Hi, Pawan, I did not undestand your aim: SDS as ionic detergent led to protein denaturation. www.ncbi.nlm.nih.gov/pubmed/19802818 If you want to comapre denaturated protein you have to avoid SDS in the isolation buffer, loading buffer and in the gel. Moreover, please, remebre that if you run gel in non-denaturating conditions, you have to used other molecular marker because protein migration rules are different from denaturating one. Fir your question, I have seen only one obvious possibility: without bioloing your protein is in precipotate, so, you can not separate in the gel. Good luck!,

The protein, without boiling is likely sitting at the top of the well as an insoluble precipitate.

So you think that 13000rpm spin is not enough to settle those precipitate.

Point of clarification: If you coomassie stain the entire gel, including leaving the wells attached, do you get any stain at the very top of the well of the non-denatured sample or is it totally unstained?

If you spin first, before solubilizing your sample, ie by heating the sample, then all of the protein may be in the precipitate before the loading. If you load everything without spinning, then it would likely remain in the well at the top of the gel.

If you have potassium in the buffer, it could precipitate everything after adding the SDS, before loading. Is this a possibility?

I have yet to do this as I got this suggestion from my colleagues too. But do you think protein will stay during staining destining process in the wells. I would also like to tell that the I have truncated this protein for my studies (his tagged at N-term as FL) and found that N-terminal of the protein (1-435aa) also behave the same way all other truncation like c-term (436-685aa) or smaller truncation of N-term itself is fine in this experiment in both condition. One more point I would like to add that protein is very less soluble so I get very little amount of both FL and N-term, other truncation's are quite soluble. So for FL and N-term I cant do gel filtration because of the limitation in yield. Full length is 685aa long.

I purify my protein in a buffer used for binding and washing (NaH2PO4- 50mM, NaCl - 300mM(17.54g, Imidazole- 20mM, β-ME- 10mM and Glycerol- 10% by volume). Then I elute in same buffer containing 500mM imidazole. pH of both buffer is 8.0. Then I dialize the protein in PBS for for 8-10hr at 4 degree. Then after dialysis I give a spin of 13000rpm for 15mint at 4 degree, then I collect the supernatant then add dye as mentioned above, then boil one sample and not the other then I do the SDS page followed by commassie staining.

I dont add DTT or beta-ME in both condition.

You can see your boiled protein because you might have denatured it with boiling.

As you said you want non-denaturing conditions, I recomend you to check your protocol reagents because some could be denaturing agents such as SDS, B-ME or DTT.

Maybe this link might help:

http://www.molecularinfo.com/MTM/G/G1/G1-1.html

As suggested by Elena, you will not be able to run your non-denatured sample on the same gel under the same conditions as your denatured sample as they require different conditions to migrate successfully.

Also, keep in mind when you do run your native gel, be sure the isoelectric point of your different constructs are less than 8 (since the protein is migrating based on intrinsic charge), otherwise you'll have to switch the anode and cathode for it to migrate into the gel, as the pH of the running buffer typically is 8.3.

Hi, Pawan, I did not undestand your aim: SDS as ionic detergent led to protein denaturation. www.ncbi.nlm.nih.gov/pubmed/19802818 If you want to comapre denaturated protein you have to avoid SDS in the isolation buffer, loading buffer and in the gel. Moreover, please, remebre that if you run gel in non-denaturating conditions, you have to used other molecular marker because protein migration rules are different from denaturating one. Fir your question, I have seen only one obvious possibility: without bioloing your protein is in precipotate, so, you can not separate in the gel. Good luck!,

If the multimers of your protein is made only by disulphide bonds, then your current method with SDS and without any reducing agent should work. If not, as said earlier, you have to avoid SDS in loading and running buffer.

The composition of the buffer has been taken from an published manuscript. The PI of my FL and N-term protein is around 6. In literature there are proteins which forms SDS resistant complexes without cysteine, so the amount of SDS 0.64% (in the dye) and also in running buffer (0.1%) is below 2% to keep it low according to literature. As I have mentioned that all other truncations are fine in both condition with some have PI around 6 similar to FL and N-term. The gel is not completely native as it contains SDS in both dye as well as running buffer, the purpose of adding it is to prove that it forms an SDS resistant complexes in condition similar to published earlier.

