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I am transiently transfecting mesenchymal stem cells with a mammalian plasmid-based expression vector that does not contain any mammalian selection marker and also the gene of interest will be cloned into the plasmid without co-expression of any reporter fusion gene.

I would like to know if selection or expression testing is good practice and mainstream in plasmid-based transient transfection of mammalian cells.

As you know, in bacterial transformation selection against non-transformed bacterial cells is mandatory so that transformed bacterial cells could be enriched for. I wonder whether this is also mandatory in plasmid-based transient transfection of mammalian cells. If not, then would not non-transfected cells outgrow transfected ones and affect downstream cell-based functional assays? Also, do I have to measure transfection efficiency beforehand? Also, how long can a mammalian cell retain a plasmid without a mammalian selection marker? Thank you.

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    $\begingroup$ Welcome to the site. This is not a tutorial site to explain every aspect of a given paper, concept, or idea. I strongly suggest you take the tour and carefully read through the help center to learn more about the site, including what is on-topic and what is not, and how to ask a good question. Please try to narrow down your question to one specific question instead of trying to get a tutorial on transfection as a whole. $\endgroup$ – MattDMo May 23 at 20:24
  • $\begingroup$ You will need to verify the quality and extent of your transfection reaction after 24-48 hours, depending on the cell line and the promotor. The most common methods in situations where you don't have any reporters are Western blot, immunofluorescence staining, and flow cytometry. The best way to do this is to have a good antibody to the target. If you don't, make sure your construct includes an N- or C-terminal epitope tag like Myc, FLAG, HA, or something similar so that you can detect your protein of interest. $\endgroup$ – MattDMo May 23 at 20:28
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No and Yes, respectively.

No, you don't need to select for and generally you can't unless you use something like FACS to sort and capture cells expressing your marker/GOI.

Transient transfections are generally not selected for, if they are it is to make a more stable line, and either requires integration of the marker site into the genome (most common) or in some cases use of a plasmid and cell system that supports propagation of the plasmid.

With transient transfection it is normally a good idea, at least for the first few times, if not every time, to confirm that the plasmid is expressing as you expect. You can do this by immunological methods, most commonly western blotting (WB) or immunocytochemistry (ICC). ICC will give you a percentage transfection, which can be useful in assessing how well you would expect an overall population to respond to a stimulus and how well you can see that effect - what if only 10% of your cells transfected compared to 50% or 90%, but it can't tell you how much protein is being expressed. ICC can also tell you how cytotoxic your transfection is - commonly used methods like the Lipofectamine family of reagents are often in the 20-50% cell-death range. WB on the other hand gives you how strong your expression is - how much protein is there, but it can't tell you how many cells were transfected or how much cell-death there is.

In theory at least FACS can do both number and strength, but it would require calibration of your antibody by both western and ICC.

You can also look at RNA expressed off the plasmid, but this may not bear a direct relationship to how much protein is produced.

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