Actually, we measured the expression of a particular protein in the adipose tissue of db/db mice. We loaded the first three samples from the normal group, then three samples from the control group--which was then followed by each three samples from the tested group at two different doses. Finally, we cropped a lane from each group, and submitted a paper to a journal. Furthermore, we calculated the band density of all three lanes. The reviewer asked us to clearly label the band, since it was not from the same image. How can I label them?
Hi Bhesh, I think what you can do is to draw a vertical dotted line between each lane that was cropped and clearly specify in the figure legend that it is a "montage" from the same blot..and submit also the original image so the reviewer can see that there is no manipolation....
Hi Bhesh,
We should not compare the blots from two gels unless there is very clear differences. It is better to run your samples in the same gel. Use 15 well combs instead of 10, if you have more than 10 samples to load.
Maybe it's easier just to show the entire blot so that people can see the replicates. Don't cut anything out. We have done that. We then also show a bar graph with the quantification. The blot takes up more room but easier to understand.
Good luck!
Do you have control bands, such as actin or GAPDH, you can do the ratio of your interest of protein with each corresponding control band to calculate stoichiometry, in this way, even your wb signal did not come from a same image but should be accepted I think.
I agree with Yunlong Qin. If you have controls like GAPDH or Actin, the expression of your protein can be represented as relative expression of your protein to control protein ( your protein:GAPDH).
You can then compare relative expression of protein of interest in treated with the control samples (presuming they are treated with the vector carrier control e,g. DMSO), and graphically present the trend with standard dev/ error bars (e.g calculated from mean of 3 samples per treatment, or 3 independent experiments (repetitions))
Quantitative western blot is very tricky. You need to find the optimal conc. of protein required to get clean quantifiable bands. I use ImageJ software, and have noticed that at least in my case, 7ug protein gave me a clean (almost error free) quantifiable bands.. . If you get real strong thick bands in your Westerns, you reach a saturation point for clean quantification and then software cannot calculate the values correctly.
I agree with Karki. The best way is to use a 15 well comb in order to analyze all your samples exactly in the same conditions.
I agree with Yunlong. Use an internal standard protein that you know is not affected by your treatment and calculate the optical density ratios between the bands related to your proteins and the control proteins. The ratios reflect the real change induced by your treatment. Even if you succeed to load all your samples on one gel, this procedure is essential.
I have noticed that when you use 15 wells comb (BioRad), especially for quatitative Westerns, it is are not really great.
As long as you have the same control (e.g. untreated) on both gels/Westerns, and since after Western you are calculating the ratio of your protein to GAPDH/Actin for each and every treatment, and then comparing the relative expression to the untreated sample, you shouldn't get a significant variation in values amongst different westerns.
Remember it is very important that you do not overload protein.... otherwise ImageJ has problems spewing out true values at the band saturation point.
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