Thanks, Pawan, you explain everyting very clear! Do you have also SDS in the gel? One of the more possible explanation that under non-boliling conditon you have a complex with high molecular weight (148 or higher) and they run very slowly. Perhaps, you can try reduce gel density to get separation of higher MW complex. Or your can try a gradient gel. May be this will help. Good luck!

That is my concern Taras. Yah I am casting SDS gel as mention by maniatis protocol.

I thought about the gradient gel but before proceeding I thought to check in a 6% gel where also the protein (FL and N-term) didn't entered in unboiled condition. So below 6% it will be difficult to handle the gel for coomassie staining. But I can try that also if it can work.

One more thing I would like to add that I have done the interaction studies with my bacterial protein (making its his tagged truncation as mentioned above) with a mammalian protein (GST tag FL) and map the domain of the interaction of bacterial protein keeping mammalian protein as bait on glutathione beads and incubating the bacterial His tagged FL and the truncation (equimolar) in binding buffer (PBS). According to my hypothesis the mapped domain of bacterial protein) should form SDS resistant multimer. So I have run the mild SDS gel (as mentioned above), the mapped domain is forming multimer according to my hypothesis (not the other truncations both in boiled as well as unboiled condition). But the main problem that I am facing that the FL and N-term is not entering in the gel in unboiled condition and I need these answer for my scientific inquisitiveness as well as during communication of my manuscript in future.

One of my colleagues also suggested that there are chances of soluble aggregates which have may be many fold higher in molecular weight (may not be a multimer) and may not enter in the gel. I have also got suggestion to do a high spin (like 100000g) of my protein samples to remove any soluble aggregates and then do mild SDS page with the supernatant in both boiled and unboiled condition. He also suggested Dynamic Light Scattering (DLS) but the protein concentration and amount that will need for this experiment is very tough for me to get by native purification.

But I am also slightly confuse with this thought. As many of us purify recombinant protein (like his tag) in native condition and do coomassie staining to check its purity and do interactions studies by keeping bait protein as GST tag and the prey protein as his-tagged.

Then Is this a possibility that many of proteins which we are purifying (as many of us don't check its multimer state if not required) are in form of soluble aggregates and we are doing a false studies say to map domain of interaction between two proteins or pull out interacting partner?

One more thought is boggling my mind that is these soluble aggregate can bind to their beads (say Ni-NTA in this case) and give u interaction results?

Can presence of imidazole in elution buffer (pH-8.0) if not dialized also affect multimerization of protein?

I would like to thank all the participants for giving their views in this discussion. I would also like to ask what should be the key experiments if u can summarize which I can do to solve these ambiguity?

Hi, Pawan, I understood your points and agree with your concern: interaction of isolated protein were strongly dependent form properties of your solutions (pH, ionic strength etc). You can try to overcome this problem: Just small suggestion: you have to try PLA assay in completely native conditions and, may be, after treatment with different detergent. This is easy, do not take too much time and will be quite effective. It is even much easy tto compare with other assays, like PAAG etc. But you need antibodies..

Hi Taras, I dont know about this PLA assay. I would appreciate if you can provide any protocol or any literature to go through to understand it well before performing. Also will it help me any way in my multimerization experiment. If not what I can try for that besides gradient gel.

I have one thought in mind for my multimerization experiment and like to share. I am analyzing two extreme condition one without boiled and other boiled at 100 degree for 10 mints. I want to try to load a temperature gradient of boiled FL or N-term on mild SDS gel. I mean I want to load 10min, 9min, 8min...........1min, unboiled on the mild SDS gel and do a coomassie staining. If I see a continuum of decrease of intensity of band from 10min at the same mol. wt as boiled at 10min (74kDa) which will mean that protein only entering its in linear state or may be I can capture the intermediate state if there is any.

