the appropriate personal protective equipment, remove the frozen vial containing hybridoma cells from the liquid nitrogen storage. To release the pressure inside the vial, loosen the cap slightly. Now, carefully incubate the vial in the water bath, making sure that the cap remains above the water surface to minimize the chances of contamination. When the cells are almost thawed, which typically takes around two minutes, move the vial to the tissue culture hood.
Then, wipe the outside of the vial with 70% ethanol before removing the cap. Using a sterile pipette, transfer the cells into the conical tube that contains 10 milliliters of complete RPMI medium. Then, centrifuge the tube for five minutes at 1200 RPM. After centrifugation, move the tube back to the tissue hood and wipe the outside of the tube with ethanol. Without disturbing the pellet, discard the supernatant and then add five milliliters of fresh complete RPMI medium and gently pipette up and down to resuspend. Next, transfer the cells to the T75 cell culture flask and place the flask inside a 5% carbon dioxide incubator at 37 degrees Celsius. Allow the cells to reach approximately 80% confluency, which usually takes about three days. Notice that hybridoma cells are nonadherent and will grow suspended in the medium. The time to reach sufficient confluency may vary based on the starting number of live cells and the type of hybridoma cell used.
Once the cells are sufficiently confluent, use a sterile 25-milliliter pipette to transfer them from the culture flask into a conical centrifuge tube. Pellet the cells by centrifugation at 1200 RPM for five minutes. While the cells are in the centrifuge, add 18 milliliters of complete RPMI into each of three new T75 cell culture flasks and set these aside. After centrifugation, remove the supernatant and gently resuspend the cell pellet in six milliliters of complete RPMI. Next, add two milliliters of the cell suspension into each of the three new cell culture flasks. Finally, place the flasks into an incubator set to 5% carbon dioxide and 37 degrees Celsius and incubate until the flasks are around 80% confluent, approximately three days.
At this point, the cells are ready to continue their growth in the serum-free medium designed for hybridoma cell lines, such as commercially-available HB Basal Liquid medium containing the HB101 supplement. Transfer the cells from each cell culture flask into conical centrifuge tubes and then pellet the cells by centrifugation at 1200 RPM for five minutes. Now, add 230 milliliters of supplemented HB101 serum-free medium into each of six 225-centimeter-squared cell culture flasks and set them aside. When centrifugation is complete, remove the supernatant and resuspend each pellet in 10 milliliters of supplemented HB101 medium. Then, into each cell culture flask, add five milliliters of the cell suspension. Place the flasks in the 5% carbon dioxide incubator at 37 degrees Celsius and continue growing the cells for about three weeks. During this time, the cells will produce and release the monoclonal antibody of interest into the culture medium and the antibody will be ready for purification when the cells start to die.
To remove the cellular debris from the antibody-containing culture media, pour the contents of the culture flasks into tubes for a fixed angle rotor. Place the tubes in the rotor and make sure it is properly balanced prior to centrifugation. Spin the tubes at 10,000 RPM for eight minutes. While the samples are centrifuging, place a two-liter plastic beaker with a stir bar into an ice bucket and then put the ice bucket on a stir plate.
Next, attach a 500-milliliter filter top to a one-liter bottle. Attach this bottle top filter unit to a house vacuum using the appropriate tubing. Then, pour the supernatant that contains the antibody into the filter top. Centrifuge the remaining media to separate the cell debris from the antibody-containing supernatant. When the filter top is full of supernatant, start the vacuum. Then, when the one-liter collection bottle is close to full, remove the filter top and pour the filtered supernatant into the two-liter beaker on ice. Repeat the filtration steps until all of the supernatant is processed.
When all of the sample has been processed, weigh 295 grams of ammonium sulfate per one liter of filtered supernatant. Start the stir plate and slowly add the ammonium sulfate to the supernatant over the next couple of hours. This prevents a localized high concentration of ammonium sulfate salt that may cause unwanted proteins to precipitate. Once all of the ammonium sulfate has been added, cover the beaker with foil and move it, along with the stir plate, to a cold room at four degrees Celsius and set it to stir the antibody solution overnight.
The next morning, pour the ammonium sulfate-containing antibody solution from the two-liter beaker into clean tubes for the fixed angle rotor. Centrifuge the tubes at 6500 RPM for 20 minutes without break to pellet the antibody at the bottom of the tubes. Next, vacuum aspirate the supernatant, using caution not to suck up the soft pellet. Continue using the same set of tubes to collect the pelleted antibody from the remainder of the ammonium sulfate-containing supernatant. After the last aspiration, re suspend each antibody pellet in approximately one milliliter of PBS.
To remove the ammonium sulfate from the antibody solution, first cut approximately one inch of dialysis tubing for each milliliter of antibody solution. Next, wipe the tubing with distilled water and tie a knot on one end of the tubing. Fill the tubing with distilled water to check for leakage from the knot. If there is no leakage after a few minutes, empty the water out of the tubing.
Next, pipette the antibody solution into the tubing. To recover as much antibody as possible, rinse the tubes with an additional 0.25 milliliters of PBS and transfer this to the tubing also. Secure the top of the tubing as close to the solution as possible with a dialysis clip. Then, tape the top of the tubing to the outside top of a four-liter beaker with the filled portion of the tubing hanging into the beaker. Now, take the beaker to the four degree Celsius cold room and place it onto a stir plate. Fill the beaker to the top with PBS and add a stir bar. Allow the tube and solution to stir overnight for approximately eight hours. The next morning, replace the PBS in the beaker with fresh PBS and then leave the beaker to stir again for approximately eight hours. Later that evening, repeat the process one final time. In the morning, open up the dialysis tube and then transfer the antibody solution from the tubing to 15-milliliter conical tubes. To remove any precipitant that may have formed during dialysis, centrifuge the tubes for five minutes at 1200 RPM. Finally, transfer the supernatant to fresh tubes.
To quantify the antibody concentration, first make a 20-fold dilution by adding five microliters from an antibody aliquot to 95 microliters of PBS. Then, pipette the diluted antibody into a cuvette and use a spectrophotometer to record the concentration at 280 nanometers. Next, calculate the antibody concentration using the formula shown. Finally, label screw cap vials with the antibody name, concentration, date of preparation, and, if applicable, batch number and experimenter name, and then aliquot the antibody into the labeled screw cap vials. These can be stored at minus 80 degrees Celsius until needed.
Example yields using the 120G8 anti-mouse CD317 or PDCA-1 hybridoma line ranged between 44 and 99.6 milligrams, which typically yields, on average, 67.3 milligrams amount. It is important to note that each production run with the same hybridoma cell line can be slightly different in the amount of monoclonal antibody available at the end.
