I am new to Western Blot analysis and I have recently done my first two. I am studying a phosphoprotein (a protein kinase) that can be both activated and inactivated via phosphorylation at a specific amino acid residue. I have labelled my membrane against the active and inactive forms of my protein of interest (using phosphospecific antibodies), as well as the total protein (using pan-specific antibodies).

I know that Western Blots are used to quantify the expression of a protein (i.e. how much of a specific protein there is in a sample). But I was wondering how they can be used to quantify the activity (e.g. how functional the protein is) of a protein, e.g. how can I measure/determine whether my protein is more active/inactive in my treatment samples vs. control?

For example, I have normalised the integrated density values for my active protein bands against the integrated density values for my total protein bands. Then I have calculated the fold change in protein level for my active protein in treatment samples relative to control.

If I have observed that the fold change for my active protein is reduced by 40% (as an example) in treatment samples relative to controls, would this be sufficient data to say that the activity of my protein of interest is reduced in treatment samples compared to control?

  • $\begingroup$ You may need to optimise your question here: 'How functional is the protein?'. Too broad, what does functional mean? Also, more importantly what are the differences between your control and treatment groups? Are you asking: Because there is more protein expression, is there more protein activity? If so, you do not have sufficient evidence yet. More protein does not prove more activity, especially since the activity is undefined. $\endgroup$
    – Andrew
    Commented Apr 8, 2021 at 10:36
  • 2
    $\begingroup$ The protein I am studying is constitutively active, so by functional I mean what proportion of the protein is in its active form compared to total protein (active + inactive forms of the protein) in my samples. The difference between my control and treatment groups is that I am comparing knockout and wild-type brain lysate samples. I am investigating whether knocking out a protein affects the activity of my protein of interest. $\endgroup$
    – ceno980
    Commented Apr 8, 2021 at 10:49
  • $\begingroup$ Thank you for your response, it adds clarity. $\endgroup$
    – Andrew
    Commented Apr 8, 2021 at 21:47

1 Answer 1


I see 3 parts to a complete answer:

  1. If you define the phosphorylated protein as active, and non-phosphorylated protein as inactive, and you have a total protein blot (as you say, via pan-specific antibodies) as you describe, then you have done everything correctly.

  2. It's simple math from there on based on radioactivity or fluorescence or mass or whatever you use to quantify your band intensity. The 3 components should match up with the inactive protein amount (active + inactive = total). If this is consistent, you have additional evidence that you have done it correctly.

  3. However, here come the hidden assumptions: this is all assuming that (i) you are not experiencing significant protein loss during your preparation steps, that (ii) your antibodies are good (specific and yield the complete protein extract), that (iii) the blotting procedure is performed correctly, and most importantly, that (iv) your initial definition is valid, that the phosphorylated protein is truly the active form of the protein, and that the unphosphorylated counterpart is the inactive form. You see, this is not always the case, though with some (most?) kinases, it is. If it has not previously been demonstrated, one may raise the reasonable objection that your assumption is wrong. However, I take it that in your specific case, the kinase activation by phosphorylation is well understood.


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