Can you share your views on this to refine it more or if any thought which I am not thinking.

Hi, Pawan,

here is a link: en.wikipedia.org/wiki/Proximity_ligation_assay_(PLA)

Yes, of course, you have to made different denaturation conditions (temperature, SDS, etc) and there experiments will be you answer about multimer stability under your given conditions. But this does not mean that under other conditions or in situ you will have same situation.

Because the sample is in PBS, the SDS, when you add it becomes insoluble because of the potassium. The heating probably helps solubilzing it. So I would not use PBS, or dilute the protein out before running the gel, because the SDS will precipitate if the potassium is at a high enough concentration.

Hi, Marcia, yes, this is a good idea! Tris buffer will be also OK.

When the SDS precipitates, because of the potassium, the protein precipitates with it, so it never enters into the gel. If you want to run the protein under non denaturant conditions, leave out the SDS. If you want to run a SDS page experiment, then you may want to add DTT to the loading dye. It depends if you want to see multimeric forms or just the monomeric form. Just run the gel minus and plus DTT in the sample if you want to see if it is forming disulfide bonds.

YES, Marcia, this is a very good suggestions, you can use even SDS, but not with PBS! However, do not forget that this experiemnt will give you answer about the probablity of protein interaction in the solution. The situation in situ may be very different.

That is true TARAS. In situ may be very different. The PLA seems very useful as you have mentioned.

Hi Marcia, You mean that when I am boiling the sample with SDS in PBS it solubilize the protein but when I am not protein is in precipitated form. But if that was the case my other truncation's should also not enter in the unboiled state but that is not happening. It can be possible if my FL and N-term are more prone to precipitation than other truncation. I can leave SDS, I am adding just because to prove that multimer formed is SDS resistant.

DTT I can add but I want to see the multimer state of the protein and may be cysteine is not involved in that..

I would also like to repeat that the protein which I am loading is the purified his-tagged proteins not in combination of any other interacting partner protein.

I think the heat just helps to solubilize everything, including the precipitated SDS. When the SDS precipitates, I think it is unclear what would remain soluble. Except that I would think the most insoluble proteins would precipitate first, or those on the border of being insoluble, would tend to come out of solution first. This seems intuitive to me although I don't have any references I can point to. So maybe the truncated proteins are more soluble in general than the full length forms. At any rate, I would avoid the possibility of precipitation, by changing the buffer or diluting the protein into buffer that does not have potassium. Then you don't have to worry about the SDS precipitation. If you want to see the multimeric state, if you are looking at intact disulfide bonds, then I would leave the DTT out. But it may be useful to run the gel with and without DTT in your loading dye. If you spin the sample after boiling, and everything is soluble, then you can also just continue doing that. But if you want to see what is SDS resistant, and your SDS is precipitating, then I would change the buffer of your protein. You can make a PBS like buffer, but just leave out the potassium.

Hi Marcia,

Thanks for your valuable suggestions. I am definitely going try it and will let you what happened. Thanks again.

Depending on the strength the band in the boiled sample,there is also the possibility that the simple concentration of the sample by boiling and evaporating off the water leads to the appearance of a faint band.

Hi Jeffrey , Can u explain the context for which you are talking , as I haven't got your point.

The focusing of the gel is not perfect. The same amount of sample (in mg) will show up as a sharper, more intense band in a more dilute sample than in a less concentrated one. Depending on how you load the sample, the volume of a boiled sample will generally be less due to evaporation of the water during heating. If the volumes are not equalized between the two samples before loading, evaporation of water during heating will decrease the volume relative to the unboiled sample and lead to a sharper and more intense band purely due to the change in concentration. This will not explain a very intense band in the boiled sample that is absent in the non-boiled sample, but if you see only a weak band only in the boiled sample this may be the reason.

As I am boiling the sample in closed eppendorf so what ever evaporation occured I again gave a small spin to settle it down before loading. But in the gel in unboiled condition I am not seeing anything, not a less intensity band also. So I don't think that should be the case

Charge issue!!