Hi there,

You should go for a WB with anti his antibody to see if the tag is present on more than one of the bands which would indicate protein degradation. The other thing you could do is MS Fingerprint or Edman sequencing on the three bands to identify the protein sequences...

As it is not clear when this possible degradation occur, you should also add protease inhibitor cocktail to your lysis buffer.

Hope this will help.

Appears as the degradation of your target protein. I would suggest to do go for mas-spec analysis of the bands, to see if they originate from your protein. 

Santosh.

That's quite a lot of other proteins. I'd expect they bound specifically to your column, so go either for blotting or sequencing as already stated.

But do not expect to have 100% pure enzyme after one column.

in case of being proteolysis of your protein, you can try adding a coctel of protease inhibitors to avoit such degradation and compare the profiles of both samples in SDS-PAGE. This is maybe easier than trying to have an antibody to check if the bands are due to degradation.

Anyway, it is very rare  that all bands are very strong which is not usual if they are due to degradation.

First you should check WB with your protein specific antibody, that would rule out degradation, which seems to be the case as elute lane on the gel doesn't look like degradation - it's more of non-specific bound protein to the resin. Try increasing the stringency of your washes. And you should also not get multiple protein over expression, try running the uninduced, induced and supernatant lanes together for appropriate comparison.

I approve Sachin's answer as the first easy and cheap step: induce a small culture (2ml) and lyse it in SDS loading buffer/Laemmli directly, loading it on gel next to an equally treated uninduced culture... you should clearly see how many bands are overexpressed.

Anyway most of the other things said are correct. Usually bands smaller than your protein after a column purification are degradation bands, and using an anti-His antibody you will see it, even if you don't have a specific antibody for your protein (otherwise, if they lacked the His-tag, they would not be on the column). But it is also pretty true that metal/his-tag columns are not the cleanest, to be gentle! Probably it is not worth to do MS just for the purpose of finding out your protein is degraded or there is uspecific binding to the column.

To solve the degradation problem most has been said, use protease inhibitors in the lysis buffer, always work on ice or at 4°C, I also think your sonication is a bit harsh, I would not go with more than 4 sonication cycles of 20 sec on ice, in the end you can extract quite some protein, the problem is that it might be degraded. And if it is dirty, since size is clearly smaller, a size exclusion as suggested will solve your problem.

I'm afraid you should verify wether the extra bands were the degraded peptides or non-specific proteins first, by WB or MS.

If they were non-pecific proteins, you should remove it by further purification with other chromatography resin, such as DEAE, Q, SP or CM.

If they were degraded peptides, protease inhibitors should be added to the buffer.

A Western blot with a specific antiserum should answer the question whether degradation is a problem. If not al purified proteins react with the anti-serum, which is the most likely, binding of host proteins to your resin or to your recombinant protein could be the culprit. In this case a more denaturing lysis buffer would do the trick. 

In my experience using doubly tagged proteins and using the corresponding columns, for example a N-terminal His-tag and a C-terminal Strep-tag, always results in the isolation of a single full-lengt recombinant protein

Many Thanks for all participants. I really appreciate your valuable advice.

Paul (above) is right. These are very often either degradation products which still have the purification tag or high affinity binding, co-purified bacterial proteins.

It seems to me that you had contaminant proteins that co-purified with the desired protein, and it can happen often. Most of the times, degradation bands are close to each other. Even if its degradation, your protein looks quite well-expressed from the gel and as mentioned in a post above, I would carry on with one or two purification steps, e.g. ion exchange/gel filtration.

To prevent degradation, use protease inhibitor kits. E. coli strains deficient in proteases like BL21 etc.

At the end, if we get enough pure protein from the purification scheme, I wouldn't worry much.

I might do some Western blots on whole cell lysates at different induction temperatures to see if you get a temperature that shows more of a single band (taking into account total yield) if you haven't done so already. Then I might drop the total bed volume of your bead bed a little bit if you suspect co-purifying proteins. Extra protease inhibitors, ion exchange and size exclusion are things to do next as suggested, but it might easier to back up a step or two to minimize breakdown products.... could be more tricky to get rid of than you expect them to be.

You can further purify your protein by subjecting it to size exclusion chromatography.  Alternatively, you can pass it through centrifugal filter columns that will concentrate your protein and also exclude small proteins (make sure you buy right filter, based on what you want to filter out).

Hi Your multiple bands could be either from the protein purification process or from the growth. You could adjust your growth conditions to minimize the extra bands and you could  lower your Inducer concentration (e,g.  lower IPTG concentration) or lower the growth temperature after induction.

It is also very common you have multiple bands after affinity column. like Paul and other people suggested that You could go further purification by ion-exchange column or gel filtration column. 

Hope it help

Weiping Yang