I'm investigating some possible substrates and for this I need to over express proteins in mammalian cells and test some inhibitors on them. I'm doing two at the same time. One gene/protein is no problem: cells transfected with the plasmid show overexpression of the protein on the western blot. The second gene/protein however shows nothing. All used material is exactly the same (primers, plasmid, cells), so that shouldn't be the problem. For both it's the full length version of the gene/protein. The first one is around 37 kD, the second (not successful one) is about 51 kD. Could this be the problem?
Does anybody have suggestions why this one gene/protein might not work?
The cells were visually all the same by the way, there didn't seem to be a toxic effect of the transfection.
Have you sequenced your plasmid? Sounds like you might have a problem with the coding region of the gene. You mention that the primers are the same, did you clone the gene using PCR? PCR can introduce mutations and this could be the problem. What is the promoter of the plasmid that you are using? Do both genes have a proper Kozak sequence? This would not explain why you are not getting any expression, but a good Kozak sequence can increase expression. Also, do you know that the assay that you are using for the second gene (I presume western blot) works? Do you have a positive control?
I do not think it is the protein size that is affecting expression. However, it could be a problem with the transfection procedure itself. What is the concentration of the plasmid you are transfecting with? Are both plasmids of the same concentration? You need a minimum of 100ng of plasmid for certain transfection reagents. Also, some proteins are harder to express in certain cell lines and more plasmid (~1000ng) needs to be added to the cells for efficient protein expression.
Have you sequenced your plasmid? Sounds like you might have a problem with the coding region of the gene. You mention that the primers are the same, did you clone the gene using PCR? PCR can introduce mutations and this could be the problem. What is the promoter of the plasmid that you are using? Do both genes have a proper Kozak sequence? This would not explain why you are not getting any expression, but a good Kozak sequence can increase expression. Also, do you know that the assay that you are using for the second gene (I presume western blot) works? Do you have a positive control?
Did you produce the plasmids yourself ?
As stated by Jonathan, there might be a problem with the plasmid itself. Did you pick up several colonies when you transformed your bacteria to produce the plasmid ? Maybe the one you chose had a problem and got resitant to whichever selecting agent you used despite having a incorrect/incomplete sequence for your gene of interest. If you picked several clone to produce your plasmid, try a few other ones. If the problem persists, check your inserted sequence.
Do you have a reporter gene on your plasmid ? Is it expressed ? (which may lead towards a problem with your gene of interest sequence/insertion within the plasmid construct). Do you have the same promoter on your two plasmids ? A different promoter may behave differently in a given cell line...
A little more detail would be helpful. I agree with the comments above regarding PCR-induced mutations, appropriateness of promoter to the cell line, concerns about detection assay, etc. Those are all highly valid concerns. In addition to sequencing your constructs (something one should always do) you might consider RT-PCR to see whether you're even getting transcription of your insert. Although size is unlikely the problem you may need to include an intron for stabilization of precursor mRNA. Depending on the nature of the gene being expressed there may be cellular mechanisms (e.g. miRNA or endogenous siRNA) limiting the stability of the transcripts or even preventing their translation. Look for sequences, especially in the 3' region of the sequence, that resemble known miRNA targets. Simply using another cell line is also another possibility. Anything special about the one you're using?
Sequencing the plasmid and detecting mRNA level by Real-time PCR would give you a rough idea about what is happening. In addition, one possibility is that this protein is toxic to cells, which develop a mechanism to limit its expression by degradation. Therefore, you can try is if you can detect protein expression when adding MG132 to inhibit proteasome mediated protein degradation pathway.
Along with all the other suggestions-here are a few more-
Are the genes codon optimized? As already suggested -are you using the same Kozak sequence for the two genes? Once you check the sequence of the 51 kD protein, you may want to try different transfection agents. How old is this plasmid? I am sure you have already done this but have you checked the integrity of the plasmid? Do you have a positive control included in your westerns? Are both the proteins cytoplamic or are they secreted? If cytoplasmic, how are you lysing the cells? Are you sterile filtering your DNA stock prior to transfection?
Good Luck
Have you tried transfecting one gene/protein (51 kD) separately instead of simultaneous transfection of two genes. Is the second gene/protein negative regulator of first?
1) sequence the plasmid. 2) test mRNA level of the second gene. There are several possible problems: a) gene has bene mutated during PCR(increasing mRNA but wrong sequence of plasmid). b) some novel regulation has been induced(mRNA doesn't increase). c) some novel post-transcriptional regulation (increasing mRNA, correct sequence of plasmid )
It may be due to wrong reading frame of protein. Insilico characterized reading frame may be not accurate always
It could be an issue of the Antibody you are using against the second overexpressed protein. Are you sure that it detects that particular protein..?
I have exactly the same problem (http://bit.ly/1g3Giy2). Particularly, one X protein I try to overexpress need a Y protein to be stable. Did you checked if there is any vital interacting protein?
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