We use 3 different elution protocols in our lab. All presume you have washed your beads with PBS-Tween (0.01-0.1%) for 3-4 times, spin down your beads and remove the washing buffer completely (we use magnetic racks to do this quick and easy).

Then for SDS-PAGE: Boil beads for 5 minutes in 30 uL 1x XT Sample buffer at 95 degrees. You can directly use this for SDS-PAGE, but your protein will be denatured.

Two milder protocols include eluting in low pH at 56 degrees centigrate for 15 minutes (make sure you vortex mildly every 2 or 3 minutes to keep your beads mixed). You can chose between 1M Glycine solution at pH=2.5 or a 0.1M citric acid solution at pH=2.5.

You can optimize the ideal pH value if you want, normally you will want it somewhere between 2 and 3.

The volume we typically use for this is 30 uL of elution buffer, but depending on your application you can change this.

The Glycine elution is the mildest of the three, but for some protocols (ie DIGE) the elution buffer interferes with your colouring dyes. In this case, you should chose for the citric acid elution. For your application though, I don't think it realy matters. Try glycine and citric acid and see which works best for you.

Something to watch out for is the co-elution of other proteins. In our lab we use DMP to crosslink our antibodies to the beads to prevent co-elution of IgGs, which has been quite a problem since they show up at 3 different spots on your gel. Albumin and other high abundant proteins also tend to co-elute and show up in your samples. If this bothers you, you can use some depletion or clean-up protocols to reduce the aspecific binding.

I hope this helps, good luck!

It depends on what you want to do with your sample afterwards.

Do you want to analyze it by Western blot?

After the final wash step, you should have almost dry beads, just add 2xLaemmli Buffer, mix and heat the sample

If you need your sample in a native state, you can try to elute with low pH buffer and then readjust the pH or try to displace your protein by adding an excess of the antigenic peptide.

I wanna do some functional assay after elution with the protein....

Have a look at these papers:

http://www.piercenet.com/files/TR0027-Elution-conditions.pdf

http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0018218

If you want just protein eluted, Pierce has a variety of reagents to covalently link antibody to bead. Then just use low ph or add peptide epitope specific to Ab (if known). For function, you probably do not want antibody present which may inhibit function?

If you want to retain functionality you can try eluting with 20 mM MES or PIPES pH 6.5, 3.5 M MgCl2, then dialyse or diafiltrate back into your chosen buffer. See Harlow and Lane - Antibodies for the exact recipe. A very good reference for working with antibodies.

For Jeffrey´s suggestion FAb fragments retain a carbohydrate moiety.

This is an ideal location for derivitising and attachment for immobilisation, in that there is no chance of modifying the all important variant regions inadvertantly.

Again Harlow and Lane is your friend for the protocols, periodate treatment allows attachment of biotin hydrazide to the carbohydrate, then simply attach to strepavidin magnetic beads.

You can then use the MgCl2 gentle elution system, to elute your target protein. Then clean up with a pH2 strip and reuse the beads. Simply store under the same conditions recommended for the antibody.

We use 3 different elution protocols in our lab. All presume you have washed your beads with PBS-Tween (0.01-0.1%) for 3-4 times, spin down your beads and remove the washing buffer completely (we use magnetic racks to do this quick and easy).

Then for SDS-PAGE: Boil beads for 5 minutes in 30 uL 1x XT Sample buffer at 95 degrees. You can directly use this for SDS-PAGE, but your protein will be denatured.

Two milder protocols include eluting in low pH at 56 degrees centigrate for 15 minutes (make sure you vortex mildly every 2 or 3 minutes to keep your beads mixed). You can chose between 1M Glycine solution at pH=2.5 or a 0.1M citric acid solution at pH=2.5.

You can optimize the ideal pH value if you want, normally you will want it somewhere between 2 and 3.

The volume we typically use for this is 30 uL of elution buffer, but depending on your application you can change this.

The Glycine elution is the mildest of the three, but for some protocols (ie DIGE) the elution buffer interferes with your colouring dyes. In this case, you should chose for the citric acid elution. For your application though, I don't think it realy matters. Try glycine and citric acid and see which works best for you.

Something to watch out for is the co-elution of other proteins. In our lab we use DMP to crosslink our antibodies to the beads to prevent co-elution of IgGs, which has been quite a problem since they show up at 3 different spots on your gel. Albumin and other high abundant proteins also tend to co-elute and show up in your samples. If this bothers you, you can use some depletion or clean-up protocols to reduce the aspecific binding.

I hope this helps, good luck!

If your protein can be refolded, you can elute them from the bead with 6 M urea.

Then remove the urea by dialysis to refold your protein.

I used 25 mM glycine-HCl buffer pH 2.5 for elution of protein from Protein A acrylic beads. Next, I neutralized the eluates with 25 mM Trizma Base to pH 7. Initially, I washed 3times beads with 25 mm Tris HCl pH 7.5 before elution.

If your protein is not the same size as your antibody, you can elute with 0.1M glycine, pH2-3 and then separate the protein of interest from the antibody using SEC (like superdex 200).

If your protein and antibody are the same size, you should crosslink your antibody to the Protein A resin using DMP before adding your protein. Then, wash the resin with your elution buffer (0.1M glycine, pH 2-3) to remove any antibody that is not crosslinked. Then, you can add your protein of interest. This time, when you elute with your glycine buffer, you will get only the protein of interest, with no, or very little, antibody. Make sure you immediately neutralize the pH of your eluted protein with a high molarity buffer at pH 7-8 if you want the protein to remain functional. Then you can use a desalting column to quickly get your protein back into a more ideal buffer for functional testing.

If all you want to do is run bound protein on gel, use PAGE gel elution buffer (Laemmli). Then run gel. If you do not want sntibody on gel, crosslink to bead.