Great video! Super informative. I would think you want the rectangle to encompass the entire band for accurate quantification. What is the rationale behind keeping the width of the rectangles the same vs expanding them to the full width of the band? Wouldn't you be underestimating the protein amount without adjusting the widths of the rectangles?
Very good question-in reality, band densitometry is estimating the "concentration" of the epitope across a fixed width, which is mostly informed by band thickness and intensity over the vertical range (less so by width). We have tested different widths (probably as narrow as half of a band) and reached pretty much the same conclusions as when we marked most or all of a lane. If the band is excessively narrow, then there are not enough horizontal pixels to estimate the concentration accurately and the quantities become highly susceptible to band artifacts.
Thank you for the imformative video. I followed the method in the video, but one problem is that the peak is negative in the Plot Lanes graph. May I know what the problem is?
Thank you very much, sir; it was really helpful. What if some of the bands have different widths. Which size do we need to choose? When we choose the biggest one when quantifying the smaller one, it will overlap with the other band, and the quantification result will be bigger and doesn't match the image at a glance. But if we choose the smallest one to apply to all bands, then the area calculation in the bigger band will reduced and also doesn't look to match the image. Thanks so much for your kind help.
To avoid double counting, there is no choice but to quantify with the smaller lane width. This will underestimate the wider lane somewhat. Experimentally, lane-width irregularities can be fixed by making sure each lane has something loaded on either side of it to minimize lateral diffusion into lanes with no protein in them.
Regarding the pick the pictures which come from the image j its really small I can't get to the both side of the picks second for some reasons when I see decrease in my band I found the analysis is high than the dark band and which I can see its significantly increased
Go to Image>Zoom>In to increase the size of the lane traces so that you can isolate peaks better. If you have not fully circumscribed the peak of interest with the line tool, it will highlight the entire trace and give non-intuitive "results". Also, note that the horizontal lane trace takes a gel from top to bottom and plots it left (= top) to right (= bottom).
@@kevinjanes2943 thank you for the information but do you have an idea why I got higher reading in the very low intensity band than the strong band in the analysis
Chemiluminescent bands may appear white on a black background, which can cause all the traces to be upside down from the video. If this is the case, select Edit>Invert.@@shadianada4296
May I know what is the setting of measurement Sir? Because if I choose Mean of Gray Value the black band will have less result than the gray one. Thanks in advance
The video does not collect regions of interest to compute density but rather Analyze>Gels. When the magic wand is used to select the integrated area of the peak, the measurement that appears is area.
Thank you for this useful guide. I'm trying to quantify the band density using the ImageJ, but after drawing the rectangles on each band and proceeding with Plot Lanes, my plot images are small (unlike yours). The peaks are not appearing in full, and I can't see the "bottom/the skirt" of the peaks if you will. So I cannot draw the straight line accurately under each peaks to quantify under the area of the peaks. Really need some advice on how I could adjust this.
Go to Analyze>Gels>Gel Analyzer Options and set Vertical Scale Factor to a number greater than one (for example, 5 = fivefold increase in vertical scale).
Thank you for the informative video! Do you have advice for a scenario where the curve of the peak is very wide, and it is unclear where exactly it levels off into background (for example, this happens sometimes for me when there is especially high background). Also, do you have any experience with analyzing total protein for normalization? Is there a particular way you might recommend to do this? Thank you!
What I have found is that it does not matter to much as long as you apply the same "rule" (e.g., easiest straight-line fit, best horizonal-line fit, etc.) to all the lanes you are analyzing. One only gets in trouble when one rule is applied to some lanes and a different one to others. For total protein, we have a cheap miniaturized version of commercial BCA described here if you have a microplate reader (systemsbioe.org/publication/microplate-bca-protein-assay/). If you are looking for an on-blot total protein stain, I quite like the idea of Revert 520 if you have one of the newest LI-COR instruments with three-color detection (unfortunately, I don't).
If I have a immature, intermediate, mature band very close together and I want to compare the amount of each band in a condition through densitometry, would it be this way? The peaks are morphed together
We had that exact problem in this paper (pubmed.ncbi.nlm.nih.gov/27273097/). We resolved it by extracting the ImageJ-equivalent information in MATLAB and then building a Gaussian mixture model of the profiles to separate the two (or, in your case, three) peaks computationally. The above reference was published before everyone was posting their code, but I located it in my laboratory archives if you want to drop me an email. If you are more inclined to use R, then you could do the same with mclust (cran.r-project.org/web/packages/mclust/index.html) although the image handling would be much more cumbersome.
Ok, currently I was using rectangle selection around the bands of each lane then "analyze the line graph" and save the data and plot on excel, find which points I need to input into OriginPro then do peak fitting, but it does not seem like the most efficient way to do it. @@kevinjanes2943 I would appreciate the email, where can I find it?
Only if you have the same sample on both gels/membranes to normalize the separate analyses. Otherwise, you cannot because the areas estimates are local to the image that is analyzed.
If you are referring to a two-color overlay (e.g., from a LICOR multicolor immunoblot), you would handle the two blots like image channels as described in this video: ru-vid.com/video/%D0%B2%D0%B8%D0%B4%D0%B5%D0%BE-inALgZ5YET8.html
As soon as you set the first rectangle and press Command+1, the rectangular box persists on the screen and you should be able to nudge it over to the next lane and press Command+2. Keep nudging and pressing Command+2 until all lanes are boxed, and then hit Command+3.
