Antibody purification typically involves two to three steps: an initial capture step (most often affinity chromatography), followed by one or more polishing steps to remove remaining contaminants. The method you choose depends on the antibody source, the species it comes from, and how pure the final product needs to be. Here’s a practical walkthrough of the major techniques, from crude preparation to final polishing.
Starting With a Crude Preparation
Before running any chromatography column, you can reduce the complexity of your sample with ammonium sulfate precipitation. Adding ammonium sulfate to serum or cell culture supernatant causes antibodies to fall out of solution while many other proteins stay dissolved. A concentration of 40% to 50% saturation precipitates IgG from most species, and 50% is the standard starting point. You collect the pellet by centrifugation, resuspend it in your working buffer, and then move to a more selective purification step.
This step isn’t strictly necessary if you’re working with a relatively clean starting material like hybridoma supernatant, but it’s useful when processing serum or ascites fluid where you want to remove the bulk of irrelevant proteins before loading an expensive affinity column.
Affinity Chromatography: The Workhorse
Affinity chromatography using Protein A, Protein G, or Protein L resins is the single most effective capture step for antibody purification. These bacterial proteins bind to specific regions on antibody molecules, letting you wash away nearly everything else before eluting your antibody in a purified form. Modern Protein A chromatography routinely achieves recovery rates of 85% to 99% with purity above 90% in a single step.
Choosing Between Protein A, G, and L
Each ligand binds a different part of the antibody, which matters depending on what species and subclass you’re working with. Protein A binds the Fc region (the “tail” of the antibody) and works best with human IgG1, IgG2, and IgG4, rabbit immunoglobulins, and mouse IgG2a. It has notably weak binding to mouse IgG1 and essentially no binding to rat IgG1 or IgG2a, which catches people off guard.
Protein G also binds the Fc region but covers some of the gaps Protein A leaves. It binds all four human IgG subclasses strongly, including IgG3, which Protein A misses entirely. It’s also a better choice for mouse IgG1 and rat IgG2a. For bovine, sheep, and goat antibodies, Protein G outperforms Protein A significantly.
Protein L takes a completely different approach: it binds certain kappa light chains rather than the Fc region. This makes it uniquely suited for purifying antibody fragments like Fab, F(ab’)2, and single-chain variable fragments (scFv) that lack an Fc region. Protein L also captures IgM, IgA, IgD, and IgE from human sources, all of which Protein A and G handle poorly. The catch is that Protein L only recognizes specific kappa subtypes. Human kappa I, III, and IV bind strongly, but kappa II does not. It also won’t bind lambda light chains at all, so you need to know your antibody’s light chain composition before relying on this ligand.
Elution and Neutralization
Elution from affinity columns uses low-pH buffers to disrupt the interaction between the ligand and your antibody. A common choice is 150 mM glycine-HCl at pH 3.5, or 50 mM sodium acetate at the same pH. The problem is that antibodies don’t love sitting at pH 3.5. Prolonged exposure causes aggregation and loss of activity, so you need to neutralize your eluted fractions quickly.
Adding a small volume of 1 M Tris base to each collection tube before elution is standard practice. Glycine-HCl buffers have a pKa of 2.34, meaning they require relatively little Tris to bring the pH back up to 7.5. You should have your neutralization buffer pre-dispensed into tubes so the antibody spends as little time as possible at low pH.
Purifying Recombinant Antibodies With Tags
If you’re expressing recombinant antibodies or antibody fragments in bacterial or mammalian cells, you may have engineered a purification tag onto the protein. His-tags (a stretch of six to ten histidine residues) are the most common. These bind to metal affinity resins like cobalt or nickel beads. You wash the column with a low concentration of imidazole (around 30 mM) to remove weakly bound contaminants, then elute your tagged antibody with a higher concentration (500 mM imidazole).
GST-tags offer another option, where the antibody is fused to glutathione S-transferase. Rather than competing the antibody off the resin with a chemical, you can use an enzyme (such as PreScission protease) to cleave the tag directly on the beads. This leaves you with a tag-free antibody in solution while the GST portion stays stuck to the resin. The cleavage step runs for three hours to overnight at 4°C.
