What is the MW of your ligand ? Is Superdex75 resolution sufficient to see the difference protein alone and protein + ligand ? If the protein itself dimerize upon ligand interaction, then it is not a resolution problem. Is the interaction stable to gel filtration buffer composition (i.e presence of salt) ?

Since your protein has an approximate size of 12 kDa you should definitely be able to see a difference on your gel filtration run using an S75 ... if your protein does indeed dimerise upon adding the ligand. Jean-Michel makes a good point regarding the buffer composition, as there is a good chance that the interaction does not occur or is not stable enough under your gel filtration buffer condition. You could repeat the gel filtration experiment using a variety of buffers changing pH and ionic strength (high salt in particular might interfere with your ligand binding), but you could probably test many different conditions faster by using other analytical methods and then, when you've established a good buffer condition, repeat the gel filtration experiment.

HI

Thanks for your suggestions.My ligand is around 2 kda and it has strong binding with the protein. according to literature it has to form dimer when it binds with the protein. So as all suggests, I will vary the buffer / PH/ salt composition in the gel filtration and repeat the experiment.

Between when i run the protien- ligand in the gel filtration column, i see an aggregate peak (peak eluted with in dead volume of the column) together with the peak of my protein (My protein elutes at 13ml in the s75 tricon column) but when I run the protein alone, I dint see any aggregation. I am not sure whether the ligand addition makes the protein to undergo aggregation

As well the UV absorbance in the gel filtration is in the negative range when I run protein and Protein-Ligand. I have loaded quite reasonable amount of sample around 3mg/ml for the experiment run. I have auto zero in the unicorn program when I start experiment. But I dont know when the peak seems in negative range. Kindly advise and your suggestions are welcome,

Thanks in advance

Do you mean one ligand molecule binds to two molecules of your protein. If this is the case then the molecular weight of the complex should be around 22kDa which is double the size of your protein alone.

But if the protein binds to 2 ligand molecules the molecular mass difference between the protein alone and the complex is just 4kDa which would be difficult to distinguish on S75 column.

Are both your protein and ligand in the same buffer as that of S75 column in which it is equilibrated. If they are in different buffer then that could also cause some aggregation that is detected. Did you check the aggregated fraction on the gel and find if it has both ligand as well as the protein?

I would suggest you to mix your protein (10k) with excess of ligand (2k) which would ensure complete saturation of your protein with the ligand. Now run this mixture on the gelfiltration column to get two peaks : one of the complex and and then very late  in the course of gel-filtration another peak of the ligand. May be if you compare this peak with the protein peak you may be able to see a difference.

Check these options out.. Be sure that your protein, ligand and the gel filtration column all are in the same buffer. All the best

Hi Anish

thanks for your suggestions. My ligand is not soluble in water. I have used DMSO to soluble it. 

But other than my own protein peak (eluted at 13ml inS75 tricon column), I always get peak at 18ml (which is around 1 kda). This 18ml peak elutes regardless of injecting protein or protein - ligand complex. Is there any possibility of degradation occurs at such a low molecular weight around 1kda ? This 18ml peak elutes even after His trap when I run gel filtration.

Appreciate your suggestions.

To follow up Anish's answer: another possibility is that your ligand dissociates during the course of the gel filtration, due to dilution that occurs and directly linked to the KD of the interaction. I have performed a number of experiments with a S200 column where  compounds were able to tetramerize a protein when bound to it. In some case for cpds with lower affinity tetramerization was not seen unless compound was added to the mobile phase preventing dissociation of the complex. Therefore what I did was to incubate my protein prior gel filtration with an excess of cpd (at least a 2:1 ratio) to ensure saturation and run the gel filtration in the presence of cpd in the mobile phase at a concentration around 10 X KD. Then it worked !

Hi all,

thanks very much for your suggestion. As Jean-Michel suggested, I always incubate my protein & ligand prior running gel filtration.Since the ligand is soluble in DMSO, when I add ligand to protein, I see some cloudy sedimentation like. I always spin down my samples before run into Gel filtration column. 

When I run protein  + ligand complex in s75 column, profile peak seems more noisy and hence I couldnot conclude whether it dimerise or not.

Is there any analytical method/protocol to check the dimerisation without running gelfiltration or MALS technique ?

Analytical ultracentrifugation is one of the most common methods for assessing the oligomerisation of proteins. The other method you can definitely use in your lab if you don't have access to an ultracentrifugation service is to run a native gel. The native gel has the advantage that you can run both your protein without ligand and the protein with ligand on the same gel and compare them. Native gels are tricky and often look terrible, but even when they look smeary and not good enough to include in a publication, they will still let you see if your protein is a monomer or a dimer, trimer, etc... under the buffer conditions you are testing. For the last few native gels I made I followed the blue native gel method and I found this works a lot better than traditional native gels.

Sorry Ya Yesh, I just realised what you mean. I thought you meant your protein (12 kDa) dimerised upon binding to the ligand (2 kDa), which would result in a 26 kDa complex and therefore should be obvious on the S75. You actually mean that the ligand dimerises upon binding the protein (is that right?), which should form a 16 kDa complex.

It is difficult to separate two peaks at 12 and 16 kDa on an S75, unless they are very pure, run nicely (no shoulders or smears on the peaks) and you load a very small amount. The more you load on the column the bigger the base of each peak will be and the more likely they are to mask each other.

You would also need to run a very good native gel to see a 4 kDa difference and that is difficult.

Nonetheless, I would still give it a shot. Optimise your buffer (dynamic light scattering is a good tool for this) and then run a very small sample on the column and also try the blue native gel if that fails.

One problem you might have is that gel filtration only measures the hydrodynamic radius, not necessarily the molecular weight. So if you have a monomeric species that has a quite extended conformation (more random coil percentage), and a dimer that is more compact (more beta-sheet/helix components) they might come out at the same volume. Are there any secondary structure changes associated with dimerization (e.g. using CD spectroscopy)? If the do have similiar radii, I would do either AUC as suggested above (very labor and time intensive though) or find a Multi-Angle-Light-Scattering (MALS) instrument. Then you can measure the MW of your peak at 12 ml online with SEC and know if it is a dimer or monomer.

Hi Ya,

I need to know some more details. What is the size of the S-75 column (bed volume). Have you loaded only your ligand on to S-75 column and checked where it elutes? Did you check the aggregated fraction on the gel and find if it has both ligand as well as the protein?

I dont think the peak at around 1kDa represents your ligand or any degradation product. It must be the imidazole that u used for elution from His trap column or even salt. If there would have been degradation you should have seen some more peaks other than just the 1kDa peak.

I understand your ligand does not dissolve in water but then can you do a dialysis (overnight in cold) after mixing your protein and ligand in to the same buffer in which your gel filtration column and then apply it to the column.  Try it.

All the best

Hi,

Thanks for suggestion. My gel filtration is 24ml Tricon column. I havenot loaded the ligand into gel filtration column. I have already run sds gel for aggregated peak, but I havenot see any bands.

My buffer has 150mM salt. Is it possible that salt elutes always at the end ?

yes ...salt elutes always in the end...

HI Anish,

Thanks for your suggestion. I agree if the salt elutes, the conductivity should increase. I dint see any raise in conductivity when this peak elutes. 

But always there comes some peaks eluting at low molecular weight, which I cannot see in the sds gel.

Use proper amount of nonionic detergent, like Tritonx100 for sample preparation as well as column pre-equilibration, and see if it works.