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
- Incubate the vial containing the frozen hybridoma cells in a 37°C water bath until just thawed (approximately 2 minutes).
- 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).
- Centrifuge for 5 min at 1200 RPM to wash out any contaminating freeze media.
- 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).
- Grow the cells in a standard incubator at 37°C with 5% CO2. Cells are non-adherent while in culture.
2. Hybridoma expansion
- 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. - 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.
- 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.
- The cells from each of the three T75 flasks (at ~80% confluency) are collected and centrifuged (1200 RPM for 5 min).
- 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).
- 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.
- 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. - Prepare a 2L plastic beaker with a stir bar in a bucket surrounded by ice. Place on a stir plate.
- Attach a 500 mL filter top on a 1L bottle. Attach bottle top filter unit to house vacuum via appropriate tubing.
- Pour the supernatant from step 4.1 (which still contains the mAb produced by the hybridoma cells) into the filter top.
- 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.
- 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.
- Repeat the centrifugation and filtration steps until all the media is processed.
- 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.
- 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.
- Wash the tubes using the fixed angle rotor.
5. Antibody Purification - Day 2
- 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.
- 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.
- After the last aspiration, resuspend each pellet (which contains antibodies) in ~1 ml PBS.
- 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.
- Pipette PBS/antibody solution into the tubing. Rinse the tubes with an additional 0.25 ml PBS, and transfer to the tubing.
- Secure the top of the tubing as close to the solution as possible with an orange dialysis clip.
- 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.
- Stir overnight for ~8 hours at 4°C.
- Replace the PBS in the beaker with fresh PBS and stir for ~8 hours two more times.
- 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.
- 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
- Aliquot the antibody into screw cap vials and store at -80°C.
Source: Frances V. Sjaastad1,2, Whitney Swanson2,3, and Thomas S. Griffith1,2,3,4
1 Microbiology, Immunology, and Cancer Biology Graduate Program, Uni…
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
- Incubate the vial containing the frozen hybridoma cells in a 37°C water bath until just thawed (approximately 2 minutes).
- 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).
- Centrifuge for 5 min at 1200 RPM to wash out any contaminating freeze media.
- 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).
- Grow the cells in a standard incubator at 37°C with 5% CO2. Cells are non-adherent while in culture.
2. Hybridoma expansion
- 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. - 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.
- 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.
- The cells from each of the three T75 flasks (at ~80% confluency) are collected and centrifuged (1200 RPM for 5 min).
- 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).
- 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.
- 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. - Prepare a 2L plastic beaker with a stir bar in a bucket surrounded by ice. Place on a stir plate.
- Attach a 500 mL filter top on a 1L bottle. Attach bottle top filter unit to house vacuum via appropriate tubing.
- Pour the supernatant from step 4.1 (which still contains the mAb produced by the hybridoma cells) into the filter top.
- 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.
- 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.
- Repeat the centrifugation and filtration steps until all the media is processed.
- 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.
- 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.
- Wash the tubes using the fixed angle rotor.
5. Antibody Purification - Day 2
- 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.
- 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.
- After the last aspiration, resuspend each pellet (which contains antibodies) in ~1 ml PBS.
- 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.
- Pipette PBS/antibody solution into the tubing. Rinse the tubes with an additional 0.25 ml PBS, and transfer to the tubing.
- Secure the top of the tubing as close to the solution as possible with an orange dialysis clip.
- 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.
- Stir overnight for ~8 hours at 4°C.
- Replace the PBS in the beaker with fresh PBS and stir for ~8 hours two more times.
- 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.
- 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
- Aliquot the antibody into screw cap vials and store at -80°C.
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.
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Q1: Why can't B-cells be cultured directly to produce monoclonal antibodies?
B-cells do not survive well in long-term culture, making it impossible to maintain them indefinitely for antibody production. To overcome this limitation, scientists fuse B-cells with immortal myeloma cells to create hybridomas, which combine the antibody-producing properties of B-cells with the indefinite growth capacity of myeloma cells.
Q2: What is the difference between polyclonal and monoclonal antibodies?
Polyclonal antibodies are a mixture of antibodies, each recognizing different epitopes on an antigen, produced by multiple B-cell clones. Monoclonal antibodies are identical antibodies produced by a single hybridoma cell line, recognizing only one specific epitope. This makes monoclonal antibodies more specific and useful for distinguishing between similar antigens.
Q3: How does limiting dilution ensure monoclonal antibody production?
Limiting dilution is a serial dilution process that statistically ensures single cells are seeded into screening plate wells. Wells containing one cell grow into monoclonal populations producing identical antibodies. This cloning method guarantees that the resulting hybridoma line releases only the desired antibody of known specificity.
Q4: What role does ammonium sulfate play in antibody purification?
Ammonium sulfate precipitation removes antibodies from culture medium by competing with proteins for water molecules through hydrophilic interactions. The highly charged ammonium and sulfate ions decrease antibody solubility, causing them to precipitate out of solution. This salting-out method is a standard technique for protein purification from culture supernatants.
Q5: Why is dialysis tubing used after ammonium sulfate precipitation?
Dialysis tubing removes residual ammonium sulfate from the antibody solution through osmotic exchange. The antibody solution is placed in dialysis tubing and immersed in PBS buffer overnight. Water and small salt ions pass through the tubing membrane while antibodies remain inside, resulting in purified antibody free of ammonium sulfate.
Q6: How is antibody concentration determined after purification?
Antibody concentration is measured using a spectrophotometer at 280 nanometers, which detects protein absorbance. A 20-fold dilution of the antibody sample is prepared in PBS, then analyzed using the spectrophotometer. The concentration is calculated using a standard formula and recorded for documentation and storage of the purified monoclonal antibody.
Q7: What screening method is used to identify hybridomas producing the desired antibody?
ELISA assays indirect sandwich and competitive methods are used to screen hybridoma culture media for the presence of the target antibody. After hybridoma fusion and limiting dilution cloning, ELISA detects which wells contain cells producing the desired monoclonal antibody. This screening step ensures only hybridomas with the correct antibody specificity are selected for expansion and antibody production.
Chapters in this video
0:01
Concepts
2:51
Hybridoma Expansion
6:04
Antibody Production in Serum-Free Medium
7:28
Antibody Purification - Day 1
9:25
Antibody Purification - Days 2-4
12:00
Analysis and Results
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