Ion Exchange Chromatography for Polishing
After affinity capture, ion exchange chromatography is the most common polishing step. It separates proteins based on their surface charge, which depends on the pH of your buffer relative to the protein’s isoelectric point (pI). Most monoclonal antibodies have a pI between 7.3 and 8.7, meaning they carry a net positive charge at neutral or slightly acidic pH.
Cation exchange chromatography (CEX) is the go-to for antibodies. You load your sample at a pH below the antibody’s pI so that the antibody binds the negatively charged resin. Contaminants with lower pI values flow through, and you then elute the antibody by increasing the salt concentration or raising the pH. CEX is particularly good at separating charge variants of the same antibody, which matters for pharmaceutical quality control. If your contaminants have a higher pI than your antibody, anion exchange at a pH above the antibody’s pI can work instead, though this is less common.
Size Exclusion Chromatography
Size exclusion chromatography (SEC) separates molecules purely by their physical size. You run your sample through a column packed with porous beads: smaller molecules enter the pores and take longer to travel through, while larger molecules pass around the beads and elute first. For antibody purification, SEC is primarily a polishing tool used to remove two problematic species: high-molecular-weight aggregates (which elute before the antibody) and small fragments or degradation products (which elute after).
Newer mixed-mode resins combine size exclusion with additional chemical interactions to improve selectivity. One such resin demonstrated 90% antibody yield while removing 37% of aggregates and 100% of Fc fragments in a single pass. SEC is gentle and doesn’t require harsh buffers, but it’s a low-throughput method, so it’s typically reserved for final polishing rather than the primary capture step.
Hydrophobic Interaction Chromatography
Hydrophobic interaction chromatography (HIC) separates proteins based on the hydrophobic patches on their surface. You load your sample in a high-salt buffer, which encourages proteins to bind to the hydrophobic resin. Typical binding conditions use ammonium sulfate at concentrations ranging from 200 to 650 mM in a sodium acetate buffer around pH 5.2 to 5.6, depending on the antibody. You then elute by gradually decreasing the salt concentration, which weakens the hydrophobic interaction and releases your antibody.
HIC is particularly useful as a second or third polishing step because its selectivity is orthogonal to ion exchange: it separates based on hydrophobicity rather than charge, catching contaminants that ion exchange misses.
Buffer Exchange and Concentration
After purification, you’ll almost always need to swap your antibody into a storage-compatible buffer and concentrate it. Three common methods exist, and they differ substantially in speed and practicality.
Dialysis is the gentlest option and the gold standard for fragile antibodies prone to degradation. The tradeoff is time: you need buffer volumes at least 500 times the sample size, with multiple exchanges, and the process takes hours to overnight. It also dilutes your sample, so you’ll need a separate concentration step afterward.
Gel filtration (desalting columns) is faster but requires you to collect and assess fractions manually, which is tedious and introduces some sample loss. Like dialysis, it dilutes your sample.
Centrifugal ultrafiltration (diafiltration) outperforms both on practical grounds. It concentrates your antibody and exchanges the buffer simultaneously in a single device, with shorter processing times, less hands-on work, and better protein recovery. For most routine purifications, this is the preferred method.
Storing Your Purified Antibody
Storage conditions matter as much as the purification itself. Purified antibodies are typically stored at 4°C for short-term use (days to weeks). For longer storage, aliquoting and freezing at -20°C or -80°C prevents degradation, though repeated freeze-thaw cycles cause aggregation and should be avoided.
High-concentration formulations above 100 mg/mL, which are increasingly important for therapeutic antibodies intended for subcutaneous injection, present special stability challenges. At these concentrations, antibodies can undergo liquid-liquid phase separation at refrigerated temperatures. Adding stabilizers like arginine-glutamate to the formulation prevents this phase separation and maintains a homogeneous solution during storage at 4°C. For research-scale antibodies at lower concentrations, adding a small amount of carrier protein or glycerol (typically 50%) to the storage buffer helps prevent surface adsorption losses and maintains activity over months of freezer storage.