I have cloned a short part of ECM protein that has a signal sequence. I checked the the instability index of this protein in expasy tool with and without signal peptide. This protein is unstable with signal peptide but stable without signal peptide. Interestingly, I transfected previously without signal peptide and none of them was detected in WB. I used pCDNA eGFP separately and with ECM insert as a positive control of my transfection. Transfection was absolutely fine but not the expression and detection. When I expressed that protein in E.coli it gave me thick band just after 1 our post induction in BL21. it has HA tag. His Tag and Myc tag as well. None of the approach was successful. How important is this instability index and how it might affect its detection in WB?
In addition to Steingrimurs remark concerning the reliability of instability prediction, stability prediction including the signal peptide is meaningless, since upon secretion the signal peptide is cleaved off before the protein folds.
In mammalian cells, a protein that will not fold properly will be degraded, while in E.Coli it will end up in inclusion bodies. If you have a thick band one hour after induction in E.coli, you are probably using a far too strong promoter to get proper folding - this is a good strategy to get a lot of protein in inclusion bodies for in vitro refolding, but not for soluble expression of properly folded material.
Whether you should use a signal peptide in E.coli depends on whether your protein contains cysteine residues that ought to form disulfide bonds - unless you use strains specifically engineered to have a more oxidizing redox potential in the cytoplasm, disulfide bridges will not form upon cytoplasmic expression. Export into the periplasm using a signal peptide is a way to allow the protein to fold in an oxidizing environment.
You are expressing only a part of your protein - whether your protein is able to fold at all may crucially depend on whether the fragment you express represents a structural domain that is able to fold in the absence of the rest of the sequence.
Hi Abir, instability index is just an algorithm on probability. Real world observations are often very different.
Proteins in SDS sample buffer are stable for a long time and this is probably not the issue you are dealing with.
Dear Prof,Steingrimur Stefansson ,
Thanks for your answer. If my protein is unstable in reality, Is there any way to make it stable so that we can detect in western blot?
Hi Abir, are you a using a protease inhibitor cocktal in your extraction buffer?
In addition to Steingrimurs remark concerning the reliability of instability prediction, stability prediction including the signal peptide is meaningless, since upon secretion the signal peptide is cleaved off before the protein folds.
In mammalian cells, a protein that will not fold properly will be degraded, while in E.Coli it will end up in inclusion bodies. If you have a thick band one hour after induction in E.coli, you are probably using a far too strong promoter to get proper folding - this is a good strategy to get a lot of protein in inclusion bodies for in vitro refolding, but not for soluble expression of properly folded material.
Whether you should use a signal peptide in E.coli depends on whether your protein contains cysteine residues that ought to form disulfide bonds - unless you use strains specifically engineered to have a more oxidizing redox potential in the cytoplasm, disulfide bridges will not form upon cytoplasmic expression. Export into the periplasm using a signal peptide is a way to allow the protein to fold in an oxidizing environment.
You are expressing only a part of your protein - whether your protein is able to fold at all may crucially depend on whether the fragment you express represents a structural domain that is able to fold in the absence of the rest of the sequence.
Dear Prof,Steingrimur Stefansson
Yes, I used cell lytic and PIC in the mixture.
Thank you prof Annemarie for your input. it explained a lot.
Lots of places where things could go wrong, but 2 issues come to mind:
1-How small is your fragment? How are you trying to detect it? Western or Coomassie staining? If it is a fragment, you may be detecting by staining...and expression level could be pretty low in mammalian systems. As for your BL21 expression, If you're using an Ab (to one of your epitope tags I assume) and you're pulling down your expressed protein prior to detection, make sure you're using an Ab of a different species for detection, or your thick band may be the IgG used for pulldown.
2-Have you checked your mammalian construct thoroughly? I assume you're using different constructs for E. Coli and mammalian expression; make sure your mammalian ORF is OK (start codon, no misframes or non sense mutation etc....).
Good luck!
Is it transient transfection or do you perform cell selection after the transfection?
Try changing the time between the transfection and cell lysis. For example, we can detect EGFP fluorescence 24 h after the transfection but for other proteins expression may take up to 72 h. And vice versa, sometimes we manage to express protein only via short transient transfection, not when we try to establish stable protein-overexpressing line.
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