Hello everyone,
I am blotting for caspase 3. I have an antibody that binds to both cleaved and full length caspase 3. However, when I expose the blots, there is much more full length caspase 3 than cleaved caspase 3, so they require two different exposure times.
For ImageJ analysis, I have first analyzed the cleaved caspase 3 on the 2 min exposure blot, then I analyzed the full length caspase 3 on the 3 sec exposure blot. Then I divided the cleaved caspase 3 values by the full length caspase 3 values. I plotted those ratios (shown in attached figure).
Shown on the attached figure is also a "parts of a whole" graph where I determined the relative percentages of each form of caspase 3. Again I used the data from the different exposures to calculate these values. The thing that bugs me about this graph is that the values shown are not really correct ... looking at the blots, it's actually a very small percentage of cleaved caspase, not 60% like the 12 hr and 24 hr timepoints show.
I did not blot for actin since I figured that the ratio of the two is all that matters for determining extent of apoptosis (although I am second guessing that assumption).
My question is ... is this a valid way to compare levels when using different exposure times for each form of the protein? Please let me know if you think there is a better way to analyze this type of data.
Thank you!
Due to the nature of the film exposure reaction, there is no linearity. Each crystal is either digitally black or not. Further exposure then does not make the individual spot blacker. In other words, the limited 2-dimensional resolution of the film in combination with the photochemical reaction prevents linearity at what to the eye appear to be relatively low exposures.
Looking at the blots, it appears that the quantities would be maybe 2 orders of magnitude apart. But this assessment requires the assumption that the antibody binds to both forms of the caspase equally well. Is this really the case?
The best way to find out would be to add lanes with different quantities of pure caspase 3 cleaved and full length forms and thus have a calibration curve for each form on the blot. But still you would require different exposure times.
Right, so if I did ensure that the binding affinity for both forms was equal, would what I described above be a valid way to compare the two forms of caspase 3?
You would still need a calibration curve of one of the two forms because the optical density is not linear over the entire concentration range (and strictly speaking isn't linear at all).
Hello,
I am having the same kind of issue. I use aldolase as normalization control (40 kDa) and when I look for a specific histone modification (15 kDa), what I do is that I expose for a short time the membrane to get nice bands in the upper part and then I expose for a much longer time to get a signal for my histone modification. From there, in order to compare, I calculate ratios between the histone modification signal obtained after over-exposing the membrane and the aldolase signal obtained after normally exposing the membrane. I've got triplicates for each experiment and I run them on the same gel.
I've tried playing around with the primary antibody concentration to no avail, I always have to over-expose to detect my lowly abundant histone modification. I was thinking that I could lower the amount of proteins I load per well (currently 10 ug). Anybody has any suggestion?
- Badges
- Science method