Try using gels containing 8M urea. I have used this effectively to avoid problems similar to yours. When making up running and stacking gel solutions you weigh out solid ultrapure urea and dissolve it in the concentrated buffers, adding water slowly with stirring to final volume to make the final concentration 8M. Other buffer components are as usual (1X at final volume). Make up samples with 8M urea as well. You will have to make up samples in concentrated sample buffer and weigh solid urea to achieve 8M. It's a bit of a hassle, and you have to do some calculations to account for the solution volume of urea, but you get used to it pretty quickly. I have run hundreds of gels like this and they are better by far than regular gels, for membrane proteins especially. One thing: you can NOT heat the sample AT ALL. Incubate at room temp for 15 min. The urea does the unfolding. If you heat you will transcarbamylate the proteins with the urea and you will have a mess. You don't need to add urea to the electrode buffer, but once you add the buffer over the gel you need to work quickly to load the samples, as the urea in the gel is diffusing into the running buffer with time. Just before you load each sample you also need to flush out the well that the sample is going into with a syringe with a long narrow gauge needle. This is to clear the urea gradient that forms in the well with time. You will see what I mean if you try this. The results you describe are counterintuitive, so I don't really know what is going on with your samples, but this might help. You might want to try also just increasing SDS in the sample buffer and incubating at lower temp (even as low as room temp, but for a longer time, even an hour). Native gels generally result in more aggregation, not less aggregation. 

Maybe the aggregate is so large it isn't even entering the gel under non-reducing conditions. The boiling is necessary to fully unfold the protein, allowing full SDS coverage so that a molecular weight can be accurately determined. That could be why you see a higher molecular weight band under reducing conditions- the protein is not fully unfolded. Have you tried other gel systems like Bis-Tris or even urea? Maybe that would help. 

Dear Muhammad,

SDS as well as reduction of protein could cause aggregation. So I would suggest instead of SDS-PAGE try native-PAGE. Moreover, since you have already purified the proteins, I would suggest try size-exclusion chromatography.

Best,

Jogender

Try reducing protein concentration while loading as you are using ourified proteins and not crude extract.

Dear Muhammed, according  to what you describe you seem to deal with high Mw proteins. Standard SDS-PAGE is restricted to about 200-250 kD proteins. You might want to try Blue-Native page. BN-PAGE allows you to go up to several hundred kD. Herman Schagger's group from Frankfurt, Germany, published quite some data and practical hints on this subject.

I think Jennifer is quite right that the larger proteins simply do not enter the gel because they are 'lumpy' and not unfolded. We normally heat our samples at 95 degrees for 5 minutes, a bit higher than the 80 degrees you use. Also, make sure your loading buffer has the right SDS concentration for the optimal coating of the proteins. In any case, your estimation of molecular weight will be inappropriate with unboiled samples resolved on an SDS-PAGE system.

Dear Muhammad,

first as a note concerning denaturation: For membrane proteins, it is often recommended to denature in sample buffer for 15 minutes at 37 °C (to avoid their aggregation at higher temperatures). Which basically means, you can go to lower tempereatures, but it might be beneficial to extend the denaturation time.

Anyway, if I understood you right, you do not see ANY bands in non-reducing conditions? Not even at lower apparent molecular weights? Do you centrifuge your samples before loading onto a gel, and do you find a precipitate? If you do not find any protein band without reduction migrating into the gel, than I agree with the previous comments that a change of PAGE system may become necessary...

Best regards,

Christian

Thank you very much for your kind answers. Well just for providing more information, I know that we normally heat (around boiling temperature) the protein samples in the presence of SDS to unfold the protein molecules to properly cover them with SDS. However, there are some proteins which are very sensitive to heat treatment and can aggregate at high temperature even in the presence of SDS. My proteins are one of them and for such kind of proteins we need a mild heat treatment (50 -60°C for 10 to 20 minutes). This I have to try yet.

Well, my proteins are small molecular proteins having a molecular weight of less then 80 kDa. The only problem that I am having is that I have some larger bands (around 80kDa) under reducing conditions but I have only small molecular bands under non-reducing conditions, which is not normal. Well, I think that my proteins are not properly covered by SDS and when they are unfolded by reduction of disulfide bonds their uncovered reactive parts make aggregates (supposition). It has already been documented that if the protein molecules are not properly covered by SDS then their aggregation or even cleavage can occur. I suppose my proteins are not properly covered by SDS.

Well, I am planning to use mild heat treatment and will increase SDS concentration. On suggestion of one of my colleague LDS can also be used which is more effective than SDS.

