Do a Blue Native PAGE.

Basically, it is like SDS-PAGE ... except that:

1) Absolutely no Sodium dodecyl sulfate (SDS), none in the buffer, none in the gel, nowhere.

2) Absolutely no boiling of protein samples

3) Use Coomossie G250 as your loading dye for your proteins. (Yeah, this is important).

Hook up your electrodes like normally black to black, red to red. Your protein will move toward the Anode(+) at the bottom of the gel.

Coomossie G250 gives proteins a negative charge, but unlike SDS, it does not linearize or denature proteins. If Blue Native PAGE works then you will get a band at double the molecular weight Indicating the dimer.

Then Coomossie stain your gel after you run it:

40% ethanol, 10% Acetic Acid, 3% Coomossie R250. Protocol in my lab conventionally uses methanol but ethanol works so much better as I realized.

You could also do Native PAGE, but if your protein is neutrally or positively charged you will get no movement into the gel, or your protein will move toward the cathode and out of the gel. You could invert the electrodes then so the cathode is at the bottom of the gel.

i guess you could western blot the gel, but you shouldn’t need to if you are good at protein purification.

Do a Blue Native PAGE.

Basically, it is like SDS-PAGE ... except that:

1) Absolutely no Sodium dodecyl sulfate (SDS), none in the buffer, none in the gel, nowhere.

2) Absolutely no boiling of protein samples

3) Use Coomossie G250 as your loading dye for your proteins. (Yeah, this is important).

Hook up your electrodes like normally black to black, red to red. Your protein will move toward the Anode(+) at the bottom of the gel.

Coomossie G250 gives proteins a negative charge, but unlike SDS, it does not linearize or denature proteins. If Blue Native PAGE works then you will get a band at double the molecular weight Indicating the dimer.

Then Coomossie stain your gel after you run it:

40% ethanol, 10% Acetic Acid, 3% Coomossie R250. Protocol in my lab conventionally uses methanol but ethanol works so much better as I realized.

You could also do Native PAGE, but if your protein is neutrally or positively charged you will get no movement into the gel, or your protein will move toward the cathode and out of the gel. You could invert the electrodes then so the cathode is at the bottom of the gel.

i guess you could western blot the gel, but you shouldn’t need to if you are good at protein purification.

The simplest option is Native PAGE as Adron Ung suggested. An alternative strategy to corroborate your data is by gluteraldehyde crosslinking.

For glutaraldehyde treatment, reaction mixtures with 50 to 100 µg of protein in 20 mM HEPES buffer (pH 7.5) in a total volume of 100µl are treated with 5 µl of 2.3% freshly prepared solution of glutaraldehyde for 2 to 5 minutes at 37C. The reaction is terminated by addition of 10 µl of 1 M Tris-HCl, pH 8.0. Cross-linked proteins are solubilized by addition of an equal volume of Laemmli sample buffer to which a few µl of 0.1% bromophenol blue is added and electrophoresis is conducted in 5% to 20% SDS-polyacrylamide gels. Your oligomers will show up as multiples of your monomer's molecular weight. Couple this run with a Western blot to validate your protein and its dimer.

Here is a link to a more detailed protocol : Article Determination of protein oligomerization state: Two approach...

Hello, see an attached paper with all descriptions in detail.

good luck!

In addition to other suggestions, you can use Size-exclusion chromatography

Suipwksiow Wjkp As I understand, you have got 60 kDa size with SDS-PAGE. Perform the gel filtration (size-exclusion) chromatography with your protein, and based on Ve/Vo values, you will know whether it is dimer or not. Thats quite state-forward.

I would recommend SEC-MALS. It will give you all its oligomeric states with their respective molecular weight.