Dear Muhammad,

as Johan already mentioned, it would be better to setup a BN-PAGE experiment to have a complete profile of your proteome in a relatively easy way. In the same blot you'll have either momomeric and multimeric state(s) of your protein. Then, if you are interested in resolving your multiprotein complex, you could perform a BN/SDS-PAGE to get more information (e.g. stoichometry, interacting proteins...).

Best, Davide

Try using gels containing 8M urea. I have used this effectively to avoid problems similar to yours. When making up running and stacking gel solutions you weigh out solid ultrapure urea and dissolve it in the concentrated buffers, adding water slowly with stirring to final volume to make the final concentration 8M. Other buffer components are as usual (1X at final volume). Make up samples with 8M urea as well. You will have to make up samples in concentrated sample buffer and weigh solid urea to achieve 8M. It's a bit of a hassle, and you have to do some calculations to account for the solution volume of urea, but you get used to it pretty quickly. I have run hundreds of gels like this and they are better by far than regular gels, for membrane proteins especially. One thing: you can NOT heat the sample AT ALL. Incubate at room temp for 15 min. The urea does the unfolding. If you heat you will transcarbamylate the proteins with the urea and you will have a mess. You don't need to add urea to the electrode buffer, but once you add the buffer over the gel you need to work quickly to load the samples, as the urea in the gel is diffusing into the running buffer with time. Just before you load each sample you also need to flush out the well that the sample is going into with a syringe with a long narrow gauge needle. This is to clear the urea gradient that forms in the well with time. You will see what I mean if you try this. The results you describe are counterintuitive, so I don't really know what is going on with your samples, but this might help. You might want to try also just increasing SDS in the sample buffer and incubating at lower temp (even as low as room temp, but for a longer time, even an hour). Native gels generally result in more aggregation, not less aggregation. 

Thank you very much Mr. Kartner for your detailed and very valuable answer. I am also highly thankful to all others who participated in this discussion.

Hi Muhammad,

I think you already got some very good advices, especially those suggesting you to use urea, urea/thiousea and detergents (and I voted them UP!). 

Using DTT instead of 2-mercaptoethanol for reduction could be an option, as DTT is rapidly active (30 minutes) at room temperature (and it is significantly less toxic). You could try to find the best conditions with increasing DTT concentrations (starting from 100 mM in 1x solubilisation buffer).

Also, you could try overnight incubation at 4 °C with 2-mercaptoethanol in your usual solubilisation buffer or in  one containing chaotropic agents or detergents. 

Good luck

Mr. Bianchi,

You are very right, thanks for the suggestion. I do remember when I left my samples overnight at 4°C I was not having large molecular species. It means I gave sufficient time for denaturing substances (SDS and beta mercaptoethanol) for their activity. However, I read in the recommendations for SDS-PAGE, it was mentioned that the SDS samples should be analyzed the same day. That's why I didn't took the results of overnight kept samples. But its true that my samples were not having large species. Could you please guide me if this is the recommended method to incubate the samples overnight at 4°C.

Waiting for your kind response

Hi......

I have gone through your result 

The principle of separation of proteins on denaturing gels and native gels is different.

On denaturing gels, the proteins in a mixture get separated on the basis of molecular weight; whereas on native (non denaturing) gels, the separation is based on net negative charge.

It is also important to note the percentage of gels.  What is the T and C of your gels?

How much protein you are loading on gels? loading too much of protein will spoil the resolution.

If tray buffer strength is high, the time for electrophoretic run will be less but then the resolution will not be satisfactory.

I have worked with a variety of gels with varying gel concentrations; from 7% to 15%.  In high percentage gels, the high molecular weight proteins will never enter the small pore gel and they remain in large pore gel.  

Denaturing with mercaptoethanol and heating the sample in water bath just prior to electrophoresis gives excellent results.

If you can let me know the answers for some of my questions, I may be able to give you some suggestions

I have run samples in regular sample buffer that were kept on ice in a refrigerator overnight (i.e. zero degrees, 16 h, no additional heating) and the results were very good. If this works for you, it is probably the easiest solution to your problem. It is problematic whether this would work with high SDS or urea, since they might precipitate out at low temp overnight (LiDS might work, whereas SDS might not). DTT is a good suggestion for reduction, but 5 mM is plenty.

Hello,

I agree with Stefano Jaconi, "For sure you do not see your High MW proteins band in Non-denaturing conditions since proteins migrate towards their net charge in such conditions.". That explains why you do not see the bands on gel.

I would advise you to increase Quante of denaturant and lower concentration protein sample before heat treatment.

Best Regards

Assuming that you are working with legume seed proteins, you might find this article useful. http://ssa.ism2.univ-cezanne.fr/SSA/UE02_files/02prot%20aggregation.pdf

It shows different SDS-PAGE gel patterns after treatment of soy proteins with heat and/or reducing agents.